Metadata of the book that will be visualized in SpringerLink
Publisher Name
Springer Berlin Heidelberg
Publisher Location
Berlin, Heidelberg
Series ID
SeriesTitle
Book ID
272454_1_En
Book Title
Polyploidy and Genome Evolution
Book DOI
10.1007/978-3-642-31442-1
Copyright Holder Name
Springer-Verlag Berlin Heidelberg
Copyright Year
2012
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 1/8
EC
TE
D
PR
OO
F
Polyploidy and Genome Evolution
CO
RR
1
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: FM
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 3/8
Pamela S. Soltis Douglas E. Soltis
•
Editors
4
Polyploidy and Genome
Evolution
CO
RR
EC
TE
D
5
PR
OO
3
6
F
2
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: FM
123
Editors
Pamela S. Soltis
Florida Museum of Natural History
University of Florida
Gainesville, FL
USA
Douglas E. Soltis
Florida Museum of Natural History
University of Florida
Gainesville, FL
USA
F
14
15
16
17
18
19
PR
OO
20
49
50
51
ISBN 978-3-642-31442-1
(eBook)
Springer Heidelberg New York Dordrecht London
EC
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
ISBN 978-3-642-31441-4
DOI 10.1007/978-3-642-31442-1
Library of Congress Control Number: 2012945474
Springer-Verlag Berlin Heidelberg 2012
This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of
the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations,
recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or
information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar
methodology now known or hereafter developed. Exempted from this legal reservation are brief
excerpts in connection with reviews or scholarly analysis or material supplied specifically for the
purpose of being entered and executed on a computer system, for exclusive use by the purchaser of the
work. Duplication of this publication or parts thereof is permitted only under the provisions of
the Copyright Law of the Publisher’s location, in its current version, and permission for use must always
be obtained from Springer. Permissions for use may be obtained through RightsLink at the Copyright
Clearance Center. Violations are liable to prosecution under the respective Copyright Law.
The use of general descriptive names, registered names, trademarks, service marks, etc. in this
publication does not imply, even in the absence of a specific statement, that such names are exempt
from the relevant protective laws and regulations and therefore free for general use.
While the advice and information in this book are believed to be true and accurate at the date of
publication, neither the authors nor the editors nor the publisher can accept any legal responsibility for
any errors or omissions that may be made. The publisher makes no warranty, express or implied, with
respect to the material contained herein.
CO
RR
23
22
24
25
26
27
28
29
TE
D
21
UN
Editor Proof
7
8
9
10
11
12
13
Printed on acid-free paper
Springer is part of Springer Science+Business Media (www.springer.com)
Book ISBN: 978-3-642-31441-4
Page: 5/8
Preface
57
58
59
60
61
62
63
64
65
66
67
68
69
70
71
72
73
74
75
76
77
78
79
80
81
82
83
D
56
TE
55
Polyploidy (whole-genome duplication; WGD) is common in plants and has long
been considered as both an important speciation mechanism and a crucial component of plant genome structure. Analyses of chromosome numbers and
hypothesized breaks between diploid and polyploid base numbers have suggested
anywhere from 30 to 80 % of all angiosperms are polyploid. While recent
polyploids may be easily detected through comparison of chromosome numbers,
various processes of diploidization or fractionation may substantially alter chromosome numbers and structure, ultimately masking the evolutionary history of
duplication events. In contrast, other footprints of ancient WGD may remain in the
genome, even when chromosome numbers no longer carry the signature of past
WGDs. Genome sequences and other sources of genomic data tell us that, in fact,
all angiosperms, as well as all seed plants, have undergone one or more rounds of
polyploidy. Furthermore, ancient WGD characterizes all vertebrates, with subsequent, more recent polyploidization in fishes and amphibians. Ancient WGD is
also evident in the genomes of yeast and other fungi. While more common in
plants than other major lineages of life, polyploidy is now recognized as a fundamental process in all crown eukaryotes. Polyploidy plays a major role in shaping
genome structure and organization and in establishing patterns and mechanisms of
gene regulation. In fact, it is now impossible to construct models of genome
evolution that do not account for genomic content and genetic interactions contributed by WGD.
It has been over 30 years since the publication of a comprehensive treatment of
polyploidy [Polyploidy: Biological Relevance, W. H. Lewis (ed.), 1980]. The
intervening years have witnessed a technological revolution with a transition from
the early days of recombinant DNA to nearly routine genome sequencing of nonmodel organisms and from limited biological computing to high-performance
computing networks for the biological sciences. These transformations in methodology and computation permit fresh perspectives on polyploidy and the ability
to ask old questions with new tools.
Over the past decade, it has been a dream of ours to publish a book that
synthesizes the rapid progress in understanding the role of polyploidy in genome
EC
54
CO
RR
53
PR
OO
F
52
Book ID: 272454_1_En
Date: 16-8-2012
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: FM
v
Layout: T1 Standard SC
Chapter No.: FM
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 6/8
Preface
106
Gainesville, April 2012
88
89
90
91
92
93
94
95
96
97
98
99
100
101
102
103
104
PR
OO
87
D
86
TE
85
F
105
evolution, and this book is now a reality. In the current volume, we have compiled
the expertise of scientists studying polyploid genome evolution from multiple
perspectives in phylogenetically diverse organisms. Topics range from the conceptual and theoretical underpinnings of polyploidy (chapters by McGrath and
Lynch, Birchler) to processes at work in polyploid genomes (Zielinski and
Mittelsten Scheid, Finigan et al., Evans et al.), to patterns of ancient polyploidy
and its detection (Burleigh, Paterson et al.), to a series of case studies that both
document attributes of genome evolution in focal species and address general
properties of polyploid genomes, from ancient polyploids [maize (Schnable and
Freeling), legumes (Doyle), vertebrates (Cañestro), fishes (Braasch and Postlethwait), yeast (Hudson and Conant)] to classic model polyploids [cotton (Wendel
et al.), tobacco (Kovarik et al.), wheat (Feldman et al.)] to very recent ones
[Spartina (Ainouche), Senecio (Hegarty et al.), and Tragopogon (Soltis et al.)].
The emerging paradigm from these studies is that polyploidy—through alterations
in genome structure and gene regulation, some of which occur shortly after
polyploid formation—generates genetic and phenotypic novelty that manifests
itself at the chromosomal, physiological, and organismal levels, with long-term
ecological and evolutionary consequences.
We thank our many colleagues, students, and postdocs for lively and challenging discussions on polyploidy and its many evolutionary consequences. We
further acknowledge the support of the U.S. National Science Foundation (Grants
9624643, 0346437, 0614421, 0919254, and 0922003).
84
CO
RR
EC
107
UN
Editor Proof
vi
Pamela S. Soltis
Douglas E. Soltis
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 7/8
Contents
109
1
Evolutionary Significance of Whole-Genome Duplication . . . . . . .
C. L. McGrath and M. Lynch
1
2
Genetic Consequences of Polyploidy in Plants . . . . . . . . . . . . . . .
James A. Birchler
21
3
Meiosis in Polyploid Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Marie-Luise Zielinski and Ortrun Mittelsten Scheid
33
4
Origins of Novel Phenotypic Variation in Polyploids . . . . . . . . . .
Patrick Finigan, Milos Tanurdzic and Robert A. Martienssen
57
5
Identifying the Phylogenetic Context of Whole-Genome
Duplications in Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
J. Gordon Burleigh
115
116
117
118
119
120
93
7
Genomic Plasticity in Polyploid Wheat . . . . . . . . . . . . . . . . . . . .
Moshe Feldman, Avraham Levy, Boulos Chalhoub
and Khalil Kashkush
109
8
Maize (Zea mays) as a Model for Studying the Impact
of Gene and Regulatory Sequence Loss Following
Whole-Genome Duplication . . . . . . . . . . . . . . . . . . . . . . . . . . . .
James C. Schnable and Michael Freeling
124
126
127
128
129
130
D
Ancient and Recent Polyploidy in Monocots . . . . . . . . . . . . . . . .
Andrew H. Paterson, Xiyin Wang, Jingping Li and Haibao Tang
123
125
77
6
121
122
TE
113
114
EC
111
112
CO
RR
110
PR
OO
F
108
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: FM
9
Polyploidy in Legumes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Jeff J. Doyle
137
147
vii
Layout: T1 Standard SC
Chapter No.: FM
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 8/8
10
132
133
140
141
142
13
143
144
145
14
146
147
148
149
150
15
151
152
153
16
154
155
PR
OO
12
139
Polyploid Evolution in Spartina: Dealing with Highly
Redundant Hybrid Genomes. . . . . . . . . . . . . . . . . . . . . . . . . . . .
M. Ainouche, H. Chelaifa, J. Ferreira, S. Bellot, A. Ainouche
and A. Salmon
Allopolyploid Speciation in Action: the Origins and Evolution
of Senecio cambrensis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Matthew J. Hegarty, Richard J. Abbott and Simon J. Hiscock
D
137
The Early Stages of Polyploidy: Rapid and Repeated
Evolution in Tragopogon . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Douglas E. Soltis, Richard J. A. Buggs, W. Brad Barbazuk,
Srikar Chamala, Michael Chester, Joseph P. Gallagher,
Patrick S. Schnable and Pamela S. Soltis
209
225
245
271
TE
136
138
Evolutionary Implications of Genome and Karyotype
Restructuring in Nicotiana tabacum L. . . . . . . . . . . . . . . . . . . . .
Ales Kovarik, Simon Renny-Byfield, Marie-Angèle Grandbastien
and Andrew Leitch
181
F
11
135
Yeast as a Window into Changes in Genome Complexity
Due to Polyploidization. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Corey M. Hudson and Gavin C. Conant
293
Two Rounds of Whole-Genome Duplication: Evidence
and Impact on the Evolution of Vertebrate Innovations . . . . . . . .
Cristian Cañestro
309
EC
134
Jeans, Genes, and Genomes: Cotton as a Model
for Studying Polyploidy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Jonathan F. Wendel, Lex E. Flagel and Keith L. Adams
CO
RR
131
Contents
17
Polyploidy in Fish and the Teleost Genome Duplication . . . . . . . .
Ingo Braasch and John H. Postlethwait
18
385
160
Polyploidization and Sex Chromosome Evolution
in Amphibians . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Ben J. Evans, R. Alexander Pyron and John J. Wiens
161
Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
411
156
157
158
159
162
UN
Editor Proof
viii
341
Metadata of the chapter that will be visualized in
SpringerLink
Book Title
Polyploidy and Genome Evolution
Series Title
Chapter Title
Evolutionary Significance of Whole-Genome Duplication
Copyright Year
2012
Copyright HolderName
Springer-Verlag Berlin Heidelberg
Corresponding Author
Family Name
Lynch
Particle
Given Name
M.
Suffix
Author
Division
Department of Biology
Organization
Indiana University
Address
Bloomington, IN, USA
Email
milynch@indiana.edu
Family Name
McGrath
Particle
Given Name
C. L.
Suffix
Division
Department of Biology
Organization
Indiana University
Address
Bloomington, IN, USA
Email
Abstract
Whole-genome duplication (WGD) appears to be a widespread phenomenon, occurring in diverse taxa
including many of the model organisms used in molecular, cellular, and developmental biology. It is therefore
essential to understand the potential evolutionary consequences for individual duplicated genes, as well as
for the lineage as a whole. For example, duplicate genes may undergo pseudogenization or may be maintained
due to neofunctionalization, subfunctionalization, or selection for increased dosage or dosage balance.
Duplicates created via WGD are maintained at higher rates than single-gene duplicates, perhaps due to dosagebalance constraints. Duplicate-gene maintenance may lead to heterodimerization of an existing homodimer
or to the divergence of an entire duplicated network or pathway. Allopolyploids and autopolyploids are likely
to undergo different evolutionary pressures due to increased divergence between allopolyploid paralogs and
an increased prevalence of multivalent formation at meiosis in autopolyploids. Perhaps most importantly,
duplicate-gene loss following a WGD may significantly increase the rate of reproductive isolation between
geographically isolated subpopulations and may therefore temporarily increase the speciation rate within
polyploid lineages.
1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 1/20
Chapter 1
4
C. L. McGrath and M. Lynch
PR
OO
F
3
Evolutionary Significance
of Whole-Genome Duplication
2
23
1.1 Introduction
11
12
13
14
15
16
17
18
19
20
24
25
26
TE
9
10
EC
8
CO
RR
7
D
21
22
Abstract Whole-genome duplication (WGD) appears to be a widespread
phenomenon, occurring in diverse taxa including many of the model organisms
used in molecular, cellular, and developmental biology. It is therefore essential to
understand the potential evolutionary consequences for individual duplicated
genes, as well as for the lineage as a whole. For example, duplicate genes may
undergo pseudogenization or may be maintained due to neofunctionalization,
subfunctionalization, or selection for increased dosage or dosage balance. Duplicates created via WGD are maintained at higher rates than single-gene duplicates,
perhaps due to dosage-balance constraints. Duplicate-gene maintenance may lead
to heterodimerization of an existing homodimer or to the divergence of an entire
duplicated network or pathway. Allopolyploids and autopolyploids are likely to
undergo different evolutionary pressures due to increased divergence between
allopolyploid paralogs and an increased prevalence of multivalent formation at
meiosis in autopolyploids. Perhaps most importantly, duplicate-gene loss following a WGD may significantly increase the rate of reproductive isolation between
geographically isolated subpopulations and may therefore temporarily increase the
speciation rate within polyploid lineages.
5
6
One of the major findings of the new field of evolutionary genomics is that
duplication events involving individual genes or multigene segments arise at rates
comparable to the rate of mutation at single-nucleotide sites (Lynch and Conery
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: 1
C. L. McGrath M. Lynch (&)
Department of Biology, Indiana University, Bloomington, IN, USA
e-mail: milynch@indiana.edu
P. S. Soltis and D. E. Soltis (eds.), Polyploidy and Genome Evolution,
DOI: 10.1007/978-3-642-31442-1_1, Springer-Verlag Berlin Heidelberg 2012
1
Layout: T1 Standard SC
Chapter No.: 1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 2/20
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
61
62
63
64
65
66
67
F
33
PR
OO
32
D
31
TE
30
EC
29
2000, 2003a, b), or possibly at even higher rates (Lipinski et al. 2011). Such
observations lend credibility to Ohno’s (1970) early speculation that gene duplication is a major resource for the origin of evolutionary novelties. Moreover, it is
now clear that whole-genome duplication (WGD) events have occurred in a wide
diversity of phylogenetic lineages, including most of the model systems relied
upon in molecular, cellular, and developmental biology. For example, budding
yeast is a descendant of an ancient genome duplication (Wolfe and Shields 1997;
see Chap. 15, this volume), as is the frog Xenopus laevis (Morin et al. 2006; see
Chap. 18, this volume) and the zebrafish (Postlethwait et al. 2000; see Chap. 17,
this volume). Many ray-finned fish lineages have experienced additional rounds of
WGD (Meyer and Van de Peer 2005; see Chap. 17, this volume), and Ohno’s
(1970) suggestion that two WGD events preceded the radiation of the vertebrate
lineage has become increasingly credible (Panopoulou and Poustka 2005; Hughes
and Liberles 2008; Putnam et al. 2008; see Chap. 16, this volume). Finally, three
WGD events are recorded within the genome of Arabidopsis thaliana (Simillion
et al. 2002), and nearly all other land-plant genomes appear to harbor a legacy of at
least one polyploidization event (Doyle et al. 2008), with a proposed WGD in the
ancestor of all seed plants and another in the ancestor of all angiosperms (Jiao
et al. 2011). Thus, it is clear that understanding the mechanisms of origin and
preservation of duplicate genes promises to reveal not only the ways in which
genes acquire new functions and organisms respond to natural selection, but also
the roots of organismal diversity across the tree of life.
Because genome duplication adds thousands of duplicate genes to the genome,
understanding the evolutionary forces that act on individual duplicate genes is
critical to our understanding of polyploidization. Processes such as neofunctionalization and subfunctionalization have the potential to influence all gene duplicates, whether created through polyploidization or smaller scale duplication
events. It has become increasingly clear, however, that duplicates that arise via
polyploidization are subject to unique evolutionary forces, such as increased
retention due to dosage-balance constraints. Further, there may be processes that
are exclusive to gene duplicates that arise via specific types of polyploidization,
such as changes in duplicate-gene expression due to the genomic merger that
occurs with allopolyploidization. The relative contributions of these evolutionary
forces that give rise to the maintenance and evolution of duplicate genes that arise
via WGD, or to the evolution of the genome or species as a whole, are currently
unknown. However, discriminating between these forces and their effects is likely
to be the subject of much research over the next several years.
CO
RR
27
28
C. L. McGrath and M. Lynch
UN
Editor Proof
2
1.2 Fates of Duplicate Genes
The fate of the vast majority of duplicate genes arising by segmental duplication is
nonfunctionalization of one member of the pair (Lynch and Conery 2000, 2003a, b),
and this is expected to occur within a few million years in the absence of any
Layout: T1 Standard SC
Chapter No.: 1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 3/20
75
76
77
78
79
80
81
82
83
84
85
86
87
88
89
90
91
92
93
94
95
96
97
98
99
100
101
102
103
104
105
106
107
108
109
110
111
112
F
PR
OO
73
74
D
72
TE
70
71
3
intrinsic advantage of a duplicate copy (Watterson 1983; Lynch et al. 2001). Despite
this, most genomes that have been studied contain a large number of duplicate
genes, some of which are clearly quite ancient (Lynch and Conery 2000). Based on
this observation, several mechanisms have been proposed for the permanent preservation of duplicate genes (Hughes 1994; Force et al. 1999; Lynch et al. 2001;
Taylor and Raes 2004; Lynch 2007; Innan and Kondrashov 2010): (1) neofunctionalization, whereby one copy acquires a novel, beneficial function at the expense
of an essential ancestral function; (2) subfunctionalization, whereby complementary
mutations lead to a partitioning of independently mutable subfunctions in the
ancestral gene; (3) selection for increased gene product; and (4) the masking of
nonfunctional alleles.
When a duplicate is maintained by selection for increased gene product, it
experiences purifying selection (and may also undergo repeated gene conversion)
in order to maintain its ancestral function; this process is likely responsible for the
multiple copies of ribosomal RNA genes present in many genomes (e.g., Pinhal
et al. 2011). Neofunctionalization, on the other hand, is thought to involve positive
selection for the mutation(s) responsible for the new function, generally arising at
the expense of an essential original function, thereby preserving both copies. There
are many examples of neofunctionalization giving rise to novel gene functions in a
variety of organisms, including Arabidopsis (Erdmann et al. 2010), fish (Ngai et al.
1993), vertebrates (Layeghifard et al. 2009), and yeast (Byrne and Wolfe 2007;
Tirosh and Barkai 2007). Because one duplicate is undergoing positive selection
for a new function while the other is under purifying selection to maintain the
ancestral function, asymmetric evolutionary rates between duplicates are often
thought to be a hallmark of neofunctionalization (Johnson and Thomas 2007; Han
et al. 2009), though purely stochastic mechanisms can also give rise to apparent
rate asymmetry (Lynch and Katju 2004).
Subfunctionalization may involve positive selection acting on both duplicates if
the partitioning of the ancestral functions leads to relaxation of pleiotropic
constraints, enabling each ancestral function to be fine-tuned and improved
through mutation independently in each copy (Piatigorsky and Wistow 1991;
Hughes 1994; Des Marais and Rausher 2008). Alternatively, subfunctionalization
may be a completely neutral process if each duplicate copy simply acquires a
degenerative mutation that renders it unable to perform one of the ancestral
functions (Force et al. 1999). At this point, both copies are needed in order to
provide the organism with all of the functionality of the original, single-copy gene,
and so both will be maintained in the genome by selection. Although identifying
definitive cases of subfunctionalization requires determining that the ancestral
gene carried multiple functions that have been partitioned in the daughter duplicates, there are nonetheless several compelling examples (e.g., Force et al. 1999;
Altschmied et al. 2002; Yu et al. 2003; Adams and Liu 2007; MacNeil et al. 2008;
Semon and Wolfe 2008; Buggs et al. 2010; Deng et al. 2010; Hickman and Rusche
2010; Colon et al. 2011; Froyd and Rusche 2011).
In addition to these cases of qualitative subfunctionalization, where duplicates
eventually come to be expressed in different tissues or at different times or carry out
EC
69
CO
RR
68
UN
Editor Proof
1 Evolutionary Significance of Whole-Genome Duplication
Layout: T1 Standard SC
Chapter No.: 1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 4/20
119
120
121
122
123
124
125
126
127
128
129
130
131
132
133
134
135
136
137
138
139
140
141
142
143
144
145
146
147
148
149
150
151
152
153
154
155
156
157
F
118
PR
OO
117
D
116
TE
115
different functions relative to each other, quantitative subfunctionalization, in which
reduction-of-expression (Force et al. 1999) or activity-reducing mutations (Stoltzfus
1999; Scannell and Wolfe 2008) affect both duplicates, is also possible. In quantitative subfunctionalization, both duplicates acquire partial loss-of-function
mutations that affect the same function, again rendering both copies essential for the
proper dosage or activity of the gene products. In this case, both copies are preserved
while retaining the ancestral gene function. Although few studies demonstrating
quantitative subfunctionalization exist, Qian et al. (2010) estimated that this process
has been responsible for the maintenance of a large proportion of duplicates in yeast
and mammals, whereas Woolfe and Elgar (2007) postulated that sequence evolution
in cis-regulatory elements may have caused quantitative subfunctionalization
among Fugu duplicates.
A related consequence of gene duplication is that it can allow for the differentiation of multimeric subunits, such as the evolution of heterodimers from
homodimers. Consider a gene whose protein product forms a homodimer. After
duplication of this gene, protein subunits produced by the two duplicates (denoted
A and B) may randomly associate to make mixtures of dimers in the ratio 1 AA: 2
AB: 1 BB. If the duplicate genes are identical initially, the AA, AB, and BB dimers
will be identical as well. However, subsequent differentiation of the duplicate
genes causes the three types of dimer to become distinct. This differentiation could
be neutral, or it could be selective. If, for example, there were pleiotropic
constraints on the form or function of the pre-duplication homodimer, duplication
could allow for escape from these constraints in the AB heterodimer, as each
subunit (A and B) can now evolve independently. This can be viewed as a special
type of subfunctionalization of duplicates. Winter et al. (2002) showed that a classB floral protein heterodimer had evolved from an ancestral homodimer via this
mechanism during the gymnosperm/angiosperm transition. In gymnosperms,
GGM2-like genes form homodimers, while the duplicated homologs in eudicots,
DEF-like genes and GLO-like genes, form heterodimers. Monocots also have
duplicated DEF-like genes and GLO-like genes, but, interestingly, it appears the
GLO-like proteins of monocots can both homodimerize and heterodimerize with
DEF-like proteins, perhaps representing the transition between the homo- and
heterodimerized states (Winter et al. 2002; Kanno et al. 2003; Soltis et al. 2006).
A similar process appears to have occurred several times in the evolution of the
DUF606 family of transmembrane proteins in bacteria (Lolkema et al. 2008). In
bacteria with a single DUF606 gene, the DUF606 proteins are able to insert into the
membrane in both orientations, and functional homodimers are formed by two
subunits in opposite (antiparallel) orientations. Other species of bacteria, however,
have duplicated DUF606 genes located tandemly in an operon. In all of these latter
cases, the two protein subunits each have a fixed but opposite orientation in the
membrane, and they heterodimerize to form the necessary antiparallel two-domain
complex. A phylogenetic analysis of the DUF606 gene family reveals that this
process of duplication followed by heterodimerization likely occurred five different
times in the history of this gene family lineage (Lolkema et al. 2008). Other
proposed examples of this mechanism include SMC proteins (Surcel et al. 2008),
EC
114
CO
RR
113
C. L. McGrath and M. Lynch
UN
Editor Proof
4
Layout: T1 Standard SC
Chapter No.: 1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 5/20
5
162
1.3 Fates of Duplicate Genes Arising via WGD
167
168
169
170
171
172
173
174
175
176
177
178
179
180
181
182
183
184
185
186
187
188
189
190
191
192
193
194
195
196
197
198
PR
OO
165
166
In addition to the general preservational processes just mentioned, paralogs
resulting from WGD events are subject to unique mechanisms of duplicate-gene
maintenance and evolution (Force et al. 1999; Lynch and Conery 2000; Yang et al.
2003; Davis and Petrov 2005; Veitia et al. 2008). Well-studied polyploid species
commonly exhibit 25–75 % retention of paralogous gene pairs from the most
recent WGD event (reviewed in Lynch 2007; Otto 2007), budding yeast being an
exception with only *8 % duplicate-gene preservation (Wolfe and Shields 1997).
These are surprisingly high preservational levels, when, as discussed above, the
fate of the vast majority of duplicate genes arising by segmental duplication is
nonfunctionalization of one duplicate (Lynch and Conery 2000, 2003a, b).
Although it is possible that many polyploid species have not yet reached equilibrium and are still in an ongoing phase of duplicate-gene loss, it has become
increasingly clear that there are likely to be additional forces acting to preserve
duplicate genes arising via WGD.
A simple explanation for the large number of preserved duplicates within
polyploids is that, unlike single-gene duplicates, WGD duplicates exhibit complete
conservation of surrounding regulatory sequences, chromosomal environments, etc.
Although this likely contributes somewhat to the pattern of higher duplicate
retention in polyploids, it does not explain the observation that different types of
genes seem to be preserved following WGD compared to smaller scale duplications.
This fact can be better explained by selection for dosage balance among proteins.
Due to stoichiometric relationships with other interacting genes (e.g., multi-subunit
complexes and numerous pathways involved in metabolism and transcriptional
regulation), the functions of a subset of protein-coding loci can be highly influenced
by dosage imbalances (Veitia 2002; Papp et al. 2003; Birchler et al. 2005; Veitia
et al. 2008). In such cases, duplication of just a single member of a gene interaction
may be detrimental and actively selected against. In contrast, following a WGD
event, most stoichiometric relationships are initially intact, and therefore
subsequent losses of interacting paralogs will be inhibited by selection for proper
dosage relationships. Thus, for dosage-dependent genes, the dosage-balance
hypothesis predicts an under-representation among duplicates created by singlegene duplications, but an over-representation among those created by WGD (Yang
et al. 2003; Davis and Petrov 2005; Veitia et al. 2008). For example, Davis and
Petrov (2005) showed that the pool of preserved duplicates from the WGD event in
S. cerevisiae is enriched for ribosomal genes (which form a large complex) and
regulatory genes encoding transferases, kinases, and transcription factors, while
D
164
TE
163
EC
160
CO
RR
159
F
161
adenylyl cyclases (Sinha et al. 2005), and mitochondrial peptidases (Brown et al.
2007), all gene families that contain duplicates that form heterodimers in eukaryotes
(or eukaryotic mitochondria) with single-copy homologs that form homodimers in
prokaryotes.
158
UN
Editor Proof
1 Evolutionary Significance of Whole-Genome Duplication
Layout: T1 Standard SC
Chapter No.: 1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 6/20
206
207
208
209
210
211
212
213
214
215
216
217
218
219
220
221
222
223
224
225
226
227
228
229
230
231
232
233
234
235
236
237
238
239
240
241
242
243
F
205
PR
OO
204
D
203
TE
202
EC
201
those involved in ion transport are under-represented. Likewise, the Paramecium
tetraurelia genome exhibits elevated retention of duplicate genes involved in known
complexes (Aury et al. 2006) and in metabolic pathways (Gout et al. 2009). As in
yeast, ribosomal genes, transferases, and kinases are over-represented among
surviving paralogs, while ion-transport genes are underrepresented. In Paramecium,
there also appears to be an additional effect whereby highly expressed genes are
over-retained in duplicate following the most recent polyploidization event (Gout
et al. 2010). That certain types of genes are maintained preferentially following a
WGD has achieved fairly convincing empirical support from other studies as well
(Papp et al. 2003; Yang et al. 2003; Barker et al. 2008; Liang et al. 2008; Qian and
Zhang 2008; Edger and Pires 2009), including studies in Arabidopsis (Blanc and
Wolfe 2004; Maere et al. 2005; Thomas et al. 2006), vertebrates (Makino and
McLysaght 2010), and across divergent species (Paterson et al. 2006). Selection to
maintain dosage balance following WGD has also been hypothesized to be the
driving force behind the original selective advantage of the WGD in the Saccharomyces cerevisiae lineage (Conant and Wolfe 2007). In this scenario, the maintenance of glycolytic genes and the loss of non-glycolytic genes following WGD
might have increased the relative dosage of glycolytic genes, thereby increasing flux
through the glycolysis pathway and providing polyploid yeast with a growth
advantage over non-polyploids due to increased glucose fermentation ability.
Duplicate genes that arise via WGD are further unique in that entire (or partial)
duplicated pathways or networks of interacting proteins can diverge in concert. For
example, Evlampiev and Isambert (2007) modeled the evolution of protein–protein
interaction networks following WGD and concluded that such networks grow under
exponential, rather than time-linear, dynamics following WGD. Interestingly, they
also found that these exponential dynamics relied on asymmetric divergence
between duplicates.
Another intriguing possibility is that following WGD, a whole ancestral
network may become neofunctionalized or subfunctionalized following polyploidization, with one set of paralogs carrying out one task or reaction and a parallel
set of paralogs carrying out a related, but largely independent, task. Obviously,
such innovations require the establishment of multiple mutations and the avoidance of pathway crosstalk. Although the essential population genetic theory
remains to be worked out, several examples of such paralog coevolution appear to
have followed the WGD in yeast: parallel paralogous networks have been identified where the expression of each gene is highly correlated with the other genes
within its network but poorly correlated with its paralog (Blanc and Wolfe 2004;
Conant and Wolfe 2006). In this way, polyploidy provides a unique mechanism for
the evolution of gene networks with new (or subdivided) functions.
A final consideration in duplicate-gene evolution is whether the forces that act
to preserve duplicates change over evolutionary time. For example, it seems
possible that following WGD, a large proportion of genes could be initially
maintained due to dosage-balance constraints. Subsequently, however, over longer
periods of evolutionary time, some duplicates might accumulate mutations that
could lead to neofunctionalization or subfunctionalization. Because these genes
CO
RR
199
200
C. L. McGrath and M. Lynch
UN
Editor Proof
6
Layout: T1 Standard SC
Chapter No.: 1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 7/20
7
271
1.4 Autopolyploidy Versus Allopolyploidy
251
252
253
254
255
256
257
258
259
260
261
262
263
264
265
266
267
268
269
272
273
274
275
276
277
278
279
280
281
282
283
284
PR
OO
249
250
D
248
TE
247
EC
246
CO
RR
245
F
270
are dosage sensitive (hence their initial preservation due to selection for dosage
balance), it is likely that such neo- or subfunctionalizing mutations would need to
be preceded or rapidly followed by mutations affecting the dosage of one or both
copies. For a more detailed example, imagine proteins A and B that must interact in
a 1:1 ratio for proper functioning. Both genes become duplicated during a WGD,
giving rise to duplicates A1 and A2 and B1 and B2. Initially, all four genes are
preserved by selection for dosage balance, as loss of any one gene interrupts the
1:1 interaction ratio. Over evolutionary time, however, slightly deleterious
mutations in the A1 promoter that decrease its expression level become fixed due
to drift. To compensate, mutations in the A2 promoter that increase its expression
level are fixed, which helps to restore the 1:1 A/B ratio. At this point, A1 is
contributing fewer products to the overall A protein pool. A subsequent mutation
that changes the function of A1, allowing it to take on a new role completely, is
now more easily accommodated, as A2 is better able to compensate and take on the
full load of the ancestral A activity. Note that, instead of A1 and A2 dosage
evolving in concert, as above, A1 and B1 dosage could also evolve in concert to
maintain the proper 1:1 A/B ratio, allowing both A1 and B1 to take on new
functions.
While still just a verbal theory, this scenario has two advantages in terms of
allowing for neofunctionalization (or subfunctionalization) of WGD duplicates.
First, there is a longer time frame in which neo- or subfunctionalizing mutations
can arise, as duplicates are maintained for longer time-scales without becoming
nonfunctionalized. This is important because neofunctionalization requires the
accumulation of beneficial mutation(s), which are thought to be rare. Second, this
process would allow for neo- or subfunctionalization of dosage-sensitive duplicates, both of which might otherwise be constrained to maintain their ancestral
function indefinitely following WGD.
244
Whether polyploidization occurs by autopolyploidy or allopolyploidy can have a
significant impact on the expression and evolution of duplicate genes. Autopolyploids arise when there is an increase in ploidy within a single species (often
within a single individual), while allopolyploids are created by hybridization
between two different species, each of which contributes a full complement of
chromosomes to the hybrid, thus doubling the genome (reviewed in Coyne and Orr
2004). Many plant and frog polyploids are the result of allopolyploidization
(Adams 2007; Evans 2008), while the yeast WGD appears to have been an
autopolyploidization event (Scannell et al. 2007), although in practice it is difficult
to ascertain the ancestral state once paralog divergence has become high.
It has long been assumed that autopolyploids would initially form multivalents
at meiosis, with all four homologous chromosomes pairing randomly, while
allopolyploids would be more likely to form bivalents, with homologous
UN
Editor Proof
1 Evolutionary Significance of Whole-Genome Duplication
Layout: T1 Standard SC
Chapter No.: 1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 8/20
291
292
293
294
295
296
297
298
299
300
301
302
303
304
305
306
307
308
309
310
311
312
313
314
315
316
317
318
319
320
321
322
323
324
325
326
327
328
329
F
290
PR
OO
289
D
288
TE
287
chromosomes from each diploid ancestor pairing independently. This would mean
that duplicate copies in autopolyploids would not represent true paralogs as the
term is usually understood, but would instead represent a doubling of the number
of homologs (i.e., four homologs instead of two). The presence of multivalents is
significant biologically, as multivalent pairing can lead to intergenomic recombination via segregation, crossing-over, and double reduction. Certain duplicates
from one diploid parent could be lost completely via this process, leaving only
duplicates from the other diploid parent. This would not represent gene silencing
as it is typically understood then, but would rather be a byproduct of multivalent
formation and segregation. Evidence from plants indicates that multivalent pairing
is indeed more prevalent among autopolyploids, though the difference between the
two forms of polyploidy is perhaps less than originally expected: a survey of plant
polyploids indicated that the mean percent occurrence of multivalents is 28.8 % in
autopolyploids and 8.0 % in allopolyploids (Ramsey and Schemske 2002).
Although multivalent formation occurs at a lower rate in allopolyploids, it may be
more biologically significant than multivalent formation in autopolyploids, as
intergenomic recombination is likely to have a greater effect when genomes are
more divergent. Over time, divergence between duplicated chromosomes would
lead to increased bivalent formation.
Because allopolyploids are the result of a genomic merger between two species,
duplicate genes in allopolyploids are already differentiated to some extent
immediately after polyploidization, while duplicates in autopolyploids are likely to
be more similar in sequence and may even be identical. Allopolyploids often
exhibit immediate changes in gene expression due to the genetic differentiation
present between homeologs. This can lead to changes in methylation (Salmon
et al. 2005; Gaeta et al. 2007), changes in heterochromatin formation and transposable element suppression (Josefsson et al. 2006), biased expression of
homeologs (Adams et al. 2003; Bottley et al. 2006; Tate et al. 2006; Udall et al.
2006; Rapp et al. 2009), and non-additive expression effects between homeologs
(Hegarty et al. 2006; Wang et al. 2006; Rapp et al. 2009). These initial expression
differences between homeologs can, in turn, impact the long-term evolution of
duplicates, as selection pressures may be expected to act differently on genes that
are differentially expressed. For example, Anderson and Evans (2009) showed that
in octoploid and dodecaploid Xenopus species, paralogs of RAG1b were more
likely to become pseudogenized than paralogs of RAG1a (the homeolog of
RAG1b from an earlier allopolyploidy event), and they inferred that this was due
to differences in ancestral expression between RAG1a and RAG1b.
Many of these effects seen in allopolyploids are believed to be due to the
hybridization between two divergent genomes, rather than genome doubling per se.
Flagel et al. (2008) estimated that of the genes with biased expression between
homeologs in the allopolyploid Gossypium hirsutum, 24 % exhibit a bias due to the
genomic merger (i.e., the bias existed immediately when the allopolyploidization
occurred, at time zero), while the bias in the remaining 76 % is due to long-term
evolutionary forces such as neofunctionalization and subfunctionalization. The
relationship between the magnitude of these alterations in gene expression and the
EC
286
CO
RR
285
C. L. McGrath and M. Lynch
UN
Editor Proof
8
Layout: T1 Standard SC
Chapter No.: 1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 9/20
9
336
1.5 Polyploidization and Speciation
338
339
340
341
342
343
344
345
346
347
348
349
350
351
352
353
354
355
356
357
358
359
360
361
362
363
364
365
366
367
368
369
370
PR
OO
337
Perhaps the most pivotal role that polyploidization plays in evolution is in the
creation of new species. The polyploidization event itself can lead to instantaneous
reproductive isolation and speciation, as the cross between a new tetraploid (4n)
and its diploid progenitor (2n) yields triploid (3n) offspring, which are often sterile
due to problems with chromosome pairing/segregation during meiosis and the
production of aneuploid gametes (reviewed in Coyne and Orr 2004). It is for this
reason that models predict that species capable of self-fertilization are more likely
to give rise to a successful polyploid lineage (Rodriguez 1996; Baack 2005;
Rausch and Morgan 2005).
Perhaps more importantly, however, once a polyploid lineage is established,
subsequent silencing of duplicate genes can lead to further reproductive isolation
among subpopulations of the polyploids themselves and, therefore, give rise to
additional daughter species (Oka 1988; Werth and Windham 1991; Lynch and
Conery 2000; Lynch and Force 2000). In this model, we assume a pair of fully
functional and redundant duplicate genes, A and B, in an ancestral population, such
that each member of the initial population has the genotype AABB (Fig. 1.1). If two
subpopulations become geographically isolated and one duplicate becomes
nonfunctionalized in each subpopulation, there is a 50 % probability that a different
duplicate copy will be lost in each of the two groups. This reciprocal gene loss (or
divergent resolution) would result in the genotypes aaBB and AAbb for the two
subpopulations, where a and b denote null alleles. Hybridization between the two
groups would then lead to offspring with the genotype AaBb. Gametes produced by
these F1 individuals would have a 1/4 probability of carrying an ab genotype and
would therefore be inviable if a functional copy of the A/B gene were essential for
gamete survival or function. Even if this were not the case, 1/16 of the F2 individuals
would have the aabb genotype and, if a functional copy were essential for zygote
viability or sterility, would be inviable or sterile, whereas another 1/4 would have
three null alleles and might experience reduced viability or sterility. Up to 50–65 %
of the genes encoding transcription factors, membrane receptors, and members of
macromolecular protein complexes are estimated to be haploinsufficient (JimenezSanchez et al. 2001; Veitia 2002), suggesting that only one functional allele of such
genes is indeed likely to be deleterious.
An appealing aspect of the divergent resolution model is that it is a natural
consequence of degenerative mutations, requiring no adaptive evolution at the
D
334
TE
333
EC
332
CO
RR
331
F
335
genetic divergence between the two parental genomes is not well understood,
however, as demonstrated by Brassica allopolyploids (Pires and Gaeta 2011). While
the parental species that gave rise to the allopolyploid Brassica napus are more
similar to each other than those that gave rise to B. juncea, resynthesized B. napus
polyploids exhibit more genomic rearrangements, changes in gene expression, and
epigenetic alterations than do resynthesized B. juncea polyploids.
330
UN
Editor Proof
1 Evolutionary Significance of Whole-Genome Duplication
Layout: T1 Standard SC
Chapter No.: 1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 10/20
C. L. McGrath and M. Lynch
Fig. 1.1 Divergent
resolution of duplicate genes
can lead to hybrid sterility/
inviability and reproductive
isolation. Red bar represents
a gene that becomes
duplicated. See text for
details
PR
OO
F
Gene Duplication
CO
RR
EC
TE
D
Population Isolation and
Divergent Resolution
UN
Editor Proof
10
Hybridization
F1 progeny
1/4 F1 gametes
(no functional copy)
1/16 F2 progeny
(no functional copy)
OR
1/4 F2 progeny
(hemizygous)
Layout: T1 Standard SC
Chapter No.: 1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 11/20
377
378
379
380
381
382
383
384
385
386
387
388
389
390
391
392
393
394
395
396
397
398
399
400
401
402
403
404
405
406
407
408
409
410
411
412
413
414
F
376
PR
OO
375
D
374
TE
373
11
molecular level for speciation to occur. Moreover, in addition to genes that are
reciprocally silenced, duplicate pairs that undergo neofunctionalization or
subfunctionalization may also contribute to hybrid sterility/inviability in a similar
fashion—for example if a different duplicate becomes neofunctionalized and loses
its ancestral function in each subpopulation, or if the two duplicates become
subfunctionalized in complementary ways in the two subpopulations (Lynch and
Force 2000).
The process of reciprocal gene loss has been shown to be responsible for male
sterility between hybrids of Drosophila melanogaster and D. simulans (Masly
et al. 2006). A gene essential for male fertility, JYAlpha, is located on the fourth
chromosome in D. melanogaster and on the third chromosome in D. simulans.
This translocation presumably occurred via duplication of the JYAlpha gene and
subsequent silencing of one copy. The difference in chromosomal location of the
gene in the two species causes a proportion of hybrids to completely lack JYAlpha,
leading to their sterility.
Two similar cases were recently identified in rice. The first involves reproductive isolation between two subspecies of Oryza sativa. The ancestral O. sativa
genome appears to have had a pair of duplicates termed DOPPELGANGER1
(DPL1) and DOPPELGANGER2 (DPL2) (Mizuta et al. 2010). The subspecies
japonica and indica have experienced independent losses of one copy each: DPL1
has become a pseudogene in indica, while DPL2 has been nonfunctionalized in
japonica. Hybrid pollen lacking a functional copy of either DPL1 or DPL2 is
nonfunctional and does not germinate, contributing to the partial reproductive
isolation present between the subspecies. This validates an earlier hypothesis by
Oka (1988) that F1 sterility between japonica and indica was caused by ‘‘duplicate
gametophytic sterility genes’’, japonica being homozygous for one nonfunctional
copy and indica being homozygous for another nonfunctional copy. In the second
rice example, reciprocal loss of one of the duplicated nuclear genes encoding
mitochondrial ribosomal protein L27 in O. sativa and O. glumaepatula again
causes a proportion of the pollen produced by F1 hybrids to be sterile (Yamagata
et al. 2010).
The final example of reproductive isolation through reciprocal gene loss comes
from A. thaliana, where the histidinol-phosphate amino-transferase gene appears
in different chromosomal locations (as in the Drosophila example, presumably via
duplication and subsequent silencing of one copy) in the Columbia and Cape
Verde Island accessions (Bikard et al. 2009). F2 offspring homozygous for both
null alleles completely lack the gene’s product, HPA, which results in arrested
embryo development and seed abortion. In addition, in at least one intermediate
heterozygote, a quantitative phenotype termed ‘‘weak root’’ was observed,
suggesting that the presence of three null alleles is somewhat deleterious in this
cross. As these four examples constitute *1/3 of the dozen or so successful
searches for the genes underlying the speciation process (most in Drosophila
species; Presgraves 2010), there now seems little question that the passive nonfunctionalization of duplicate genes is a major mechanism of speciation.
EC
372
CO
RR
371
UN
Editor Proof
1 Evolutionary Significance of Whole-Genome Duplication
Layout: T1 Standard SC
Chapter No.: 1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 12/20
C. L. McGrath and M. Lynch
439
1.6 Unsolved Problems
422
423
424
425
426
427
428
429
430
431
432
433
434
435
436
437
440
441
442
443
444
445
446
447
448
449
450
451
452
453
454
455
PR
OO
420
421
D
419
TE
418
EC
417
The maintenance of duplicate genes via selection for increased gene product,
neofunctionalization, and subfunctionalization has been hypothesized for nearly
40 years (Ohno 1970). Recent genetic and genomic data have now identified
compelling examples of these processes and have further contributed to our
understanding of the prevalence of whole-genome duplications and the dosagebalance theory of duplicate maintenance. However, a number of unresolved
questions related to WGDs and duplicate maintenance merit further scrutiny.
The first avenue for future study involves a more comprehensive understanding
of the relative importance of the forces behind duplicate-gene maintenance,
including maintenance for increased dosage, dosage-balance constraints,
neofunctionalization, and subfunctionalization. All of these mechanisms have been
demonstrated to be responsible for duplicate maintenance in certain cases, but it
remains unclear which, if any, is responsible for maintaining the majority of
duplicate genes or how such contributions vary among phylogenetic lineages. Most
likely, there will be no single driving force for duplicate maintenance but the relative
strength of these forces will differ among taxonomic groups or among functional
CO
RR
416
F
438
These examples demonstrate that the divergent resolution of even one duplicated gene can lead to detectable reproductive isolation. However, genetic
incompatibility between two populations can be magnified substantially when
reciprocal gene loss occurs at hundreds or thousands of duplicated loci simultaneously, as is the case in polyploid lineages. The probability that an F2 offspring
obtained by outcrossing will be double null for at least one of n pairs of divergently resolved loci is 1-(15/16)n, which takes on values of 0.063, 0.276, 0.476,
and 0.998 for n = 1, 5, 10, and 100, respectively. Moreover, in species that
undergo autogamy or selfing, such as Paramecium, this probability can be as high
as 1-(3/4)n, giving probabilities of 0.250, 0.763, 0.944, and & 1 for n = 1, 5, 10,
and 100. Speciation events will continue to occur as long as duplicates are still
being resolved between subpopulations, leading to nested rounds of speciation,
and, because a large number of duplicates are thought to be silenced quickly
following WGD (Scannell et al. 2006), a cluster of speciation events might occur
within a brief period of time. The net result is the expected generation of a species
radiation following a WGD event.
It has been suggested that this nested speciation process might be responsible
for the radiations of the polyploid yeast species (Scannell et al. 2006), teleost fishes
(Semon and Wolfe 2007; see Chap. 15, this volume), angiosperms (Soltis et al.
2009; though see Mayrose et al. 2011), and the Paramecium aurelia species
complex (Aury et al. 2006). This mechanism may also be responsible for
reproductive isolation between mutagenized lines of an experimentally derived
allotetraploid created by hybridizing two species of Saccharomyces (Maclean and
Greig 2010).
415
UN
Editor Proof
12
Layout: T1 Standard SC
Chapter No.: 1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 13/20
463
464
465
466
467
468
469
470
471
472
473
474
475
476
477
478
479
480
481
482
483
484
485
486
487
488
489
490
491
492
493
494
495
496
497
498
499
F
462
PR
OO
461
D
459
460
TE
458
13
classes of genes. For example, subfunctionalization of duplicates may be more
likely within species that have evolved a modular (and therefore independently
mutable) regulatory structure. Such modular systems are predicted to arise more
easily within species with smaller population sizes (Force et al. 2005), demonstrating how species-level features of an organism may influence the evolutionary
forces acting upon duplicate genes. More comprehensive studies of large numbers
of duplicates from a variety of organisms are required to address what other features
might influence the relative strengths of mechanisms of duplicate maintenance.
Such studies must not only detail what genes remain duplicated vs. single-copy, but
must also detail whether existing duplicates have the same function as each other
(to assess rates of neofunctionalization) or share functions with the pre-duplicated
ancestor (to assess rates of subfunctionalization) (Fig. 1.2).
There are few data on the rate of duplicate-gene loss over time following a
WGD, though data from polyploid yeast species suggest that the rate changes over
time (Scannell et al. 2006). Data from additional taxa would aid in determining
whether this is a general pattern among all WGDs (Fig. 1.3). A related unresolved
question is whether the evolutionary forces controlling duplicate maintenance
change over time following a WGD, e.g., whether a dosage-sensitive gene may
initially be preserved due to selection for dosage balance, but then evolve a new
function concurrent with its release from such dosage constraints. An analysis of
this question could be made by comparing the fates of duplicate genes in multiple
lineages descended from a single WGD event. Such an analysis might identify
duplicates that had been maintained due to dosage constraints in the majority of
daughter lineages but that had become neofunctionalized in one lineage, perhaps
suggesting a secondary mechanism of retention.
The unsolved question that promises to be the hardest to answer is why certain
lineages or taxonomic groups appear to contain more WGD events than others.
This is not the same as asking why certain groups contain more polyploid species,
as this may be a simple reflection of the fact that WGD may promote subsequent
reproductive isolation and speciation. The pattern remains, however, that some
phylogenetic groups seem to contain more independent WGD events than others in
their evolutionary pasts. For example, a recent analysis estimated that among
ferns, 31 % of speciation events involve polyploidization, while the value for
angiosperms is only 15 % (Wood et al. 2009). Similarly, in the history of the
Xenopus lineage, there are many more instances of WGD events than compared to,
say, mammals. Several factors could contribute to such patterns, such as the ability
to hybridize and form allopolyploids, or the ability to self-fertilize (at least transiently) or undergo asexual reproduction, which helps a polyploid lineage become
abundant in a surrounding world of diploids. It is not even understood whether
mechanistic reasons (at meiosis, say) or differences in developmental programs
would facilitate or hinder creation of a viable polyploid in certain lineages, or
whether discrepancies in ecological persistence of polyploid species alone are able
to explain the patterns that we see. Perhaps the best way to approach such a
question is to study closely related lineages where one exhibits several WGD
EC
457
CO
RR
456
UN
Editor Proof
1 Evolutionary Significance of Whole-Genome Duplication
Layout: T1 Standard SC
Chapter No.: 1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 14/20
14
C. L. McGrath and M. Lynch
Expression Level
PR
OO
F
A
Conditions/Tissues
B
or
D
Editor Proof
Fig. 1.2 Distinguishing
between the evolutionary
forces that maintain duplicate
genes. Panel A shows the
expression level of a gene
(purple) across different
conditions or tissues before
duplication. The bottom six
panels (B–G) show patterns
that might be seen for the two
copies (red and blue) once the
gene has been duplicated and
what evolutionary processes
these patterns would indicate.
Note that panel D might
indicate maintenance for
increased dosage in the case
of a single-gene duplicate or
maintenance for dosage
balance in the case of a
duplicate arising via WGD
TE
Nonfunctionalization
EC
D
500
501
UN
CO
RR
Dosage/dosage balance
F
Qualitative
Subfunctionalization
C
Nonfunctionalization
E
Neofunctionalization
G
Quantitative
Subfunctionalization
events and the other does not, though teasing apart the mechanistic and ecological
differences is certain to remain a challenge for decades to come.
Layout: T1 Standard SC
Chapter No.: 1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 15/20
1 Evolutionary Significance of Whole-Genome Duplication
(a)
D
C
B
Outgroup
A
D
% of duplicates lost
(b)
505
506
507
508
509
510
511
512
513
514
515
516
517
518
EC
504
F
Species 4
B
C
D
Time
Whole-genome duplications are widespread across the tree of life and appear in
the evolutionary history of a large number of model organisms. Processes such as
neo- and subfunctionalization affect retention of individual gene duplicates, and
dosage-balance constraints promote the retention of large sets of genes following
polyploidization. Allopolyploidization, through hybridization and subsequent
changes or biases in homeolog expression, has the ability to instantaneously create
a population of individuals that are ecologically and epigenetically unique from
either parent lineage, providing a new lineage upon which natural selection can
act. Both allo- and autopolyploidization provide a unique opportunity for the
differentiation of new gene networks and pathways through concerted evolution of
duplicated, interacting proteins. Most importantly, WGD can lead to reproductive
isolation through divergent resolution of duplicated genes, thus creating new
species and species groups. Further understanding of the relative importance and
the temporal properties of the forces acting on polyploid species and the duplicate
genes within their genomes promises to enhance our knowledge of the origins of
species as well as genetic, protein network, and organismal complexity.
CO
RR
503
1.7 Conclusions
UN
502
Species 3
PR
OO
WGD
Species 1
Species 2
A
TE
Editor Proof
Fig. 1.3 Determining the
rate of duplicate-gene loss
over time. a An example of a
tree for four species that share
a whole-genome duplication
(WGD). Duplicate-gene
presence/absence information
for each of the four species
could be used to infer the
number of duplicate genes
lost on each of the branches
labeled A, B, C, and D. b The
data on gene retention/loss
gathered in A could be used to
plot the percentage of
duplicate genes lost per unit
time (or divergence).
Depending on where points
A, B, C, or D land on the
graph, the data may indicate
that the rate of gene loss
remains constant (top dashed
line) or changes (bottom
dotted line) over time
15
Layout: T1 Standard SC
Chapter No.: 1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 16/20
C. L. McGrath and M. Lynch
References
520
521
522
523
524
525
526
527
528
529
530
531
532
533
534
535
536
537
538
539
540
541
542
543
544
545
546
547
548
549
550
551
552
553
554
555
556
557
558
559
560
561
562
563
564
565
566
567
568
569
570
Adams KL (2007) Evolution of duplicate gene expression in polyploid and hybrid plants. J Hered
98(2):136–141
Adams KL, Cronn R et al (2003) Genes duplicated by polyploidy show unequal contributions to
the transcriptome and organ-specific reciprocal silencing. Proc Nat Acad Sci U S A 100:4649–
4654
Adams KL, Liu Z (2007) Expression partitioning between genes duplicated by polyploidy under
abiotic stress and during organ development. Curr Biol 17(19):1669–1674
Altschmied J, Delfgaauw J et al (2002) Subfunctionalization of duplicate mitf genes associated
with differential degeneration of alternative exons in fish. Genetics 161(1):259–267
Anderson DW, Evans BJ (2009) Regulatory evolution of a duplicated heterodimer across species
and tissues of allopolyploid clawed frogs (Xenopus). J Mol Evol 68:236–247
Aury J-M, Jaillon O et al (2006) Global trends of whole-genome duplications revealed by the
ciliate Paramecium tetraurelia. Nature 444:171–178
Baack EJ (2005) To succeed globally, disperse locally: effects of local pollen and seed dispersal
on tetraploid establishment. Heredity 94:538–546
Barker MS, Kane NC et al (2008) Multiple paleopolyploidizations during the evolution of the
compositae reveal parallel patterns of duplicate gene retention after millions of years. Mol
Biol Evol 25(11):2445–2455
Bikard D, Patel D et al (2009) Divergent evolution of duplicate genes leads to genetic
incompatibilities within A. thaliana. Science 323:623–626
Birchler JA, Riddle NC et al (2005) Dosage balance in gene regulation: biological implications.
Trends Genet 21(4):219–226
Blanc G, Wolfe KH (2004) Functional divergence of duplicated genes formed by polyploidy
during Arabidopsis evolution. Plant Cell 16:1679–1691
Bottley A, Xia GM et al (2006) Homoeologous gene silencing in hexaploid wheat. Plant J
47:897–906
Brown MT, Goldstone HMH et al (2007) A functionally divergent hydrogenosomal peptidase
with protomitochondrial ancestry. Mol Microbiol 64(5):1154–1163
Buggs RJA, Elliott NM et al (2010) Tissue-specific silencing of homoeologs in natural
populations of the recent allopolyploid Tragopogon mirus. New Phytol 186:175–183
Byrne KP, Wolfe KH (2007) Consistent patterns of rate asymmetry and gene loss indicate
widespread neofunctionalization of yeast genes after whole-genome duplication. Genetics
175(3):1341–1350
Colon M, Hernandez F et al (2011) Saccharomyces cerevisiae Bat1 and Bat2 aminotransferases
have functionally diverged from the ancestral-like Kluyveromyces lactis orthologous enzyme.
PLoS ONE 6(1):e16099
Conant GC, Wolfe KH (2006) Functional partitioning of yeast co-expression networks after
genome duplication. PLoS Biol 4(4):0545–0554
Conant GC, Wolfe KH (2007) Increased glycolytic flux as an outcome of whole-genome
duplication in yeast. Mol Syst Biol 3:129
Coyne JA, Orr HA (2004) Speciation. Sinauer Associates, Inc, Sunderland, MA
Davis JC, Petrov DA (2005) Do disparate mechanisms of duplication add similar genes to the
genome? Trends Genet 21(10):548–551
Deng C, Cheng CHC et al (2010) Evolution of an antifreeze protein by neofunctionalization
under escape from adaptive conflict. Proc Nat Acad Sci U S A 107(50):21593–21598
Des Marais DL, Rausher MD (2008) Escape from adaptive conflict after duplication in an
anthocyanin pathway gene. Nature 454(7205):762–765
Doyle JJ, Flagel LE et al (2008) Evolutionary genetics of genome merger and doubling in plants.
Annu Rev Genet 42:443–461
Edger PP, Pires JC (2009) Gene and genome duplications: the impact of dosage-sensitivity on the
fate of nuclear genes. Chromosome Res 17:699–717
CO
RR
EC
TE
D
PR
OO
F
519
UN
Editor Proof
16
Layout: T1 Standard SC
Chapter No.: 1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 17/20
17
EC
TE
D
PR
OO
F
Erdmann R, Gramzow L et al (2010) GORDITA (AGL63) is a young paralog of the Arabidopsis
thaliana B (sister) MADS box gene ABS (TT16) that has undergone neofunctionalization.
Plant J 63(6):914–924
Evans BJ (2008) Genome evolution and speciation genetics of clawed frogs (Xenopus and
Silurana). Front Biosci 13:4687–4706
Evlampiev K, Isambert H (2007) Modeling protein network evolution under genome duplication
and domain shuffling. BMC Syst Biol 1:49
Flagel LE, Udall JA et al (2008) Duplicate gene expression in allopolyploid gossypium reveals
two temporally distinct phases of expression evolution. BMC Biol 6:16
Force A, Cresko WA et al (2005) The origin of subfunctions and modular gene regulation.
Genetics 170(1):433–446
Force A, Lynch M et al (1999) The preservation of duplicate genes by complementary,
degenerative mutations. Genetics 151:1531–1545
Froyd CA, Rusche LN (2011) The duplicated deacetylases sir2 and hst1 subfunctionalized by
acquiring complementary inactivating mutations. Mol Cell Biol 31(16):3351–3365
Gaeta RT, Pires JC et al (2007) Genomic changes in resynthesized brassica napus and their effect
on gene expression and phenotype. Plant Cell 19(11):3403–3417
Gout J-F, Duret L, et al. (2009) Differential retention of metabolic genes following wholegenome duplication. Mol Biol Evol 26(5):1067–1072
Gout J-F, Kahn D, et al. (2010) The relationship among gene expression, the evolution of gene
dosage, and the rate of protein evolution. PLoS Genetics 6(5):e1000944
Han MV, Demuth JP et al (2009) Adaptive evolution of young gene duplicates in mammals.
Genome Res 19(5):859–867
Hegarty MJ, Barker GL et al (2006) Transcriptome shock after interspecific hybridization in
Senecio is ameliorated by genome duplication. Curr Biol 16:1652–1659
Hickman MA, Rusche LN (2010) Transcriptional silencing functions of the yeast protein Orc1/
Sir3 subfunctionalized after gene duplication. Proc Nat Acad Sci U S A 107(45):19384–19389
Hughes AL (1994) The evolution of functionally novel proteins after gene duplication. Proc R
Soc Lond B Biol Sci 256:119–124
Hughes T, Liberles DA (2008) Whole-genome duplications in the ancestral vertebrate are
detectable in the distribution of gene family sizes of tetrapod species. J Mol Evol 67(4):343–357
Innan H, Kondrashov F (2010) The evolution of gene duplications: classifying and distinguishing
between models. Nat Rev Genet 11:97–108
Jiao YN, Wickett NJ et al (2011) Ancestral polyploidy in seed plants and angiosperms. Nature
473(7345):97–100
Jimenez-Sanchez G, Childs B et al (2001) Human disease genes. Nature 409:853–855
Johnson DA, Thomas MA (2007) The monosaccharide transporter gene family in Arabidopsis
and rice: a history of duplications, adaptive evolution, and functional divergence. Mol Biol
Evol 24(11):2412–2423
Josefsson C, Dilkes B et al (2006) Parent-dependent loss of gene silencing during interspecies
hybridization. Curr Biol 16:1322–1328
Kanno A, Saeki H et al (2003) Heterotopic expression of class B floral homeotic genes supports a
modified ABC model for tulip (Tulipa gesneriana). Plant Mol Biol 52:831–841
Layeghifard M, Pirhaji L et al (2009) Adaptive evolution in the Per gene family of vertebrates:
neofunctionalization by positive Darwinian selection after two major gene duplications. Biol
Rhythm Res 40(6):433–444
Liang H, Plazonic KR et al (2008) Protein under-wrapping causes dosage sensitivity and
decreases gene duplicability. PLoS Genet 4(1):0072–0077
Lipinski KJ, Farslow JC et al (2011) High spontaneous rate of gene duplication in Caenorhabditis
elegans. Curr Biol 21:306–310
Lolkema JS, Dobrowolski A et al (2008) Evolution of antiparallel two-domain membrane proteins:
tracing multiple gene duplication events in the DUF606 family. J Mol Biol 378:596–606
Lynch M (2007) The origins of genome architecture. Sinauer Associates, Sunderland, MA
CO
RR
571
572
573
574
575
576
577
578
579
580
581
582
583
584
585
586
587
588
589
590
591
592
593
594
595
596
597
598
599
600
601
602
603
604
605
606
607
608
609
610
611
612
613
614
615
616
617
618
619
620
621
622
623
UN
Editor Proof
1 Evolutionary Significance of Whole-Genome Duplication
Layout: T1 Standard SC
Chapter No.: 1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 18/20
EC
TE
D
PR
OO
F
Lynch M, Conery JS (2000) The evolutionary fate and consequences of duplicate genes. Science
290(5494):1151–1155
Lynch, M, Conery JS (2003a). The evolutionary demography of duplicate genes. In: Meyer A,
Van de Peer Y (eds) Genome evolution. Kluwer Academic Publishers, Dordrecht, 35–44
Lynch M, Conery JS (2003) The origins of genome complexity. Science 302:1401–1404
Lynch M, Force AG (2000) The origin of interspecific genomic incompatibility via gene
duplication. Am Naturaliste 156(6):590–605
Lynch M, Katju V (2004) The altered evolutionary trajectories of gene duplicates. Trends Genet
20:544–549
Lynch M, O’Hely M et al (2001) The probability of preservation of a newly arisen gene
duplicate. Genetics 159:1789–1804
Maclean CJ, Greig D (2010) Reciprocal gene loss following experimental whole-genome
duplication causes reproductive isolation in yeast. Evolution 65(4):932–945
MacNeil AJ, McEachern LA et al (2008) Gene duplication in early vertebrates results in tissuespecific subfunctionalized adaptor proteins: CASP and GRASP. J Mol Evol 67(2):168–178
Maere S, De Bodt S et al (2005) Modeling gene and genome duplications in eukaryotes. Proc Nat
Acad Sci U S A 102(15):5454–5459
Makino T, McLysaght A (2010) Ohnologs in the human genome are dosage balanced and
frequently associated with disease. Proc Nat Acad Sci U S A 107:9270–9274
Masly JP, Jones CD et al (2006) Gene transposition as a cause of hybrid sterility in Drosophila.
Science 313:1448–1450
Mayrose I, Zhan SH et al (2011) Recently formed polyploid plants diversify at lower rates.
Science 333(6047):1257
Meyer A, Van de Peer Y (2005) From 2R to 3R: evidence for a fish-specific genome duplication
(FSGD). BioEssays 27:937–945
Mizuta Y, Harushima Y et al (2010) Rice pollen hybrid incompatibility caused by reciprocal gene
loss of duplicated genes. Proc Nat Acad Sci U S A 107(47):20417–20422
Morin RD, Chang E et al (2006) Sequencing and analysis of 10,967 full-length cDNA clones
from Xenopus laevis and Xenopus tropicalis reveals post-tetraploidization transcriptome
remodeling. Genome Res 16:796–803
Ngai J, Dowling MM et al (1993) The family of genes encoding odorant receptors in the channel
catfish. Cell 72:657–666
Ohno S (1970) Evolution by gene duplication. Springer, Berlin
Oka HI (1988). Functions and genetic bases of reproductive barriers. Origin of cultivated rice.
Japan Scientific Societies Press/Elsevier, HI Oka, Tokyo, pp 181–209
Otto SP (2007) The evolutionary consequences of polyploidy. Cell 131:452–462
Panopoulou G, Poustka AJ (2005) Timing and mechanism of ancient vertebrate genome
duplications—the adventure of a hypothesis. Trends Genet 21:559–567
Papp B, Pal C et al (2003) Dosage sensitivity and the evolution of gene families in yeast. Nature
424:194–197
Paterson AH, Chapman BA et al (2006) Many gene and domain families have convergent fates
following independent whole-genome duplication events in Arabidopsis, Oryza, Saccharomyces and Tetraodon. Trends Genet 22(11):597–602
Piatigorsky J, Wistow G (1991) The recruitment of crystallins—new functions precede gene
duplication. Science 252(5009):1078–1079
Pinhal D, Yoshimura TS et al (2011) The 5S rDNA family evolves through concerted and birthand-death evolution in fish genomes: an example from freshwater stingrays. BMC Evol Biol
11:151
Pires JC, Gaeta RT (2011) Structural and functional evolution of resynthesized polyploids. In:
Schmidt R, Bandcroft I (eds) Genetics and genomics of the brassicaceae, Springer, New York,
9:195–214
Postlethwait JH, Woods IG et al (2000) Zebrafish comparative genomics and the origins of
vertebrate chromosomes. Genome Res 10:1890–1902
CO
RR
624
625
626
627
628
629
630
631
632
633
634
635
636
637
638
639
640
641
642
643
644
645
646
647
648
649
650
651
652
653
654
655
656
657
658
659
660
661
662
663
664
665
666
667
668
669
670
671
672
673
674
675
676
C. L. McGrath and M. Lynch
UN
Editor Proof
18
Layout: T1 Standard SC
Chapter No.: 1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 19/20
19
EC
TE
D
PR
OO
F
Presgraves DC (2010) The molecular evolutionary basis of species formation. Nat Rev Genet
11:175–180
Putnam NH, Butts T et al (2008) The amphioxus genome and the evolution of the chordate
karyotype. Nature 453:1064–1071
Qian W, Zhang J (2008) Gene dosage and gene duplicability. Genetics 179:2319–2324
Qian WF, Liao BY et al (2010) Maintenance of duplicate genes and their functional redundancy
by reduced expression. Trends Genet 26(10):425–430
Ramsey J, Schemske DW (2002) Neopolyploidy in flowering plants. Annu Rev Ecol Syst
33:589–639
Rapp RA, Udall JA et al (2009) Genomic expression dominance in allopolyploids. BMC Biol
7:18
Rausch JH, Morgan MT (2005) The effect of self-fertilization, inbreeding depression, and
population size on autopolyploid establishment. Evolution 59(9):1867–1875
Rodriguez DJ (1996) A model for the establishment of polyploidy in plants. Am Naturalist
147(1):33–46
Salmon A, Ainouche ML et al (2005) Genetic and epigenetic consequences of recent
hybridization and polyploidy in Spartina (Poaceae). Mol Ecol 14:1163–1175
Scannell DR, Byrne KP et al (2006) Multiple rounds of speciation associated with reciprocal gene
loss in polyploid yeasts. Nature 440:341–345
Scannell DR, Frank AC et al (2007) Independent sorting-out of thousands of duplicated gene
pairs in two yeast species descended from a whole-genome duplication. Proc Nat Acad Sci U
S A 104(20):8397–8402
Scannell DR, Wolfe KH (2008) A burst of protein sequence evolution and a prolonged period of
asymmetric evolution follow gene duplication in yeast. Genome Res 18(1):137–147
Semon M, Wolfe KH (2007) Reciprocal gene loss between Tetraodon and zebrafish after whole
genome duplication in their ancestor. Trends Genet 23(3):108–112
Semon M, Wolfe KH (2008) Preferential subfunctionalization of slow-evolving genes after
allopolyploidization in Xenopus laevis. Proc Nat Acad Sci U S A 105(24):8333–8338
Simillion C, Vandepoele K et al (2002) The hidden duplication past of Arabidopsis thaliana. Proc
Nat Acad Sci U S A 99:13627–13632
Sinha SC, Wetterer M et al (2005) Origin of asymmetry in adenylyl cyclases: structures of
Mycobacterium tuberculosis Rv1900c. EMBO J 24:663–673
Soltis DE, Albert VA et al (2009) Polyploidy and angiosperm diversification. Am J Bot
96(1):336–348
Soltis PS, Soltis DE et al (2006) Expression of floral regulators in basal angiosperms and the
origin and evolution of ABC-function. Adv Bot Res 44:483–506
Stoltzfus A (1999) On the possibility of constructive neutral evolution. J Mol Evol 49:169–181
Surcel A, Zhou X et al (2008) Long-term maintenance of stable copy number in the eukaryotic
SMC family: origin of a vertebrate meiotic SMC1 and fate of recent segmental duplicates.
J Syst Evol 46(3):405–423
Tate JA, Ni Z et al (2006) Evolution and expression of homeologous loci in Tragopogon
miscellus (Asteraceae), a recent and reciprocally formed allopolyploid. Genetics 173:1599–
1611
Taylor JS, Raes J (2004) Duplication and divergence: The evolution of new genes and old ideas.
Annu Rev Genet 38:615–643
Thomas BC, Pedersen B et al (2006) Following tetraploidy in an Arabidopsis ancestor, genes
were removed preferentially from one homeolog leaving clusters enriched in dose-sensitive
genes. Genome Res 16:934–946
Tirosh I, Barkai N (2007) Comparative analysis indicates regulatory neofunctionalization of yeast
duplicates. Genome Biol 8(4):R50
Udall JA, Swanson JM et al (2006) A novel approach for characterizing expression levels of
genes duplicated by polyploidy. Genetics 173:1823–1827
Veitia RA (2002) Exploring the etiology of haploinsufficiency. BioEssays 24(2):175–184
CO
RR
677
678
679
680
681
682
683
684
685
686
687
688
689
690
691
692
693
694
695
696
697
698
699
700
701
702
703
704
705
706
707
708
709
710
711
712
713
714
715
716
717
718
719
720
721
722
723
724
725
726
727
728
729
UN
Editor Proof
1 Evolutionary Significance of Whole-Genome Duplication
Layout: T1 Standard SC
Chapter No.: 1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 20/20
EC
TE
D
PR
OO
F
Veitia RA, Bottani S et al (2008) Cellular reactions to gene dosage imbalance: genomic,
transcriptomic, and proteomic effects. Trends Genet 24(8):390–397
Wang J, Tian L et al (2006) Genomewide nonadditive gene regulation in Arabidopsis
allotetraploids. Genetics 172:507–517
Watterson GA (1983) On the time for gene silencing at duplicate loci. Genetics 105(3):745–766
Werth CR, Windham MD (1991) A model for divergent, allopatric speciation of polyploid
Pteridophytes resulting from silencing of duplicate-gene expression. Am Naturalist
137(4):515–526
Winter KU, Weiser C et al (2002) Evolution of class B floral homeotic proteins: obligate
heterodimerization orginated from homodimerization. Mol Biol Evol 19(5):587–596
Wolfe KH, Shields DC (1997) Molecular evidence for an ancient duplication of the entire yeast
genome. Nature 387:708–713
Wood TE, Takebayashi N et al (2009) The frequency of polyploid speciation in vascular plants.
Proc Nat Acad Sci US A 106(33):13875–13879
Woolfe A, Elgar G (2007) Comparative genomics using Fugu reveals insights into regulatory
subfunctionalization. Genome Biol 8(4):R53
Yamagata Y, Yamamoto E et al (2010) Mitochondrial gene in the nuclear genome induces
reproductive barrier in rice. Proc Nat Acad Sci U S A 107(4):1494–1499
Yang J, Lusk R et al (2003) Organismal complexity, protein complexity, and gene duplicability.
Proc Nat Acad Sci U S A 100(26):15661–15665
Yu W-P, Brenner S et al (2003) Duplication, degeneration and subfunctionalization of the nested
synapsin-Timp genes in Fugu. Trends Genet 19:180–183
CO
RR
730
731
732
733
734
735
736
737
738
739
740
741
742
743
744
745
746
747
748
749
750
751
C. L. McGrath and M. Lynch
UN
Editor Proof
20
Metadata of the chapter that will be visualized in
SpringerLink
Book Title
Polyploidy and Genome Evolution
Series Title
Chapter Title
Genetic Consequences of Polyploidy in Plants
Copyright Year
2012
Copyright HolderName
Springer-Verlag Berlin Heidelberg
Corresponding Author
Family Name
Birchler
Particle
Given Name
James A.
Suffix
Abstract
Division
Division of Biological Sciences
Organization
University of Missouri
Address
117 Tucker Hall, 65211, Missouri, Colombia
Email
birchlerj@missouri.edu
Most eukaryotes have a history of whole-genome multiplication events followed by a progressive return to
a more diploid state. The initial state of polyploidization, in which more than two copies of the genome are
present, is considered here and the various types of genetic consequences that occur depending on the nature
of the polyploid formed. The degree of association of chromosomes in meiosis is determined by the relative
homology and will affect the segregation of the chromosome which determines the genetic properties. If all
the chromosomes are quite similar and form associations of like type, this situation is referred to as
autopolyploidy. If the different sets of multiple chromosomes are sufficiently dissimilar to each other, then
the homologs will pair in meiosis with themselves and segregate independently of the different but related
chromosome pair. This situation is referred to as allopolyploidy. Gene expression in ploidal series typically
follows a per cell level correlated more or less with the number of sets of chromosomes present. Variation of
individual chromosomes, or aneuploidy, produces a greater number of modulations of gene expression in
parallel to classical studies noting that aneuploids have greater impact on the phenotype than changes in the
copy number of the whole genome. The genetic properties of odd-number ploidies, such as triploids, are also
described as well as higher ploidal levels such as hexaploidy and octoploidy.
1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 21/32
Chapter 2
4
James A. Birchler
11
12
13
14
15
16
17
18
19
20
21
22
23
24
PR
OO
D
9
10
TE
8
EC
7
Abstract Most eukaryotes have a history of whole-genome multiplication events
followed by a progressive return to a more diploid state. The initial state of
polyploidization, in which more than two copies of the genome are present, is
considered here and the various types of genetic consequences that occur
depending on the nature of the polyploid formed. The degree of association of
chromosomes in meiosis is determined by the relative homology and will affect the
segregation of the chromosome which determines the genetic properties. If all the
chromosomes are quite similar and form associations of like type, this situation is
referred to as autopolyploidy. If the different sets of multiple chromosomes are
sufficiently dissimilar to each other, then the homologs will pair in meiosis with
themselves and segregate independently of the different but related chromosome
pair. This situation is referred to as allopolyploidy. Gene expression in ploidal
series typically follows a per cell level correlated more or less with the number of
sets of chromosomes present. Variation of individual chromosomes, or aneuploidy,
produces a greater number of modulations of gene expression in parallel to classical studies noting that aneuploids have greater impact on the phenotype than
changes in the copy number of the whole genome. The genetic properties of oddnumber ploidies, such as triploids, are also described as well as higher ploidal
levels such as hexaploidy and octoploidy.
CO
RR
5
6
F
3
Genetic Consequences of Polyploidy
in Plants
2
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: 2
J. A. Birchler (&)
Division of Biological Sciences, University of Missouri, 117 Tucker Hall,
65211 Missouri, Colombia
e-mail: birchlerj@missouri.edu
P. S. Soltis and D. E. Soltis (eds.), Polyploidy and Genome Evolution,
DOI: 10.1007/978-3-642-31442-1_2, Springer-Verlag Berlin Heidelberg 2012
21
Layout: T1 Standard SC
Chapter No.: 2
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 22/32
25
J. A. Birchler
2.1 Introduction
56
2.2 Allopolyploids
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
57
58
59
60
61
62
63
PR
OO
32
D
31
TE
29
30
EC
28
CO
RR
27
F
55
Most eukaryotes have a history of polyploidization followed by fractionation back
to a near diploid level (Wolfe and Shields 1997; Simillion et al. 2002; Bowers
et al. 2003; Blanc and Wolfe 2004; Chapman et al. 2006; Maere et al. 2005;
Blomme et al. 2006; Freeling and Thomas 2006; Barker et al. 2008). Thus, at the
least, polyploidy in essence is a matter of degree, and it has played an important
role in the composition of the gene repertoire of many species. Typically, it is
defined as the presence of more copies of the whole genome than the normal two
that constitute a diploid (Stebbins 1947). However, from the standpoint of gene
content, the determination of whether a species is a polyploid is somewhat arbitrary and dependent on the time before the present when the copy number of the
genome was increased. Nevertheless, for evolutionarily ‘‘recent’’ events, certain
principles can apply which will be summarized in this chapter.
In the polyploidy literature, the basic chromosome number is designated by
x and consists of the complete set of chromosomes, or a genome. The number of
chromosomes in the gametophyte generation and hence the gametes is referred to
as the gametic chromosome number or n. In diploids, x = n, but at higher levels of
polyploidy, this is not the case.
Polyploidy is typically divided into at least two categories that are determined
by the type of chromosome pairing in meiosis I and the distribution of chromosomes during this process. Indeed, the type of chromosome pairing that occurs in
meiosis affects the genetic properties of the species so such classifications have
value. If the increase in genome copy number results from the combination of
chromosome sets from divergent species, the different types of chromosomes will
usually not pair with each other in prophase of meiosis I. In the case of tetraploids,
if both divergent genomes are doubled by whatever means, those sets of chromosomes that are similar or identical will preferentially pair with each other to the
exclusion of the other genome. This type of pairing is referred to as ‘‘disomic’’ in
analogy with the situation in a diploid. A species with this type of scenario is
referred to as an allopolyploid because the contributing genomes are different from
each other.
26
Genetic ratios in an allotetraploid depend on the constitution of each genome
(Clausen and Goodspeed 1925; Clausen 1941; Clausen and Cameron 1944). The
different sets of related chromosomes are referred to as homoeologues. If both the
homoeologues possess the homoeologous gene copies that are expressed similarly,
then both would need to be mutant in order to express a recessive phenotype.
Under these circumstances, duplicate gene ratios would typically be observed. In
other words, recessive phenotypes would be found in 1/16 (1/4 9 1/4) of the F2
UN
Editor Proof
22
Layout: T1 Standard SC
Chapter No.: 2
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 23/32
23
70
2.3 Autopolyploids
73
74
75
76
77
78
79
80
81
82
83
84
85
86
87
88
89
90
91
92
93
94
95
96
97
98
99
100
101
102
103
104
PR
OO
72
If on the other hand the increase in genome copy number in a polyploid results
from the same species such that the chromosomes are all quite similar, the pairing
in prophase of meiosis I forms conglomerates that switch pairing partners along
the length of the chromosome (Fig. 2.1). This type of pairing is referred to as
‘‘quadrivalent’’ pairing because all four chromosomes present can be involved
with each other. However, 3:1 and 2:2 associations are also observed. The segregation in this case will depend on the position of the locus in question in the
chromosome and relative to the respective centromere (Blakeslee et al. 1923;
Haldane 1930; Bartlett and Haldane 1934; Mather 1935, 1936; Randolph 1935;
Little 1945, 1958; Doyle 1973). Those genes near the centromere will be distributed to the diploid gametes based on the usual case that pairs of centromeres
will separate from each other at meiosis I and that the two sets from each chromosome of the complement will do so at random. A homozygous dominant
autotetraploid (AAAA) is referred to as a quadruplex and the homozygous recessive
(aaaa) as a nulliplex. There are three types of heterozygotes: AAAa (triplex), AAaa
(duplex), and Aaaa (simplex). If one designates a hybrid autotetraploid as AA0 aa0 ,
then there are six types of possible gametes that will be formed: AA0 , Aa, Aa0 , aA,
a0 A0 , aa0 . The frequency of diploid homozygous gametes under these circumstances is 1/6 (0.167). A self-pollination will produce 2.77 % of the progeny that
are homozygous for the recessive allele (Fig. 2.2). However, as the distance of a
gene from the centromere increases, recombination between the locus and the
centromere will randomize the distribution of the different alleles into the diploid
gametes to the point that the frequency of homozygous diploid gametes will be
(4/8 9 3/7 = 0.21) as a maximum. In this case, a self-pollination will produce
4.41 % of the progeny that are homozygous.
Recombination between the monitored locus and the centromere can also
produce homozygous spores from a triplex heterozygote (AAAa) to produce aa
gametes (Catcheside 1956). This process is called double reduction. Again, this
result is affected by the position of the locus under consideration from the centromere with greater double reduction increasing with distance. Another factor
affecting segregation in autotetraploids is aneuploidy, i.e., altered copy number of
individual chromosomes. This circumstance would change the pairing and segregation properties of individual chromosomes. Autotetraploids can also generate
spontaneous diploid progeny via parthenogenesis (Randolph and Fischer 1939).
D
71
TE
67
68
EC
66
CO
RR
65
F
69
from a self of an F1 between parental types that are dominant and recessive.
However, if one of the gene copies is missing or expressed in other tissues from
one of the homoeologous chromosomes, then genetic ratios typical of a diploid
will be found because only one genome will have different alleles in an F1, and
they will segregate to produce a 3:1 ratio because the single genome will behave as
a diploid.
64
UN
Editor Proof
2 Genetic Consequences of Polyploidy in Plants
Layout: T1 Standard SC
Chapter No.: 2
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 24/32
J. A. Birchler
PR
OO
F
Editor Proof
24
Aa
Aa
Aa
TE
AA
D
Fig. 2.1 Comparison of meiotic anaphase I in matched diploid and tetraploid maize plants.
a Gray value image of anaphase I of diploid inbred line B73. Note that membrers of each pair of
homologs separate from each other. Modified from (Birchler 2011). b Gray value image of
anaphase I of a tetraploid derivative of inbred line B73. Note the multivalent associations of
chromosomes. Photos by Zhi Gao and Fangpu Han
Aa
aa
AA
AA
AA
Aa
AA
Aa
AA
Aa
AA
Aa
AA
aa
Aa
AA
Aa
Aa
Aa
Aa
Aa
Aa
Aa
Aa
Aa
Aa
aa
Aa
AA
Aa
Aa
Aa
Aa
Aa
Aa
Aa
Aa
Aa
Aa
aa
Aa
AA
Aa
Aa
Aa
Aa
Aa
Aa
Aa
Aa
Aa
Aa
aa
Aa
AA
Aa
Aa
Aa
Aa
Aa
Aa
Aa
Aa
Aa
Aa
aa
aa
AA
aa
aa
Aa
aa
Aa
aa
Aa
aa
Aa
aa
aa
CO
RR
EC
AA
UN
Fig. 2.2 Genotypes in a self-pollination of an autotetraploid heterozygote AAaa. In an
autotetraploid, the gametes are diploid. With two chromosomes carrying the dominant A allele
and two carrying the recessive a allele, the distribution of gametes from the heterozygote is
shown across the top and along the side. The combinations of these gametes to produce the
tetraploid progeny are shown in the grid. Only one out of 36 are homozygous for A, and one out
of 36 are homozygous for the recessive a. Other combinations of A and a are shown. These
conditions hold for genes closely linked to centromeres as described in the text
Layout: T1 Standard SC
Chapter No.: 2
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 25/32
105
25
2.4 Segmental Allopolyploids
118
2.5 Heterosis and Ploidy
113
114
115
116
119
120
121
122
123
124
125
126
127
128
129
130
131
132
133
134
135
136
137
138
139
140
141
142
143
PR
OO
112
D
111
Because of the chromosome pairing considerations noted above, allopolyploids
will have a diversity of gene products ‘‘fixed’’ in their genomic structure. This
circumstance will basically maintain the essence of hybrid vigor even though
technically an otherwise high degree of homozygosity might be present. Thus,
allopolyploids typically exhibit robust biomass and excellent fertility compared to
the diploid progenitor species. Nevertheless, crosses between different isolates of
allopolyploids can show even greater heterotic effects when each genome is heterozygous as well (Gustafson 1946). Autopolyploids, which can have up to four
different alleles at one locus, also exhibit hybrid vigor and with increasing
diversity of alleles present, a phenomenon known as progressive heterosis, there is
increasing biomass and fertility (Busbice and Wilsie 1966; Levings et al. 1967;
Mok and Peloquin 1975; Groose et al. 1989). However, autotetraploids can be
subject to inbreeding depression in which the potential exists for all copies of a
chromosome to become homozygous (Busbice and Wilsie 1966; Sockness and
Dudley 1989a, b). Polyploids that are entirely homozygous exhibit extreme
depression and reduction of stature and fertility (Busbice and Wilsie 1966; Riddle
et al. 2006; Abel and Becker 2007; Stupar et al. 2007; Redei 1964; d’Erfurth et al.
2009; Yao et al. 2011). This situation is unlikely under natural circumstances.
A discussion of the vigor of polyploids needs to consider the intersection with
heterosis or hybrid vigor. In recent years, it has been possible to produce ploidy
series for completely or highly homozygous materials (Riddle et al. 2006; Abel
and Becker 2007; Stupar et al. 2007; d’Erfurth et al. 2009; Yao et al. 2011). The
general rule that emerges from these studies is that with increasing ploidy and the
maintenance of homozygosity, there is usually a decline in stature and fertility.
The cell and pollen size increases with ploidy and the plants typically take on a
TE
109
110
EC
108
CO
RR
107
F
117
A third classification based upon empirical chromosome associations is segmental
allopolyploid (Stebbins 1947). In this case, some chromosomes exhibit bivalent
pairing and others show quadrivalent pairing. The basis of such behavior was not
clear until recently. Xiong and colleagues (Xiong et al. 2011) found that in
resynthesized Brassica napus derived from the diploid progenitors, B. oleracea and
B. rapa, different lineages could form compensating nullisomic-tetrasomic
configurations for different chromosomes. In this case, the tetraploid will have some
chromosomes that are basically identical and other members of the set that will be
divergent. Similar results were reported for naturally occurring tetraploid
Tragopogon miscellus (Chester et al. 2012). Such a species will be a composite of
allo- and autotetraploid chromosomes for different members of the karyotype and
would be expected to exhibit the pairing characterized by a segmental allopolyploid.
106
UN
Editor Proof
2 Genetic Consequences of Polyploidy in Plants
Layout: T1 Standard SC
Chapter No.: 2
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 26/32
J. A. Birchler
153
2.6 Aneuploidy Relative to Ploidy
151
154
155
156
157
158
159
160
161
162
163
164
165
166
167
168
169
170
171
172
173
174
175
176
177
178
179
180
181
182
183
PR
OO
150
In contrast to a ploidy series, changes in dosage of individual chromosomes (or
substantial parts of chromosomes) have a more dramatic effect on the phenotype
(Blakeslee et al. 1920; Blakeslee 1934). Typically, the removal of a chromosome
or chromosomal segment has the strongest effects and is lethal in some cases
(Kush and Rick 1968; Vizir and Mulligan 1999). All of the monosomics for each
of the ten chromosomes in maize have been recovered and studied (Weber 1983),
but this is not the case in other species in which this issue has been examined such
as tomato (Kush and Rick 1968) and Arabidopsis (Vizir and Mulligan 1999). The
addition of a chromosome to produce a trisomic usually also has a detrimental
effect on plant vigor but the usual circumstance is that the impact is much less than
monosomics (Lee et al. 1996). Indeed, full sets of trisomics have been produced
for many plant species (Singh 1993). Tetrasomics for whole chromosome arms,
otherwise called secondary trisomics, have been produced in Datura by recovery
of extra chromosomes that are duplicated for one or the other arm of the progenitor
chromosome (Blakeslee 1934). These secondary trisomics usually have more
intensified phenotypic effects and are more intensified when present in haploids
(Satina et al. 1937a, b).
Extra or missing chromosomes in higher ploidies have less severe phenotypic
effects. A comprehensive set of aneuploids was generated in hexaploid wheat
(Sears 1944; Sears 1953, 1954). Monosomics and trisomics are regularly produced, and because of the high ploidy state, nullisomics, which are missing both
copies of a chromosome, can be produced (Sears 1953, 1954). Nullisomics have a
more severe effect than the corresponding monosomic. Tetrasomics can be produced and have a more severe effect than the respective trisomic. Compensating
nullisomics for one homoeologue and tetrasomics for another return to a more
normal phenotype than exhibited by the nullisomic or tetrasomic alone (Sears
1953, 1954). Newly synthesized B. napus (Xiong et al. 2011) and natural neopolyploids of T. miscellus (Chester et al. 2012) will exhibit aneuploidy that
resolves into compensating 4:0 or 3:1 contributions from different progenitor
genomes illustrating that the compensating balanced condition is favored in
D
149
TE
148
EC
146
147
CO
RR
145
F
152
‘‘stocky’’ appearance but with extreme ploidies, the plants are depauperate
(Blakeslee 1941; Randolph 1942; Rhoades and Dempsey 1966; d’Erfurth et al. 2009;
Yao et al. 2011). In contrast, hybrids with increasing ploidy tend to exhibit greater
biomass and in the species in which it has been examined closely, there is an increase
in heterosis with increasing diversity of alleles, i.e., progressive heterosis (Busbice
and Wilsie 1966; Mok and Peloquin 1975; Levings et al. 1967; Groose et al. 1989;
Bingham et al. 1994; Riddle and Birchler 2008). The common view that polyploids
exhibit more robust stature derives from experience with allopolyploids or with
heterotic autopolyploids, which are the situations most commonly encountered.
144
UN
Editor Proof
26
Layout: T1 Standard SC
Chapter No.: 2
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 27/32
27
187
2.7 Gene Expression Studies
PR
OO
185
F
186
laboratory or natural selection. Together, these results further illustrate that the
greater the deviation from the standard set of chromosomes, the more severe the
impact on the phenotype.
184
205
2.8 Genomic Balance
192
193
194
195
196
197
198
199
200
201
202
203
206
207
208
209
210
211
212
213
214
215
216
217
218
219
TE
191
EC
190
CO
RR
189
D
204
Studies on gene expression in ploidy and aneuploid series parallel the phenotypic
results. When individual genes are sampled in a ploidy series, the expression level
is more or less proportional to the ploidal level, although there are examples of
genes whose expression deviates from this trend both positively and negatively
(Birchler and Newton 1981; Guo et al. 1996). Genome-wide studies of gene
expression in ploidy series demonstrate a similar pattern (Wang et al. 2004;
Albertin et al. 2005; Stupar et al. 2007; Riddle et al. 2010; Yu et al. 2010). In
contrast, sampling of individual genes or protein patterns in aneuploids reveals a
greater set of changes from the diploid level of expression (Birchler 1979; Birchler
and Newton 1981; Guo and Birchler 1994). A dosage series for a particular
chromosomal region would alter the amount of expression of a portion of the total
gene products encoded across the genome. The effects could be positive or negative correlations with the change in dosage. The more common effect especially
with trisomics was a negative correlation between the dosage and the target gene
expression (Birchler 1979; Birchler and Newton 1981; Guo and Birchler 1994).
Thus, the gene expression patterns show changes in a ploidy series but aneuploid
series exhibit greater effects in parallel with the phenotypic relationships.
188
This gene expression relationship led to the suggestion that the stoichiometry of
regulatory genes affected the outcome of gene expression (Birchler and Newton
1981) and ultimately the phenotype (Guo and Birchler 1994). Studies to identify
single genes that would mimic the aneuploid effects using a partial loss of function
mutation in the white eye color gene in Drosophila produced single-gene mutations that would modulate the target’s expression either positively or negatively
(Rabinow et al. 1991; Birchler et al. 2001). The molecular identification of many
of these genes revealed them to be transcription factors, chromatin modifiers, and
components of signal transduction (Birchler et al. 2001).
Interestingly, these same classes of genes are typical of those that exhibit
preferential retention following a polyploidization event (Blanc and Wolfe, 2004;
Freeling and Thomas, 2006) and underrepresentation in segmental duplications
(Maere et al. 2005; Freeling et al. 2008). Thus, this evidence suggests that if these
classes of genes are out of register with each other, there is a negative fitness
UN
Editor Proof
2 Genetic Consequences of Polyploidy in Plants
Layout: T1 Standard SC
Chapter No.: 2
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 28/32
J. A. Birchler
226
2.9 Triploids
222
223
224
PR
OO
221
F
225
consequence. Thus, the phenotypic, gene expression and evolutionary studies form
a coherent picture that these types of genes form a balance. When individual
components exhibit a dosage effect, this will ultimately produce a fitness consequence due to the impact of the altered gene expression on the phenotype (Birchler
et al. 2001; Veitia 2002; Veitia 2004; Birchler et al. 2005, 2007; Veitia et al. 2008;
Birchler and Veitia 2007, 2010).
220
240
2.10 Higher Ploidal Levels
231
232
233
234
235
236
237
238
241
242
243
244
245
246
247
248
249
250
251
252
253
254
TE
230
EC
229
Ploidal levels above the tetraploid level most commonly involve hexaploids and
octoploids although much higher levels have been documented. Chromosome
pairing in allohexaploids has been studied in detail, for example in wheat, which is
ordinarily disomic in nature (Kihara 1919; Lilienfeld 1951; Dvorak et al. 1988).
The Ph system insures pairing of homologs and against pairing of homoeologues
but homoeologues can pair in mutant plants (Yousafzai et al. 2010; see Chap. 7,
this volume). The wheat genome is composed of three different slightly diverged
genomes tracing back through the joining of an allotetraploid composed of two
genomes with the third. The Ph system maintains disomic pairing and hence
excellent fertility. Octoploids, using sugar cane as an example, have variable
chromosome numbers due to the minimal detrimental effects of aneuploidy at this
level (Piperidis et al. 2010). In contrast, triticale, which is an octoploid consisting
of hexaploid wheat with the addition of a rye genome, exhibits faithful chromosomes numbers.
CO
RR
228
D
239
Triploids are a polyploid level between diploid and tetraploid. They arise from
crosses between diploid and tetraploids of the same or related species or from
unreduced gametes from one diploid parent. In meiosis, the chromosomes associate in trivalents, which consists of pairing of any two chromosomes at any one
point (McClintock 1929; Punyasingh 1947; Upcott 1935). The distribution of
chromosomes is nearly random, resulting in spores that range from 1x to 2x. As a
consequence, the gametophytes are mostly highly aneuploid and in some cases
abort (Satina and Blakeslee 1937a, b). Fertilization involving gametes of different
chromosome numbers in the endosperm will often cause endosperm abortion
(Satina et al. 1938; Punyasingh 1947; Brink and Cooper 1947; Cooper 1951). The
gametes that are successful tend to be those at or near the 1x or 2x level. Because
of the variability of the chromosome numbers in gametes from triploid individuals,
this ploidal level is not stable.
227
UN
Editor Proof
28
Layout: T1 Standard SC
Chapter No.: 2
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 29/32
255
29
2.11 Concluding Remarks
270
Acknowledgments Research supported by National Science Foundation grant DBI 0733857.
271
References
272
273
274
275
276
277
278
279
280
281
282
283
284
285
286
287
288
289
290
291
292
293
294
295
296
297
Abel S, Becker HC (2007) The effect of autopolyploidy on biomass production in homozygous
lines of Brassica rapa and Brassica oleracea. Plant Breed 126:642–643
Albertin W, Brabant P, Catrice O, Eber F, Jenczewski E, Chèvre AM, Thiellement H (2005)
Autopolyploidy in cabbage (Brassica oleracea L.) does not alter significantly the proteomes
of green tissues. Proteomics 5:2131–2139
Barker MS, Kane NC, Matvienko M, Kozik A, Michelmore RW, Knapp SJ, Rieseberg LH (2008)
Multiple paleopolyploidizations during the evolution of the compositae reveal parallel
patterns of duplicate gene retention after millions of years. Mol Biol Evol 25:2445–2455
Bartlett MS, Haldane JBS (1934) The theory of inbreeding in Autotetraploids. J Genet 29:175–180
Bingham ET, Groose RW, Woodfield DR, Kidwell KK (1994) Complementary gene interactions
in alfalfa are greater in autotetraploids than diploids. Crop Sci 34:823–829
Birchler JA (1979) A study of enzyme activities in a dosage series of the long arm of
chromosome one in maize. Genetics 92:1211–1229
Birchler JA (2011) Epigenetic aspects of centromere function in plants. Curr Opin Plant Biol
14:217–222
Birchler JA, Newton KJ (1981) Modulation of protein levels in chromosomal dosage series of
maize: the biochemical basis of aneuploid syndromes. Genetics 99:247–266
Birchler JA, Bhadra U, Pal Bhadra M, Auger DL (2001) Dosage dependent gene regulation in
multicellular eukaryotes: implications for dosage compensation, aneuploid syndromes and
quantitative traits. Dev Biol 234:275–288
Birchler JA, Riddle NC, Auger DL, Veitia RA (2005) Dosage balance in gene regulation:
biological implications. Trends Genet 21:219–226
Birchler JA, Veitia RA (2007) The gene balance hypothesis: from classical genetics to modern
genomics. Plant Cell 19:395–402
Birchler JA, Veitia RA (2010) The gene balance hypothesis: implications for gene regulation,
quantitative traits and evolution. New Phytol 186:54–62
262
263
264
265
266
267
268
PR
OO
261
D
260
TE
259
EC
258
CO
RR
257
F
269
The genetics of polyploids depends essentially on the pairing properties of the
multiple chromosomes in meiosis. If the multiple copies of a genome are sufficiently dissimilar from each other, they tend to pair among themselves and
maintain the genetic variation within each genome. If the multiple copies of a
genome are similar to each other, then all copies are free to pair and recombine
among themselves. In this circumstance, the genetic behavior of a particular gene
is dependent on its position on the chromosome and the fidelity of the pairing of
homologs. Aneuploidy, i.e., the variation of a single chromosome or chromosomal
segment, can have more severe consequences than varying the whole genome.
However, as the background ploidy increases, the effect of the same chromosome
change of aneuploidy becomes less. The phenotypic effects, gene expression
patterns and the evolutionary results of differential gene retention following
whole-genome duplications versus segmental duplication suggest the importance
of genomic balance.
256
UN
Editor Proof
2 Genetic Consequences of Polyploidy in Plants
Layout: T1 Standard SC
Chapter No.: 2
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 30/32
EC
TE
D
PR
OO
F
Birchler JA, Yao H, Chudalayandi S (2007) Biological consequences of dosage dependent gene
regulatory systems. Biochem Biophys Acta-Gene Struct Expr 1769:422–428
Blakeslee AF (1934) New Jimson weeds from old chromosomes. J Hered 24:80–108
Blakeslee AF (1941) Effect of induced polyploidy in plants. Am Nat 75:117–135
Blakeslee AF, Belling J, Farnham ME (1920) Chromosomal duplication and Mendelian
phenomena in Datura mutants. Science 52:388–390
Blakeslee AF, Belling J, Farnham ME (1923) Inheritance in tetraploid Daturas. Bot Gaz
76:329–373
Blanc G, Wolfe KH (2004) Functional divergence of duplicated genes formed by polyploidy
during Arabidopsis evolution. Plant Cell 16:1679–1691
Blomme T, Vandepoele K, De Bodt S, Simillion C, Maere S, Van de Peer Y (2006) The gain and
loss of genes during 600 million years of vertebrate evolution. Genome Biol 7:43
Bowers JE, Chapman BA, Rong J, Paterson AH (2003) Unraveling angiosperm genome evolution
by phylogenetic analysis of chromosomal duplication events. Nature 422:433–438
Busbice TH, Wilsie CP (1966) Inbreeding depression and heterosis in autotetraploids with
application to Medicago sativa L. Euphytica 15:52–67
Brink RA, Cooper DC (1947) The endosperm in seed development. Bot Rev 13:423–541
Catcheside DG (1956) Double reduction and numerical non-disjunction in tetraploid maize.
Heredity 10:205–218
Chapman BA, Bowers JE, Feltus FA, Paterson AH (2006) Buffering of crucial functions by
paleologous duplicated genes may contribute cyclicality to angiosperm genome duplication.
Proc Natl Acad Sci USA 103:2730–2735
Chester M, Gallagher JP, Symonds VV, Cruz da Silva AV, Mavrodiev EV, Leitch AR, Soltis PS,
Soltis DE (2012) Extensive chromosomal variation in a recently formed natural allopolyploid,
Tragopogon miscellus (Asterasceae). Proc Natl Acad Sci USA (in press)
Clausen RE (1941) Polyploidy in Nicotiana. Am Nat 75:291–306
Clausen RE, Cameron DR (1944) Inheritance in Nicotiana tabacum. XVIII. Monosomic analysis.
Genetics 29:447–477
Clausen RE, Goodspeed TH (1925) Interspecific hybridization in Nicotiana II. A tetraploid
glutinosa-tabacum hybrid. An experimental verification of Winge’s hypothesis. Genetics
10:278–284
Cooper DC (1951) Caryopsis development following matings between diploid and tetraploid
strains of Zea mays. Am J Bot 38:702–708
Doyle GG (1973) Autotetraploid segregation. Theor Appl Genet 43:139–146
d’Erfurth I, Jolivet S, Froger N, Catrice O, Novatchkova M, Mercier R (2009) Turning meiosis
into mitosis. PLoS Biol 7:1000124
Dvorak J, McGuire PE, Cassidy B (1988) Apparent sources of the a genomes of wheats inferred
from the polymorphism in abundance and restriction fragment length of repeated nucleotide
sequences. Genome 30:680–689
Freeling M, Lyons E, Pedersen B, Alam M, Ming R, Lisch D (2008) Many or most genes in
Arabidopsis transposed after the origin of the order Brassicales. Genome Res 18:1924–1937
Freeling M, Thomas BC (2006) Gene-balanced duplications, like tetraploidy, provide predictable
drive to increase morphological complexity. Genome Res 16:805–814
Guo M, Birchler JA (1994) Trans-acting dosage effects on the expression of model gene systems
in maize aneuploids. Science 266:1999–2002
Guo M, Davis D, Birchler JA (1996) Dosage effects on gene expression in a maize ploidy series.
Genetics 142:1349–1355
Groose RW, Talbert LE, Kojis WP, Bingham ET (1989) Progressive heterosis in autotetraploid
alfalfa: studies using two types of inbreds. Crop Sci 29:1173–1177
Gustafson A (1946) The effect of heterozygosity on viability and vigor. Hereditas 32:263–286
Haldane JBS (1930) The theoretical genetics of autopolyploids. J Genet 22:359–372
Kihara H (1919) Uber zytologische Studien bei einigen Getreidearten. Bot Mag Tokyo 33
Kush GS, Rick CM (1968) Cytogenetic analysis of the tomato genome by means of induced
deficiencies. Chromosoma 23:452–484
CO
RR
298
299
300
301
302
303
304
305
306
307
308
309
310
311
312
313
314
315
316
317
318
319
320
321
322
323
324
325
326
327
328
329
330
331
332
333
334
335
336
337
338
339
340
341
342
343
344
345
346
347
348
349
350
351
J. A. Birchler
UN
Editor Proof
30
Layout: T1 Standard SC
Chapter No.: 2
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 31/32
31
EC
TE
D
PR
OO
F
Lee EA, Darrah LL, Coe EH (1996) Dosage effects on morphological and quantitative traits in
maize aneuploids. Genome 39:898–908
Levings CS, Dudley JW, Alexander DE (1967) Inbreeding and crossing in autotetraploid maize.
Crop Sci 7:72–73
Lilienfeld FA (1951) H. Kihara: Genome analysis in Triticum and Aegilops. Concluding review.
Cytologia 16:101–123
Little TM (1945) Gene segregation in autotetraploids. Bot Rev 11:60–85
Little TM (1958) Gene segregation in autotetraploids. II. Bot Rev 24:319–339
Maere S, DeBodt S, Raes J, Casneuf T, Van Montagu M, Kuiper M, Van de Peer Y (2005) Modeling
gene and genome duplications in eukaryotes. Proc Natl Acad Sci USA 102:5454–5459
Mather K (1935) Reductional and equational separation of the chromosomes in bivalents and
tetravalents. J Genet 30:53–78
Mather K (1936) Segregation and linkage in autotetraploids. J Genet 32:287–314
McClintock B (1929) A cytological and genetical study of triploid maize. Genetics 14:180–227
Mok DWS, Peloquin SJ (1975) Breeding value of 2n pollen (diploandroids) in tetraploid x diploid
crosses in potato. Theor Appl Genet 46:307–314
Piperidis G, Piperidis N, D’Hont A (2010) Molecular cytogenetic investigation of chromosome
composition and transmission in sugarcane. Mol Genet Genomics 284:65–73
Punyasingh K (1947) Chromosome numbers in crosses of diploid, triploid and tetraploid maize.
Genetics 32:541–554
Rabinow L, Nguyen-Huynh AT, Birchler JA (1991) A trans-acting regulatory gene that inversely
affects the expression of the white, brown and scarlet loci in Drosophila melanogaster.
Genetics 129:463–480
Randolph L (1935) Cytogenetics of tetraploid maize. J Agri Res 50:591–605
Randolph LF (1942) The influence of heterozygosis on fertility and vigor in autotetraploid maize.
Genetics 27:163
Randolph LF, Fischer HE (1939) The occurrence of parthenogenetic diploids in tetraploid maize.
Proc Natl Acad Sci USA 25:161–164
Redei G (1964) Crossing experiences with polyploids. Arabidopsis Inf Serv 1:13
Rhoades MM, Dempsey E (1966) Induction of chromosome doubling at meiosis by the elongate
gene in maize. Genetics 54:505–522
Riddle NC, Birchler JA (2008) Comparative analysis of inbred and hybrid maize at the diploid
and tetraploid levels. Theor Appl Genet 116:563–576
Riddle NC, Kato A, Birchler JA (2006) Genetic variation for the response to ploidy change in Zea
mays L. Theor Appl Genet 114:101–111
Riddle NC, Jiang H, An L, Doerge RW, Birchler JA (2010) Gene expression analysis at the
intersection of ploidy and hybridity in maize. Theor Appl Genet 120:341–353
Satina S, Blakeslee AF (1937a) Chromosome behavior in triploid Datura stramonium I. The male
gametophyte. Am J Bot 24:518–527
Satina S, Blakeslee AF (1937b) Chromosome behavior in triploid Datura stramonium II. The
female gametophyte. Am J Bot 24:621–627
Satina S, Blakeslee AF, Avery AG (1937) Balanced and unbalanced haploids in Datura. J Hered
28:192–202
Satina S, Blakeslee AF, Avery AG (1938) Chromosome behavior in triploid Datura. III. The seed.
Am J Bot 25:595–602
Sears ER (1944) Cytogenetic studies with polyploid species of wheat. II. Additional chromosome
aberrations in Triticum vulgare. Genetics 29:232–246
Sears ER (1953) Nullisomic analysis in common wheat. Am Nat 87:245–252
Sears ER (1954) The aneuploids of common wheat. Univ Mo Res Bull 572:1–58
Simillion C, Vandepoele K, Montagu MC, Zabeau M, Van de Peer Y (2002) The hidden
duplication past of Arabidopsis thaliana. Proc Natl Acad Sci USA 99:13627–13632
Singh RJ (1993) Plant cytogenetics. CRC Press Inc, Boca Raton
Sockness BA, Dudley JW (1989a) Performance of single and double cross autotetraploid maize
hybrids with different levels of inbreeding. Crop Sci 29:875–879
CO
RR
352
353
354
355
356
357
358
359
360
361
362
363
364
365
366
367
368
369
370
371
372
373
374
375
376
377
378
379
380
381
382
383
384
385
386
387
388
389
390
391
392
393
394
395
396
397
398
399
400
401
402
403
404
405
UN
Editor Proof
2 Genetic Consequences of Polyploidy in Plants
Layout: T1 Standard SC
Chapter No.: 2
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 32/32
EC
TE
D
PR
OO
F
Sockness BA, Dudley JW (1989b) Morphology and yield of isogenic diploid and tetraploid maize
inbreds and hybrids. Crop Sci 29:1029–1032
Stebbins GL Jr (1947) Types of polyploids: their classification and significance. Adv Genet
1:403–429
Stupar RM, Bhaskar PB, Yandell BS, Rensink WA, Hart AL, Ouyang S, Veilleux RE, Busse JS,
Erhardt RJ, Cr Buell, Jiang J (2007) Phenotypic and transcriptomic changes associated with
potato autopolyploidazation. Genetics 176:2055–2067
Upcott M (1935) The cytology of triploid and tetraploid Lycopersicum esculentum. J Genet
27:105–132
Veitia RA (2002) Exploring the etiology of haploinsufficiency. BioEssays 24:175–184
Veitia RA (2004) Gene dosage balance in cellular pathways: implications for dominance and
gene duplicability. Genetics 168:569–574
Veitia RA, Bottani S, Birchler JA (2008) Cellular reactions to gene dosage imbalance: genomic,
transcriptomic and proteomic effects. Trends Genet 24:390–397
Vizir IY, Mulligan BJ (1999) Genetics of gamma-irradiation-induced mutations in Arabidopsis
thaliana: large chromosomal deletions can be rescued through fertilization of diploid eggs.
J Hered 90:412–417
Wang J, Tina L, Madlung A, Lee HS, Chen M, Lee JJ, Watson B, Kagochi T, Comai L, Chen ZJ
(2004) Stochastic and epigenetic changes of gene expression in Arabidopsis polyploids.
Genetics 167:1961–1973
Weber DF (1983) Monosomic analysis in diploid crop plants. In: Swaminathan MS, Gupta PK,
Sinha U (eds) Cytogenetics of crop plants. Macmillan India Limited, New Delhi, pp 351–378
Wolfe KH, Shields DC (1997) Molecular evidence for an ancient duplication of the entire yeast
genome. Nature 387:708–713
Xiong Z, Gaeta RT, Pires JC (2011) Homoeologous shuffling and chromosome compensation
maintain genome balance in resynthesized allopolyploid Brassica napus. Proc Natl Acad Sci
USA 108:7908–7913
Yao H, Kato A, Mooney B, Birchler JA (2011) Phenotypic and gene expression analysis of a
ploidy series of maize inbred Oh43. Plant Mol Biol 75:237–251
Yousafzai FK, Al-Kaff N, Moore G (2010) The molecular features of chromosome pairing at
meiosis: the polyploidy challenge using wheat as a reference. Funct Integr Genomics
10:147–156
Yu Z, Haberer G, Matthes M, Rattei T, Mayer KF, Gierl A, Torres-Ruiz RA (2010) Impact of
natural genetic variation on the transcriptome of autotetraploid Arabidopsis thaliana. Proc
Natl Acad Sci USA 107:17809–17814
CO
RR
406
407
408
409
410
411
412
413
414
415
416
417
418
419
420
421
422
423
424
425
426
427
428
429
430
431
432
433
434
435
436
437
438
439
440
J. A. Birchler
UN
Editor Proof
32
Metadata of the chapter that will be visualized in
SpringerLink
Book Title
Polyploidy and Genome Evolution
Series Title
Chapter Title
Meiosis in Polyploid Plants
Copyright Year
2012
Copyright HolderName
Springer-Verlag Berlin Heidelberg
Corresponding Author
Family Name
Mittelsten Scheid
Particle
Given Name
Ortrun
Suffix
Division
Author
Organization
Gregor Mendel Institute of Molecular Plant Biology
Address
Vienna, Austria
Email
ortrun.mittelsten_scheid@gmi.oeaw.ac.at
Family Name
Zielinski
Particle
Given Name
Marie-Luise
Suffix
Division
Organization
Gregor Mendel Institute of Molecular Plant Biology
Address
Vienna, Austria
Email
Abstract
Meiosis is an obligate process during sexual reproduction, which involves the combination of parental
genomes and the coordinated segregation of the recombined chromosomes to the gametes. Polyploidy has
direct and fundamental consequences on meiosis, which are gradually and individually different between the
extreme cases of auto- and allopolyploids. Multiple chromosome complements have a major impact,
especially on chromosome pairing during pachytene and on the segregation of genotypes and phenotypes in
progeny. At the same time, irregularities during meiosis are a major source of naturally occurring
polyploidization events by the formation of unreduced gametes. Although individuals originating from
nonhaploid gametes may suffer from reduced vigor and fecundity, their gametogenesis can produce many
more chromosomal combinations than regular diploids, and thereby expose more diversity to natural selection.
A more relaxed control of pairing and segregation in polyploids, possibly also with increased recombination
rates, might be an important contribution to evolution and adaptation potential, especially under drastic or
frequent changes in environmental conditions.
Book ISBN: 978-3-642-31441-4
Page: 33/54
Chapter 3
2
Meiosis in Polyploid Plants
3
Marie-Luise Zielinski and Ortrun Mittelsten Scheid
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
D
8
TE
7
EC
6
Abstract Meiosis is an obligate process during sexual reproduction, which involves
the combination of parental genomes and the coordinated segregation of the
recombined chromosomes to the gametes. Polyploidy has direct and fundamental
consequences on meiosis, which are gradually and individually different between the
extreme cases of auto- and allopolyploids. Multiple chromosome complements have
a major impact, especially on chromosome pairing during pachytene and on the
segregation of genotypes and phenotypes in progeny. At the same time, irregularities
during meiosis are a major source of naturally occurring polyploidization events by
the formation of unreduced gametes. Although individuals originating from
nonhaploid gametes may suffer from reduced vigor and fecundity, their gametogenesis can produce many more chromosomal combinations than regular diploids,
and thereby expose more diversity to natural selection. A more relaxed control of
pairing and segregation in polyploids, possibly also with increased recombination
rates, might be an important contribution to evolution and adaptation potential,
especially under drastic or frequent changes in environmental conditions.
CO
RR
4
5
PR
OO
1
F
Book ID: 272454_1_En
Date: 16-8-2012
3.1 Introduction
Polyploidization can affect the size of the nucleus and the cell, stimulate genetic
rearrangements, or modify gene expression patterns, but these changes very likely
operate by the same mechanistic principles of nuclear organization, recombination,
or gene regulation, as in diploid cells. In contrast, reducing the number of
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: 3
M.-L. Zielinski O. Mittelsten Scheid (&)
Gregor Mendel Institute of Molecular Plant Biology, Vienna, Austria
e-mail: ortrun.mittelsten_scheid@gmi.oeaw.ac.at
P. S. Soltis and D. E. Soltis (eds.), Polyploidy and Genome Evolution,
DOI: 10.1007/978-3-642-31442-1_3, Springer-Verlag Berlin Heidelberg 2012
33
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
61
62
63
64
65
66
67
68
F
30
PR
OO
29
D
28
TE
27
chromosomes during the formation of gametes requires a more elementary
modification in polyploids. At the same time, irregular gamete formation is a
major trigger for polyploidization. Both processes converge in meiosis. This is a
sequence of usually two special, subsequent cell divisions in all sexually reproducing eukaryotic organisms in which the number of chromosomes is reduced to
half, before two gametes of different parental origin fuse to form a zygote to
produce the next generation.
Like mitosis, the cell division of somatic cells, meiosis is also preceded by the
replication of the nuclear DNA into duplicate sister chromatids which are attached at
the centromeres. While in mitosis the replicated chromosomes are arranged individually on the metaphase plate, and chromatids become separated in the subsequent
anaphase, each replicated chromosome in meiotic cells is first aligned with its
respective homologous partner. This pairing is followed by programed induction of
DNA double-strand breaks (DSB), from which at least one per chromosome becomes
converted into a recombination event between non-sister chromatids (crossover, CO).
Later, these COs become cytologically visible as chiasmata. Recombination within
chromosome pairs results in new and unique combinations of paternal and maternal
genetic information per chromosome, while it is also prerequisite for subsequent
organized chromosome segregation. Paired chromosomes become physically
connected by the synaptonemal complex (SC). The process of pairing, synapsis, and
recombination takes most of the time during prophase I, which is subdivided into
leptotene, zygotene, pachytene, diplotene, and diakinesis according to chromosome
condensation and configuration. In the subsequent metaphase I, the recombined
meiotic chromosome pairs are arranged on the equatorial plane, sister kinetochores of
each pair attach to opposite spindles and are pulled apart in anaphase I. Correct and
synchronous pairing is a prerequisite for equal distribution of the chromosomes to the
two daughter cells. After telophase I and prophase II, sister chromatids become
separated during meta-, ana-, and telo-phase II, resulting in a classical meiosis with
four nuclei. Faultless meiosis is an essential factor for fertility and thereby decisive for
evolutionary success. Therefore, it is not surprising that it is a tightly controlled
process and intensely studied in many organisms.
The majority of meiosis research is performed with diploid organisms, and
there are excellent reviews available that describe morphological, genetic, and
mechanistic aspects of meiosis in many different systems (Dawe 1998; Bhatt et al.
2001; Armstrong et al. 2003; Mezard et al. 2007; Mercier and Grelon 2008;
Harrison et al. 2010; Pawlowski 2010). However, characteristic differences in all
stages of meiosis in many different polyploids have been observed repeatedly
(reviewed in Ramsey and Schemske 2002). Therefore, we will focus on those
aspects that are known or expected to be different between meiosis in diploids and
polyploids. Although mechanistically intertwined, we will separate three aspects,
(1) chromosome pairing, (2) recombination and crossover, and (3) segregation.
Whenever relevant, we will distinguish between autopolyploids, with multiples of
similar chromosomes, and allopolyploids, with chromosomes of different origin,
and therefore higher divergence.
EC
26
CO
RR
25
M.-L. Zielinski and O. Mittelsten Scheid
UN
Editor Proof
34
Book ISBN: 978-3-642-31441-4
Page: 34/54
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 35/54
35
76
3.2 Recognition, Pairing, and Synapsis
77
78
79
80
81
82
83
84
85
86
87
88
89
90
91
92
93
94
95
96
97
98
99
100
101
102
103
104
105
106
PR
OO
74
Early in meiosis, homologous chromosomes have to find and recognize each other.
Interphase chromosomes are thought to occupy nonrandom territories (Schubert
and Shaw 2011), and nonrandom spatial organization of homologous chromosomes has been recognized (reviewed in Avivi and Feldman 1980). In many
eukaryotes, telomeres and centromeres can cluster at opposite poles of the nucleus
(Rabl 1885; Cowan et al. 2001). This Rabl-configuration (Fig. 3.1a) is found in
wheat, rye, oats, and barley, but not in maize or Arabidopsis (reviewed in Schubert
and Shaw 2011). It is assumed to be related to large genome size and/or
chromosome length (Dong and Jiang 1998). Rabl-configuration may support gene
expression control and the onset of meiosis (Cowan et al. 2001).
After the premeiotic DNA replication, the two sister chromatids are held together
by sister chromatid cohesion and gradually become condensed. During leptotene, their
chromatin is folded into loops attached to a protein fiber core, the axial element (AE)
(Harper et al. 2004). During transition to zygotene, the so-called telomere bouquet is
formed, in which the chromosome ends attach to the inner nuclear envelope and cluster
(Fig. 3.1b). This association is likely to be actively regulated (Scherthan 2007).
Recognition between the homologous chromosomes appears to occur between subtelomeric regions (Corredor et al. 2007). The telomere bouquet, found in almost all
studied organisms except Drosophila and C. elegans (Harper et al. 2004), is not
absolutely required for the following steps but is proposed to make progression of
meiosis more efficient. Both Rabl-configuration and telomere bouquet bundle chromosomes and thereby help reduce the spatial distance between them. After telomere
bouquet formation in yeast, the centromeric regions are distributed throughout the
nucleus and oscillate to enhance finding homologous sequences (Bass et al. 1997;
Scherthan 2007). Chromosome movements are extremely dynamic at this stage
(Sheehan and Pawlowski 2009), which increases physical encounters and makes
meeting and recognition of the homologs more likely (Pawlowski and Cande 2005).
D
73
TE
72
EC
71
CO
RR
70
F
75
Since polyploidy in animal germ lines is restricted to relatively few groups, and
the research literature about meiotic mechanisms is scarce, most of the evidence
summarized here stems from investigations in fungi and plants. There, polyploidy is
by far most common among angiosperms, but genome research provides growing
evidence for ancient polyploidization events in other groups (Sundstrom et al. 2008;
Jiao et al. 2011). Independent of how and when polyploids originate, mastering
meiosis is an important checkpoint for survival and evolutionary success.
69
UN
Editor Proof
3 Meiosis in Polyploid Plants
3.2.1 Homology Recognition in Polyploids
It seems plausible to expect that higher chromosome numbers, and larger nuclei in
general, but especially more potential homologous partner chromosomes in
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
Editor Proof
36
Book ISBN: 978-3-642-31441-4
Page: 36/54
M.-L. Zielinski and O. Mittelsten Scheid
(b)
PR
OO
F
(a)
Fig. 3.1 Chromosome arrangement (red circles centromeres; orange triangles telomeres).
a Rabl-configuration in interphase, with centromeres clustered at one pole of the nucleus and
telomeres oriented toward the other. b Bouquet formation during meiosis at the onset of zygotene,
with telomeres clustered at the inner nuclear envelope
125
3.2.2 Pairing in Autopolyploids
113
114
115
116
117
118
119
120
121
122
123
126
127
128
129
130
131
TE
112
EC
111
CO
RR
109
110
There is one important difference that distinguishes the pachytene stage in
autopolyploids: the possibility of pairing between more than two chromosomes,
resulting in the formation of multivalents. This requires a certain degree of
homology between pairing partners, and this is reflected in different frequency of
multivalent occurrence depending on the type of polyploidy, species, individual
chromosomes, and chromosome segments (Sybenga 1996). While allopolyploids
UN
108
D
124
autopolyploids, could delay progression through meiosis, since the early stages of
recognition and pairing can be more complex. Indeed, meiosis in autotetraploid
Saccharomyces cerevisiae strains is delayed in comparison to its diploid counterpart (Trelles-Sticken et al. 2003). While higher C-values (genomic DNA content) in
different plant species also increase the duration of meiosis when compared at the
same ploidal level, the comparison of plants with different ploidy within the same
species revealed a surprising, negative correlation: higher polyploidy seems to
shorten the time needed for completing meiosis (Bennett 1977). These discrepancies might be created by defining the duration only by cytologically visible events.
Analysis with higher resolution techniques and consideration of molecular events
(Carlton et al. 2006) at the onset of meiosis might provide more accurate information on meiosis kinetics in polyploids. In spite of individual differences, it is
remarkable that a wide range of genome and nuclear size variation does not modify
the length of the recognition process (Moore and Shaw 2009). It appears that
homology recognition is not the limiting step, due to supportive elements like Rablconfiguration and telomere bouquet , specialized chromosome pairing sites as found
in D. melanogaster and C. elegans (McKee 1996; McKim 2007), and intensive
chromosome movements that are effective in diploids and polyploids.
107
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 37/54
140
141
142
143
144
145
146
147
148
149
150
151
152
153
154
155
156
157
158
159
160
161
162
163
164
165
166
167
168
169
170
171
172
F
138
139
PR
OO
136
137
D
135
TE
134
with genomes composed of genetically divergent parents often form only bivalents
aligned over the whole length of the chromosome, autopolyploids may have
multivalents. These are generated by simultaneous alignment of different partners
and at different chromosome ends. The probability depends on the degree of ploidy
and homology (random-end pairing model by John and Henderson 1962) and is
controlled by genetic factors (see below), but it is independent of chromosome
length (Morrison and Rajhathy 1960). Although multivalent formation is more
often associated with autopolyploidy, many exceptions delimit the universality of
this correlation: newly formed autopolyploids often have a lower rate of multivalents than expected, and allopolyploids may form multivalents to some extent
(Ramsey and Schemske 2002). A few of these cases might be due to tri- or
tetrasomy compensated by lack of other chromosomes, resulting in apparent
euploidy (Mestiri et al. 2010).
Progressive pairing starting simultaneously from opposite ends can result in
multivalents with pairing partner switches (PPS), so that one chromosome can be
aligned with two or more others in different segments (Fig. 3.2a). The distribution
of PPS is variable but probably not totally random. More than one switch per
chromosome (Fig. 3.2b) indicates the existence of additional, autonomous pairing
sites (APS) along the chromosomes, each with a uniform and low probability of
generating a PPS (Jones and Vincent 1994). The number of switches indicates the
minimal number of pairing sites (Loidl 1995). These are likely not only determined genetically, since a higher number of PPS in autotriploid Crepis capillaris,
compared to autotetraploid (Jones 1994), indicates other factors than just APS
distribution, but probably some interference between pairing initiation sites.
Pairing with one partner at one pairing site preferentially promotes continuation of
pairing in a zipper-like manner, due to steric constraints. However, this preference
is valid only once synapsis has been initiated.
According to the random-end pairing model (John and Henderson 1962),
alignment of two chromosome ends enhances the two remaining to pair, although
the opposite ends of the chromosome still can pair randomly, resulting in 2/3
multivalents and 1/3 bivalents in tetraploids (only chromosome ends are considered under this model). Exceptions with fewer than expected multivalents (Weiss
and Maluszynska 2000; Santos et al. 2003; Carvalho et al. 2010) or bivalent
formation due to selective pairing (Simioni and do Valle 2011, and references
within) reflect genetic and/or epigenetic heterozygosity between parental chromosome sets, due to ongoing diploidization. This can be different between individual plants or individual chromosomes (Santos et al. 2003) and extremely
divergent in complex polyploids, as deduced from genetic maps of sugarcane
(Jannoo et al. 2004). Higher than expected ratios of multivalents can also occur, as
in newly generated autotetraploids of Arabidopsis thaliana, indicating multiple
active pairing initiation sites even in small chromosomes (Santos et al. 2003).
EC
133
37
CO
RR
132
UN
Editor Proof
3 Meiosis in Polyploid Plants
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
Editor Proof
38
Book ISBN: 978-3-642-31441-4
Page: 38/54
M.-L. Zielinski and O. Mittelsten Scheid
(a)
TE
D
PR
OO
F
(b)
175
176
177
178
179
180
181
182
183
184
185
186
187
188
CO
RR
174
3.2.3 Pairing in Allopolyploids
Although newly formed hybrids combining different genomes and different chromosome numbers and shapes can exhibit erroneous pairing or lack of alignment
(Ozkan and Feldman 2009), established allopolyploids show usually diploid-like
pairing and formation of bivalents. Nevertheless, multivalents or segmental multivalent pairing can be found in allopolyploids, depending on external or genetic factors. In many polyploids, a genetic control system has evolved that regulates the
pairing behavior during meiosis (Watanabe 1981; Gupta and Fedak 1985; MartinezPerez et al. 2001; Comai et al. 2003; Martinez-Perez et al. 2003; Jenczewski and Alix
2004). Probably the best studied example was discovered in the hexaploid bread
wheat (Triticum aestivum, composed of genomes A, B, and D). In the presence of the
Ph1 locus, pairing and recombination occur preferentially between homologous
chromosomes (Riley and Chapman 1958); lack of Ph1 leads to a substantial increase
of homoeologous pairing. The Ph1 locus from wheat can also affect pairing after
introgression into rye (Lukaszewski and Kopecky 2010). The absence of Ph1 in
diploids suggests its emergence through polyploidization (Griffiths et al. 2006). The
UN
173
EC
Fig. 3.2 Pairing partner switches during multivalent formation in polyploids. Alignment and
partial synapsis between four chromosomes with one (a) or two (b) pairing partner switches
(PPS) on the long arm
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 39/54
39
197
3.2.4 Synapsis
198
199
200
201
202
203
204
205
206
207
208
209
210
211
212
213
214
215
216
217
218
219
220
221
222
223
224
225
226
PR
OO
195
As soon as meiotic chromosomes align, they become joined by a stable proteinaceous
structure, the SC. Its formation starts mainly at telomeres (Stack and Anderson 2002)
and is intimately connected with DSBs and recombination (see next paragraph). The
SC consists of three components: the two parallel AEs, the connecting central element, and periodically occurring recombination nodules (RNs) (Lohmiller et al.
2008). The RNs are multiprotein complexes thought to be involved in synapsis and
recombination and can be distinguished by their structure and appearance as early or
late RNs (Stack and Anderson 2002; Anderson and Stack 2005).
Synapsis is expected to follow once pairing is established (Loidl 1995). However,
unequal chromosome numbers can make a difference in polyploids: in autotriploid,
but not in autotetraploid, yeast, trivalent SC formation was observed (Loidl 1995).
Such fixed multivalents are prone to nondisjunction at metaphase I. Multivalents with
equal numbers seem to resolve more often into regular bivalents, although SC
connections between several AEs can occur and become fixed by crossovers. Without
crossover at the region of the PPS, the central region is twisted and bivalents are
formed (von Wettstein et al. 1984). When, and how, are interesting questions,
especially for meiosis in autopolyploids.
In summary, recognition, pairing, and synapsis of homologous chromosomes in
diploids and polyploids are different in several aspects and determined by various
parameters. The high rate of multivalent formation in autopolyploids depends on
the type and history of the polyploidization, the number of pairing initiation sites,
and odd or even chromosome multiplication. Allopolyploids rather generally form
bivalents, but segment-, chromosome-, or genome-specific multivalent formation,
and genetic and environmental influence, can also result in meiotic progression
different from that in diploids.
D
194
TE
193
EC
191
192
CO
RR
190
F
196
Ph1 locus is a complex rearrangement between subtelomeric heterochromatin
translocated within a cluster of cdk2-like genes localized on one of the chromosomes
of the B genome (Griffiths et al. 2006).
Ph1 controls meiosis at several levels. While telomere pairing is not affected,
Ph1 prevents pairing between centromeres of nonhomologous chromosomes, keeps
centromeres at the nuclear periphery, synchronizes chromatin condensation, and
controls the expression of synapsis and recombination components (Naranjo and
Corredor 2004; Moore and Shaw 2009; Knight et al. 2010; Yousafzai et al. 2010).
189
UN
Editor Proof
3 Meiosis in Polyploid Plants
3.3 Recombination and Crossover
Meiotic recombination in plants is initiated by multiple DSBs. Some of the breaks
are repaired by homologous recombination using the non-sister chromatid as a
template, resulting in reciprocal strand exchange (CO events). As the number of
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
234
235
236
237
238
239
240
241
242
243
244
245
246
247
248
249
250
251
252
253
254
255
256
257
258
259
260
261
262
263
264
265
266
267
268
269
270
271
F
232
233
PR
OO
231
D
230
TE
229
initial DSBs exceeds the number of COs, the majority of DSBs are processed
otherwise, either by separating double Holliday junction into non-crossover (NCO)
products or by the synthesis-dependent strand annealing (SDSA) pathway. There
are excellent recent reviews that describe details of DSB processing and components of meiotic recombination (Hamant et al. 2006; Mercier and Grelon 2008;
Sanchez-Moran et al. 2008; De Muyt et al. 2009; Edlinger and Schlogelhofer
2011; Osman et al. 2011), and there is no reason to assume that the principal
mechanisms are different in polyploids. However, there is growing evidence that
the frequency with which DSBs give rise to COs can be regionally or generally
modified by polyploidy. Therefore, we will describe some factors that influence
crossover frequencies.
Due to the role of recombination in the regulation of chromosome pairing and
segregation, there appears to be a minimum of one CO per chromosome in many
species (Youds and Boulton 2011). Beyond this, and in spite of variation between
different species, there seems to be no correlation between the length of chromosomes and the number of COs (Brubaker et al. 1999; Mezard 2006). The genomes
are rather composed of ‘hot spots’ and ‘cold spots’, with high and low probabilities
for meiotic recombination (Drouaud et al. 2006; Mezard 2006; Kim et al. 2007).
Even small local sequence divergence, like a transgene insertion, can modify
meiotic recombination locally (Sun et al. 2008). Variation in recombination or CO
frequency among Arabidopsis accessions (Barth et al. 2001; Sanchez-Moran et al.
2002) also indicates genetic components that regulate the meiotic recombination
frequency in trans. Further, temperature and age of flowers within the plant can
modify CO events (Francis et al. 2007). Another aspect to consider is CO interference, the inhibition of additional recombination events by COs in their proximity
or even along the whole chromosome (Holliday 1977; van Veen and Hawley 2003;
Baudat and de Massy 2007; Youds and Boulton 2011).
Differences in the number of CO events in polyploids have been described in
several systems. The A and D genome components of allotetraploid Gossypium
(cotton) are very different in size but marker pairs have, nevertheless, comparable
genetic distances if compared between diploid or allopolyploid mapping populations, respectively. Surprisingly, the comparison for the same markers between
diploids and allotetraploids indicated a higher recombination rate in the latter
(Brubaker et al. 1999; Desai et al. 2006). F1 hybrids between the diploid parents
Brassica oleracea and Brassica rapa, compared with F1 plants derived from the
same hybrid with doubled chromosome numbers after colchicine treatment,
produced more progeny with intergenome recombination (Szadkowski et al.
2010, 2011). A comparison of the CO number among diploid, allotriploid, and
allotetraploid hybrids from crosses between Brassica oleracea and Brassica rapa
revealed the highest number for allotriploids, intermediate values for allotetraploids, and the lowest numbers in diploids (Nicolas et al. 2008; Leflon et al. 2010).
This indicates stimulation of recombination by hybridity and/or polyploidy,
however, in a nonlinear correlation with the number of homologous chromosomes. This might be coupled with the occurrence of univalents, in addition to
bivalents, in the triploids, whereas the tetraploids exclusively form bivalents
EC
228
CO
RR
227
M.-L. Zielinski and O. Mittelsten Scheid
UN
Editor Proof
40
Book ISBN: 978-3-642-31441-4
Page: 40/54
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 41/54
41
292
3.4 Chromosome and Allele Segregation
293
3.4.1 Chromosome Segregation
280
281
282
283
284
285
286
287
288
289
290
294
295
296
297
298
299
300
301
302
303
304
305
306
307
308
309
310
311
PR
OO
279
D
278
TE
277
EC
276
Following pairing and recombination in prophase I, chromosomes are arranged at
the equatorial plane during metaphase I. The following anaphase I is distinct from
that in mitosis since both centromeres of sister chromatids attach to spindles of the
same pole and retain their cohesion when the chromosome pairs get dragged apart,
moving to opposite poles. This ‘reductional’ division, therefore, halves the
chromosome number. It is followed by metaphase II with the formation of two new
equatorial planes, and anaphase II, in which the sister chromatids now become
separated and distributed to the resulting four postmeiotic nuclei. A tightly
controlled order of maintenance and stepwise release of cohesion specifies all stages
of meiosis (Sakuno and Watanabe 2009), but there is no evidence that an increased
chromosome number in polyploids affects this control.
In contrast, due to the effects of polyploidy on pairing and/or recombination,
there is a much higher probability of unequal segregation of chromosomes into the
postmeiotic nuclei. Unequal distribution of the chromosomes can occur in many
different ways (Pagliarini 2000). It can decrease the regular nuclear DNA content, if
unresolved multivalents, unpaired or laggard chromosomes, or parts of chromosomes are not included or translocated to other chromosomes (Madlung et al. 2005;
Charles et al. 2010; Gaeta and Pires 2010; Wang et al. 2010). Extreme cases of
CO
RR
274
275
F
291
(Leflon et al. 2010), indicating a control mechanism to prevent multivalents. A
good candidate is the PrBn locus (Jenczewski et al. 2003; Nicolas et al. 2009;
Cifuentes et al. 2010). Exploring an assay system for meiotic recombination
based on fluorescent proteins expressed in seeds (Melamed-Bessudo et al. 2005),
a comparison among diploids, isogenic autotetraploids, and allotetraploids generated by interspecies hybridization revealed an unexpected increase of recombination between the markers for both types of polyploids (Pecinka et al. 2011).
The presence of mainly multivalent formation in one, and bivalent formation in
the other, argue against a correlation with the pairing behavior.
In summary, the frequency of recombination between any two markers is
determined by interplay of physical distance, local cis- and trans-acting genetic
control elements, the degree of overall genetic divergence, and sex-specific differences (Vizir and Korol 1990; De Vicente and Tanksley 1991; Drouaud et al.
2007; Nelson et al. 2005; Pecinka et al. 2011), regardless of ploidy. The few briefly
described examples of increased recombination frequency in polyploid plants
suggest that pairing behavior, potential modification of crossover interference by
PPS, distinctive segregation patterns (see below), or other still unknown factors
add to the complexity. However, it is tempting to speculate that high(er) recombination rates, at least in part, contributed to the prevalence of polyploidy among
angiosperms and crop plants bred within the last 12,000 years.
272
273
UN
Editor Proof
3 Meiosis in Polyploid Plants
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
319
320
321
322
323
324
325
326
327
328
329
330
331
332
333
334
335
336
337
338
339
340
341
342
343
344
345
346
347
348
349
350
351
F
PR
OO
317
318
D
316
TE
314
315
systematic chromosome elimination were observed in the allopolyploid Paspalum
subciliatum (Adamowski et al. 1998) and in pentaploid Brachiaria decumbens
(Ricci et al. 2010), due to kinetic asynchrony of the two different genomes after
diakinesis. Alternatively, the nuclear DNA may be increased if restitution of nuclei
occurs prior to division or around incompletely separated chromosomes (Ramsey
2007; Brownfield and Kohler 2011). This can occur either after the first or the
second meiotic division, differing in the degree of heterozygosity transmitted to the
progeny (Peloquin et al. 2008).
While loss of chromosomal material is often deleterious for the resulting cell and
leads to cell death or reduced viability and fertilization, a gain, and optimally a
balanced multiplication of all chromosomes, is much less detrimental. Accordingly,
unreduced gametes occur in numerous species and are a major source of polyploidization events (Leitch and Leitch 2008; Koehler et al. 2010). The frequency of
unreduced gametes varies significantly between hybrids (e.g., Ortiz 1997; Lim et al.
2004), between different ploidal levels (e.g., Burton and Husband 2001; Ramsey
2007), and also depends on genetic factors (reviewed in Brownfield and Kohler
2011), external conditions, and stress factors (reviewed in Ramsey and Schemske
1998). The frequent occurrence of unreduced gametes has been explained by the
lack of, or less effective, ‘pachytene checkpoint’ in plants (Li et al. 2009), but may
be an inherent element of circumventing meiotic problems in interspecies hybrids
by polyploidization.
The occurrence of diploid or other unreduced post-meiotic cells also has
consequences for the subsequent phase of the life cycle. Haploid gametophytes
from diploid parents depend on functionality of each single genomic copy that
encodes factors for proper gametophyte viability and fertilization potential.
Gametophytes from polyploid species contain more than one copy, providing
backups for defective alleles and the potential for heterozygosity even during the
two and three postmeiotic cell divisions forming the male and female gametophytes, respectively. Since at least pollen selection is efficient at several levels
(Ottaviano et al. 1990), avoiding haploidy might provide reduced selection
pressure and increased variability of transmitted genetic information.
Not directly related to meiosis but with consequences for gamete formation and
fertility is the phenomenon of cytomixis, the fragmentation of chromatin and
transfer of fragments among cells through cytoplasmic channels. Observed in
many species, and especially in pollen mother cells (e.g., Mursalimov and Deineko
2011), one published comparison between diploid and tetraploid varieties of
Withania somnifera indicated that the resulting reduction of fertility is much less
pronounced in the tetraploids, along with a reduced extent of intercellular
connections (Singhal and Kumar 2008). However, a correlation with polyploidy in
other species awaits investigation.
EC
313
CO
RR
312
M.-L. Zielinski and O. Mittelsten Scheid
UN
Editor Proof
42
Book ISBN: 978-3-642-31441-4
Page: 42/54
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 43/54
358
359
360
361
362
363
364
365
366
367
368
369
370
371
372
373
374
375
376
377
378
379
380
381
382
383
384
385
386
387
388
389
390
391
392
393
394
F
357
PR
OO
356
D
355
The genetic segregation of traits in polyploids can be quite different from that in
diploids. Heterozygosity and multiple alleles per gamete increase the combinatorial
possibilities in the progeny population substantially and make the segregation patterns
deviate from the Mendelian ratios for diploids. This makes QTL analysis or any
mapping process complex, as in the extreme case of highly polyploid and partially
aneuploid sugarcane (Andru et al. 2011). Naturally, analysis of inheritance requires
consideration of pairing behavior and the resulting chromosome segregation. Variation among hybrids, individuals, chromosomes, and chromosome segments does not
allow establishing general rules. Therefore, we will depict the extremes: autotetraploids with quadrivalent formation, unbiased chromosome segregation, and tetrasomic
inheritance on one hand, and allopolyploids with strict bivalent formation and disomic
inheritance on the other. A Punnett square for segregation of one trait in the progeny of
heterozygous autotetraploid parents with two copies of two different alleles has 36
fields (Fig. 3.3b), rather than four in the case of a heterozygous diploid (Fig. 3.3a).
Deleterious recessive alleles become apparent as affected phenotypes in only 1:35,
rather than 1:3 in diploids. Each individual tetraploid can have one, two, three, or four
identical alleles (termed simplex, duplex, triplex, or quadruplex) at each locus, and the
expected segregation of the different genotypes upon crossing with a partner of the
same genotype is 1:8:18:8:1 (Fig. 3.3b). In the case of allopolyploids with strict
bivalent formation between the chromosomes of common origin, each gamete carries
one of each chromosome type so that the progeny are uniform (Fig. 3.3c). If the
chromosomes carry different alleles, autotetraploids can produce 19 different genotypes at one locus (Fig. 3.4a), while disomic inheritance in allotetraploids is limited to
9 combinations (Fig. 3.4b). While exclusive tetrasomic or disomic inheritance in
polyploids does occur, intermediate forms and changes in both directions over time
are frequently observed in nature and need to be considered in genetic and population
studies (Stift et al. 2008).
A genetic peculiarity of polyploid meiosis that does not occur in diploid
organisms is the chance of double reduction (Darlington 1929a; Butruille and
Boiteux 2000). This term describes the possibility that regions from two sister
chromatids become combined in the same gamete. Double reduction results from
recombination events between the locus under observation and the centromere. If
the two chromosomes that have recombined move to the same pole in anaphase I,
there is a high probability that the distal end of the recombined chromatid will be
included in the same nucleus as the distal end of the non-recombined chromatid,
after anaphase II (locus B in 2 gametes in Fig. 3.5). The frequency of double
reduction for a genetic locus depends on its distance from the centromere and the
frequency of multivalent formation.
Since segregation of traits in polyploids is so complex, but of high importance
for population geneticists and plant breeders, there were many attempts to
approach the problem via mathematical modeling. Ground-breaking theoretical
work (Mather 1936; Fisher 1947) has more recently stimulated several
TE
354
3.4.2 Allelic Segregation
EC
353
43
CO
RR
352
UN
Editor Proof
3 Meiosis in Polyploid Plants
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
44
M.-L. Zielinski and O. Mittelsten Scheid
PR
OO
F
(a)
(b)
EC
TE
D
Fig. 3.3 Comparison of
segregation for an individual
locus with two alleles upon
bivalent or quadrivalent
formation. Two different
alleles (red, blue) in diploid
parents are transmitted with
equal probability, resulting in
three genotypes, forming
twice as many heterozygotes
as each homozygote (a). Two
different alleles with two
copies each in tetraploids
with random pairing between
each of the four
chromosomes result in five
genotypes, with only one
homozygote among 36
individuals (b). The same
situation in tetraploids with
strict bivalent formation
between the more similar
chromosome pairs renders
only one combination (c)
Editor Proof
Book ISBN: 978-3-642-31441-4
Page: 44/54
395
396
397
398
UN
CO
RR
(c)
computational methods based on maximum-likelihood and other statistical
approaches to estimate double reduction frequency and recombination and to
support mapping of quantitative trait loci in polyploids with multivalent formation
(Doerge and Craig 2000; Ridout et al. 2001; Wu et al. 2001a, b; Luo et al. 2004,
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 45/54
3 Meiosis in Polyploid Plants
PR
OO
F
(a)
D
Editor Proof
Fig. 3.4 Comparison of
segregation for an individual
locus with four alleles upon
bivalent or quadrivalent
formation. Four different
alleles (red, orange, dark
blue, light blue) in tetraploids
with random pairing between
each of the four
chromosomes result in
nineteen genotypes (a). The
same situation in tetraploids
with strict bivalent formation
between the more similar
chromosome pairs renders
only nine combinations (b)
45
400
401
402
403
404
405
406
407
408
2006; Li et al. 2010). However, different modeling approaches do not always
support the same conclusions (Ma et al. 2002; Cao et al. 2004), and it is likely that
we need to extend and refine experimental data collection as well as modeling
approaches to provide satisfying tools. Already difficult for strict autopolyploids
with mostly polysomic inheritance, the situation is even more complex for
polyploids with intermediate types of inheritance, as mentioned above. These can
be caused either by segment- or chromosome-specific homology differences, or by
stochastic pairing differences. Not even the situation in tetraploid yeast strains,
where tetrad analysis provides excellent resolution of segregation analysis, is easy
to interpret (Albertin et al. 2009; Stift et al. 2010). Allelic segregation in
UN
399
CO
RR
EC
TE
(b)
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
M.-L. Zielinski and O. Mittelsten Scheid
AA
BB
PR
OO
BB
AA
F
Editor Proof
46
Book ISBN: 978-3-642-31441-4
Page: 46/54
A
B
Meiosis I
AA
AA
BB
BB
Meiosis II
TE
D
Fig. 3.5 Double reduction. Limited to polyploids, there is a certain probability that distal regions
(around locus B) from two sister chromatids are included in the same gamete. This exceptional
configuration occurs if two recombined chromosomes move to the same pole in anaphase I,
which is excluded in diploids
allopolyploid plants, as described, depends very much on the ratio of multivalent
formation at the genetic locus under investigation.
411
3.5 External Influence
414
415
416
417
418
419
420
421
422
423
424
425
426
427
As for other developmental and physiological processes, progress and efficiency of
meiosis can be modified by environmental influences. Temperature has been most
extensively studied, with a range of different responses. Higher temperatures
reduced regular meiosis in diploid Allium ursinum (Loidl 1989) and Rosa (Pecrix
et al. 2011) and increased the number of unreduced pollen (reviewed in Ramsey
and Schemske 1998). This could enhance the formation of polyploids under heat
stress and thereby support the described adaptive role of polyploidization under
adverse conditions. However, data for polyploids are rare, and partially divergent.
In allopolyploid Brassica hybrids, more unreduced pollen was observed at lower
temperatures (Mason et al. 2011), and meiosis in wheat was not affected within a
physiological range of temperatures (Bayliss and Riley 1972).
A more uniform effect across species (yeast, worms, and plants) appears to be a
stimulation of meiotic recombination rates at higher temperatures (Rose and
Baillie 1979; Borner et al. 2004; Francis et al. 2007). It is likely that adaptation to
habitats with more or less drastic environmental challenges has selected a
matching range of meiotic responses. Further, the more recently established
CO
RR
413
UN
412
EC
410
409
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 47/54
47
polyploid hybrids generated by breeders might respond quite differently than
polyploids originating from spontaneous events long ago.
430
3.6 Bypassing Meiosis
437
438
439
440
441
442
443
444
445
446
447
448
449
450
451
452
453
454
455
456
457
458
459
460
461
462
463
464
465
466
PR
OO
435
436
D
434
TE
433
Ancient polyploidization events and the prevalence of polyploids among living
plants leave no doubt that the ecological and metabolic advantages of multiple
chromosome complements preponderate over possible disadvantages (Soltis and
Soltis 1999; Ramsey and Schemske 2002; Comai 2005; Otto 2007; Parisod et al.
2010; Jiao et al. 2011). As described, successful meiosis and gamete formation are
mastered in different ways. Nevertheless, the survival and success of each new
hybridization or polyploidization event depends on multiple factors during
meiosis. Besides the requirements to adapt chromosome pairing and segregation,
asynchrony, incompatibility of protein complex subunits, dosage effects, or
differences in gene regulation can present meiotic barriers.
One solution in plants to avoid such a decrease in reproduction is to avoid
meiosis by apomixis. Apomixis, the asexual production of seeds, leads to offspring
that is genetically identical to the mother plant and originate via different modifications of the double fertilization pathway (Gustaffson 1946; Asker and Jerling
1992). Gametophytic apomixis (Nogler 1984) is divided into apospory, where the
unreduced embryo sac is formed by a cell of the nucellus, and diplospory, where
the unreduced embryo sac is formed by bypassing meiosis I during megasporogenesis. Both result in an embryo sac that contains an unreduced egg cell, which
develops into an embryo independent of fertilization. Although apomicts are found
in diploids and polyploids, they are more common among polyploids (Asker and
Jerling 1992). Apomixis is thought to have evolved several times from sexual
ancestors in over 400 plant species (Nogler 1984). It is clearly associated with
changes in gene expression, kinetics, or epigenetic regulation (Carman 1997;
Sharbel et al. 2010). Whether polyploidization is a driving force toward apomixis,
or a consequence of apomictic propagation, is a matter of debate. Apomixis
harbors the potential to avoid sexual sterility caused by multivalent formation in
polyploids (Horandl et al. 2011). On the other hand, and as described, polyploidy
promotes the formation of unreduced gametes (Ramsey and Schemske 1998) and
might thereby foster the establishment of higher ploidal levels. Polyploidization of
apomicts could also protect them against extinction, by buffering against the
irreversible accumulation of deleterious mutations in the absence of recombination
as depicted by the metaphor of ‘‘Muller’s ratchet’’ (Darlington 1929b). Nevertheless, it is likely that these conditions are relevant in varying and individual
combinations, with additional components such as highly efficient DNA repair
system in asexually reproducing species (Schoen and Martens 1998) or occasional
or conditional sexual reproduction (D’Souza et al. 2004).
EC
432
CO
RR
431
F
429
428
UN
Editor Proof
3 Meiosis in Polyploid Plants
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
467
M.-L. Zielinski and O. Mittelsten Scheid
3.7 Manipulating Meiosis
500
References
501
502
503
504
505
Adamowski ED, Pagliarini MS, Batista LAR (1998) Chromosome elimination in paspalum
subciliatum (Notata group). Sex Plant Reprod 11(5):272–276
Agashe B, Prasad CK, Siddiqi I (2002) Identification and analysis of DYAD: a gene required for
meiotic chromosome organisation and female meiotic progression in Arabidopsis. Development 129(16):3935–3943
475
476
477
478
479
480
481
482
483
484
485
486
487
488
489
490
491
492
493
494
495
496
497
498
PR
OO
473
474
D
472
TE
471
EC
470
CO
RR
469
F
499
While natural selection in sexually reproducing species rapidly eradicates failures
in meiosis, these are of interest for plant breeders. If not the seed, but the
surrounding fruit is the wanted product, for example in Citrus species or melons,
customers prefer products with small, reduced, or no seeds. Besides exploiting
developmental mutations, one effective and popular way to achieve such fruits is
the generation of triploids (Sanchez-Moran et al. 2002) by crossing tetraploids
with diploids. Triploid embryos are prone to abort subsequently (Kamiri et al.
2011). Since fruit development is often regulated by phytohormones originating
from the developing seeds (Dorcey et al. 2009), this strategy depends on their
seed-independent substitution (Pandolfini 2009). However, once generated, triploid plants are often seedless, due to a high number of univalents in meiosis, a
resulting low number of balanced gametes, and poor pollination and fertilization
rates. This principle is applied in banana (Heslop-Harrison and Schwarzacher
2007) and watermelon (Beaulieu and Lea 2006), or is combined with selection for
developmental mutants as for squash (Menezes et al. 2005).
Once a beneficial combination of genetic traits by crossing is achieved, breeders
want to fix this for further progeny (Wijnker and de Jong 2008; Chan 2010).
Programed switches between crossing and apomixis would, therefore, be desirable
(Spillane et al. 2004), especially for polyploids with their complex segregation
patterns. Recent advances in provoking apomixis have been made by strategies to
disturb meiosis. A mutation in the Arabidopsis gene SWI/DYAD (Mercier et al. 2001;
Agashe et al. 2002) leads to the formation of unreduced egg cells, albeit at low
frequencies. This is due to failure in female meiosis and maintains complete maternal
heterozygosity in the triploid progeny (Ravi et al. 2008). Combining three mutations
affecting different steps of meiosis (osd1/Atspo11-1/Atrec8) turned meiosis into
mitosis and resulted in unreduced male and female gametes (d’Erfurth et al. 2009).
Consequently, ploidy in subsequent generations was doubled, accompanied by
reduced fertility. Nevertheless, the achievement of fertilization-independent seed
development is a step toward introducing apomixis into crop plants, and of special
interest for polyploid plants.
The work in the OMS lab was supported by grants from the Austrian Science
Fund FWF P18986 and I489.
468
UN
Editor Proof
48
Book ISBN: 978-3-642-31441-4
Page: 48/54
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 49/54
49
EC
TE
D
PR
OO
F
Albertin W, Marullo P, Aigle M, Bourgais A, Bely M, Dillmann C, De Vienne D, Sicard D
(2009) Evidence for autotetraploidy associated with reproductive isolation in Saccharomyces
cerevisiae: towards a new domesticated species. J Evol Biol 22(11):2157–2170
Anderson LK, Stack SM (2005) Recombination nodules in plants. Cytogenet Genome Res 109
(1–3):198–204
Andru S, Pan YB, Thongthawee S, Burner DM, Kimbeng CA (2011) Genetic analysis of the
sugarcane (Saccharum spp.) cultivar LCP 85-384’. I. Linkage mapping using AFLP, SSR, and
TRAP markers. Theor Appl Genet 123(1):77–93
Armstrong SJ, Franklin FCH, Jones GH (2003) A meiotic time-course for Arabidopsis thaliana.
Sex Plant Reprod 16(3):141–149
Asker S, Jerling L (1992) Apomixis in plants. CRC Press, Boca Raton
Avivi L, Feldman M (1980) Arrangement of chromosomes in the interphase nucleus of plants.
Hum Genet 55(3):281–295
Barth S, Melchinger AE, Devezi-Savula B, Lubberstedt T (2001) Influence of genetic background
and heterozygosity on meiotic recombination in Arabidopsis thaliana. Genome 44(6):
971–978
Bass HW, Marshall WF, Sedat JW, Agard DA, Cande WZ (1997) Telomeres cluster de novo
before the initiation of synapsis: a three-dimensional spatial analysis of telomere positions
before and during meiotic prophase. J Cell Biol 137(1):5–18
Baudat F, de Massy B (2007) Regulating double-stranded DNA break repair towards crossover or
non-crossover during mammalian meiosis. Chromosome Res 15(5):565–577
Bayliss MW, Riley R (1972) Analysis of temperature-dependent asynapsis in Triticum aestivum.
Genet Res 20(2):193
Beaulieu JC, Lea JM (2006) Characterization and semiquantitative analysis of volatiles in
seedless watermelon varieties using solid-phase microextraction. J Agric Food Chem
54(20):7789–7793
Bennett MD (1977) The time and duration of meiosis. Philos Trans R Soc Lond B Biol Sci
277(955):201–226
Bhatt AM, Canales C, Dickinson HG (2001) Plant meiosis: the means to 1 N. Trends Plant Sci
6(3):114–121
Borner GV, Kleckner N, Hunter N (2004) Crossover/noncrossover differentiation, synaptonemal
complex formation, and regulatory surveillance at the leptotene/zygotene transition of
meiosis. Cell 117(1):29–45
Brownfield L, Kohler C (2011) Unreduced gamete formation in plants: mechanisms and
prospects. J Exp Bot 62(5):1659–1668
Brubaker CL, Paterson AH, Wendel JF (1999) Comparative genetic mapping of allotetraploid
cotton and its diploid progenitors. Genome 42(2):184–203
Burton TL, Husband BC (2001) Fecundity and offspring ploidy in matings among diploid,
triploid and tetraploid Chamerion angustifolium (Onagraceae): consequences for tetraploid
establishment. Heredity 87:573–582
Butruille DV, Boiteux LS (2000) Selection-mutation balance in polysomic tetraploids: impact of
double reduction and gametophytic selection on the frequency and subchromosomal
localization of deleterious mutations. Proc Nat Acad Sci USA 97(12):6608–6613
Cao DC, Osborn TC, Doerge RW (2004) Correct estimation of preferential chromosome pairing
in autotetraploids. Genome Res 14(3):459–462
Carlton PM, Farruggio AP, Dernburg AF (2006) A link between meiotic prophase progression
and crossover control. PLoS Genet 2:119–128
Carman JG (1997) Asynchronous expression of duplicate genes in angiosperms may cause
apomixis, bispory, tetraspory, and polyembryony. Biol J Linn Soc 61(1):51–94
Carvalho A, Delgado M, Baroa A, Frescatada M, Ribeiro E, Pikaard CS, Viegas W, Neves N
(2010) Chromosome and DNA methylation dynamics during meiosis in the autotetraploid
Arabidopsis arenosa. Sex Plant Reprod 23(1):29–37
Chan SWL (2010) Chromosome engineering: power tools for plant genetics. Trends Biotechnol
28(12):605–610
CO
RR
506
507
508
509
510
511
512
513
514
515
516
517
518
519
520
521
522
523
524
525
526
527
528
529
530
531
532
533
534
535
536
537
538
539
540
541
542
543
544
545
546
547
548
549
550
551
552
553
554
555
556
557
558
559
UN
Editor Proof
3 Meiosis in Polyploid Plants
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
EC
TE
D
PR
OO
F
St Charles J, Hamilton ML, Petes TD (2010) Meiotic chromosome segregation in triploid strains
of Saccharomyces cerevisiae. Genetics 186(2):537–550
Cifuentes M, Grandont L, Moore G, Chevre AM, Jenczewski E (2010) Genetic regulation of
meiosis in polyploid species: new insights into an old question. New Phytol 186(1):29–36
Comai L (2005) The advantages and disadvantages of being polyploid. Nat Rev Genet 6(11):836–
846
Comai L, Tyagi AP, Lysak MA (2003) FISH analysis of meiosis in Arabidopsis allopolyploids.
Chromosome Res 11(3):217–226
Corredor E, Lukaszewski AJ, Pachon P, Allen DC, Naranjo T (2007) Terminal regions of wheat
chromosomes select their pairing partners in meiosis. Genetics 177(2):699–706
Cowan CR, Carlton PM, Cande WZ (2001) ‘The polar arrangement of telomeres in interphase
and meiosis. Rabl organization and the bouquet. Plant Physiol 125(2):532–538
d’Erfurth I, Jolivet S, Froger N, Catrice O, Novatchkova M, Mercier R (2009) Turning meiosis
into mitosis. PLoS Biol 7(6):e1000124
D’Souza TG, Storhas M, Schulenburg H, Beukeboom LW, Michiels NK (2004) Occasional sex in
an ‘asexual’ polyploid hermaphrodite. Proc Biol Sci 271(1543):1001–1007
Darlington CD (1929a) Chromosome behaviour and structural hybridity in the tradescantiae.
J Genet 21(2):207–286
Darlington CD (1929b) Polyploids and polyploidy. Nature 124:62–64
Dawe RK (1998) Meiotic chromosome organization and segregation in plants. Annu Rev Plant
Physiol Plant Mol Biol 49:371–395
De Muyt A, Mercier R, Mezard C, Grelon M (2009) Meiotic recombination and crossovers in
plants. Genome Dyn 5:14–25
Desai A, Chee PW, Rong J, May OL, Paterson AH (2006) Chromosome structural changes in
diploid and tetraploid A genomes of Gossypium. Genome 49(4):336–345
De Vicente MC, Tanksley SD (1991) Genome-wide reduction in recombination of backcross
progeny derived from male versus female gametes in an interspecific cross of tomato. Theor
Appl Genet 83(2):173–178
Doerge RW, Craig BA (2000) Model selection for quantitative trait locus analysis in polyploids.
Proc Nat Acad Sci USA 97(14):7951–7956
Dong F, Jiang J (1998) Non-Rabl patterns of centromere and telomere distribution in the
interphase nuclei of plant cells. Chromosome Res 6(7):551–558
Dorcey E, Urbez C, Blazquez MA, Carbonell J, Perez-Amador MA (2009) Fertilizationdependent auxin response in ovules triggers fruit development through the modulation of
gibberellin metabolism in Arabidopsis. Plant J 58(2):318–332
Drouaud J, Camilleri C, Bourguignon PY, Canaguier A, Berard A, Vezon D, Giancola S, Brunel
D, Colot V, Prum B et al (2006) Variation in crossing-over rates across chromosome 4 of
Arabidopsis thaliana reveals the presence of meiotic recombination ‘‘hot spots’’. Genome Res
16(1):106–114
Drouaud J, Mercier R, Chelyshev L, Bérard A, Falque M, Martin O, Zanni V, Brunel D, Mézard
C (2007) Sex-specific crossover distributions and variations in interference level along
Arabidopsis thaliana chromosome 4. PLoS Genet 3(6):e106
Edlinger B, Schlogelhofer P (2011) Have a break: determinants of meiotic DNA double strand
break (DSB) formation and processing in plants. J Exp Bot 62(5):1545–1563
Fisher RA (1947) The theory of linkage in polysomic inheritance. Philos Trans R Soc Lond B
Biol Sci 233(594):55–87
Francis KE, Lam SY, Harrison BD, Bey AL, Berchowitz LE, Copenhaver GP (2007) Pollen
tetrad-based visual assay for meiotic recombination in Arabidopsis. Proc Nat Acad Sci USA
104(10):3913–3918
Gaeta RT, Pires JC (2010) Homologous recombination in allopolyploids: the polyploid ratchet.
New Phytol 186(1):18–28
Griffiths S, Sharp R, Foote TN, Bertin I, Wanous M, Reader S, Colas I, Moore G (2006)
Molecular characterization of Ph1 as a major chromosome pairing locus in polyploid wheat.
Nature 439(7077):749–752
CO
RR
560
561
562
563
564
565
566
567
568
569
570
571
572
573
574
575
576
577
578
579
580
581
582
583
584
585
586
587
588
589
590
591
592
593
594
595
596
597
598
599
600
601
602
603
604
605
606
607
608
609
610
611
612
613
M.-L. Zielinski and O. Mittelsten Scheid
UN
Editor Proof
50
Book ISBN: 978-3-642-31441-4
Page: 50/54
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 51/54
51
EC
TE
D
PR
OO
F
Gupta PK, Fedak G (1985) Genetic control of meiotic chromosome pairing in polyploids in the
genus Hordeum. Can J Genet Cytol 27(5):515–530
Gustaffson A (1946) Apomixis in higher plants. Lunds Universitets Arsskrift 42(2):1–370
Hamant O, Ma H, Cande WZ (2006) Genetics of meiotic prophase I in plants. Annu Rev Plant
Biol 57:267–302
Harper L, Golubovskaya I, Cande WZ (2004) A bouquet of chromosomes. J Cell Sci 117(Pt 18):
4025–4032
Harrison CJ, Alvey E, Henderson IR (2010) Meiosis in flowering plants and other green
organisms. J Exp Bot 61(11):2863–2875
Heslop-Harrison JS, Schwarzacher T (2007) Domestication, genomics and the future for banana.
Ann Bot 100(5):1073–1084
Holliday R (1977) Recombination and meiosis. Philos Trans R Soc Lond B Biol Sci 277(955):
359–370
Horandl E, Cosendai AC, Rodewald J (2011) Origin and distribution of autopolyploids via
apomixis in the alpine species Ranunculus kuepferi (Ranunculaceae). Taxon 60(2):355–364
Jannoo N, Grivet L, David J, D’Hont A, Glaszmann JC (2004) Differential chromosome pairing
affinities at meiosis in polyploid sugarcane revealed by molecular markers. Heredity
93(5):460–467
Jenczewski E, Alix K (2004) From diploids to allopolyploids: the emergence of efficient pairing
control genes in plants. Crit Rev Plant Sci 23(1):21–45
Jenczewski E, Eber F, Grimaud A, Huet S, Lucas MO, Monod H, Chevre AM (2003) PrBn, a
major gene controlling homologous pairing in oilseed rape (Brassica napus) haploids.
Genetics 164(2):645–653
Jiao Y, Wickett NJ, Ayyampalayam S, Chanderbali AS, Landherr L, Ralph PE, Tomsho LP, Hu
Yi, Liang H, Soltis PS et al (2011) Ancestral polyploidy in seed plants and angiosperms.
Nature 473(7345):97–100
John B, Henderson SA (1962) Asynapsis and polyploidy in Schistocerca paranensis.
Chromosoma 13(2):111–147
Jones GH (1994) Meiosis in autopolyploid crepis-capillaris 3. Comparison of triploids and
tetraploids: evidence for nonindependence of autonomous pairing sites. Heredity 73:215–219
Jones GH, Vincent JE (1994) Meiosis in autopolyploid crepis capillaris 2. Autotetraploids.
Genome 37(3):497–505
Kamiri M, Stift M, Srairi I, Costantino G, Moussadik AE, Hmyene A, Bakry F, Ollitrault P,
Froelicher Y (2011) Evidence for non-disomic inheritance in a citrus interspecific tetraploid
somatic hybrid between C. reticulata and C. limon using SSR markers and cytogenetic
analysis. Plant Cell Rep 30(8):1415–1425
Kim S, Plagnol V, Hu TT, Toomajian C, Clark RM, Ossowski S, Ecker JR, Weigel D, Nordborg
M (2007) Recombination and linkage disequilibrium in Arabidopsis thaliana. Nat Genet
39(9):1151–1155
Knight E, Greer E, Draeger T, Thole V, Reader S, Shaw P, Moore G (2010) Inducing
chromosome pairing through premature condensation: analysis of wheat interspecific hybrids.
Funct Integr Genomics 10(4):603–608
Koehler C, Mittelsten OS, Erilova A (2010) The impact of the triploid block on the origin and
evolution of polyploid plants. Trends Genet 26(3):142–148
Leflon M, Grandont L, Eber F, Huteau V, Coriton O, Chelysheva L, Jenczewski E, Chevre AM
(2010) Crossovers get a boost in brassica allotriploid and allotetraploid hybrids. Plant Cell
22(7):2253–2264
Leitch AR, Leitch IJ (2008) Genomic plasticity and the diversity of polyploid plants. Science
320(5875):481–483
Li J, Das K, Fu G, Tong C, Li Y, Tobias C, Wu R (2010) Em algorithm for mapping quantitative
trait Loci in multivalent tetraploids. Int J Plant Genomics 2010:216547
Li XC, Barringer BC, Barbash DA (2009) The pachytene checkpoint and its relationship to
evolutionary patterns of polyploidization and hybrid sterility. Heredity 102(1):24–30
CO
RR
614
615
616
617
618
619
620
621
622
623
624
625
626
627
628
629
630
631
632
633
634
635
636
637
638
639
640
641
642
643
644
645
646
647
648
649
650
651
652
653
654
655
656
657
658
659
660
661
662
663
664
665
666
UN
Editor Proof
3 Meiosis in Polyploid Plants
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
EC
TE
D
PR
OO
F
Lim KB, Shen TM, Barba-Gonzalez R, Ramanna MS, Van Tuyl JM (2004) Occurrence of SDR
2 N-gametes in Lilium hybrids. Breed Sci 54(1):13–18
Lohmiller LD, De Muyt A, Howard B, Offenberg HH, Heyting C, Grelon M, Anderson LK
(2008) Cytological analysis of MRE11 protein during early meiotic prophase I in Arabidopsis
and tomato. Chromosoma 117(3):277–288
Loidl J (1989) Effects of elevated temperature on meiotic chromosome synapsis in Allium
ursinum. Chromosoma 97(6):449–458
Loidl J (1995) Meiotic chromosome pairing in triploid and tetraploid Saccharomyces cerevisiae.
Genetics 139(4):1511–1520
Lukaszewski AJ, Kopecky D (2010) The Ph1 locus from wheat controls meiotic chromosome
pairing in autotetraploid rye (Secale cereale L.). Cytogenet Genome Res 129(1–3):117–123
Luo ZW, Zhang RM, Kearsey MJ (2004) Theoretical basis for genetic linkage analysis in
autotetraploid species. Proc Nat Acad Sci USA 101(18):7040–7045
Luo ZW, Zhang Z, Zhang RM, Pandey M, Gailing O, Hattemer HH, Finkeldey R (2006)
Modeling population genetic data in autotetraploid species. Genetics 172(1):639–646
Ma CX, Casella G, Shen ZJ, Osborn TC, Wu RL (2002) A unified framework for mapping
quantitative trait loci in bivalent tetraploids using single-dose restriction fragments: a case
study from alfalfa. Genome Res 12(12):1974–1981
Madlung A, Tyagi AP, Watson B, Jiang HM, Kagochi T, Doerge RW, Martienssen R, Comai L
(2005) Genomic changes in synthetic Arabidopsis polyploids. Plant J 41(2):221–230
Martinez-Perez E, Shaw P, Aragon-Alcaide L, Moore G (2003) Chromosomes form into seven
groups in hexaploid and tetraploid wheat as a prelude to meiosis. Plant J 36(1):21–29
Martinez-Perez E, Shaw P, Moore G (2001) The Ph1 locus is needed to ensure specific somatic
and meiotic centromere association. Nature 411(6834):204–207
Mason AS, Nelson MN, Yan G, Cowling WA (2011) Production of viable male unreduced
gametes in Brassica interspecific hybrids is genotype specific and stimulated by cold
temperatures. BMC Plant Biol 11:103
Mather K (1936) Segregation and linkage in autotetraploids. J Genetics 32(2):287–314
McKee BD (1996) The license to pair: identification of meiotic pairing sites in Drosophila.
Chromosoma 105(3):135–141
McKim KS (2007) Meiotic pairing: a place to hook up. Curr Biol 17(5):R165–R168
Melamed-Bessudo C, Yehuda E, Stuitje AR, Levy AA (2005) A new seed-based assay for
meiotic recombination in Arabidopsis thaliana. Plant J 43(3):458–466
Menezes CB, Maluf WR, Azevedo SM, Faria MV, Nascimento IR, Nogueira DW, Gomes LA,
Bearzoti E (2005) Inheritance of parthenocarpy in summer squash (Cucurbita pepo L.). Genet
Mol Res 4(1):39–46
Mercier R, Grelon M (2008) Meiosis in plants: ten years of gene discovery. Cytogenet Genome
Res 120(3–4):281–290
Mercier R, Vezon D, Bullier E, Motamayor JC, Sellier A, Lefevre F, Pelletier G, Horlow C
(2001) SWITCH1 (SWI1): a novel protein required for the establishment of sister chromatid
cohesion and for bivalent formation at meiosis. Genes Dev 15(14):1859–1871
Mestiri I, Chague V, Tanguy AM, Huneau C, Huteau V, Belcram H, Coriton O, Chalhoub B,
Jahier J (2010) Newly synthesized wheat allohexaploids display progenitor-dependent meiotic
stability and aneuploidy but structural genomic additivity. New Phytol 186(1):86–101
Mezard C (2006) Meiotic recombination hotspots in plants. Biochem Soc Trans 34:531–534
Mezard C, Vignard J, Drouaud J, Mercier R (2007) The road to crossovers: plants have their say.
Trends Genet 23(2):91–99
Moore G, Shaw P (2009) Improving the chances of finding the right partner. Curr Opin Genet
Dev 19(2):99–104
Morrison JW, Rajhathy T (1960) Frequency of quadrivalents in autotetraploid plants. Nature
187(4736):528–530
Mursalimov SR, Deineko EV (2011) An ultrastructural study of cytomixis in tobacco pollen
mother cells. Protoplasma 248(4):717–724
CO
RR
667
668
669
670
671
672
673
674
675
676
677
678
679
680
681
682
683
684
685
686
687
688
689
690
691
692
693
694
695
696
697
698
699
700
701
702
703
704
705
706
707
708
709
710
711
712
713
714
715
716
717
718
719
M.-L. Zielinski and O. Mittelsten Scheid
UN
Editor Proof
52
Book ISBN: 978-3-642-31441-4
Page: 52/54
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 53/54
53
EC
TE
D
PR
OO
F
Naranjo T, Corredor E (2004) Clustering of centromeres precedes bivalent chromosome pairing
of polyploid wheats. Trends Plant Sci 9(5):214–217
Nelson MN, Nixon J, Lydiate DJ (2005) Genome-wide analysis of the frequency and distribution
of crossovers at male and female meiosis in Sinapis alba L. (white mustard). Theor Appl
Genet 111(1):31–43
Nicolas SD, Leflon M, Liu Z, Eber F, Chelysheva L, Coriton O, Chevre AM, Jenczewski E
(2008) Chromosome ‘speed dating’ during meiosis of polyploid Brassica hybrids and
haploids. Cytogenet Genome Res 120(3–4):331–338
Nicolas SD, Leflon M, Monod H, Eber F, Coriton O, Huteau V, Chevre AM, Jenczewski E (2009)
Genetic regulation of meiotic cross-overs between related genomes in Brassica napus
haploids and hybrids. Plant Cell 21(2):373–385
Nogler GA (1984) Gametophytic apomixis. In: Johri BM (ed) Embryology of angiosperms.
Springer, Berlin
Ortiz R (1997) Occurrence and inheritance of 2n pollen in Musa. Ann Bot 79(4):449–453
Osman K, Higgins JD, Sanchez-Moran E, Armstrong SJ, Franklin F, Chris H (2011) Pathways to
meiotic recombination in Arabidopsis thaliana. New Phytol 190(3):523–544
Ottaviano E, Sari Gorla M, Mulcahy DL (1990) Pollen selection: efficiency and monitoring. In:
Ogita ZI, Markert CL (eds) Isozymes: structure, function, and use in biology and medicine.
Wiley-Liss, New York, pp 575–588
Otto SP (2007) The evolutionary consequences of polyploidy. Cell 131(3):452–462
Ozkan H, Feldman M (2009) Rapid cytological diploidization in newly formed allopolyploids of
the wheat (Aegilops-Triticum) group. Genome 52(11):926–934
Pagliarini MS (2000) Meiotic behavior of economically important plant species: the relationship
between fertility and male sterility. Genet Mol Biol 23(4):997–1002
Pandolfini T (2009) Seedless fruit production by hormonal regulation of fruit set. Nutrients
1(2):168–177
Parisod C, Holderegger R, Brochmann C (2010) Evolutionary consequences of autopolyploidy.
New Phytol 186(1):5–17
Pawlowski WP, Cande WZ (2005) Coordinating the events of the meiotic prophase. Trends Cell
Biol 15(12):674–681
Pawlowski WP (2010) Chromosome organization and dynamics in plants. Curr Opin Plant Biol
13(6):640–645
Pecinka A, Fang W, Rehmsmeier M, Levy Avraham A, Mittelsten Scheid O (2011)
Polyploidization increases meiotic recombination frequency in Arabidopsis. BMC Biol 9:24
Pecrix Y, Rallo G, Folzer H, Cigna M, Gudin S, Le Bris M (2011) Polyploidization mechanisms:
temperature environment can induce diploid gamete formation in Rosa sp. J Exp Bot 62(10):
3587–3597
Peloquin SJ, Boiteux LS, Simon PW, Jansky SH (2008) A chromosome-specific estimate of
transmission of heterozygosity by 2n gametes in potato. J Hered 99(2):177–181
Ramsey J (2007) Unreduced gametes and neopolyploids in natural populations of Achillea
borealis (Asteraceae). Heredity 98(3):143–150
Ramsey J, Schemske DW (1998) Pathways, mechanisms, and rates of polyploid formation in
flowering plants. Annu Rev Ecol Syst 29:467–501
Ramsey J, Schemske DW (2002) Neopolyploidy in flowering plants. Annu Rev Ecol Syst
33:589–639
Ravi M, Marimuthu MP, Siddiqi I (2008) Gamete formation without meiosis in Arabidopsis.
Nature 451(7182):1121–1124
Ricci GCL, Pagliarini MS, Valle CB (2010) Genome elimination during microsporogenesis in
two pentaploid accessions of Brachiaria decumbens (Poaceae). Genet Mol Res 9(4):
2364–2371
Ridout MS, Bell JA, Simpson DW (2001) Analysis of segregation data from selfed progeny of
allopolyploids. Heredity 87:537–543
Riley R, Chapman V (1958) Genetic control of the cytologically diploid behaviour of hexaploid
wheat. Nature 182(4637):713–715
CO
RR
720
721
722
723
724
725
726
727
728
729
730
731
732
733
734
735
736
737
738
739
740
741
742
743
744
745
746
747
748
749
750
751
752
753
754
755
756
757
758
759
760
761
762
763
764
765
766
767
768
769
770
771
772
773
UN
Editor Proof
3 Meiosis in Polyploid Plants
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
EC
TE
D
PR
OO
F
Rose AM, Baillie DL (1979) Effect of temperature and parental age on recombination and
nondisjunction in Caenorhanditis elegans. Genetics 92(2):409–418
Sakuno T, Watanabe Y (2009) Studies of meiosis disclose distinct roles of cohesion in the core
centromere and pericentromeric regions. Chromosome Res 17(2):239–249
Sanchez-Moran E, Armstrong SJ, Santos JL, Franklin FCH, Jones GH (2002) Variation in
chiasma frequency among eight accessions of Arabidopsis thaliana. Genetics 162(3):
1415–1422
Sanchez-Moran E, Osman K, Higgins JD, Pradillo M, Cunado N, Jones GH, Franklin FCH (2008)
ASY1 coordinates early events in the plant meiotic recombination pathway. Cytogenet
Genome Res 120(3–4):302–312
Santos JL, Alfaro D, Sanchez-Moran E, Armstrong SJ, Franklin FC, Jones GH (2003) Partial
diploidization of meiosis in autotetraploid Arabidopsis thaliana. Genetics 165(3):1533–1540
Scherthan H (2007) Telomere attachment and clustering during meiosis. Cell Mol Life Sci 64(2):
117–124
Schoen I, Martens K (1998) DNA repair in ancient asexuals: a new solution to an old problem? J
Nat Hist 32:943–948
Schubert I, Shaw P (2011) Organization and dynamics of plant interphase chromosomes. Trends
Plant Sci 16(5):273–281
Sharbel TF, Voigt ML, Corral JM, Galla G, Kumlehn J, Klukas C, Schreiber F, Vogel H, Rotter B
(2010) Apomictic and sexual ovules of Boechera display heterochronic global gene
expression patterns. Plant Cell 22(3):655–671
Sheehan MJ, Pawlowski WP (2009) Live imaging of rapid chromosome movements in meiotic
prophase I in maize. Proc Natl Acad Sci USA 106(49):20989–20994
Simioni C, do Valle CB (2011) Meiotic analysis in induced tetraploids of Brachiaria decumbens
Stapf. Crop Breed Appl Biotech 11(1):43–49
Singhal VK, Kumar P (2008) Cytomixis during microsporogenesis in the diploid and tetraploid
cytotypes of Withania somnifera (L.) Dunal, 1852 (Solanaceae). Comp Cytogenet 2(1):85–92
Soltis DE, Soltis PS (1999) Polyploidy: recurrent formation and genome evolution. Trends Ecol
Evol 14(9):348–352
Spillane C, Curtis MD, Grossniklaus U (2004) Apomixis technology development-virgin births in
farmers fields? Nat Biotechnol 22(6):687–691
Stack SM, Anderson LK (2002) Crossing over as assessed by late recombination nodules is
related to the pattern of synapsis and the distribution of early recombination nodules in maize.
Chromosome Res 10(4):329–345
Stift M, Reeve R, van Tienderen PH (2010) Inheritance in tetraploid yeast revisited: segregation
patterns and statistical power under different inheritance models. J Evol Biol 23(7):1570–1578
Stift Marc, Berenos C, Kuperus P, van Tienderen PH (2008) Segregation models for disomic,
tetrasomic and intermediate inheritance in tetraploids: a general procedure applied to Rorippa
(Yellow cress) microsatellite data. Genetics 179(4):2113–2123
Sun X, Zhang Y, Yang S, Chen JQ, Hohn B, Tian D (2008) Insertion DNA promotes ectopic
recombination during meiosis in Arabidopsis. Mol Biol Evol 25(10):2079–2083
Sundstrom G, Larsson TA, Larhammar D (2008) Phylogenetic and chromosomal analyses of
multiple gene families syntenic with vertebrate Hox clusters. BMC Evol Biol 8:254
Sybenga J (1996) Chromosome pairing affinity and quadrivalent formation in polyploids: do
segmental allopolyploids exist? Genome 39(6):1176–1184
Szadkowski E, Eber F, Huteau V, Lode M, Coriton O, Jenczewski E, Chevre AM (2011)
Polyploid formation pathways have an impact on genetic rearrangements in resynthesized
Brassica napus. New Phytol 191(3):884–894
Szadkowski E, Eber F, Huteau V, Lode M, Huneau C, Belcram H, Coriton O, ManzanaresDauleux MJ, Delourme R, King GJ et al (2010) The first meiosis of resynthesized Brassica
napus, a genome blender. New Phytol 186(1):102–112
Trelles-Sticken E, Loidl J, Scherthan H (2003) Increased ploidy and KAR3 and SIR3 disruption
alter the dynamics of meiotic chromosomes and telomeres. J Cell Sci 116(12):2431–2442
CO
RR
774
775
776
777
778
779
780
781
782
783
784
785
786
787
788
789
790
791
792
793
794
795
796
797
798
799
800
801
802
803
804
805
806
807
808
809
810
811
812
813
814
815
816
817
818
819
820
821
822
823
824
825
826
M.-L. Zielinski and O. Mittelsten Scheid
UN
Editor Proof
54
Book ISBN: 978-3-642-31441-4
Page: 54/54
Layout: T1 Standard SC
Chapter No.: 3
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 55/54
55
EC
TE
D
PR
OO
F
van Veen JE, Hawley RS (2003) Meiosis: when even two is a crowd. Curr Biol 13(21):
R831–R833
Vizir IY, Korol AB (1990) Sex difference in recombination frequency in Arabidopsis. Heredity
65(3):379–383
von Wettstein D, Rasmussen SW, Holm PB (1984) The synaptonemal complex in genetic
segregation. Annu Rev Genet 18:331–413
Wang Y, Jha AK, Chen R, Doonan JH, Yang M (2010) Polyploidy-associated genomic instability
in Arabidopsis thaliana. Genesis 48(4):254–263
Watanabe K (1981) Studies on the control of diploid-like meiosis in polyploid taxa of
Chrysanthemim japonese. Cytologia 46(3):459–498
Weiss H, Maluszynska J (2000) Chromosomal rearrangement in autotetraploid plants of
Arabidopsis thaliana. Hereditas 133(3):255–261
Wijnker E, de Jong H (2008) Managing meiotic recombination in plant breeding. Trends Plant
Sci 13(12):640–646
Wu RL, Gallo-Meagher M, Littell RC, Zeng ZB (2001a) A general polyploid model for analyzing
gene segregation in outcrossing tetraploid species. Genetics 159(2):869–882
Wu SS, Wu RL, Ma CX, Zeng ZB, Yang MC, Casella G (2001b) A multivalent pairing model of
linkage analysis in autotetraploids. Genetics 159(3):1339–1350
Youds JL, Boulton SJ (2011) The choice in meiosis: defining the factors that influence crossover
or non-crossover formation. J Cell Sci 124(Pt 4):501–513
Yousafzai FK, Al-Kaff N, Moore G (2010) The molecular features of chromosome pairing at
meiosis: the polyploid challenge using wheat as a reference. Funct Integr Genomics
10(2):147–156
CO
RR
827
828
829
830
831
832
833
834
835
836
837
838
839
840
841
842
843
844
845
846
847
848
849
UN
Editor Proof
3 Meiosis in Polyploid Plants
Metadata of the chapter that will be visualized in
SpringerLink
Book Title
Polyploidy and Genome Evolution
Series Title
Chapter Title
Origins of Novel Phenotypic Variation in Polyploids
Copyright Year
2012
Copyright HolderName
Springer-Verlag Berlin Heidelberg
Corresponding Author
Family Name
Martienssen
Particle
Given Name
Robert A.
Suffix
Division
Author
Organization
Cold Spring Harbor Laboratory
Address
One Bungtown Road, 11724, Cold Spring Harbor, NY, USA
Email
martiens@cshl.edu
Family Name
Finigan
Particle
Given Name
Patrick
Suffix
Division
Organization
Cold Spring Harbor Laboratory
Address
One Bungtown Road, 11724, Cold Spring Harbor, NY, USA
Email
Author
Family Name
Tanurdzic
Particle
Given Name
Milos
Suffix
Division
Organization
Cold Spring Harbor Laboratory
Address
One Bungtown Road, 11724, Cold Spring Harbor, NY, USA
Email
Abstract
Polyploid species represent a special type of organism in nature, one that can survive and compete with three
or more full sets of homologous chromosomes. While less common in the animal and fungal kingdoms,
polyploid species are highly prevalent in the plant kingdom. Indeed, most agricultural crops are polyploids,
typically because polyploidy confers greater robustness and therefore higher yields. Among many examples
of novel phenotypic variation exhibited by polyploids are the production of larger fruits, reduced tillering,
delays in the reproductive transition, and even the creation of visually stunning flower pigmentation patterns
coveted by gardeners. The source of this novel variation in polyploids is still largely unclear. However,
multiple cellular mechanisms have been proposed, with some supporting evidence, to explain novel variation.
We review some of these mechanisms here.
Book ISBN: 978-3-642-31441-4
Page: 57/75
Chapter 4
4
Patrick Finigan, Milos Tanurdzic and Robert A. Martienssen
9
10
11
12
13
14
15
16
D
8
TE
7
Abstract Polyploid species represent a special type of organism in nature, one
that can survive and compete with three or more full sets of homologous
chromosomes. While less common in the animal and fungal kingdoms, polyploid
species are highly prevalent in the plant kingdom. Indeed, most agricultural crops
are polyploids, typically because polyploidy confers greater robustness and
therefore higher yields. Among many examples of novel phenotypic variation
exhibited by polyploids are the production of larger fruits, reduced tillering, delays
in the reproductive transition, and even the creation of visually stunning flower
pigmentation patterns coveted by gardeners. The source of this novel variation in
polyploids is still largely unclear. However, multiple cellular mechanisms have
been proposed, with some supporting evidence, to explain novel variation. We
review some of these mechanisms here.
EC
5
6
PR
OO
3
Origins of Novel Phenotypic Variation
in Polyploids
2
19
20
21
22
23
24
25
CO
RR
17
18
F
1
Book ID: 272454_1_En
Date: 16-8-2012
4.1 Prevalence and Significance of Polyploids
Polyploid organisms have three or more complete sets of homologous chromosomes (Winge 1917 #1369; Ramsey 2002 #1334). Polyploidy, or whole-genome
duplication (WGD), can arise through multiple ways—most often from unreduced
gametes following meiosis (Ahloowalia 1961 #1252; Harlan 1975 #1285;
Jørgensen 1928 #1290; Newton 1929 #1318; Skalinska 1946 #1344; Ramsey 1998
#1333; Bretagnolle 1995 #1258). In mammals, WGD events are considered to be
rare and typically lethal, presumably due to deleterious effects associated with
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: 4
P. Finigan M. Tanurdzic R. A. Martienssen (&)
Cold Spring Harbor Laboratory, One Bungtown Road, Cold Spring Harbor, NY 11724, USA
e-mail: martiens@cshl.edu
P. S. Soltis and D. E. Soltis (eds.), Polyploidy and Genome Evolution,
DOI: 10.1007/978-3-642-31442-1_4, Springer-Verlag Berlin Heidelberg 2012
57
Layout: T1 Standard SC
Chapter No.: 4
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 58/75
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
61
62
63
64
65
66
67
68
69
70
F
32
33
PR
OO
31
D
30
TE
29
EC
28
dosage (Bertrand 2010 #1255; Van de Peer 2009 #1361) see also Chap. 18, this
volume). However, genomic sequencing has revealed evidence for a few widespread ancient WGD events in the animal kingdom (Maere 2005 #1308; Dehal
2005 #1270; Kellis 2004 #1293; Ohno 1970 #1322; Meyer 2005 #1314, 1999
#1313) (see also Chaps. 16 and 17, this volume). Extant vertebrate genomes in
particular are believed to be the result of two or three separate WGD events during
evolution (Maere 2005 #1308; Dehal 2005 #1270; Kellis 2004 #1293; Ohno 1970
#1322; Meyer 2005 #1314, 1999 #1313) (Chap. 16, this volume). In plants,
polyploidy is generally tolerated, and most plant species are recent or ancient
polyploids (Lexer 2003 #1302; Soltis et al. 2009 #1346; Jiao 2011 #1289).
Polyploidy is thought to play a major role in speciation (Clausen 1945 #1267;
Grant 1981 #1282; Lumaret 1988 #1306), as polyploids are often reproductively
isolated from their progenitors—crosses between diploids and tetraploids generally yield triploid progeny with unbalanced meiosis, leading to polyploid infertility
(Darlington 1963 #1269; Grant 1981 #1282).
On a simplified level, Polyploid organisms can be either autopolyploids or
allopolyploids (Kihara 1926 #1296). Following one definition, an autopolyploid is
a Polyploid organism in which all of the chromosome sets are derived from the
same species, whereas an allopolyploid organism has chromosome sets derived
from different species (Kihara 1926 #1296). Allopolyploids are therefore permanent hybrids, but with a complete chromosome set from each parental species
(Kihara 1926 #1296). The complete parental chromosome sets in allopolyploids
allow for proper pairing of homologous chromosomes during meiosis (disomic
segregation), rather than missegregation of trivalents and quadrivalents as may be
found in autopolyploids (multisomic segregation) (Stebbins 1971 #1355; Ramsey
2002 #1334). Thus, allopolyploids have often been considered more stable than
autopolyploids, and ‘‘fix’’ hybrid genotypes in successive generations, along with
any beneficial or detrimental phenotypes that result (Winge 1932 #1370). However, it has been recognized that autopolyploids may have functional (either
disomic or multisomic) chromosome pairing and be highly fertile (Soltis et al.
2007 #1349; Soltis and Rieseberg 1986 #1347).
Many important agricultural crops are polyploids (Eigsti 1957 #1275). Polyploid crops typically demonstrate increased growth, including flower and fruit size,
and novel variation compared to their diploid counterparts that can make them
better suited as agricultural products (Stebbins 1971 #1355; Ramsey 2002 #1334;
Grant 1981 #1282). The modern-day bread wheat, Triticum aestivum, is an allohexaploid that is the combination of three different diploid species (Dubcovsky
2007 #1272) (see also Chap. 7, this volume). T. aestivum has largely replaced its
diploid progenitors and now accounts for about 95 % of the entire wheat crop
produced around the world (Dubcovsky 2007 #1272). In addition to wheat, many
staple crops (including corn, cotton, coffee, oat, canola, rye, apple, banana,
watermelon, potato, sugar cane, and soybean) are also recent or ancient polyploids
(Gaut 1997 #1281; Stebbins 1971 #1355; Ohno 1970 #1322; Shoemaker 1996
#1341; Lagercrantz 1996 #1298; Eigsti 1957 #1275). The reduced seed set of
certain polyploids (odd ploidal levels) can make them more desirable for
CO
RR
26
27
P. Finigan et al.
UN
Editor Proof
58
Layout: T1 Standard SC
Chapter No.: 4
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 59/75
59
79
4.2 Origin of Novel Variation in Neopolyploids
77
80
81
82
83
84
85
86
87
88
89
90
91
92
93
94
95
96
97
98
99
100
101
102
103
104
105
106
107
PR
OO
76
The prevalence of polyploids in the plant kingdom despite the rarity of their
formation (Ramsey 1998 #1333) indicates that polyploids may have a fitness
advantage compared to their progenitors (Stebbins 1950 #1354). The explanation
for this fitness advantage stems from the observation that many polyploids display
novel phenotypic variation compared to their progenitors (Randolph 1941 #1335;
Levin 1983 #1300, 2002 #1301; Lumaret 1988 #1306; Ramsey 2002 #1334;
Müntzing 1936 #1316). This novel phenotypic variation is believed to enable
polyploids to exploit different environmental niches better than their progenitors;
this variation bestows a fitness advantage on the polyploids (Clausen 1945 #1267;
Stebbins 1950 #1354). One caveat of this hypothesis is that studies documenting
the geographic distribution of polyploids and their diploid progenitors used extant
diploid relatives instead of the exact diploid progenitors (usually unknown)
(Stebbins 1971 #1355; Ramsey 2002 #1334). Thus, it is possible the real progenitors had the same geographic range distribution as the current polyploids. To
address this problem, many current studies of polyploids use synthetic polyploids
that can be compared to their exact diploid progenitors (Stebbins 1971 #1355;
Ramsey 2002 #1334). Future studies should help to clarify if range expansion
relative to their progenitors is a general characteristic of polyploids.
The emergence of novel phenotypes in polyploids, especially neoallopolyploids
(defined as early-generation polyploids), compared to their progenitors has been of
great interest because of its application in plant breeding and relevance to biodiversity and evolution (Eigsti 1957 #1275; Stebbins 1971 #1355; Ramsey 2002
#1334; Grant 1981 #1282). Genetic mechanisms encompassing gene dosage/allelic
combinations, novel gene interactions, genomic alterations, and epigenomic
reorganization have all been proposed to explain the origin of novel phenotypes of
polyploids (Fig. 4.1). The relevance of these different mechanisms for causing
novel variation is still largely unclear, with important distinctions between autopolyploids and allopolyploids.
D
75
TE
74
EC
73
CO
RR
72
F
78
consumption, like seedless watermelons and bananas. Moreover, allopolyploids
can overcome hybrid incompatibility between different species, allowing production of new varieties and introgression of favorable traits into important crops
(Eigsti 1957 #1275). Allohexaploid Nicotiana plants were used as a bridge to
transfer the gene responsible for tobacco mosaic virus resistance from wild to
commercial varieties of Nicotiana species (Eigsti 1957 #1275). This was only
possible through a Polyploid intermediate, as the interspecies Nicotiana hybrids
were sterile (Eigsti 1957 #1275) (see also Chap. 11, this volume).
71
UN
Editor Proof
4 Origins of Novel Phenotypic Variation in Polyploids
Layout: T1 Standard SC
Chapter No.: 4
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 60/75
Editor Proof
60
P. Finigan et al.
G
ag
os
en
D
PR
OO
lic
om
ic
lle
Al
/A
te
ic
en
G
F
(b)
ra
tio
ns
(a)
e
R
e
en
TE
G
g
in
(c)
el
od
eg
ul
em
D
at
R
or
ic
y
et
In
en
te
ig
ra
ct
Ep
io
ns
Novel Phenotypic Variation
(d)
109
110
111
112
113
114
4.2.1 Gene Dosage and Allelic Combinations May Result in Novel
Variation in Neopolyploids
Genome-wide increases in DNA content have been proposed to contribute to novel
phenotypes in neopolyploids (Randolph 1941 #1335). The ‘‘gigas effect’’ is a welldocumented phenotype associated with polyploid formation, where certain organs
are larger in the polyploids than their progenitors—but not necessarily the entire
plant (Randolph 1941 #1335; Stebbins 1971 #1355). The gigas effect is believed to
UN
108
CO
RR
EC
Fig. 4.1 Molecular mechanisms that may contribute to novel phenotypic variation. Four distinct
molecular mechanisms have been suggested to contribute to changes in gene expression and
function that may result in novel phenotypic variation in neoallopolyploids compared to their
progenitors—adapted from (Osborn 2003 #1324). The figure diagrams each mechanism by
showing the two progenitor states in comparison with their resultant polyploid offspring.
a Changes in genic or allelic dosage are diagramed by the inheritance of novel chromosome sets.
b Genomic alterations are shown by the inheritance of recombined chromosomes. c Novel gene
regulatory interactions are diagrammed by unique protein–protein interactions. d Epigenomic
reorganization is shown as the inheritance of large-scale changes in chromatin compaction. The
progenitors and their polyploid offspring are diagrammed by ellipses, with the progenitors
represented by the ellipses furthest from the central ‘‘novel phenotypic variation’’ ellipse. The
polyploids are represented by the ellipses closest to the central ellipse.
Layout: T1 Standard SC
Chapter No.: 4
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 61/75
121
122
123
124
125
126
127
128
129
130
131
132
133
134
135
136
137
138
139
140
141
142
143
144
145
146
147
148
149
150
151
152
153
154
155
156
157
158
F
120
PR
OO
119
D
118
TE
117
61
be a consequence of increasing the genome size, leading to the increase in cell size
and reduction of cell divisions that are commonly associated with neopolyploids
compared to their progenitors (Noggle 1946 #1321; Stebbins 1971 #1355).
Additionally, some evidence suggests that neopolyploids have a protracted
reproductive growth phase, with a delay in the onset of flowering and longer
reproductive growth (Stebbins 1971 #1355). While these changes in morphology
and growth may be the result of simply increasing the amount of DNA in neopolyploids, they may also stem from increasing the relative level of gene and
protein expression.
Gene and protein expression may be sensitive to gene dosage effects, whereby
increases in ploidy result in changes in expression that are not proportional to the
increase in gene dosage. Gene and protein expression studies in Zea, Solanum,
Helianthus, and Arabidopsis autoploid series have suggested that the vast majority
of changes in expression are proportional to gene dosage (Yu 2010 #1376; Pignatta
2010 #1327; Wang 2006a, b #1366; Stupar 2007 #1357; Guo 1996 #1283; Yao
2011 #1375; Riddle 2010 #1337; Church 2009 #1266). Pignatta et al. (2010)
created independent neo-autotetraploid lines of Arabidopsis thaliana (Columbia)
from the same homogeneous diploid progenitor. The researchers performed genome-wide gene expression studies to identify reproducible changes in gene
expression in the autotetraploids compared to the diploid progenitor. Only a few
genes, out of 26,107, were found that potentially displayed changes in gene
expression that were not proportional to the ploidal levels. Of note, resolution or
sampling limitations may have missed some dosage-sensitive genes (Pignatta 2010
#1327). A recent study with Columbia and Landsberg accessions of Arabidopsis
ploidal series found similar results to those above, with slightly higher numbers of
genes whose expression was disproportionately affected by changes in ploidy (Yu
2010 #1376). A greater role for gene dosage was identified in potato and maize
studies, as increasing the ploidal levels resulted in roughly 10 % of genes with
disproportionate effects on expression (Stupar 2007 #1357; Guo 1996 #1283; Yao
2011 #1375; Riddle 2010 #1337; Church 2009 #1266). Taken together, these
studies suggest that most gene expression is proportional to ploidal level. None of
these studies identified dosage-sensitive genes whose increase in expression correlated with any ploidy-dependent phenotype. One unanswered question is: what
effect does increasing the overall level of gene expression, without changing the
relative abundance of gene products, have on biological pathways?
In theory, increased gene dosage and mixtures in polyploids could result in
unique allelic combinations that would expand phenotypic diversity (Bingham
1979 #1256; Grant 1981 #1282; Stebbins 1971 #1355). Instead of possessing only
two alleles for a given gene as in a diploid, a tetraploid could possess four alleles
that may allow additional variability for a phenotypic trait. This mechanism of
allelic interaction is a natural extension of phenotypic variation in diploids
attributed to combinations of different alleles that affect quantitative traits (Guo
1994 #1377; Birchler 2001 #1257; Osborn 2003 #1324). However, it remains
unclear what allelic combinations contribute to novel variation in polyploids
EC
116
CO
RR
115
UN
Editor Proof
4 Origins of Novel Phenotypic Variation in Polyploids
Layout: T1 Standard SC
Chapter No.: 4
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 62/75
165
166
167
168
169
170
171
172
173
174
175
176
177
178
179
180
181
182
183
184
185
186
187
188
189
190
191
192
193
194
F
PR
OO
164
Hybrid gene interactions may contribute to phenotypic variation in allopolyploids
(Osborn 2003 #1324; Ramsey 2002 #1334). Similar to interspecies hybrids, allopolyploids have divergent genetic contributions from their progenitors. This
heterogeneity could result in perturbations in regulatory networks that could dramatically alter their outcomes (Ramsey 2002 #1334; Stebbins 1947 #1353;
Stebbins 1950 #1354; Osborn 2003 #1324). In synthetic Arabidopsis allopolyploids, altered regulatory interactions between the two progenitor-derived chromosome sets are thought to be responsible for delaying flowering time (Wang
2006a, b #1365). The A. arenosa FRIGIDA (FRI) gene is a transcription factor that
trans-activates the A. thaliana (Landsberg) Flowering Locus C (FLC) gene (Wang
2006a, b #1365). FLC is a major negative regulator of the reproductive transition in
Arabidopsis species, and its over-expression is predicted to delay flowering in the
Arabidopsis neoallopolyploids (Shindo 2005 #1340; Simpson 2002 #1342). While
FRI and FLC genes are present in both the progenitors, A. thaliana (Landsberg) FRI
and A. arenosa FLC alleles have reduced expression or function (Wang 2006a, b
#1365). Therefore, it is possible that the flowering delay might be explained by the
novel genic interaction between A. arenosa FRI and A. thaliana FLC in the neoallopolyploids (Wang 2006a, b #1365).
D
163
TE
162
4.2.2 Hybrid Regulatory Interactions Can Cause Novel
Phenotypic Variation
EC
161
(Osborn 2003 #1324), and indeed allelic variation in neopolyploids may be limited
due to genetic ‘‘bottlenecking’’ in their formation.
4.2.3 Chromosome Missegregation Can Lead to Novel Variation
in Neopolyploids
CO
RR
159
160
P. Finigan et al.
Perturbations in chromosome segregation have been proposed to be a major source
of novel phenotypic variation in polyploids (Stebbins 1971 #1355; Müntzing 1937
#1317; Ramsey 2002 #1334; Song 1993 #1352, 1995 #1351; Soltis and Soltis 1999
#1348). During meiotic prophase, the increased chromosome sets in polyploids
will often lead to multivalent chromosome pairing that can result in slower cell
division, unequal chromosome partitioning to daughter cells, and even homeologous recombination (Storchova 2006 #1356; Stebbins 1971 #1355; Ramsey 2002
#1334). For example, autotetraploids have four homologous chromosomes instead
of two, like their diploid counterparts. These four homologous chromosomes may
form univalents, trivalents, or quadrivalents during prophase that will cause lagging or unequal chromosome partitioning during anaphase and result in aneuploid
gametes (Ramsey 2002 #1334). The prevalence of multivalent formation in
UN
Editor Proof
62
Layout: T1 Standard SC
Chapter No.: 4
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 63/75
201
202
203
204
205
206
207
208
209
210
211
212
213
214
215
216
217
218
219
220
221
222
223
224
225
226
227
228
229
230
231
232
233
234
235
236
237
238
239
F
200
PR
OO
199
D
198
TE
197
63
polyploids is widespread; a survey of the available literature by Ramsey et al.
(2002) found that the mean multivalent frequency for auto- and allopolyploids was
estimated to be 28.8 and 8.0 %, respectively. The lower percentage of allopolyploids with multivalent pairing versus autopolyploids, is most likely the result of
sequence divergence between homeologous chromosomes in allopolyploids that
favors disomic pairing.
Multivalent pairing is expected to result in a much higher degree of aneuploidy
than disomic pairing and ultimately reduce the fertility of polyploids (Ramsey
2002 #1334; Stebbins 1971 #1355). Ramsey et al. (2002) compared the occurrence
of aneuploidy in gametic and sporophytic cells in different Polyploid species. They
found that the mean frequency of aneuploid pollen was approximately 40 %, while
the mean frequency of aneuploid progeny was an estimated 29 %. Importantly,
there were no studies on the frequency of aneuploid ovules, but if one were to
assume a similar ratio of aneuploidy as found in pollen, it would imply that 64 %
of all zygotes would be aneuploids. These differences in the expected and observed
percentages of aneuploid progeny suggest that some aneuploid gametes or progeny
do not survive and that chromosome segregation defects correlate with the reduced
fertility of polyploid. In contrast to the very different multivalent frequencies
observed at meiosis, the frequency of aneuploid gametes and progeny was similar
in auto- and allopolyploids. If aneuploidy requires multivalent pairing, it is unclear
how allopolyploids could have a similar level of aneuploidy as autopolyploids.
Either there is a much higher level of multivalent pairing in allopolyploids than
reported, or other factors contribute to the unexpected level of aneuploidy in
allopolyploids (Ramsey 2002 #1334; Chester 2012 #1265).
In addition to aneuploidy, multivalent pairing of homeologous chromosomes
can also result in genetic recombination that can severely disrupt genome organization and contribute to phenotypic variation (Stebbins 1971 #1355; Ramsey
2002 #1334; Song 1993 #1352, 1995 #1351). In allopolyploids, unequal recombination between chromosomes can result from synteny between homeologous
chromosomes. Changes in gene dosage, gene expression, the epigenetic landscape,
and even gene conversion are all possible outcomes of homeologous recombination in allopolyploids. Homeologous recombination has been observed in many
different resynthesized allopolyploid species (Wendel 2000 #1368; Osborn 2003
#1324; Doyle 2008 #1271; Hegarty 2008 #1287; Leitch 2008 #1299; Soltis and
Soltis 2009 #1346; Stebbins 1971 #1355; Grant 1981 #1282; Ramsey 2002 #1334;
Chester 2012 #1265; Pires 2004 #1328; Buggs 2009 #1259), suggesting it may be
a major mechanism of genome alteration and evolution. In synthetic Arabidopsis
allopolyploids derived from A. thaliana and A. arenosa progenitors, the nucleolus
organizer region (NOR) and 5S rDNA genetic regions have been shown to be lost
or recombined between the chromosomes derived from A. thaliana and A. arenosa
by the F3 generation (Pontes 2004 #1329). Rearrangements of the NOR and 5S
rDNA regions were also demonstrated in the naturally occurring Arabidopsis
allopolyploid, A. suecica. Genetic rearrangements in resynthesized Brassica napus
allopolyploids have been linked to phenotypic variation (Gaeta 2007 #1280; Pires
2004 #1328; Xiong 2011 #1372; Song 1995 #1351). In one striking example,
EC
196
CO
RR
195
UN
Editor Proof
4 Origins of Novel Phenotypic Variation in Polyploids
Layout: T1 Standard SC
Chapter No.: 4
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 64/75
247
248
249
250
251
252
253
254
255
256
257
258
259
260
261
262
263
264
265
266
267
268
269
270
271
272
273
274
275
276
277
278
F
PR
OO
246
4.2.4 Epigenetic Remodeling Can Result in Novel Variation
in Neopolyploids
Epigenetic remodeling of polyploid genomes has also been suggested to play a role
in the origin of novel phenotypes (Osborn 2003 #1324). Epigenetic changes
involving DNA methylation, histone post-translational modifications (PTMs),
histone replacement, and sRNA-mediated silencing have all been demonstrated to
affect gene expression levels in diploids (Wolffe 1999 #1371; Calarco 2011
#1378). Moreover, the dynamic nature of these epigenetic modifications in combination with the instability of neopolyploid genomes would suggest that there is
the potential for rampant epigenetic remodeling in polyploids that could affect
gene expression (Osborn 2003 #1324; Wolffe 1999 #1371; Ramsey 2002 #1334;
Matzke 1999 #1310).
Concurrently, there have been many studies documenting novel changes in
DNA methylation (Li 2010 #1303; Chen 2008 #1264; Lukens 2006 #1305; Xu
2009 #1373; Wang 2009 #1364, 2004 #1367; Liu 2003 #1304; Madlung 2002
#1307; Kenan-Eichler 2011 #1294; Parisod 2009 #1325; Yaakov 2011 #1374) as
well as changes in sRNA profiles (Kenan-Eichler 2011 #1294; Ha 2009 #1284;
Preuss 2008 #1331) within allopolyploid genomes compared to their progenitors,
but there has been limited data demonstrating links of these epigenetic changes to
phenotypic consequences. One classic example of epigenetic reprogramming in
allopolyploids is the occurrence of nucleolar dominance among interspecies
hybrids, including allopolyploids (McStay 2006 #1311; Preuss 2007 #1330).
Nucleolar dominance occurs when rRNA genes from one parent, or progenitor
species, are preferentially silenced in a hybrid; this silencing is neither the result of
random inactivation, nor correlated with imprinting and sexual parent of origin
(McStay 2006 #1311; Preuss 2007 #1330). In Arabidopsis allopolyploids, small
interfering RNAs (sRNA), DNA methylation, and histone deacetylation result in
the silencing of A. thaliana-derived rRNA genes in A. suecica (Preuss 2007 #1330,
2008 #1331; Earley 2006 #1274). While no clear phenotype is associated with
nucleolar dominance, it is believed to be important for genome stability and
prevention of premature aging, which has been linked to perturbations in rDNA
genic regions (Finigan 2008 #1277).
D
245
TE
243
244
EC
242
variation for flowering time between two different B. napus allopolyploid lines was
found to correlate with a genomic rearrangement disrupting the expression of FLC
(Pires 2004 #1328). These allopolyploid lines also displayed phenotypic variation
in flowering time that exceeded the range of their progenitors (Gaeta 2007 #1280;
Pires 2004 #1328). Together, these results provide powerful evidence for the role
of genomic rearrangements in neopolyploids driving the creation of novel phenotypic variation that is not present in their diploid progenitors.
CO
RR
240
241
P. Finigan et al.
UN
Editor Proof
64
Layout: T1 Standard SC
Chapter No.: 4
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 65/75
286
287
288
289
290
291
292
293
294
295
296
297
298
299
300
301
302
303
304
305
306
307
308
309
310
311
312
313
314
315
316
317
318
319
320
321
322
323
F
PR
OO
284
285
D
283
TE
282
EC
281
65
For histone PTMs, a few studies in Arabidopsis neoallopolyploids demonstrate
changes in histone modification patterns that correlate with changes in gene
expression (Wang 2006a, b #1365; Ni et al. 2009 #1320). Interestingly, these
changes in gene expression patterns were linked to phenotypic changes in flowering time (Wang 2006a, b #1365) and chlorophyll and sugar content (Ni et al.
2009 #1320). However, these perturbations are most likely the downstream consequences of novel regulatory interactions (Ni et al. 2009 #1320; Wang 2006a, b
#1365). Additionally, a tentative link between seed death and perturbations in
histone H3 lysine 27 methylation (H3K27me) has been identified in Arabidopsis
neoallopolyploids (Josefsson 2006 #1291; Walia 2009 #1363). Josefsson et al.
(2006) reported that the expression of PHERES1, MEDEA, and other imprinted
genes were deregulated in developing Arabidopsis neoallopolyploid seeds. In
addition to the imprinted genes, Athila retrotransposons from the pollen parent
were also deregulated in Arabidopsis allotriploid seed, which are inviable. This
deregulation of imprinted genes and Athila elements was suggested to result from
a lack of silencing of their pollen parent copies by the POLYCOMB REPRESSIVE
COMPLEX (PRC), which is responsible for H3K27me in Arabidopsis plants
(Josefsson 2006 #1291). Subsequent work suggested that the PRC complex was
not functioning in these neoallopolyploids because of a 2–10-fold down-regulation
in the expression of FERTILIZATION-INDEPENDENT SEED 2 (FIS2), one of the
subunits of the PRC (Walia 2009 #1363). Why the PRC complex, and FIS2, are
misregulated in the developing neoallopolyploids is still an unanswered question,
but major disruptions of at least H3K27 methylation patterns in these neoallopolyploids would be predicted by this model (Josefsson 2006 #1291; Walia 2009
#1363). Future studies to analyze H3K27 methylation patterns in the neoallopolyploids will be important to validate this hypothesis. This misregulation of the
PRC complex is only present in the neoallopolyploids but absent in its autopolyploid progenitors and depends on both the ploidy level and genetic diversity of
the progenitors (Josefsson 2006 #1291; Walia 2009 #1363).
So far, the limited evidence for epigenetic remodeling has been primarily
identified in allo-, and not autopolyploids. While allopolyploids inherit epigenetic
and genomic divergence from their progenitors, one might still expect to see
epigenomic remodeling in autopolyploids due to their dynamic nature. One role
for epigenetic mechanisms in autopolyploids is in the origin of unreduced gametes
from which they arise. In the ovules of most sexual flowering plants, female
gametogenesis is initiated from a single surviving gametic cell, the functional
megaspore, formed after meiosis of the somatically derived megaspore mother cell
(MMC). The Arabidopsis small RNA binding protein ARGONAUTE 9 (AGO9)
controls female gamete formation by restricting the specification of gametophyte
precursors to the MMC (Olmedo-Monfil 2010 #1323; Durán-Figueroa 2010
#1273). Mutations in AGO9 lead to the differentiation of diploid (unreduced)
gametic cells from the surrounding ovule that are able to initiate gametogenesis.
Mutations in the maize AGO9 homolog also lead to unreduced gametes, through a
related but distinct mechanism (Singh 2011 #1343). AGO9 preferentially interacts
with 24-nucleotide sRNAs derived from transposable elements (TEs), and its
CO
RR
279
280
UN
Editor Proof
4 Origins of Novel Phenotypic Variation in Polyploids
Layout: T1 Standard SC
Chapter No.: 4
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 66/75
330
331
332
333
334
335
336
337
338
339
340
341
342
343
344
345
346
347
348
349
350
351
352
353
354
355
356
357
358
359
360
361
362
363
F
329
PR
OO
328
4.2.5 Nonadditive Gene Expression and Novel Phenotypic
Variation
Mechanisms encompassing gene dosage/allelic combination, novel gene interactions, genomic alterations, and epigenomic reorganization must all converge on
gene expression and function to induce novel phenotypic variation. While little is
known about perturbations in gene function in neopolyploids, changes in gene
expression (Wang 2006a, b #1366, #1365; Akhunova 2010 #1253; Pumphrey 2009
#1332; Chaudhary 2009 #1263; Chagué 2010 #1261; Flagel 2008 #1278, 2010
#1279; Hegarty 2006 #1286) and protein levels (Albertin 2006 #1254; Ng 2011
#1319) have been demonstrated in allopolyploids compared to their progenitors.
Transcript profiling experiments in synthetic Arabidopsis allopolyploids were
the first demonstrations of genome-wide nonadditive gene expression in allopolyploids (Wang 2006a, b #1366). Nonadditive gene expression occurs when the
expression level of genes in hybrids differs from the average expression level of
the orthologous genes in the parental species (i.e. midparent level). In Arabidopsis
neoallopolyploids, an estimated 5–35 % of all genes were nonadditively expressed—depending on the statistical method employed (Wang 2006a, b #1366).
Subsequent research in other neoallopolyploid species has suggested that nonadditive gene expression is a general trend found associated with recent allopolyploidization; however, the amount of nonadditive gene expression in specific
allopolyploids is highly variable (Hegarty 2006 #1286; Chaudhary 2009 #1263;
Flagel 2008 #1278, 2010 #1279; Akhunova 2010 #1253; Pumphrey 2009 #1332;
Chagué 2010 #1261). For example, in bread wheat allohexaploids nonadditive
gene expression has been reported as anywhere from 7 to 40 % (Pumphrey 2009
#1332; Chagué 2010 #1261; Kashkush 2002 #1292; Akhunova 2010 #1253).
The discovery of nonadditive gene expression in neoallopolyploids implies that
the progenitor-derived chromosome sets must be interacting to result in novel gene
expression patterns. How these chromosome sets interact is just beginning to be
unraveled; but, by definition, these novel interactions must stem directly from
changes in cis- and trans-regulatory divergence between the progenitor-derived
genomes. Cis- and/or trans-regulatory differences could result in changes in the
expression of homeologous genes (homeoalleles) from each progenitor-derived
chromosome set. Cis-regulatory divergence can act directly on single genes or on
D
327
TE
326
activity is necessary to silence TEs in female gametes and their accessory cells
(Olmedo-Monfil 2010 #1323; Durán-Figueroa 2010 #1273). That AGO9-dependent sRNA silencing is also crucial to specify ploidy in the gametes indicates that
epigenetic reprogramming may link transposon silencing to germ cell fate (Slotkin
2009 #1345). This may reflect the coevolution of transposon activity and sexual
reproduction, as well as the increased tolerance of polyploids to mutations caused
by TE insertions (Martienssen 2010 #1309).
EC
325
CO
RR
324
P. Finigan et al.
UN
Editor Proof
66
Layout: T1 Standard SC
Chapter No.: 4
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 67/75
371
372
373
374
375
376
377
378
379
380
381
382
383
384
385
386
387
388
389
390
391
392
393
394
395
396
397
398
399
400
401
402
403
F
370
PR
OO
368
369
D
367
TE
366
localized chromatin domains, such as promoters or enhancers, resulting in
asymmetric accumulation of homeologous transcripts in allopolyploids. Transregulatory divergence between the parental genomes will affect homeologous
genes equally and result in equal accumulation of homeologous transcripts. Thus,
both cis- and trans-regulatory differences could result in nonadditive homeologous
gene expression, but only cis-regulatory differences could lead to biased homeoallele-specific gene expression. The relative contribution of cis- and trans-regulatory divergence to nonadditive gene expression is still unclear, but some
evidence has arisen recently implicating a major role of cis-regulatory differences
for this phenomenon.
Transcriptome profiling studies in Gossypium (Flagel 2010 #1279; Doyle 2008
#1271), Arabidopsis (Wang 2006a, b #1366), and maize (Schnable 2011 #1339),
and small-scale analyses in Tragopogon (Buggs 2011 #1260; Tate 2006 #1359)
polyploids have suggested that there is a common theme of ‘‘genome dominance’’
in homeologous gene expression in allopolyploids (Rapp 2009 #1336). In the
genome dominance model, one of the progenitor transcription profiles outcompetes the other to shift the overall expression in the allopolyploids toward a progenitor-specific expression profile. In Gossypium allopolyploids, five different
allopolyploids have been suggested to have a 54–60 % transcriptional bias in favor
of the D genome versus the A genome (Flagel 2010 #1279). A similar phenomenon has been reported in Arabidopsis allopolyploids, where a 55 % bias toward
A. arenosa over A. thaliana homeoallele expression has been reported for natural
allopolyploids (Chang 2010 #1262) and suggested for synthetic allopolyploids
(Wang 2006a, b #1366). In contrast, the evidence for genome dominance has been
inconclusive in Triticum (Akhunova 2010 #1253; Pumphrey 2009 #1332; Chagué
2010 #1261). However, this phenomenon is still un-resolved, because the majority
of the evidence for genome dominance are based on comparing the expression
profiles between the progenitors and their allopolyploid offspring, without distinguishing between homeoalleles in the allopolyploids (Flagel 2008 #1278, 2010
#1279; Wang 2006a, b #1366). Further, in cases where genome-wide profiles of
homeologous gene expression have suggested biased expression of one genome
over the other (Akhunova 2010 #1253; Chang 2010 #1262; Schnable 2011 #1339),
the exact progenitors of these established allopolyploids were extinct and true
comparisons could not be made. Future studies should address these discrepancies
and help to unravel the phenotypic consequences of nonadditive gene expression.
EC
365
67
CO
RR
364
4.3 Technical Considerations When Comparing Gene
Expression Studies in Polyploids
UN
Editor Proof
4 Origins of Novel Phenotypic Variation in Polyploids
Many of the controversies surrounding genome dominance and the variability of
nonadditive gene expression in neoallopolyploids may stem from the differences in
the methodologies employed and the organisms studied. One important distinction
Layout: T1 Standard SC
Chapter No.: 4
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 68/75
Editor Proof
68
P. Finigan et al.
(a)
Species α‘ (4x = 2n)
Polyploidization
Species α‘ β‘ (4x = 2n)
Hybridization
Polyploidization
(b)
Species α (2x = 2n)
‘
Species α‘ β (2x = 1n)
PR
OO
Species β‘ (4x = 2n)
Species β (2x = 2n)
F
Species α (2x = 2n)
Species α‘ β‘ (4x = 2n)
Polyploidization
Hybridization
TE
D
Species β (2x = 2n)
405
406
407
408
409
410
411
412
413
414
415
416
417
418
is whether polyploidization preceded hybridization or vice versa (Hegarty 2008
#1287). If polyploidization precedes hybridization then the resulting neoallopolyploid will have duplicate chromosomes that can accurately pair at meiosis
(Fig. 4.2). Synthetic Arabidopsis allopolyploids are an example, and natural
Arabidopsis allopolyploids are believed to have arisen this way (Hegarty 2008
#1287; Josefsson 2006 #1291). On the other hand, if polyploidization follows
hybridization, then the resulting hybrid will first transit through an unbalanced
chromosome state, most likely severely infertile, before restoration with a polyploidization event to create a neoallopolyploid (Fig. 4.2) (Hegarty 2006 #1286,
2008 #1287). Gossypium, Triticum, Tragopogon, Spartina, and Senecio allopolyploids are examples of the latter type (Hegarty 2006 #1286). Transcriptome
profiling studies in Senecio (Hegarty 2006 #1286) and Triticum (Feldman 2005
#1276) hybrids before and after polyploidization suggested that the homologous
gene expression levels were more perturbed (vs. the midparent) in the sterile
hybrids before chromosome doubling. These results may suggest that gene
UN
404
CO
RR
EC
Fig. 4.2 The origin of allopolyploids. Two types of allopolyploids are distinguished by the
relative order of polyploidization and hybridization. a Type I allopolyploids arise when
polyploidization precedes hybridization, typically via unreduced gametes. Type I neoallopolyploids will have duplicate chromosomes that properly pair in meiosis. b Type II allopolyploids are
created when hybridization precedes polyploidization. Type II allopolyploids transit through a
hybrid state, which is usually sterile, as chromosomes do not pair during meiosis. A subsequent
polyploidization event restores 2n chromosome number in allopolyploids
Layout: T1 Standard SC
Chapter No.: 4
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 69/75
69
455
4.4 Conclusions
426
427
428
429
430
431
432
433
434
435
436
437
438
439
440
441
442
443
444
445
446
447
448
449
450
451
452
453
456
457
458
459
PR
OO
425
D
423
424
TE
422
EC
421
CO
RR
420
F
454
expression profiles in the resulting allopolyploid may be influenced by the creation
of a hybrid intermediate.
Another important distinction is the nature of the progenitor species used to
compare with allopolyploid expression profiles. Many allopolyploids do not have
extant progenitors, and relatives must be used instead (Chang 2010 #1262; Rapp
2009 #1336; Flagel 2010 #1279). These relatives may not be exact genetic matches (Chang 2010 #1262; Rapp 2009 #1336; Flagel 2010 #1279) and may contain
epigenetic differences as well, as is the case for different accessions of A. thaliana
(Vaughn 2007 #1362; Kliebenstein 2006 #1297). A recent study illustrates this
point. An estimated 6,790 homologous genes were differentially expressed
between A. suecica and synthetic Arabidopsis allopolyploids (Chang 2010 #1262),
and comparative genome sequencing revealed that 938 homeologous genes were
missing from A. suecica compared to the synthetic allopolyploid (Chang 2010
#1262). These genomic differences are comparable to those between different
Arabidopsis thaliana accessions (Vaughn 2007 #1362; Kliebenstein 2006 #1297)
and may represent differences between the A. suecica progenitors and their extant
relatives.
Additionally, some studies employ progenitor species that have been severely
genetically manipulated (Akhunova 2010 #1253; Chang 2010 #1262; Kerber 1964
#1295; Wang 2006a, b #1366). Previous analysis of fifth-generation Arabidopsis
allopolyploids have relied on a tetraploidized A. thaliana (Landsberg accession) line
produced spontaneously from root explant regenerants that underwent a callus phase
and cell culture treatment (Chang 2010 #1262; Valvekens 1988 #1360; Wang 2006a,
b #1366; Comai 2000 #1268). Plant lines regenerated from callus/cell cultures
typically display heritable changes in genetic and epigenetic regulation—namely,
somaclonal variation (Phillips 1994 #1326; Meins 2003 #1312; Mohan Jain 2001
#1315; Tanurdzic 2008 #1379). These changes could have a profound effect on gene
expression and epigenetic regulation in neoallopolyploids derived from such progenitors. Classical approaches based on colchicine treatment have also been applied
to the generation of tetraploid A. thaliana lines (Henry 2005 #1288; Santos 2003
#1338). Studies in Gossypium (Rapp 2009 #1336; Chaudhary 2009 #1263), Triticum
(Akhunova 2010 #1253; Pumphrey 2009 #1332; Chagué 2010 #1261), and Tragopogon (Tate et al. 2009) have also relied on colchicine treatments to polyploidize
sterile triploid hybrids after fertilization. Importantly, there was no difference in
genetic or DNA methylation changes between spontaneous or colchicine produced
Brassica polyploids (Gaeta 2007 #1280).
419
UN
Editor Proof
4 Origins of Novel Phenotypic Variation in Polyploids
The prevalence of polyploidy in nature suggests it has been positively selected for
during evolution. Indeed, the higher yields of polyploids among agricultural crops
and their usefulness to bridge species barriers have led to their widespread use and
creation for the agricultural industry (Eigsti 1957 #1275). Understanding the
Layout: T1 Standard SC
Chapter No.: 4
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 70/75
P. Finigan et al.
472
References
473
474
475
476
477
478
479
480
481
482
483
484
485
486
487
488
489
490
491
492
493
494
495
496
497
498
499
500
501
502
503
504
505
Ahloowalia B, Garber F (1961) The genus Collinsia. XIII. Cytogenetic studies of interspecific
hybrids involving species with pediceled flowers. Bot Gaz 122:219
Akhunova A, Matniyazov R, Liang H, Akhunov E (2010) Homoeolog-specific transcriptional
bias in allopolyploid wheat. BMC Genomics 11(1):505
Albertin W, Balliau T, Brabant P, Chevre A-M, Eber F, Malosse C, Thiellement H (2006)
Numerous and rapid Nonstochastic modifications of Gene products in newly synthesized
10.1534/
Brassica
napus
Allotetraploids.
Genetics
173(2):1101–1113.
doi:
genetics.106.057554
Bertrand D, Gagnon Y, Blanchette M, El-Mabrouk N (2010) Reconstruction of ancestral genome
subject to whole genome duplication, speciation, rearrangement and loss. Paper presented at
the Proceedings of the 10th international conference on algorithms in bioinformatics,
Liverpool, UK
Bingham E (1979) Maximizing heterozygosity in autopolyploids. Basic Life Sci 13:471–489
Birchler JA, Bhadra U, Bhadra MP, Auger DL (2001) Dosage-dependent Gene regulation in
multicellular Eukaryotes: implications for dosage compensation, Aneuploid syndromes, and
quantitative traits. Dev Biology 234(2):275–288. doi: 10.1006/dbio.2001.0262
Bretagnolle F, Thompson J (1995) Tansley review no. 78. Gametes with the somatic chromosome
number: mechanisms of their formation and role in the evolution of Autopolyploid plants.
New Phytol 129:1
Buggs RJA, Doust AN, Tate JA, Koh J, Soltis K, Feltus FA, Paterson AH, Soltis PS, Soltis DE
(2009) Gene loss and silencing in Tragopogon miscellus (Asteraceae): comparison of natural
and synthetic Allotetraploids. Heredity 103 (1):73–81. doi: http://www.nature.com/hdy/
journal/v103/n1/suppinfo/hdy200924s1.html
Buggs Richard JA, Zhang L, Miles N, Tate Jennifer A, Gao L, Wei W, Schnable Patrick S,
Barbazuk WB, Soltis Pamela S, Soltis Douglas E (2011) Transcriptomic shock generates
evolutionary novelty in a newly formed. Natural allopolyploid plant. Curr Biol
21(7):551–556. doi: 10.1016/j.cub.2011.02.016
Calarco JP, Martienssen RA (2011) Genome reprogramming and small interfering RNA in the
Arabidopsis germline. Curr Opin Genet Dev 21(2):134–139. doi: 10.1016/j.gde.2011.01.014
Chagué V, Just J, Mestiri I, Balzergue S, Tanguy A-M, Huneau C, Huteau V, Belcram H, Coriton
O, Jahier J, Chalhoub B (2010) Genome-wide gene expression changes in genetically stable
synthetic and natural wheat allohexaploids. New Phytol 187(4):1181–1194. doi: 10.1111/
j.1469-8137.2010.03339.x
466
467
468
469
470
PR
OO
465
D
464
TE
463
EC
462
CO
RR
461
F
471
emergent properties of neopolyploids, such as novel phenotypic variation and
reproductive barriers, is vitally important to further advance the usefulness of
polyploids for crop breeding. While we are beginning to understand the molecular
mechanisms that contribute to novel variation in polyploids, there is still a lack of
specific links between genes, proteins, and phenotypes. We also do not understand
the interplay between these different mechanisms and the important drivers of
phenotypic diversity. What role do genomic alterations or allelic diversity play in
the origin of this novel phenotypic variation? Many homeologous genes are
nonadditively expressed in neoallopolyploids, but what is the source of this differential expression? If nonadditive gene expression is a result of genome dominance, than how is such a mechanism established? The answers to these questions
are likely to emerge in the coming years.
460
UN
Editor Proof
70
Layout: T1 Standard SC
Chapter No.: 4
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 71/75
71
EC
TE
D
PR
OO
F
Chang P, Dilkes B, McMahon M, Comai L, Nuzhdin S (2010) Homoeolog-specific retention and
use in allotetraploid Arabidopsis suecica depends on parent of origin and network partners.
Genome Biol 11(12):R125
Chaudhary B, Flagel L, Stupar R, Udall J, Verma N, Springer N, Wendel J (2009) Reciprocal
silencing, transcriptional bias and functional divergence of homeologs in polyploid cotton
(Gossypium). Genetics 182:503–517
Chen M, Ha M, Lackey E, Wang J, Chen Z (2008) RNAi of met1 reduces DNA methylation and
induces genome-specific changes in gene expression and centromeric small RNA accumulation in Arabidopsis Allopolyploids. Genetics 178:1845–1858
Chester M, Gallagher JP, Symonds VV, Cruz da Silva AV, Mavrodiev EV, Leitch AR, Soltis PS,
Soltis DE (2012) Extensive chromosomal variation in a recently formed natural allopolyploid
species, Tragopogon miscellus (Asteraceae). Proc Nat Acad Sci 109(4):1176–1181. doi:
10.1073/pnas.1112041109
Church SA, Spaulding EJ (2009) Gene expression in a wild Autopolyploid sunflower series.
J Hered 100(4):491–495. doi: 10.1093/jhered/esp008
Clausen J, Keck D, Hiesey W (1945) Experimental studies on the nature of species. II. Plant
evolution through Amphiploidy and autoploidy, with examples from the Madiinae. Carnegie
Inst Wash Publ 564
Comai L, Tyagi AP, Winter K, Holmes-Davis R, Reynolds SH, Stevens Y, Byers B (2000)
Phenotypic instability and rapid Gene silencing in newly formed Arabidopsis Allotetraploids.
Plant Cell 12(9):1551–1568. doi: 10.1105/tpc.12.9.1551
Darlington CD (1963) Chromosome botany, and the origins of cultivated plants. Allen and
Unwin, London
Dehal P, Boore JL (2005) Two rounds of whole genome duplication in the Ancestral Vertebrate.
PLoS Biol 3(10):e314
Doyle JJ, Flagel LE, Paterson AH, Rapp RA, Soltis DE, Soltis PS, Wendel JF (2008)
Evolutionary genetics of genome merger and doubling in plants. Annu Rev Genet
42(1):443–461. doi: 10.1146/annurev.genet.42.110807.091524
Dubcovsky J, Dvorak J (2007) Genome plasticity a key factor in the success of polyploid wheat
under domestication. Science 316(5833):1862–1866. doi: 10.1126/science.1143986
Durán-Figueroa N, Vielle-Calzada J-P (2010) ARGONAUTE9-dependent silencing of transposable elements in pericentromeric regions of Arabidopsis. Plant Signal Behav 5(11):1476–1479
Earley K, Lawrence RJ, Pontes O, Reuther R, Enciso AJ, Silva M, Neves N, Gross M, Viegas W,
Pikaard CS (2006) Erasure of histone acetylation by Arabidopsis HDA6 mediates large-scale
gene silencing in nucleolar dominance. Genes Dev 20(10):1283–1293. doi: 10.1101/
gad.1417706
Eigsti OJ (1957) Induced Polyploidy. Am J Bot 44(3):272–279
Feldman M, Levy AA (2005) Allopolyploidy—a shaping force in the evolution of wheat
genomes. Cytogenet Genome Res 109(1–3):250–258
Finigan P, Martienssen RA (2008) Nucleolar dominance and DNA methylation directed by small
interfering RNA. Mol Cell 32(6):753–754
Flagel L, Udall J, Nettleton D, Wendel J (2008) Duplicate gene expression in allopolyploid
Gossypium reveals two temporally distinct phases of expression evolution. BMC Biol 6:16
Flagel LE, Wendel JF (2010) Evolutionary rate variation, genomic dominance and duplicate gene
expression evolution during allotetraploid cotton speciation. New Phytol 186(1):184–193.
doi: 10.1111/j.1469-8137.2009.03107.x
Gaeta RT, Pires JC, Iniguez-Luy F, Leon E, Osborn TC (2007) Genomic changes in
Resynthesized Brassica napus and their effect on Gene expression and Phenotype. Plant
Cell Online 19(11):3403–3417. doi: 10.1105/tpc.107.054346
Gaut BS, Doebley JF (1997) DNA sequence evidence for the segmental allotetraploid origin of
maize. Proc Nat Acad Sci U S A 94(13):6809–6814
Grant V (1981) Plant speciation, 2nd edn. Columbia University Press, New York
Guo M, Birchler JA (1994) Trans-acting dosage effects on the expression of model Gene systems
in Maize Aneuploids. Science 266(5193):1999–2002. doi: 10.1126/science.266.5193.1999
CO
RR
506
507
508
509
510
511
512
513
514
515
516
517
518
519
520
521
522
523
524
525
526
527
528
529
530
531
532
533
534
535
536
537
538
539
540
541
542
543
544
545
546
547
548
549
550
551
552
553
554
555
556
557
558
559
UN
Editor Proof
4 Origins of Novel Phenotypic Variation in Polyploids
Layout: T1 Standard SC
Chapter No.: 4
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 72/75
EC
TE
D
PR
OO
F
Guo M, Davis D, Birchler JA (1996) Dosage effects on Gene expression in a Maize Ploidy series.
Genetics 142(4):1349–1355
Ha M, Lu J, Tian L, Ramachandran V, Kasschau KD, Chapman EJ, Carrington JC, Chen X, Wang
X-J, Chen ZJ (2009) Small RNAs serve as a genetic buffer against genomic shock in
Arabidopsis interspecific hybrids and allopolyploids. Proc Nat Acad Sci 106(42):17835–17840.
doi: 10.1073/pnas.0907003106
Harlan J, deWet J (1975) On Ö. Winge and a prayer: the origins of polyploidy. Bot Rev 41
(4):361–390. doi: 10.1007/bf02860830
Hegarty MJ, Barker GL, Wilson ID, Abbott RJ, Edwards KJ, Hiscock SJ (2006) Transcriptome
shock after interspecific hybridization in senecio is ameliorated by genome duplication. Curr
Biol 16(16):1652–1659. doi: 10.1016/j.cub.2006.06.071
Hegarty MJ, Hiscock SJ (2008) Genomic clues to the evolutionary success of polyploid plants.
Curr Biol: CB 18(10):R435–R444
Henry IM, Dilkes BP, Young K, Watson B, Wu H, Comai L (2005) Aneuploidy and genetic
variation in the Arabidopsis thaliana Triploid response. Genetics 170(4):1979–1988. doi:
10.1534/genetics.104.037788
Jiao Y, Wickett NJ, Ayyampalayam S, Chanderbali AS, Landherr L, Ralph PE, Tomsho LP, Hu
Y, Liang H, Soltis PS, Soltis DE, Clifton SW, Schlarbaum SE, Schuster SC, Ma H, LeebensMack J, dePamphilis CW (2011) Ancestral polyploidy in seed plants and angiosperms. Nature
473 (7345):97–100. doi: http://www.nature.com/nature/journal/v473/n7345/abs/10.1038nature09916-unlocked.html#supplementary-information
Jørgensen C (1928) The experimental formation of heteroploid plants in the genus Solanum.
J Genet 11:133
Josefsson C, Dilkes B, Comai L (2006) Parent-dependent loss of Gene silencing during
interspecies hybridization. Curr Biol 16(13):1322–1328. doi: 10.1016/j.cub.2006.05.045
Kashkush K, Feldman M, Levy A (2002) Gene loss, silencing and activation in a newly
synthesized wheat Allotetraploid. Genetics 160:1651–1659
Kellis M, Birren BW, Lander ES (2004) Proof and evolutionary analysis of ancient genome
duplication in the yeast Saccharomyces Cerevisiae. Nature 428 (6983):617–624. doi: http://
www.nature.com/nature/journal/v428/n6983/suppinfo/nature02424_S1.html
Kenan-Eichler M, Leshkowitz D, Tal L, Noor E, Melamed-Bessudo C, Feldman M, Levy AA
(2011) Wheat hybridization and Polyploidization results in deregulation of small RNAs.
Genetics 188(2):263–272. doi: 10.1534/genetics.111.128348
Kerber E (1964) Wheat: reconstitution of the tetraploid component (AABB) of hexaploids.
Science 143:53–255
Kihara H, Ono T (1926) Chromosomenzahlen und systematische Gruppierung der Rumex-Arten.
Z Zellforsch Mikr Anat 4:475
Kliebenstein DJ, West MAL, van Leeuwen H, Kim K, Doerge RW, Michelmore RW, St. Clair
DA (2006) Genomic survey of Gene expression diversity in Arabidopsis thaliana. Genetics
172(2):1179–1189. doi: 101534/genetics.105.049353
Lagercrantz U, Lydiate DJ (1996) Comparative genome mapping in Brassica. Genetics
144(4):1903–1910
Leitch AR, Leitch IJ (2008) Genomic plasticity and the diversity of polyploid plants. Science
320(5875):481–483. doi: 10.1126/science.1153585
Levin DA (1983) Polyploidy and novelty in flowering plants. Am Nat 122(1):1–25
Levin DA (2002) The role of chromosomal change in plant evolution. Oxford University Press,
Oxford
Lexer C, Welch ME, Raymond O, Rieseberg LH (2003) The origin of ecological divergence in
Helianthus paradoxus (Asteraceae): selection on transgressive characters in a novel hybrid
habitat. Evolution 57(9):1989–2000. doi: 10.1111/j.0014-3820.2003.tb00379.x
Li X, Guo W, Wang B, Li X, Chen H, Wei L, Wang Y, Wu J, Long H (2010) Instability of
chromosome number and DNA methylation variation induced by hybridization and
amphidiploid formation between Raphanus sativus L. and Brassica Alboglabra Bailey.
BMC Plant Biol 10(1):207
CO
RR
560
561
562
563
564
565
566
567
568
569
570
571
572
573
574
575
576
577
578
579
580
581
582
583
584
585
586
587
588
589
590
591
592
593
594
595
596
597
598
599
600
601
602
603
604
605
606
607
608
609
610
611
612
613
P. Finigan et al.
UN
Editor Proof
72
Layout: T1 Standard SC
Chapter No.: 4
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 73/75
73
EC
TE
D
PR
OO
F
Liu B, Wendel JF (2003) Epigenetic phenomena and the evolution of plant allopolyploids. Mol
Phylogenet Evol 29(3):365–379. doi: 10.1016/s1055-7903(03)00213-6
Lukens LN, Pires JC, Leon E, Vogelzang R, Oslach L, Osborn T (2006) Patterns of sequence loss
and cytosine methylation within a population of newly Resynthesized Brassica napus
allopolyploids. Plant Physiol 140(1):336–348. doi: 10.1104/pp.105.066308
Lumaret R (1988) Adaptive strategies and ploidy levels. Acta Oecol Oecol Plant 9:83
Madlung A, Masuelli RW, Watson B, Reynolds SH, Davison J, Comai L (2002) Remodeling of
DNA Methylation and Phenotypic and transcriptional changes in synthetic Arabidopsis
Allotetraploids. Plant Physiol 129(2):733–746. doi: 10.1104/pp.003095
Maere S, De Bodt S, Raes J, Casneuf T, Van Montagu M, Kuiper M, Van de Peer Y (2005)
Modeling gene and genome duplications in Eukaryotes. Proc Nat Acad Sci U S A
102(15):5454–5459. doi: 10.1073/pnas.0501102102
Martienssen RA (2010) Heterochromatin, small RNA and post-fertilization dysgenesis in
allopolyploid and interploid hybrids of Arabidopsis. New Phytol 186(1):46–53. doi: 10.1111/
j.1469-8137.2010.03193.x
Matzke MA, Scheid OM, Matzke AJM (1999) Rapid structural and epigenetic changes in
polyploid and aneuploid genomes. BioEssays 21(9):761–767. doi: 10.1002/(sici)15211878(199909)21:9\761:aid-bies7[3.0.co;2-c
McStay B (2006) Nucleolar dominance: a model for rRNA gene silencing. Genes Dev
20(10):1207–1214. doi: 10.1101/gad.1436906
Meins F, Thomas M (2003) Meiotic transmission of epigenetic changes in the cell-division factor
requirement of plant cells. Development 130(25):6201–6208. doi: 10.1242/dev.00856
Meyer A, Schartl M (1999) Gene and genome duplications in vertebrates: the one-to-four (-toeight in fish) rule and the evolution of novel gene functions. Curr Opin Cell Biol
11(6):699–704. doi: 10.1016/s0955-0674(99)00039-3
Meyer A, Van de Peer Y (2005) From 2R to 3R: evidence for a fish-specific genome duplication
(FSGD). BioEssays 27(9):937–945. doi: 10.1002/bies.20293
Mohan Jain S (2001) Tissue culture-derived variation in crop improvement. Euphytica
118(2):153–166. doi: 10.1023/a:1004124519479
Müntzing A (1936) The evolutionary significance of Autopolyploidy. Hereditas
21(2–3):363–378. doi: 10.1111/j.1601-5223.1936.tb03204.x
Müntzing A (1937) The effects of chromosomal variation in Dactylis. Hereditas 23:113
Newton W, Pellew C (1929) Primula kewensis and its derivatives. J Genet 20:405
Ng DWK, Zhang C, Miller M, Shen Z, Briggs SP, Chen ZJ (2011) Proteomic divergence in
Arabidopsis autopolyploids and allopolyploids and their progenitors. Heredity. doi: http://
www.nature.com/hdy/journal/vaop/ncurrent/suppinfo/hdy201192s1.html
Ni Z, Kim E-D, Ha M, Lackey E, Liu J, Zhang Y, Sun Q, Chen ZJ (2009) Altered circadian
rhythms regulate growth vigour in hybrids and Allopolyploids. Nature 457 (7227):327–331.
doi: http://www.nature.com/nature/journal/v457/n7227/suppinfo/nature07523_S1.html
Noggle G (1946) The physiology of polyploidy in plants. Lloydia 9:153
Ohno S (1970) Evolutoin by gene duplication. Springer, Berlin
Olmedo-Monfil V, Duran-Figueroa N, Arteaga-Vazquez M, Demesa-Arevalo E, Autran D,
Grimanelli D, Slotkin RK, Martienssen RA, Vielle-Calzada J-P (2010) Control of female
gamete formation by a small RNA pathway in Arabidopsis. Nature 464 (7288):628–632. doi:
http://www.nature.com/nature/journal/v464/n7288/suppinfo/nature08828_S1.html
Osborn T, Pires J, Birchler J, Auger D, Chen Z, Lee H, Comai L, Madlung A, Doerge R, Colot V,
Martienssen R (2003) Understanding mechanisms of novel gene expression in polyploids.
Trends Genet 19:141–147
Parisod C, Salmon A, Zerjal T, Tenaillon M, Grandbastien M-A, Ainouche M (2009) Rapid
structural and epigenetic reorganization near transposable elements in hybrid and allopolyploid
genomes in Spartina. New Phytol 184(4):1003–1015. doi: 10.1111/j.1469-8137.2009.03029.x
Phillips RL, Kaeppler SM, Olhoft P (1994) Genetic instability of plant tissue cultures: breakdown
of normal controls. Proc Nat Acad Sci U S A 91(12):5222–5226
CO
RR
614
615
616
617
618
619
620
621
622
623
624
625
626
627
628
629
630
631
632
633
634
635
636
637
638
639
640
641
642
643
644
645
646
647
648
649
650
651
652
653
654
655
656
657
658
659
660
661
662
663
664
665
666
UN
Editor Proof
4 Origins of Novel Phenotypic Variation in Polyploids
Layout: T1 Standard SC
Chapter No.: 4
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 74/75
EC
TE
D
PR
OO
F
Pignatta D, Dilkes BP, Yoo S-Y, Henry IM, Madlung A, Doerge RW, Jeffrey Chen Z, Comai L
(2010) Differential sensitivity of the Arabidopsis thaliana transcriptome and enhancers to the
effects of genome doubling. New Phytol 186(1):194–206. doi: 10.1111/j.1469-8137.2010.
03198.x
Pires JC, Zhao J, Schranz ME, Leon EJ, Quijada PA, Lukens LN, Osborn TC (2004) Flowering
time divergence and genomic rearrangements in resynthesized Brassica polyploids (Brassicaceae). Biol J Linn Soc 82(4):675–688. doi: 10.1111/j.1095-8312.2004.00350.x
Pontes O, Neves N, Silva M, Lewis MS, Madlung A, Comai L, Viegas W, Pikaard CS (2004)
Chromosomal locus rearrangements are a rapid response to formation of the allotetraploid
Arabidopsis suecica genome. Proc Nat Acad Sci U S A 101(52):18240–18245. doi: 10.1073/
pnas.0407258102
Preuss S, Pikaard CS (2007) rRNA gene silencing and nucleolar dominance: insights into a
chromosome-scale epigenetic on/off switch. Biochimica et Biophysica Acta (BBA)—Gene
structure and expression 1769 (5–6):383–392. doi: 10.1016/j.bbaexp.2007.02.005
Preuss SB, Costa-Nunes P, Tucker S, Pontes O, Lawrence RJ, Mosher R, Kasschau KD,
Carrington JC, Baulcombe DC, Viegas W, Pikaard CS (2008) Multimegabase silencing in
nucleolar dominance involves siRNA-directed DNA methylation and specific methylcytosinebinding proteins. Mol Cell 32(5):673–684. doi: 10.1016/j.molcel.2008.11.009
Pumphrey M, Bai J, Laudencia-Chingcuanco D, Anderson O, Gill B (2009) Nonadditive
expression of homoeologous genes is established upon polyploidization in hexaploid wheat.
Genetics 181:1147–1157
Ramsey J, Schemske DW (1998) Pathways, mechanisms, and rates of polyploid formation in
flowering plants. Annu Rev Ecol Syst 29(1):467–501. doi: 10.1146/annurev.ecolsys.29.1.467
Ramsey J, Schemske DW (2002) Neopolyploidy in flowering plants. Annu Rev Ecol Syst
33(1):589–639. doi: 10.1146/annurev.ecolsys.33.010802.150437
Randolph L (1941) An evaluation of induced polyploidy as a method of breeding crop plants. Am
Nat 75:347
Rapp R, Udall J, Wendel J (2009) Genomic expression dominance in allopolyploids. BMC Biol
7:18
Riddle N, Jiang H, An L, Doerge R, Birchler J (2010) Gene expression analysis at the intersection
of ploidy and hybridity in maize. Theor Appl Genet 120(2):341–353. doi: 10.1007/s00122009-1113-3
Santos JL, Alfaro D, Sanchez-Moran E, Armstrong SJ, Franklin FCH, Jones GH (2003) Partial
diploidization of meiosis in Autotetraploid Arabidopsis thaliana. Genetics 165(3):1533–1540
Schnable JC, Springer NM, Freeling M (2011) Differentiation of the maize subgenomes by
genome dominance and both ancient and ongoing gene loss. Proc Nat Acad Sci
108(10):4069–4074. doi: 10.1073/pnas.1101368108
Shindo C, Aranzana MJ, Lister C, Baxter C, Nicholls C, Nordborg M, Dean C (2005) Role of
FRIGIDA and flowering locus c in determining variation in flowering time of Arabidopsis.
Plant Physiol 138(2):1163–1173. doi: 10.1104/pp.105.061309
Shoemaker RC, Polzin K, Labate J, Specht J, Brummer EC, Olson T, Young N, Concibido V,
Wilcox J, Tamulonis JP, Kochert G, Boerma HR (1996) Genome duplication in soybean
(Glycine subgenus soja). Genetics 144(1):329–338
Simpson GG, Dean C (2002) Arabidopsis, the rosetta stone of flowering time? Science
296(5566):285–289. doi: 10.1126/science.296.5566.285
Singh M, Goel S, Meeley RB, Dantec C, Parrinello H, Michaud C, Leblanc O, Grimanelli D
(2011) Production of viable gametes without Meiosis in Maize deficient for an ARGONAUTE
protein. Plant Cell Online 23(2):443–458. doi: 10.1105/tpc.110.079020
Skalinska M (1946) Polyploidy in valeriana officinalis Linn. In relation to its ecology and
distribution. J Linn Soc London, Bot 53:159
Slotkin RK, Vaughn M, Borges F, Tanurdzic M, Becker JD, Feijo JA, Martienssen RA (2009)
Epigenetic reprogramming and small RNA silencing of transposable elements in pollen. Cell
136 (3):461–472. doi: S0092-8674(08)01644-9 [pii] 10.1016/j.cell.2008.12.038
CO
RR
667
668
669
670
671
672
673
674
675
676
677
678
679
680
681
682
683
684
685
686
687
688
689
690
691
692
693
694
695
696
697
698
699
700
701
702
703
704
705
706
707
708
709
710
711
712
713
714
715
716
717
718
719
P. Finigan et al.
UN
Editor Proof
74
Layout: T1 Standard SC
Chapter No.: 4
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 75/75
75
EC
TE
D
PR
OO
F
Soltis DE, Albert VA, Leebens-Mack J, Bell CD, Paterson AH, Zheng C, Sankoff D, dePamphilis
CW, Wall PK, Soltis PS (2009) Polyploidy and angiosperm diversification. Am J Bot
96(1):336–348. doi: 10.3732/ajb.0800079
Soltis DE, Rieseberg LH (1986) Autopolyploidy in Tolmiea menziesii (Saxifragaceae): genetic
insights from enzyme electrophoresis. Am J Bot 73(2):310–318
Soltis DE, Soltis PS (1999) Polyploidy: recurrent formation and genome evolution. Trends Ecol
Evol 14(9):348–352. doi: 10.1016/s0169-5347(99)01638-9
Soltis DE, Soltis PS, Schemske DW, Hancock JF, Thompson JN, Husband BC, Judd WS (2007)
Autopolyploidy in angiosperms: have we grossly underestimated the number of species?
Taxon 56(1):13–30
Soltis PS, Soltis DE (2009) The role of hybridization in plant speciation. Annu Rev Plant Biol
60(1):561–588. doi: 10.1146/annurev.arplant.043008.092039
Song K, Lu P, Tang K, Osborn TC (1995) Rapid genome change in synthetic polyploids of
Brassica and its implications for polyploid evolution. Proc Nat Acad Sci 92(17):7719–7723
Song K, Tang K, Osborn T (1993) Development of synthetic Brassica amphidiploids by
reciprocal hybridization and comparison to natural amphidiploids. Theor Appl Genet 86:811
Stebbins G (1947) Types of polyploids: their classification and significance. Adv Genet 1:403
Stebbins GL (1950) Variation and evolution in plants. Columbia University Press, New York
Stebbins GL (1971) Chromosomal evolution in higher plants [by] G. Ledyard Stebbins.
Contemporary biology, vol Accessed from http://nla.gov.au/nla.cat-vn1859678. Edward
Arnold, London
Storchova Z, Breneman A, Cande J, Dunn J, Burbank K, O’Toole E, Pellman D (2006) Genomewide genetic analysis of polyploidy in yeast. Nature 443 (7111):541–547. doi: http://
www.nature.com/nature/journal/v443/n7111/suppinfo/nature05178_S1.html
Stupar RM, Bhaskar PB, Yandell BS, Rensink WA, Hart AL, Ouyang S, Veilleux RE, Busse JS,
Erhardt RJ, Buell CR, Jiang J (2007) Phenotypic and Transcriptomic changes associated with
Potato Autopolyploidization. Genetics 176(4):2055–2067. doi: 10.1534/genetics.107.074286
Tanurdzic M, Vaughn MW, Jiang H, Lee T-J, Slotkin RK, Sosinski B, Thompson WF, Doerge
RW, Martienssen RA (2008) Epigenomic consequences of immortalized plant cell suspension
culture. PLoS Biol 6(12):e302. doi: 10.1371/journal.pbio.0060302
Tate JA, Symonds VV, Doust AN, Buggs RJA, Mavrodiev E, Koh J, Soltis PS, Soltis DE (2009)
Synthetic polyploids of Tragopogon miscellus and T. mirus (Asteraceae): 50 ? years after
Ownbey’s discovery. Am J Bot 96:979–988
Tate JA, Ni Z, Scheen A-C, Koh J, Gilbert CA, Lefkowitz D, Chen ZJ, Soltis PS, Soltis DE
(2006) Evolution and expression of Homeologous Loci in Tragopogon miscellus (Asteraceae), a recent and reciprocally formed allopolyploid. Genetics 173(3):1599–1611. doi:
10.1534/genetics.106.057646
Valvekens D, Montagu MV, Lijsebettens MV (1988) Agrobacterium tumefaciens-mediated
transformation of Arabidopsis thaliana root explants by using kanamycin selection. Proc Nat
Acad Sci U S A 85(15):5536–5540
Van de Peer Y, Maere S, Meyer A (2009) The evolutionary significance of ancient genome
duplications. Nat Rev Genet 10(10):725–732
Vaughn MW, Tanurdzic M, Lippman Z, Jiang H, Carrasquillo R, Rabinowicz PD, Dedhia N,
McCombie WR, Agier N, Bulski A, Colot V, Doerge RW, Martienssen RA (2007) Epigenetic
natural variation in Arabidopsis thaliana. PLoS Biol 5 (7):e174. doi: 06-PLBI-RA-2115 [pii]
10.1371/journal.pbio.0050174
Walia H, Josefsson C, Dilkes B, Kirkbride R, Harada J, Comai L (2009) Dosage-dependent
deregulation of an AGAMOUS-LIKE gene cluster contributes to interspecific incompatibility.
Curr Biol: CB 19(13):1128–1132
Wang H, Chai Y, Chu X, Zhao Y, Wu Y, Zhao J, Ngezahayo F, Xu C, Liu B (2009) Molecular
characterization of a rice mutator-phenotype derived from an incompatible cross-pollination
reveals transgenerational mobilization of multiple transposable elements and extensive
epigenetic instability. BMC Plant Biol 9(1):63
CO
RR
720
721
722
723
724
725
726
727
728
729
730
731
732
733
734
735
736
737
738
739
740
741
742
743
744
745
746
747
748
749
750
751
752
753
754
755
756
757
758
759
760
761
762
763
764
765
766
767
768
769
770
771
772
UN
Editor Proof
4 Origins of Novel Phenotypic Variation in Polyploids
Layout: T1 Standard SC
Chapter No.: 4
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 76/75
EC
TE
D
PR
OO
F
Wang J, Tian L, Lee H-S, Chen ZJ (2006a) Nonadditive regulation of FRI and FLC Loci
mediates flowering-time variation in Arabidopsis Allopolyploids. Genetics 173(2):965–974.
doi: 10.1534/genetics.106.056580
Wang J, Tian L, Lee H-S, Wei NE, Jiang H, Watson B, Madlung A, Osborn TC, Doerge RW,
Comai L, Chen ZJ (2006b) Genomewide nonadditive Gene regulation in Arabidopsis
Allotetraploids. Genetics 172(1):507–517. doi: 10.1534/genetics.105.047894
Wang J, Tian L, Madlung A, Lee H-S, Chen M, Lee JJ, Watson B, Kagochi T, Comai L, Chen ZJ
(2004) Stochastic and Epigenetic changes of Gene expression in Arabidopsis Polyploids.
Genetics 167(4):1961–1973. doi: 10.1534/genetics.104.027896
Wendel JF (2000) Genome evolution in polyploids. Plant Mol Biol 42(1):225–249. doi: 10.1023/
a:1006392424384
Winge O (1917) The chromosomes: their number and general importance. C.R. Trav Lab,
Carlsberg
Winge Ö (1932) On the origin of constant species-hybrids. Sven Bot Tidskr 26:107
Wolffe AP, Matzke MA (1999) Epigenetics: regulation through repression. Science
286(5439):481–486. doi: 10.1126/science.286.5439.481
Xiong Z, Gaeta RT, Pires JC (2011) Homoeologous shuffling and chromosome compensation
maintain genome balance in resynthesized allopolyploid Brassica napus. Proc Nat Acad Sci
108(19):7908–7913. doi: 10.1073/pnas.1014138108
Xu Y, Zhong L, Wu X, Fang X, Wang J (2009) Rapid alterations of gene expression and cytosine
methylation in newly synthesized Brassica napus allopolyploids. Planta 229(3):471–483. doi:
10.1007/s00425-008-0844-8
Yaakov B, Kashkush K (2011) Massive alterations of the methylation patterns around DNA
transposons in the first four generations of a newly formed wheat allohexaploid. Genome
54(1):42–49. doi: 10.1139/g10-091
Yao H, Kato A, Mooney B, Birchler J (2011) Phenotypic and gene expression analyses of a
ploidy series of maize inbred Oh43. Plant Mol Biol 75(3):237–251. doi: 10.1007/s11103-0109722-4
Yu Z, Haberer G, Matthes M, Rattei T, Mayer KFX, Gierl A, Torres-Ruiz RA (2010) Impact of
natural genetic variation on the transcriptome of autotetraploid Arabidopsis thaliana. Proc Nat
Acad Sci 107(41):17809–17814. doi: 10.1073/pnas.1000852107
CO
RR
773
774
775
776
777
778
779
780
781
782
783
784
785
786
787
788
789
790
791
792
793
794
795
796
797
798
799
800
801
802
803
P. Finigan et al.
UN
Editor Proof
76
Metadata of the chapter that will be visualized in
SpringerLink
Book Title
Polyploidy and Genome Evolution
Series Title
Chapter Title
Identifying the Phylogenetic Context of Whole-Genome Duplications in Plants
Copyright Year
2012
Copyright HolderName
Springer-Verlag Berlin Heidelberg
Corresponding Author
Family Name
Burleigh
Particle
Given Name
J. Gordon
Suffix
Abstract
Division
Department of Biology
Organization
University of Florida
Address
32611, Gainesville, FL, USA
Email
gburleigh@ufl.edu
Although evolutionary biologists have long recognized the transformative evolutionary potential of wholegenome duplications (WGDs) in plants, identifying the precise phylogenetic location of WGDs presents many
challenges. This chapter reviews some new approaches to map WGDs on a phylogeny, the first step for
understanding the large-scale evolutionary and ecological consequences of WGDs in plants. Specifically, it
examines approaches for using chromosome and gene copy number data, gene trees, and other genomic
insights to identify the evolutionary location of WGDs. The abundance of genomic sequence data and
advances in phylogenetic methods present unprecedented opportunities to place WGDs within the plant tree
of life. Still, there exist few direct tests to identify and place WGDs, and analyses of complex data are often
susceptible to error.
1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 77/91
Chapter 5
4
J. Gordon Burleigh
9
10
11
12
13
14
15
16
18
19
20
21
22
23
24
25
26
27
28
29
A central challenge in evolutionary biology is to determine the genetic mechanisms
that generate species diversity as well as new traits, functions, and adaptations. Plant
evolutionary biologists have long recognized the transformative evolutionary potential of polyploidy or whole-genome duplication (WGD) (e.g., Stebbins 1950; Grant
1963; Levin 2002; Soltis and Soltis 2000; Soltis et al. 2009). However, to study the
evolutionary consequences of whole-genome duplications (WGDs) and link WGDs to
phenotypic changes or diversification, the WGDs must first be placed in a phylogenetic context. Unfortunately, the rapid gene loss and genome rearrangements that
frequently follow WGDs erase evidence of WGD (e.g., Wolfe 2001; Doyle et al.
2008). This process of diploidization brings about a paradox of the study of polyploidy; despite its apparent pervasiveness throughout the evolutionary history of land
plants and its evolutionary importance, clear, unambiguous evidence of ancient
WGDs can be remarkably difficult to locate within a phylogeny.
CO
RR
17
PR
OO
D
8
TE
7
Abstract Although evolutionary biologists have long recognized the transformative evolutionary potential of whole-genome duplications (WGDs) in plants,
identifying the precise phylogenetic location of WGDs presents many challenges.
This chapter reviews some new approaches to map WGDs on a phylogeny, the first
step for understanding the large-scale evolutionary and ecological consequences of
WGDs in plants. Specifically, it examines approaches for using chromosome and
gene copy number data, gene trees, and other genomic insights to identify the
evolutionary location of WGDs. The abundance of genomic sequence data and
advances in phylogenetic methods present unprecedented opportunities to place
WGDs within the plant tree of life. Still, there exist few direct tests to identify and
place WGDs, and analyses of complex data are often susceptible to error.
EC
5
6
F
3
Identifying the Phylogenetic Context
of Whole-Genome Duplications in Plants
2
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: 5
J. G. Burleigh (&)
Department of Biology, University of Florida, Gainesville, FL 32611, USA
e-mail: gburleigh@ufl.edu
P. S. Soltis and D. E. Soltis (eds.), Polyploidy and Genome Evolution,
DOI: 10.1007/978-3-642-31442-1_5, Springer-Verlag Berlin Heidelberg 2012
77
Layout: T1 Standard SC
Chapter No.: 5
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 78/91
J. Gordon Burleigh
48
5.1 Chromosome Evolution
32
33
34
35
36
37
38
39
40
41
42
43
44
45
51
52
53
54
55
56
57
58
59
60
61
62
63
64
65
66
67
68
69
70
Many early surveys of polyploidy in plants used chromosome counts to estimate
the percentage of polyploid species (e.g., Grant 1963, 1982; Stebbins 1938, 1950,
1971; Goldblatt 1980). This work laid the foundation for understanding the role of
WGDs in plant evolution. However, lacking a phylogenetic context, it is difficult
to estimate the frequency of WGD events, let alone the evolutionary placement of
WGDs simply from the percentage of polyploids (although see Levin and Wilson
1976; Otto and Whitton 2002). For example, if a plant family has 100 species, 50
of which are recent polyploids, this could be the result of as few as one WGD or as
many as 50 WGDs (assuming at most one WGD per lineage). Understanding the
relationships among species is necessary to infer the history of WGD. Furthermore, chromosome number alone is not necessarily indicative of polyploidy. Zea
mays, with a haploid chromosome number of 10, has had multiple WGDs in the
last *20 million years (Gaut and Doebley 1997; Gaut 2001), and Arabidopsis
thaliana, with a haploid chromosome number of five, has experienced 3–5 WGDs
since the origin of seed plants (Vision et al. 2000; Blanc et al. 2003; Bowers et al.
2003; Jiao et al. 2011). Yet few chromosome number surveys would not have
considered either polyploid.
Mapping chromosome numbers on a phylogenetic hypothesis can help reveal
the frequency and evolutionary placement of WGD events. Informal phylogenetic
observations were first used to deduce ancestral chromosome numbers. For
example, numerous studies surmised a low base chromosome number for angiosperms based on the chromosome counts of the ‘‘basal’’ angiosperm lineages (e.g.,
EC
50
CO
RR
49
TE
D
46
PR
OO
31
F
47
WGD in plants has been and studied for more than 100 years (e.g., Digby 1912;
Winge 1917). Although sometimes cryptic, evidence of WGDs may be gleaned
from such disparate sources as chromosome counts (e.g., Stebbins 1971), guard
cell size (Masterson 1994), age distributions of gene copies (e.g., Blanc and Wolfe
2004), or large segmental duplications within genomes (e.g., Vision et al. 2000).
Still, it is difficult to infer the phylogenetic location of the WGDs with any precision from these observations alone. To place WGDs in an evolutionary context
requires a robust, and ideally well–sampled, phylogenetic hypothesis, large-scale
comparative data indicating WGDs, and models and methods to map these data
onto a phylogeny. Only recently have advances in phylogenetics, comparative
methods, and genome sequencing made this possible on a large scale. These
advances have enabled the first studies addressing some of the basic evolutionary
questions about polyploidy, such as the frequency of polyploid speciation (Wood
et al. 2009) or the effect of WGDs on diversification (Vamosi and Dickinson 2006;
Soltis et al. 2009; Mayrose et al. 2011), in a rigorous phylogenetic framework.
This chapter reviews some new approaches to map WGDs on a phylogeny, the first
step for understanding the large-scale evolutionary and ecological consequences of
WGDs in plants.
30
UN
Editor Proof
78
Layout: T1 Standard SC
Chapter No.: 5
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 79/91
77
78
79
80
81
82
83
84
85
86
87
88
89
90
91
92
93
94
95
96
97
98
99
100
101
102
103
104
105
106
107
108
109
110
111
112
113
114
115
F
76
PR
OO
75
D
74
TE
73
79
Ehrendorfer et al. 1968; Stebbins 1971; Walker 1972; Raven 1975). These
observations implied a history of WGD near the root of angiosperms, although
they could not map this precisely; indeed, the relationships among these lineages
were unknown. With the growth of phylogenetic methods and data, more formal
maximum parsimony approaches were used to reconstruct ancestral chromosome
number on the inner nodes of phylogenetic trees (e.g., Stace et al. 1997; Schultheis
2001; Mishima et al. 2002; Guggisberg et al. 2006; Hipp et al. 2007). To do this,
chromosome number can be treated as a discrete variable, and the ancestral states
can be reconstructed using linear or squared change parsimony. If the ancestral
states are far higher than the base chromosome number, then that ancestral node
may represent a polyploid. It is possible to construct elaborate chromosome
number transition matrices for parsimony analyses, for example, allowing chromosome doubling as well as single chromosome changes, or to weight different
changes, like down-weighting chromosome losses, but these analyses are rarely
performed. In any case, parsimony reconstructions often have difficulty accounting
for multiple transitions on a single branch or quantifying uncertainty in ancestral
state reconstructions.
More recently, probabilistic models of chromosome number evolution have
been developed (Meyers and Levin 2006; Mayrose et al. 2010). In a simple formulation, the chromosome models allow transitions that add a chromosome,
remove a chromosome, or double the number of chromosomes (Mayrose et al.
2010). Thus, although the transition matrix among chromosome states (chromosome numbers) may be extremely large, the evolutionary process can be modeled
with only a few parameters. The performance of these models has not been
characterized in detail; however, they appear to infer more ancient WGDs than
parsimony methods (Wood et al. 2009) and may also provide quite different
ancestral state reconstructions of chromosome number (Cusimano et al. 2012). In
the future, these models may link chromosome evolution to diversification rates or
phenotypes related to the frequency of WGDs to obtain even more accurate
estimates of WGDs in a phylogeny.
Studies of plant chromosome numbers have provided a wealth of insight into
polyploidy in plants and have contributed substantially to canonical views of plant
speciation and evolution. Yet chromosome number is a sort of summary statistic
for WGD, a simple observation that is meant to represent a complex, large-scale
genomic change, and chromosome number alone may not be sufficient to detect
WGDs. A small chromosome number, as in Arabidopsis thaliana, does not necessarily imply the absence of historical WGDs, and high chromosome numbers are
not necessarily evidence of WGDs. Without additional cytological or genetic data,
it is impossible to distinguish between a WGD and increasing dysploidy, a change
in the chromosome number that is not associated with a change in the amount of
genetic material, based solely on chromosome counts.
Also, as with any phenotype, there are limitations and biases associated with
ancestral state reconstruction (e.g., Schluter et al. 1997; Ané 2008). Often
reconstruction is most difficult for characters with high rates of evolution or high
degrees of homoplasy. Chromosome numbers may be unstable following a WGD
EC
72
CO
RR
71
UN
Editor Proof
5 Identifying the Phylogenetic Context of Whole-Genome
Layout: T1 Standard SC
Chapter No.: 5
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 80/91
J. Gordon Burleigh
133
5.2 Gene Copy Models
122
123
124
125
126
127
128
129
130
131
134
135
136
137
138
139
140
141
142
143
144
145
146
147
148
149
150
151
152
153
154
155
PR
OO
121
D
120
TE
119
Form availability of large-scale genomic data from an increasing number of plant
species, estimates of copy numbers for gene families are increasingly available for
many plant species. Since WGD should change not only the chromosome numbers
but also the copy numbers for all gene families, gene family copy number should
provide more data to estimate a WGD than simply a single chromosome number.
Like chromosome number evolution, ancestral gene copy numbers can be reconstructed using parsimony methods (Snel et al. 2002; Kunin and Ouzounis 2003;
Mirkin et al. 2003; Csürös 2010; Ames et al. 2012; Librado et al. 2012). The
parsimony models can be implemented in numerous ways, including weighting
gains and losses differently. It is not necessarily easy to find evidence of WGDs
based on the number of gene gains or losses on a branch in the species tree, but we
might expect WGDs will result in far more gains, and subsequently losses, per unit
time than are found on other branches. Hahn et al. (2005) developed a stochastic
birth and death model that assumes a homogeneous process of duplication and loss
throughout the species tree. The ML implementation of this model in CAFÉ
(De Bie et al. 2006), as well as a similar Bayesian approach (Liu et al. 2011),
estimates gene family gain and loss rates across the tree and can identify
anomalous gene families and branches on the tree. These branches may reflect the
effects of WGDs. More complex models that account for heterogeneity in the rates
of duplications and losses across lineages, and in some cases also allowing gains of
genes or gene families by lateral transfer, also have been proposed (Iwasaki and
Takagi 2007; Csurös 2010; Ames et al. 2012; Librado et al. 2012).
EC
118
CO
RR
117
F
132
(see Lim et al. 2008; Chester et al. 2012) and can decrease quickly following a
WGD. Thus, modeling approaches likely will have difficulty for identifying
ancient WGDs. In fact, it appears that the frequency of chromosome loss and
diploidization was not always appreciated in studies that only examined chromosome numbers, and even with large-scale genomic data, the mechanisms for
rapid chromosome loss are not clear (Doyle et al. 2008). This lack of appreciation
for the lability of chromosome numbers may have contributed to the failure to
detect, or even surmise, the extent of ancient WGDs, and also may have
encouraged the idea that WGDs were evolutionary dead ends (e.g., Stebbins 1950).
Despite the limitations of chromosome numbers alone, data are available for
many thousands of plant species, for example on the online Index to Plant
Chromosome Numbers (IPCN) database (http://www.tropicos.org/Project/IPCN).
Thus, until large-scale genomic sequence data sets become available for thousands
of phylogenetically diverse taxa, chromosome number may provide the best
opportunity to identify putative WGD events, especially recent events, with
phylogenetic precision and to examine the macroevolutionary consequences of
WGD throughout the history of all plants.
116
UN
Editor Proof
80
Layout: T1 Standard SC
Chapter No.: 5
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 81/91
162
163
164
165
166
167
168
169
170
171
172
173
174
175
176
177
178
179
180
181
182
183
184
185
186
187
188
189
190
191
192
193
194
195
196
F
161
PR
OO
160
D
159
TE
158
In spite of much recent work on developing models of gene family copy number,
all of the gene copy models assume that gene duplications or gene gains are
independent. Thus, a WGD in a plant might be viewed as 20,000 gene duplications
rather than a single duplication event. Consequently, while an increased duplication
rate on a branch or increased loss rates on subsequent branches may suggest a
WGD, there is no definitive test of WGD, and it may be difficult to distinguish
WGD from simply an elevated duplication rate or a large-scale duplication. One
approach may be to create a model that could estimate a rate of doubling for all gene
family numbers. This transition matrix could be applied to different branches to test
for either a rate of WGD [0 or different rates between clades or branches.
Although gene copy number provides more detailed assessment of genomic
content than chromosome numbers, inferring the histories of gene copy number
and chromosome number have similar limitations. For example, gene copies
appear to be rapidly silenced and lost immediately following a WGD (e.g., Tate
et al. 2006; Buggs et al. 2009, 2012; see Chap. 14, this volume), which may
quickly obscure the evidence for WGDs. However, simply obtaining accurate
estimates of gene copy number for extant taxa may be a challenge. Without
complete genome sequencing, it can be difficult to distinguish a gene loss from
simply a failure to sample a gene. In fact, the lack of complete sequencing across a
broad range of plant species may explain the lack of studies of gene copy number
evolution in plants. Even with complete genome sequences, estimates of gene copy
number depend on the vagaries of the extremely complex genome annotation and
gene family circumscription problems. Furthermore, there are high levels of
intraspecific variation in gene copy number in some plants (e.g., Springer et al.
2009; Debolt 2010; Zheng et al. 2011). It is possible to account for uncertainty in
the gene copy numbers or incomplete sampling in a likelihood model, although
such approaches have not been implemented.
Still, gene copy number does not always provide direct evidence for the location
of gene or WGDs. For example, take the case in Fig. 5.1, in which an outgroup has a
single gene copy, and two sister taxa each have two gene copies. The parsimonious
explanation for these data would be that a gene duplication preceded the most
recent common ancestor of the sister taxa, although it is possible that there were
independent duplications in each sister lineage. In this case, the gene topologies can
provide much more insight into the history of duplication than simply looking at
copy number and can easily distinguish between the two duplication scenarios in
Fig. 5.1. The additional information from evolutionary history of the genes can
further help identify the placement of historical duplications and WGDs.
EC
157
81
CO
RR
156
UN
Editor Proof
5 Identifying the Phylogenetic Context of Whole-Genome
5.3 Gene Tree Reconciliation
The general problem of gene tree reconciliation is based on the observation that
population-level processes, such as coalescence (lineage sorting), as well as
evolutionary events, such as gene duplications and loss, recombination,
Layout: T1 Standard SC
Chapter No.: 5
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 82/91
J. Gordon Burleigh
D
PR
OO
F
Editor Proof
82
TE
Fig. 5.1 Gene copy number data and corresponding gene trees. The gene copy number data
mapped on a species tree implies at least one gene duplication, but do not specify the location of the
duplication(s). Gene tree 1 implies a single duplication preceding the most recent common ancestor
of A and B. Gene tree 2 implies independent duplications in the lineages leading to A and B
209
5.3.1 Parsimony Approaches
200
201
202
203
204
205
206
207
210
211
212
CO
RR
199
UN
198
EC
208
hybridization, or lateral gene transfer can result in gene topologies that differ from
the phylogeny of the species in which the genes evolve (e.g., Maddison 1997). The
general challenge of gene tree reconciliation is to find the evolutionary scenario
that best explains the gene tree topologies. For the case of WGD, we might ask if a
collection of gene trees is consistent with a WGD event, or at least a large number
of duplications, at a particular point in the species phylogeny. While this approach
seeks to directly map the location of gene duplications, and thus provide more
direct evidence of WGD than simply examining changes in chromosome or gene
copy numbers, in practice it often is complicated by the number of different
scenarios that can cause gene tree incongruence and the inherent difficulty of
accurately inferring a gene tree from only a gene sequence alignment (for more
detailed, general reviews see Eulenstein et al. 2010; Doyon et al. 2011b).
197
Much of the initial work in gene tree reconciliation was based on optimizing a parsimony criterion; that is, finding a mapping of a gene tree topology onto the species tree
that implies the fewest evolutionary events. The gene duplication model was first
Layout: T1 Standard SC
Chapter No.: 5
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 83/91
5 Identifying the Phylogenetic Context of Whole-Genome
216
217
218
219
220
221
222
223
224
225
226
227
228
229
230
231
232
233
234
F
PR
OO
EC
215
introduced by Goodman et al. (1979; also see Page 1994 and Guigó et al. 1996) to find
the minimum number of gene duplications needed to explain the incongruence
between a gene tree and species tree. To do this, a gene tree can be embedded into a
species tree through least common ancestor mapping (lca-mapping), which maps
every node in the gene tree (tips and internal nodes) to the most recent node in the
species tree that could have contained the gene node (Fig. 5.2). A duplication occurred
if a parent and child node in the gene tree share the same lca-mapping in the species
tree. The lca-mapping identifies the most recent possible location of the gene duplication in the species tree, but this often is not the only possible location of the gene
duplication. In many cases, the duplication could have preceded the lca-mapping,
although the earlier placement of the duplication may imply additional gene losses.
However, as in the gene copy number analyses, gene losses are often difficult to
distinguish from incomplete sampling.
This simple gene reconciliation can provide a direct estimate of the phylogenetic location of gene duplication events with a precision that is impossible with
chromosome or gene copy number data. Also, it does not even require the presence
of duplicated, paralogs genes; incongruence between a single-copy gene tree
topology and the species phylogeny may be evidence of a hidden history of gene
duplication and loss. On the other hand, gene duplications and losses are not the
only explanation of gene tree incongruence. For example, incomplete lineage
sorting or reticulation also can confound gene tree topologies. In this case, the
gene duplication model may mistakenly imply a large number of duplications
CO
RR
214
UN
213
TE
D
Editor Proof
Fig. 5.2 An example of
lca-mapping of gene
duplications. The parent and
child nodes in the gene tree
(h and t) map to the same
node (X) in the species tree.
This implies a single
duplication occurring prior to
or at node X in the species
tree
83
Layout: T1 Standard SC
Chapter No.: 5
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 84/91
241
242
243
244
245
246
247
248
249
250
251
252
253
254
255
256
257
258
259
260
261
262
263
264
265
266
267
268
269
270
271
272
273
274
275
F
240
PR
OO
239
D
238
TE
237
preceding rapid cladogenesis in the species tree, where we might expect high
levels of incomplete lineage sorting.
Perhaps the most difficult problem underlying the gene reconciliation approaches
is that, in many cases, the incongruence between a gene tree and the species phylogeny may simply be due to error. The gene tree model will interpret any topological
error as evidence of duplications. Consequently, this approach often implies far more
duplications rather than biologically plausible (e.g., Rasmussen and Kellis 2011).
The errors in the gene tree tend to place erroneously large numbers of duplications
near the root of the species tree, which may falsely suggest large-scale duplication
events at the origins of major clades (Hahn 2007; Burleigh et al. 2010). In the
parsimony context, several strategies have been proposed to ameliorate the problems
of gene tree error. First, poorly supported clades in the gene tree may be collapsed.
Several algorithmic approaches have extended the gene duplication model to deal
with reconciling nonbinary trees (Berglund-Sonnhammer et al. 2006; Chang and
Eulenstein 2006; Durand et al. 2006). Also, several approaches have been developed
to allow minor modifications of the gene tree topology if they reduce the number of
implied duplications (e.g., Chen et al. 2000; Chaudhury et al. 2011, 2012; Gorecki
and Eulenstein 2012). For example, Chaudhury et al. (2012) introduced an algorithm
that, given a gene tree and a species tree, finds a gene topology in a subtree pruning
and regrafting (SPR) neighborhood of the original gene tree that minimizes the
number of implied duplications. These local rearrangements can massively reduce
the number of estimated gene duplication events.
In spite of the many issues related to gene tree reconciliation, simple and
informal gene tree reconciliations have been effective at helping to identify the
phylogenetic location of WGDs in plants. These approaches are usually limited to
small gene trees with paralogs, that is, gene trees in which at least one duplication
must have occurred. In a simple three-taxon approach, a gene tree is constructed
with a pair of paralogs genes from a test taxon, and the best homologs from a sister
taxon and from an outgroup taxon (e.g., Bowers et al. 2003). If paralogs from the
test taxon form a clade, they diverged after the common ancestor with the sister
taxon; if they do not, they diverged before the most recent common ancestor. This
three-taxon phylogenetic approach provides only a limited phylogenetic context
for the duplications, but it has been used to determine the timing of WGDs in
Arabidopsis relative to its divergence from pines, rice, and other eudicots (Bowers
et al. 2003) and rice relative to its divergence from pines, Arabidopsis, and other
monocots (Vandepoele et al. 2003; Chapman et al. 2004). More recently, Jiao et al.
(2011) counted the gene trees that were consistent with different scenarios of
WGD to infer WGDs at the root of angiosperms and seed plants.
EC
236
CO
RR
235
J. Gordon Burleigh
UN
Editor Proof
84
5.3.2 Parsimony Methods to Identify WGDs
The gene duplication problem described above treats each duplication independently. Although it may identify places in the species tree with high numbers of
Layout: T1 Standard SC
Chapter No.: 5
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 85/91
85
306
5.3.3 Likelihood-Based Approaches
282
283
284
285
286
287
288
289
290
291
292
293
294
295
296
297
298
299
300
301
302
303
304
307
308
309
310
311
312
313
314
315
316
PR
OO
281
D
280
TE
279
EC
278
CO
RR
277
F
305
gene duplications, it does not attempt to find large-scale duplication events.
Several proposed approaches attempt to identify the minimum number of gene
duplication events, where an event may include duplications of many or all genes,
rather than simply the number of duplications. One indirect approach is to examine
all the possible locations of each gene duplication and find a mapping that minimizes the number of locations (nodes) on the species tree where gene duplications
occur (Guigó et al. 1996; Page and Cotton 2002; Burleigh et al. 2009; Luo et al.
2009). This approach does not directly infer WGDs; but ideally, it can identify
places in the species tree that are possible locations of clusters of many duplications. With a limited number of gene trees, this approach appears to be effective at
identifying some locations of ancient WGDs in plants (Burleigh et al. 2009).
Unfortunately, with a large number of gene trees, it is likely that all possible
duplication mappings will require duplication events at every node in the species
tree. In this case, every possible mapping of gene duplications will be equally
optimal, and this approach will be uninformative.
Another approach seeks to find a gene duplication mapping that implies the
fewest gene duplication episodes (Guigó et al. 1996; Page and Cotton 2002;
Bansal and Eulenstein 2008; Luo et al. 2009). Given a single gene tree and species
tree, any set of gene duplications, from the same or different gene trees, that occur
on the same node in a species tree can be explained by a single gene duplication
episode (or event) as long as none of the gene duplications in the set have an
ancestor–descendant relationship with each other (Fig. 5.3). This approach appears
to help identify WGDs, which should be very large episodes, but the largest
episode is simply the largest episode found on any single gene tree (Page and
Cotton 2002; Burleigh et al. 2010). In practice, the mapping that minimizes the
number of episodes is determined by only the largest gene trees. Furthermore,
randomizing the leaf labels (taxon names) on the gene trees can result in gene tree
mappings that imply fewer episodes (Burleigh et al. 2010). Thus, although the
notion of finding gene tree mappings that are consistent with large-scale duplications is desirable, it is not clear that this problem has been properly formulated.
276
If the gene trees are accurate, the parsimony criterion for mapping gene duplications appears to perform well when the rates of duplication and loss are low
(Åkerborg et al. 2009; Doyon et al. 2009). However, these approaches do not
consider branch lengths in the gene or species trees, and they have a limited ability
to allow multiple duplications and losses on a single branch. Perhaps more
important, it is difficult to incorporate the parsimony criterion into a rigorous
statistical framework to examine evolutionary hypotheses associated with gene
duplication. Numerous likelihood-based models of gene duplication and loss for
reconciling gene trees and species trees have been proposed (e.g., Arvestad et al.
2003, 2004, 2009; Åkerborg et al. 2009; Doyon et al. 2009, 2011a; Rasmussen and
UN
Editor Proof
5 Identifying the Phylogenetic Context of Whole-Genome
Layout: T1 Standard SC
Chapter No.: 5
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 86/91
J. Gordon Burleigh
PR
OO
F
Editor Proof
86
321
322
323
324
325
326
327
328
329
330
331
332
333
334
335
336
337
338
EC
319
320
Kellis 2011; Gorecki et al. 2011). Most modeling approaches are based on using
birth–death processes to model duplications and losses of genes as they evolve
within a species tree (e.g., Arvestad et al. 2003, 2004, 2009; Åkerborg et al. 2009;
Rasmussen and Kellis 2011). Also, Doyon et al. (2009, 2011a) introduced a reconciliation algorithm that calculates the likelihood of possible reconciliations
based on a constant duplication and loss rate model, and this appears to produce
similar reconciliations as a parsimony approach. Gorecki et al. (2011) used a
Poisson model to identify the most likely distribution of reconciliations across
branches in the tree based on the species branch lengths and the gene tree
topologies.
Most of the likelihood model-based approaches require as input a species tree
with branch lengths. The method of Gorecki et al. (2011) also requires gene tree
topologies. Thus, it may be susceptible to the same problems with gene tree error
as parsimony approaches. Several approaches, however, take a gene sequence
alignment as input and use a Markov chain Monte Carlo (MCMC) approach to
simultaneously obtain the posterior distributions of gene tree topologies and gene
duplication and loss mappings (e.g., Arvestad et al. 2004, 2009; Åkerborg et al.
2009; Rasmussen and Kellis 2011). This approach provides an elegant, although
computationally difficult, way to incorporate gene tree uncertainty into the gene
tree reconciliation. Rasmussen and Kellis (2011) demonstrated that this approach
can produce more accurate gene trees and consequently greatly reduce the number
of implied duplication events compared to a parsimony approach.
CO
RR
318
UN
317
TE
D
Fig. 5.3 Examples of gene duplication episodes. Duplications in the gene trees are noted with a
* followed by their location in the species tree. Both gene trees have three duplications, one at the
root node and two at species node C. In gene tree 1, the two duplications at species node C can be
explained by a single duplication episode (since there is not a parent–child relationship between
the duplication nodes). However, in gene tree 2, the two duplication episodes must have occurred
at species node C since one duplication preceded the other
Layout: T1 Standard SC
Chapter No.: 5
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 87/91
87
352
5.4 Other Genomic Data
347
348
349
350
353
354
355
356
357
358
359
360
361
362
363
364
365
366
367
368
369
370
371
372
373
374
375
376
377
378
379
PR
OO
345
346
D
344
From availability of large-scale genomic data, evidence of cryptic ancient WGDs
often comes from either identifying large, syntenic (duplicated) blocks within a
single genome, or by looking at the age distribution of duplicated genes within a
chromosome (see Van de Peer 2004). Since these approaches use only data from
a single species and are focused more on identifying ancient WGDs than placing
the WGDs in an evolutionary context, I will not cover them in detail. The presence
of duplicated chromosomal segments may provide direct, unambiguous evidence
of WGDs that may be difficult to obtain from simply gene copy numbers or gene
trees (Vision et al. 2000). However, in practice, rapid gene losses and rearrangements after polyploidy can make it extremely difficult to detect such duplications, and different methods of detecting duplicated blocks and using different
criteria for defining a syntenic block can greatly affect interpretations of the history
of large-scale duplications (see Durand and Hoberman 2006). Although simply
examining the genome of a single species cannot reveal the phylogenetic context
of a WGD, the dates of the ancient divergences can be estimated based on the
molecular divergence of paralogs. It may be possible to map the evolution of large
duplicated segments on a tree, but in plants, this may require extending the taxonomic sampling of species with adequate genomic mapping data. Perhaps the
greater contribution of these duplicated regions is that they can define sets of
paralogs that originated from WGDs, and this information can be used to validate
mappings of duplications from gene copy number or gene reconciliation analyses.
The rapidly increasing abundance of large-scale transcriptome data sets for
plants provides an opportunity to define WGDs based on the age distribution of
duplicated genes (see Cui et al. 2006). The methods first can define pairs of most
recent gene duplicates using methods such as an all-by-all BLAST. If gene
duplication and loss occur at a constant rate, the frequency of duplicated genes in a
genome will decrease exponentially with time. In contrast, a large-scale duplication
TE
342
343
EC
341
CO
RR
340
F
351
The computational complexity of these likelihood-based approaches raises concerns that they may have difficulty exploiting the magnitude of new genomic data.
Yet, in many ways, they still greatly simplify the complexity of genome evolution.
Rasmussen and Kellis (2012) have described the first models of both duplication and
coalescence, and other modeling approaches estimate the effects of hybridization and
coalescence, but not duplication and loss (e.g., Meng and Kubatko 2009; Gerard et al.
2011). These represent important steps in simultaneously accounting for the many
processes that affect gene tree topologies. Still, all of the modeling approaches for
duplications and losses assume that the genes and all gene duplications are independent. Thus, they may detect branches in the species tree with high rates of
duplication or loss, but they do not directly assess the likelihood of WGDs. The
development of such models that allow for simultaneous duplications across many
genes can allow for rigorous statistical tests of the placement of WGD events.
339
UN
Editor Proof
5 Identifying the Phylogenetic Context of Whole-Genome
Layout: T1 Standard SC
Chapter No.: 5
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 88/91
J. Gordon Burleigh
391
5.5 Conclusions
382
383
384
385
386
387
388
389
PR
OO
381
F
390
event, like a WGD, should result in an overrepresentation of duplicated gene pairs
at the time corresponding to the large-scale duplication event. In practice, in a plot
of the age distribution, usually represented by synonymous substitution distance, of
duplicated genes, peaks in the age distribution curves or evidence of multiple
distributions, may indicate WGDs. Again, it is difficult to precisely place a WGD
just from the pairwise divergence of duplicated sequences, but with data available
from many taxa, comparisons of these age plots from related species can be
informative. In some cases, analyses of the age distribution plots have failed to
detect known WGDs (e.g., Blanc and Wolfe 2004; Paterson et al. 2004). However,
unlike gene tree reconciliation methods, they will not be misled by incomplete
lineage sorting or gene tree error.
380
407
408
409
410
411
Acknowledgments This chapter was developed and written in parts with support from the Gene
Tree Reconciliation Working Group at NIMBioS through NSF award EF-0832858, with additional support from the University of Tennessee. Discussions with members of the working group
including Cecile Ané, Oliver Eulenstein, Pawel Gorecki, and Brian O’Meara were helpful for this
manuscript.
412
References
396
397
398
399
400
401
402
403
404
405
413
414
415
416
TE
395
EC
394
CO
RR
393
D
406
New genomic sequence data and advances in phylogenetic methods presents
unprecedented opportunities to place WGDs within the plant tree of life. Still, there is
much work to do. Although numerous data sources and methods may provide evidence of WGDs, there exist few statistical tests of WGD. A rigorous statistical
framework still must be developed to examine hypotheses about the locations of
WGDs. Also, examinations of the phylogenetic placement of WGDs often are based
on available data; data sets are rarely generated solely for the purpose of placing the
location of WGDs. Thus, there has been little discussion about the optimal methods
or optimal data sets for mapping WGDs. Indeed, this is a complex issue. For example,
gene trees may allow direct observations of the patterns of gene duplication and loss,
but they also are susceptible to many errors and biases that may not be problems with
simpler data, such as gene copy number. Ideally, learning more about the evolutionary context and implications of WGDs in plants (e.g., their effect on diversification rates and their relationship to phenotypes such as life history or mating system)
will also help to identify and place WGDs in plants.
392
UN
Editor Proof
88
Åkerborg Ö, Sennlad B, Arvestad L, Lagergren J (2009) Simultaneous bayesian gene tree
reconstruction and reconciliation analysis. Proc Natl Acad Sci USA 106:5714–5719
Ames RM, Money D, Ghatge VP, Whelan S, Lovell SC (2012) Determining the evolutionary
history of gene families. Bioinformatics (In press)
Layout: T1 Standard SC
Chapter No.: 5
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 89/91
89
EC
TE
D
PR
OO
F
Ané C (2008) Analysis of comparative data with hierarchical autocorrelation. Ann Appl Stat
2:107–1102
Arvestad L, Berglund A-C, Lagergren J, Sennblad B (2003) Bayesian gene/species tree
reconciliation and orthology analysis using MCMC. Bioinformatics 19:i7–i15
Arvestad L, Berglund A-C, Lagergren J, Sennblad B (2004) Gene tree reconstruction and
orthology analysis based on an integrated model for duplications and sequence evolution.
RECOMB 2004:326–335
Arvestad L, Lagergren J, Sennblad B (2009) The gene evolution model and computing its
associated probabilities. J ACM 56:7
Bansal MS, Eulenstein O (2008) The multiple gene duplication problem revisited. Bioinformatics
24:i132–i138
Berglund-Sonnhammer A-C, Steffansson P, Betts MJ, Liberles DA (2006) Optimal gene-trees
from sequences and species trees using a soft interpretation of parsimony. J Mol Evol
63:240–250
Blanc G, Wolfe KH (2004) Widespread paleopolyploidy in model plant species inferred from age
distributions of duplicate genes. Plant Cell 16:1093–1101
Blanc G, Hokamp K, Wolfe KH (2003) A recent polyploidy superimposed on older large-scale
duplications in the Arabidopsis genome. Genome Res 13:137–144
Bowers JE, Chapman BA, Rong J, Paterson AH (2003) Unravelling angiosperm genome eolution
by phylogenetic analysis of chromosomal duplication events. Nature 422:433–438
Buggs RJA, Doust AN, Tate JA, Koh J, Soltis K, Feltus FA, Paterson AH, Soltis PS, Soltis DE
(2009) Gene loss and silencing in Tragopogon miscellus (Asteraceae): comparison of natural
and synthetic allotetraploids. Heredity 103:73–81
Buggs RJA, Chamala S, Wu W, Tate JA, Schnable PS, Soltis DE, Soltis PS, Barbazuk WB (2012)
Rapid, repeated, and clustered loss of duplicated genes in allopolyploid plant populations of
independent origin. Curr Biol 22:1–5
Burleigh JG, Bansal MS, Wehe A, Eulenstein O (2009) Locating large-scale gene duplication
events through reconciled trees: implications for identifying ancient polyploidy in plants.
J Comput Biol 16:1071–1083
Burleigh JG, Bansal M, Eulenstein O, Vision TJ (2010) Inferring species trees from gene
duplication episodes. Proc BCB 2010:198–203
Chang W-C, Eulenstein O (2006) Reconciling gene trees with apparent polytomies. COCOON
2006. LNCS 4112:235–244
Chapman BA, Bowers JE, Schulze SR, Paterson AH (2004) A comparative phylogenetic
approach for dating whole genome duplication events. Bioinformatics 20:180–185
Chaudhary R, Burleigh JG, Eulenstein O (2011) Algorithms for rapid error correction for the
gene duplication problem (ISBRA) 2011. LNCS 6674:184-196
Chaudhary R, Burleigh JG, Eulenstein O (2012) Efficient error correction algorithms for gene tree
reconciliation based on duplication, duplication and loss, and deep coalescence. BMC
Bioinformatics 13:s11
Chen K, Durand D, Farach-Colton M (2000) Notung: a program for dating gene duplications and
optimizing gene family trees. J Comput Biol 7:429–447
Chester M, Gallagher JP, Symonds VV, Cruz da Silva AV, Mavrodiev EV, Leitch AR, Soltis PS,
Soltis DE (2012) Extensive chromosomal variation in a recently formed natural allopolyploid
species, Tragapogon miscellus (Asteraceae). Proc Nat Acad Sci USA 109:1176–1181
Csurös M (2010) Count: evolutionary analysis of phylogenetic profiles with parsimony and
likelihood. Bioinformatics 26:1910–1912
Cui L, Wall PK, Leebens-Mack JH, Lindsay BG, Soltis DE, Doyle JJ, Soltis PS, Carlson JE,
Arumuganathan K, Barakat A, Albert VA, Ma H, de Pamphilis CW (2006) Widespread
genome duplications throughout the history of flowering plants. Genome Res 16:738–749
Cusimano N, Sousa A, Renner SS (2012) Maximum likelihood inference implies a high, not a
low, ancestral haploid chromosome number in araceae, with a critique of the bias introduced
by ‘x’. Ann Bot 109:681-692
CO
RR
417
418
419
420
421
422
423
424
425
426
427
428
429
430
431
432
433
434
435
436
437
438
439
440
441
442
443
444
445
446
447
448
449
450
451
452
453
454
455
456
457
458
459
460
461
462
463
464
465
466
467
468
469
UN
Editor Proof
5 Identifying the Phylogenetic Context of Whole-Genome
Layout: T1 Standard SC
Chapter No.: 5
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 90/91
EC
TE
D
PR
OO
F
De Bie T, Cristianini N, Demuth JD, Hahn MW (2006) CAFÉ: a computational tool for the study
of gene family evolution. Bioinformatics 22:1269–1271
DeBolt S (2010) Copy number variation shapes genome diversity in Arabidopsis over immediate
family generational scales. Genome Biol Evol 2:441–453
Digby L (1912) The cytology of Primula kewensis and of other related Primula hybrids. Ann Bot
26:357–388
Doyle JJ, Flagel LE, Paterson AH, Rapp RA, Soltis DE, Soltis PS, Wendel JF (2008)
Evolutionary genetics of genome merger and doubling in plants. Annu Rev Genet 42:443–461
Doyon J-P, Chauve C, Hamel S (2009) Space of gene/species tree reconciliations and
parsimonious models. J Comput Biol 16:1399–1418
Doyon J-P, Hamel S, Chauve C (2011a) An efficient method for exploring the space of gene tree/
species tree reconciliations in a probabilistic framework. IEEE/ACM Trans. Comput Biol
Bioinform 99: (In press)
Doyon J-P, Ranwez V, Daubin V, Berry V (2011b) Models, algorithms and programs for
phylogeny reconciliation. Briefings Bioinform 12:392–400
Durand D, Halldórsson B, Vernot B (2006) A hybrid micro-macroevolutionary approach to gene
tree reconstruction. J Comput Biol 13:320–335
Durand D, Hoberman R (2006) Diagnosing duplications—can it be done? Trends Genet
22:156–164
Ehrendorfer F, Krendl F, Habeler E, Sauer W (1968) Chromosome numbers and evolution in
primitive angiosperms. Taxon 17:337–468
Eulenstein O, Huzurbazar S, Liberles DA (2010) Reconciling phylogenetic trees. In: Dittmar K,
Liberles D (eds) Evolution after gene duplication. Wiley, Hoboken, pp 185–206
Gaut BS (2001) Patterns of chromosomal duplication in maize and their implications for
comparative maps of the grasses. Genome Res 11:55–66
Gaut BS, Doebley JF (1997) DNA sequence evidence for the segmental allotetraploid origin of
maize. Proc Natl Acad Sci USA 94:6809–6814
Gerard D, Gibbs HL, Kubatko L (2011) Estimating hybridization in the presence of coalescence
using phylogenetic intraspecific sampling. BMC Evol Biol 11:291
Goldblatt P (1980) Polyploidy in angiosperms: monocotyledons. In: Lewis WH (ed) Polyploidy:
biological relevance. Plenum Press, New York, pp 219–239
Goodman M, Czelusniak J, Moore GW, Romero-Herrera AE, Matsuda G (1979) Fitting the gene
lineage into its species lineage, a parsimony strategy illustrated by cladograms constructed by
globin sequences. Syst Zool 28:132–163
Gorecki P, Eulenstein O (2012) Simultaneous error correction and rooting for gene tree
reconciliation and the gene duplication problem. BMC Bioinformatics (In press)
Gorecki P, Eulenstein O, Burleigh JG (2011) Maximum likelihood models and algorithms for
gene tree evolution with duplications and losses. BMC Bioinform 12:S15
Grant V (1963) The origin of adaptations. Columbia University Press, New York
Grant V (1982) Periodicities in the chromosome numbers of the angiosperms. Bot Gaz
143:379–389
Guggisberg A, Mansion G, Kelso S, Conti E (2006) Evolution of biogeographic patterns, ploidy
levels, and breeding systems in a diploid-polyploid species complex in primula. New Phytol
171:617–632
Guigó R, Muchnik I, Smith TF (1996) Reconstruction of ancient molecular phylogeny. Mol
Phylogenet Evol 6:189–213
Hahn MW (2007) Bias in phylogenetic tree reconciliation methods: implications for vertebrate
genome evolution. Genome Biol 8:R141
Hahn MW, De Bie T, Stajich JE, Nguyen C, Cristianni N (2005) Estimating the tempo and mode
of gene family evolution from comparative data. Genome Res 15:1153–1160
Hipp AL, Rothrock PE, Reznicek AA, Berry PE (2007) Chromosome number changes associated
with speciation in sedges: a phylogenetic study in Carex section Ovales (Cyperaceae) using
AFLP data. Aliso 23:193–203
CO
RR
470
471
472
473
474
475
476
477
478
479
480
481
482
483
484
485
486
487
488
489
490
491
492
493
494
495
496
497
498
499
500
501
502
503
504
505
506
507
508
509
510
511
512
513
514
515
516
517
518
519
520
521
522
J. Gordon Burleigh
UN
Editor Proof
90
Layout: T1 Standard SC
Chapter No.: 5
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 91/91
91
EC
TE
D
PR
OO
F
Iwasaki W, Takagi T (2007) Reconstruction of highly heterogeneous gene-content evolution
across the three domains of life. Bioinformatics 23:i230–i239
Jiao Y, Wickett NJ, Ayampalayam S, Chanderbali AS, Landherr L, Ralph PE, Tomsho LP, Hu Y,
Liang H, Soltis PS, Soltis DE, Clifton SW, Schlarbaum SE, Schuster SC, Ma H, LeebensMack J, de Pamphilis CW (2011) Ancestral polyploidy in seed plants and angiosperms.
Nature 473:97–102
Kunin V, Ouzounis CA (2003) GeneTRACE-reconstruction of gene content of ancestral species.
Bioinformatics 19:1412–1416
Levin DA (2002) The role of chromosomal change in plant evolution. Oxford University Press,
New York
Levin DA, Wilson AC (1976) Rates of evolution in seed plants: net increase in diversity of
chromosome numbers and species numbers through time. Proc Natl Acad Sci 73:2086–2090
Librado P, Vieira FG, Rozas J (2012) BadiRate: estimating family turnover rates by likelihoodbased methods. Bioinformatics 28:279–281
Lim KY, Soltis DE, Soltis PS, Tate J, Matyasek R, Srubarova H, Kovarik A, Pires JC, Xiong Z,
Leitch AR (2008) Rapid chromosome evolution in recently formed polyploids in Tragapogon
(Asteraceae). PLoS ONE 3:e3353
Liu L, Yu L, Kalavacharla V, Liu Z (2011) A bayesian model for gene family evolution. BMC
Bioinform 12:426
Luo CW, Chen MC, Chen YC, Yang RWL, Liu HF, Chao KM (2009) Linear-time algorithms for
the multiple gene duplication problems. IEEE/ACM Trans Comput Biol Bioinform 99:5555
Maddison WP (1997) Gene trees in species trees. Syst Biol 46:523–536
Masterson J (1994) Stomatal size in fossil plants: evidence for polyploidy in majority of
angiosperms. Science 264:421–424
Mayrose I, Barker MS, Otto SP (2010) Probabilistic models of chromosome evolution and the
inference of polyploidy. Syst Biol 59:132–144
Mayrose I, Zhan SH, Rothfels CJ, Magnus-Ford K, Barker MS, Rieseberg LH, Otto SP (2011)
Recently formed polyploidy plants diversify at lower rates. Science 333:1257
Meng C, Kubatko LS (2009) Detecting hybrid speciation in the presence of incomplete lineage
sorting using gene tree incongruence: a model. Theor Popul Biol 75:35–45
Meyers LA, Levin DA (2006) On the abundance of polyploids in flowering plants. Evolution
60:1198–1206
Mirkin BG, Fenner TI, Galperin MY, Koonin EV (2003) Algorithms for computing parsimonious
evolutionary scenarios for genome evolution, the last universal common ancestor and
dominance of horizontal gene transfer in the evolution of prokaryotes. BMC Evol Biol 3:2
Mishima M, Ohmido N, Fukui K, Yahara T (2002) Trends in site-number change of rDNA loci
during polyploidy evolution in Sanguisorba (Rosaceae). Chromosoma 110:550–558
Page RDM (1994) Maps between trees and cladistic analysis of historical associations among
genes, organisms, and areas. Syst Biol 43:58–77
Page RDM, Cotton JA (2002) Vertebrate phylogenomics: reconciled trees and gene duplication.
Pac Symp Biocomput, 536–547
Paterson AH, Bowers JE, Chapman BA (2004) Ancient polyploidization predating divergence of
the cereals, and its consequences for comparative genomics. Proc Natl Acad Sci
101:9903–9908
Rasmussen MD, Kellis M (2011) A Bayesian approach for fast and accurate gene tree
reconstruction. Mol Biol Evol 28:273–290
Rasmussen MD, Kellis M (2012) Unified modeling of gene duplication, loss and coalescence
using a locus tree. Genome Res 22:755-765
Raven PH (1975) The bases of angiosperm phylogeny: cytology. Ann Mo Bot Gard 62:724–764
Schluter D, Price T, Mooers AØ, Ludwig D (1997) Likelihood of ancestor states in adaptive
radiation. Evolution 41:1239–1251
Schultheis LM (2001) Systematics of Downingia (Campanulaceae) based on molecular sequence
data: implications for floral and chromosome evolution. Syst Bot 26:603–621
CO
RR
523
524
525
526
527
528
529
530
531
532
533
534
535
536
537
538
539
540
541
542
543
544
545
546
547
548
549
550
551
552
553
554
555
556
557
558
559
560
561
562
563
564
565
566
567
568
569
570
571
572
573
574
575
UN
Editor Proof
5 Identifying the Phylogenetic Context of Whole-Genome
Layout: T1 Standard SC
Chapter No.: 5
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 92/91
EC
TE
D
PR
OO
F
Soltis DE, Albert VA, Leebens-Mack J, Bell CD, Patterson AH, Zheng C, Sankoff D, de
Pamphilis CW, Wall PK, Soltis PS (2009) Polyploidy and angiosperm diversification. Am J
Bot 96:336–348
Soltis PS, Soltis DE (2000) The role of genetic and genomic attributes in the success of
polyploids. Proc Natl Acad Sci 97:7051–7057
Snel B, Bork P, Huynen MA (2002) Genomes in flux: the evolution of archael and proteobacterial
gene content. Genome Res 12:17–25
Springer NM, Ying K, Fu Y, Ji T, Yeh C-T, Jia Y, Wu W, Richmond T, Kitzman J, Rosenbaum
H, Iniguez AL, Barbazuk WB, Jeddeloh JA, Nettleton D, Schnable PS (2009) Maize inbreds
exhibit high levels of copy number variation (CNV) and presence/absence (PAV) in genome
content. PLoS Genet 5:e1000734
Stace HM, Chapman AR, Lemson KL, Powell JM (1997) Cytoevolution, phylogeny, and
taxonomy in Epacridaceae. Ann Bot 79:283–290
Stebbins GL (1938) Cytological characteristics associated with the different growth habits in the
dicotyledons. Am J Bot 25:189–198
Stebbins GL (1950) Variation and evolution in plants. Columbia University Press, New York
Stebbins GL (1971) Chromosomal evolution in higher plants. Addison-Wesley, London
Tate JA, Ni Z, Scheen A-C, Koh J, Gilbert CA, Lefkowitz D, Chen ZJ, Soltis PS, Soltis DE
(2006) Evolution and expression of homeologous loci in Tragopogon miscellus (Asteraceae),
a recent and reciprocally formed allopolyploid. Genetics 173:1599–1611
Vamosi JC, Dickinson TA (2006) Polyploidy and diversification: a phylogenetic investigation in
Rosaceae. Int J Plant Sci 167:349–358
Vandepoele K, Simillion C, Vande Peer Y (2003) Evidence that rice and other cereals are ancient
aneuploids. Plant Cell 15:2192–2202
Van de Peer Y (2004) Computational approaches to unveiling ancient genome duplications. Nat
Rev Genet 5:752–763
Vision TJ, Brown DG, Tanksley SD (2000) The origins of genomic duplication in arabidopsis.
Science 290:2114–2117
Walker JW (1972) Chromosome numbers, phylogeny, phytogeography of the Annonaceae and
their bearing on the (original) basic chromosome number of angiosperms. Taxon 21:57–65
Winge Ö (1917) The chromosomes. Their numbers and general importance. Comptes Rendus des
Travaux Laboratoire Carlsberg 13:131–275
Wolfe KH (2001) Yesterday’s polyploids and the mystery of diploidization. Nat Rev Genet
2:333–341
Wood TE, Takebayashi N, Barker MS, Mayrose I, Greenspoon PB, Rieseberg LH (2009) The
frequency of polyploidy speciation in vascular plants. Proc Natl Acad Sci USA
106:13875–13879
Zheng L-Y, Guo X-S, He B, Sun L-J, Peng Y, Dong S-S, Liu T-F, Jiang S, Ramachandran S, Liu C-M,
Jing H-C (2011) Genome-wide patterns of genetic variation in sweet and grain sorghum (Sorghum
bicolor). Genome Biol 12:R114
CO
RR
576
577
578
579
580
581
582
583
584
585
586
587
588
589
590
591
592
593
594
595
596
597
598
599
600
601
602
603
604
605
606
607
608
609
610
611
612
613
614
615
J. Gordon Burleigh
UN
Editor Proof
92
Metadata of the chapter that will be visualized in
SpringerLink
Book Title
Polyploidy and Genome Evolution
Series Title
Chapter Title
Ancient and Recent Polyploidy in Monocots
Copyright Year
2012
Copyright HolderName
Springer-Verlag Berlin Heidelberg
Corresponding Author
Family Name
Paterson
Particle
Given Name
Andrew H.
Suffix
Author
Division
Plant Genome Mapping Laboratory
Organization
University of Georgia
Address
Athens, GA, USA
Email
paterson@plantbio.uga.edu
Family Name
Wang
Particle
Given Name
Xiyin
Suffix
Author
Division
Plant Genome Mapping Laboratory
Organization
University of Georgia
Address
Athens, GA, USA
Division
Center for Genomics and Computational Biology
Organization
Hebei United University
Address
Tangshan, People’s Republic of China
Email
wang.xiyin@gmail.com
Family Name
Li
Particle
Given Name
Jingping
Suffix
Author
Division
Plant Genome Mapping Laboratory
Organization
University of Georgia
Address
Athens, GA, USA
Email
jingpingli@gmail.com
Family Name
Tang
Particle
Given Name
Haibao
Suffix
Division
Plant Genome Mapping Laboratory
Organization
University of Georgia
Address
Athens, GA, USA
Division
Abstract
Organization
J. Craig Venter Institute
Address
Rockville, MD, USA
Email
tanghaibao@gmail.com
At least two whole-genome duplications (WGD) have profoundly influenced the evolution of most, if not all,
grass (Poaceae) genomes, with the most recent of these predating the divergence of these lineages by 20
million or more years. Taxa within each major lineage of Poaceae (e.g., Panicoideae, Ehrhartoideae, Pooideae)
have independently experienced additional polyploidizations that have been of central importance to the
evolution and productivity of some of our most important crop plants [for example, sugarcane (Saccharum
spp.), and durum and bread wheat (Triticum spp.)]. Following polyploidy, adaptation to the duplicated state
is evident at the levels of transmission genetics, chromosome structure, and gene repertoire. While most
duplicated chromosomal regions re-establish largely independent evolution within a few million years, 70million-year-old duplicated chromosome segments in one unusual region of the rice genome and its orthologs
in other grasses have continued to exhibit concerted evolution more recently than the divergence of rice
subspecies japonica and indica an estimated 400,000 years ago.
1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 93/107
Chapter 6
4
Andrew H. Paterson, Xiyin Wang, Jingping Li and Haibao Tang
11
12
13
14
15
16
17
19
18
PR
OO
D
9
10
TE
8
EC
7
Abstract At least two whole-genome duplications (WGD) have profoundly influenced the evolution of most, if not all, grass (Poaceae) genomes, with the most recent
of these predating the divergence of these lineages by 20 million or more years. Taxa
within each major lineage of Poaceae (e.g., Panicoideae, Ehrhartoideae, Pooideae)
have independently experienced additional polyploidizations that have been of central
importance to the evolution and productivity of some of our most important crop
plants [for example, sugarcane (Saccharum spp.), and durum and bread wheat
(Triticum spp.)]. Following polyploidy, adaptation to the duplicated state is evident at
the levels of transmission genetics, chromosome structure, and gene repertoire. While
most duplicated chromosomal regions re-establish largely independent evolution
within a few million years, 70-million-year-old duplicated chromosome segments in
one unusual region of the rice genome and its orthologs in other grasses have continued to exhibit concerted evolution more recently than the divergence of rice subspecies japonica and indica an estimated 400,000 years ago.
CO
RR
5
6
F
3
Ancient and Recent Polyploidy
in Monocots
2
A. H. Paterson (&) X. Wang J. Li H. Tang
Plant Genome Mapping Laboratory, University of Georgia, Athens, GA, USA
e-mail: paterson@plantbio.uga.edu
X. Wang
e-mail: wang.xiyin@gmail.com
J. Li
e-mail: jingpingli@gmail.com
H. Tang
e-mail: tanghaibao@gmail.com
X. Wang
Center for Genomics and Computational Biology, Hebei United University, Tangshan,
People’s Republic of China
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: 6
H. Tang
J. Craig Venter Institute, Rockville, MD, USA
P. S. Soltis and D. E. Soltis (eds.), Polyploidy and Genome Evolution,
DOI: 10.1007/978-3-642-31442-1_6, Springer-Verlag Berlin Heidelberg 2012
93
Layout: T1 Standard SC
Chapter No.: 6
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 94/107
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
61
62
F
25
26
PR
OO
24
D
23
Monocotyledons, also known as monocots, are one of the major clades of
angiosperms. Most recent phylogenetic analyses (based largely on plastid
sequence data) reveal a successive series of basal angiosperm lineages, with
monocots often sister to Ceratophyllum plus eudicots (e.g., Soltis et al. 2011).
According to the IUCN there are 59,300 species of monocots (http://
cmsdata.iucn.org/). The most species-rich family in this clade (and indeed, one
of the largest of all angiosperms) are the orchids (family Orchidaceae), with more
than 20,000 species (Raven et al. 2005). Among the best-studied monocots are the
grasses, family Poaceae (Gramineae), which provide much of the world’s food and
plant biomass and include economically important grains, such as rice (Oryza),
wheat (Triticum), maize (Zea), barley (Hordeum), and sorghum (Sorghum), turf
and forage/pasture grasses, sugar cane, and the bamboos. Other economically
important monocot families are the palms (Arecaceae), bananas (Musaceae),
gingers (Zingiberaceae), and the onion family Alliaceae.
As of this writing, the vast majority of monocot genetic and genomic information, including all whole-genome-scale DNA sequences of sufficient contiguity
for synteny analysis, are for members of Poaceae, necessarily constraining the
focus of this chapter. However, this constraint is expected to be relieved in the
very near future, with genome sequences in progress for several non-grass
monocots (e.g. Phoenix dactylifera, Musa acuminata, Ananas comosus, Asparagus
officinalis, Phalaenopsis equestris, Zostera marina), adding important new
dimensions to understanding of monocot gene and genome evolution. A draft of
the date palm (P. dactylifera) genome is publicly available, but is presently of too
low contiguity for robust synteny analysis.
High-quality sequences for representatives of all three major grass clades have
been published, including rice (International Rice Genome Sequencing Project
2005; Yu et al. 2005) (Ehrhartoideae), sorghum (Paterson et al. 2009b) and maize
(Schnable et al. 2009) (Panicoideae), and Brachypodium (The International
Brachypodium Initiative 2010) (Pooideae) (Fig. 6.1). Draft genome sequence of
barley (Hordeum) and wheat (Triticum) group 1 chromosomes were recently made
available (Mayer et al. 2011; Wicker et al. 2011).
Initial analyses of the available genome sequences available for Poaceae have
shown at least two whole-genome duplications (WGD) influencing most, if not all,
grass genomes, with the most recent of these predating the divergence of these
lineages by an estimated 20 million or more years. Taxa within each lineage have
independently experienced additional polyploidizations and readaptation to the
duplicated state (for example, sugarcane and durum and bread wheat), with the
model genomes constituting a good starting point for accelerating progress in the
study and improvement of many additional taxa. Although chloridoid and arundinoid grasses are explored only at the EST level to date, these data show that at
least the Chloridoideae experienced an evolutionary history similar to those of the
other major grass clades, including both of the two WDGs shared by other
TE
22
EC
21
6.1 Monocot Comparative Genomics
CO
RR
20
A. H. Paterson et al.
UN
Editor Proof
94
Layout: T1 Standard SC
Chapter No.: 6
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 95/107
95
CO
RR
EC
TE
D
PR
OO
F
Editor Proof
6 Ancient and Recent Polyploidy in Monocots
63
64
65
66
UN
Fig. 6.1 Phylogenetic relationships among major lineages of monocots and estimated positions
of early paleopolyploidy events (modified from Paterson et al. 2009a). Notice that some lack
sufficient contiguity (Phoenix) or data (Ananas, Zostera) to infer genome duplication, and
accordingly the position of sigma (r) (Tang et al. 2010) cannot be precisely determined at this
time; this uncertainty is indicated by arrows
Poaceae. Chloridoids also have an additional lineage-specific genome duplication
(Kim et al. 2009).
By developing and using comparative genomics tools, research starting from
the sequenced genomes may transitively shed light on monocots that are still
Layout: T1 Standard SC
Chapter No.: 6
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 96/107
A. H. Paterson et al.
73
6.2 Rho (q), the Most Recent Pan-Poaceae Polyploidy
75
76
77
78
79
80
81
82
83
84
85
86
87
88
89
90
91
92
93
94
95
96
97
98
99
100
101
102
103
104
105
PR
OO
74
The notion that even relatively small and structurally simple grass genomes may
be paleopolyploid is of long-standing interest. Indeed, secondary associations of
grass chromosomes have been known for 80 years (Lawrence 1931), and genetic
mapping nearly 2 decades ago suggested duplication of scattered chromosome
segments in both rice (Kishimoto et al. 1994; Nagamura et al. 1995) and sorghum
(Chittenden et al. 1994).
Paleopolyploidy in the grasses was demonstrated beyond a reasonable doubt with
the first availability of the whole-genome sequence of rice, Oryza sativa ssp.
japonica (Goff et al. 2002). A brief controversy about whether the scope of duplication was confined to specific chromosomes (Vandepoele et al. 2003) or the whole
genome (Paterson et al. 2003, 2004) was soon reconciled, by analyzing the genome
sequence of another rice cultivar, O. sativa ssp. indica (Yu et al. 2005), with the
controversy attributed to differences in the approaches used to infer the duplicated
blocks (Wang et al. 2005). This polyploidy event (q) was dated to *70 million years
ago (mya) based on putatively neutral DNA substitution rates between duplicated
genes, and suggested to be shared by all main lineages of grasses.
The genome sequences of sorghum (Sorghum bicolor) and Brachypodium (The
International Brachypodium Initiative 2010) confirmed that these taxa share the
paleoduplication first discerned in rice, and that neither has been affected by
additional polyploidization after the evolutionary split of their respective lineages.
Moreover, Brachypodium and rice have preserved a very high level of gene collinearity (Paterson et al. 2009b; The International Brachypodium Initiative 2010),
making it possible to take them as a single genetic system to perform transitive
genetics research across different grasses (Freeling 2001). Only a small fraction of
genes shows differential gene losses after the split of rice (1.8 %) and sorghum
(3.1 %). These findings suggest that, after the 70-mya polyploidization event, the
genome of the last common ancestor of grasses had already experienced most gene
loss and reached a relatively stable state prior to the divergence of the extant major
grass lineages about 50 mya (Paterson 2008; The International Brachypodium
Initiative 2010). Some prior reports of deviations from collinearity in the grasses
may be accounted for by these low rates of differential gene loss on homoeologous
chromosomes.
D
71
TE
70
EC
69
CO
RR
68
F
72
lacking de novo genomic data (Van de Peer 2004; Wang et al. 2006; Lohithaswa
et al. 2007; Tang et al. 2008b). The sequences of additional grasses and non-grass
monocots such as Elaeis (Arecaceae), Musa (Musaceae), and Zostera (Zosteraceae) will clarify the functional innovation of their gene sets, further elucidating
the structural and functional evolution of this ecologically and economically
important plant family.
67
UN
Editor Proof
96
Layout: T1 Standard SC
Chapter No.: 6
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 97/107
112
113
114
115
116
117
118
119
120
121
122
123
124
125
126
127
128
129
130
131
132
133
134
135
136
137
138
139
140
141
142
143
144
145
146
147
148
F
111
PR
OO
110
Rho accounted for much, but not all, of the paleo-duplication that could be discerned
in modern cereal genomes (Paterson et al. 2004). Shortly after the discovery of rho,
an early report (Zhang et al. 2005), later supported by two independent studies
(Jaillon et al. 2007; Salse et al. 2008) hinted at the presence of additional, earlier
monocot duplications.
Detailed elucidation of sigma (r), an additional duplication event preceding rho
in the grass lineage, utilized a ‘bottom–up’ approach, first collapsing 15,640 rice
genes and 15,636 sorghum genes into 13,308 rho-nodes that computationally
reverse post-rho gene loss, increasing the sensitivity of subsequent analysis.
These nodes (genes) were compared among themselves, revealing collinear
patterns of correspondence that involve all nine major synteny blocks resulting
from the rho duplication. Some collinear patterns between pairs of rho-blocks are
one-to-one, while others are higher order, suggesting that multiple events may
have been identified. The eight largest synteny blocks that retained collinearity
following sigma contain a total of 4,168 sigma-nodes, covering 5,747 rice genes
and 5,738 sorghum genes (*20 % of the rice and sorghum transcriptomes).
Further study of these sigma-duplicated regions highlights an important
constraint in monocot genomics, albeit one that soon will be relieved. There is
little remaining intragenomic correspondence between sigma-derived rice
segments, although relationships between some of these duplicated segments can
still be identified through transitive comparisons of cereal genomes to outgroups.
At present, however, the only available outgroups are eudicots such as grape
(Vitis). Similarities between monocot and eudicot genomes resulting from
common ancestry have been obscured by many paleo-polyploidy events and
numerous genome rearrangements (Liu et al. 2001; Jaillon et al. 2007). The
availability of non-cereal monocot outgroup genomes would significantly increase
power to study these segments, particularly outgroups with genomes that have not
experienced lineage-specific duplication and associated fractionation of ancestral
gene orders.
The lack of an ideal (non-cereal non-reduplicated monocot) outgroup
notwithstanding, better understanding of monocot paleopolyploidy has improved
our ability to compare monocot and eudicot genomes, a long-standing goal
(Paterson et al. 1996) that has been complicated by paleopolyploidy events. Toward
this goal, we applied a hierarchical clustering approach, first dividing the chromosomes into small bins and then comparing all pairs of rice and grape bins.
Duplicated segments retained in grape following the eudicot ‘gamma’ hexaploidy
event (3), and homologous segments retained in rice following at least two rounds of
duplication (rho and sigma), contain 38 ‘‘putative ancestral regions’’ (PAR) that
collectively explain 19.1 % of all observed homolog pairs and 31.0 % of reciprocal
best hits between grape and rice genes, *10-fold more than would be explicable by
chance. The PARs interleave multiple grape and rice genomic regions collectively
D
109
TE
108
EC
107
97
6.3 Sigma (r), and Clarifying the Genome Composition
of the Last Universal Common Ancestor of Monocots
CO
RR
106
UN
Editor Proof
6 Ancient and Recent Polyploidy in Monocots
Layout: T1 Standard SC
Chapter No.: 6
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 98/107
156
157
158
159
160
161
162
163
164
165
166
167
168
169
170
171
172
173
174
175
176
177
178
179
180
181
182
183
184
185
186
187
188
189
F
155
PR
OO
153
154
D
152
TE
151
covering *70 % of each genome. By consolidating much of the redundancy in each
genome, the PARs create syntenic blocks with much less ambiguity and in most
cases show association between one gamma block and one sigma block. We did not
find any PAR that simultaneously mapped to two different gamma or sigma blocks
(Tang et al. 2010). Some ‘‘ghost duplications’’ (Van de Peer 2004) in rice that we
failed to identify through intragenomic comparisons (due to reciprocal gene losses
in largely complementary fashion) are much clearer in cross-species comparisons
(PARs).
Compared to the WGD events in grape where 94.5 % of the genome appears
duplicated (Jaillon et al. 2007), the rice WGDs are more complicated and
degenerate. The 38 grape–rice PARs are a qualitative advance toward a global
view of monocot-eudicot synteny. Collinearity appears to be disrupted around the
peri-centromeric regions in 10 of the 12 rice chromosomes, suggesting dynamic
reorganization of heterochromatic portions of the rice genome that may result from
massive transpositions or gene losses (Bowers et al. 2005).
Our unique approach to synteny analysis provides new insight into the number(s)
of WGD events experienced by modern cereal genomes. In many lineages, the
existence and the numbers of WGD events have been contentious. In 22 of the 38
PARs, grape–rice collinearity is clear, with 12 PARs being threefold redundant in
grape, consistent with hexaploidy (Jaillon et al. 2007). The level of redundancy in
rice is less clear, ranging from as little as twofold (1 PAR) to sevenfold (3 PARs) and
eightfold (5 PARs). In line with the intragenomic evidence from our bottom–up
analysis, these high redundancies suggest that the rice lineage experienced more
than two, perhaps three, rounds of ancient WGD that have collectively been lumped
into the single event that we are presently referring to as ‘sigma’ (r). Once again, the
availability of non-cereal and non-reduplicated monocot outgroup genomes would
significantly improve our understanding of monocot evolutionary history.
EC
150
6.4 Lineage-Specific Monocot Polyploidies and Their
Consequences
CO
RR
149
A. H. Paterson et al.
Polyploidy in monocots is, of course, not limited to ancient events, and indeed
more recent events have been integral to the evolution and productivity of many
major crops. A classical textbook example of a more recent polyploidization is the
evolution of wheat (see also Chap. 7, this volume), with formation of a tetraploid
and subsequently a hexaploid leading to independent crops, durum wheat used for
pasta (Triticum turgidum) and bread wheat (T. aestivum), respectively (Feldman
and Levy 2005). Indeed, the synthesis by humans of octoploid triticale (9 Triticosecale a hybrid between Triticum and Secale) in the past century illustrates that
there may be further gains to be made by mimicking the natural tendency of some
plant lineages to form polyploids.
Polyploidy has been particularly closely associated with productivity in
autopolyploids such as many forage and biomass grasses. One of the most extreme
UN
Editor Proof
98
Layout: T1 Standard SC
Chapter No.: 6
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 99/107
197
198
199
200
201
202
203
204
205
206
207
208
209
210
211
212
213
214
215
216
217
218
219
220
221
222
223
224
225
226
227
228
229
230
231
232
233
F
PR
OO
196
D
194
195
TE
193
EC
192
99
cases of a successful autoploid is sugar cane (Saccharum spp.), in which cultivated
forms are typically octoploid—dodecaploid (i.e., with 8–12 chromosome sets, and
sometimes variations in between!). These are interspecific hybrids (and therefore
not strict autopolyploids, but a combination of auto- and allopolyploidy) composed
of about 85–90 % chromatin from Saccharum officinarum and 10–15 % from
S. spontaneum, a wild relative. Studies using DNA markers to quantify ‘dosages’ of
homologous genomic regions in sugar cane show clearly nonlinear consequences of
allele dosage, with one or two copies of favorable alleles at homologous loci usually
having favorable effects, but additional copies yielding diminishing or even reduced
returns (Ming et al. 2001, 2002a, b). Comparative genomic studies, in particular
with sorghum, show that the Saccharum lineage experienced at least one WGD
since its divergence from the Sorghum lineage something less than 10 mya (Ming
et al. 1998; Jannoo et al. 2007).
Ploidy in Saccharum is particularly interesting, in that it has been further
complicated by a high frequency of chromosome non-reduction, yielding
2n ? n progeny in S. officinarum (female) 9 S. spontaneum (male) crosses (Bremer
1923). In efforts to broaden genetic variability in ‘noble canes’ that were the foundation of sugar cane production until the early part of the twentieth century, F1
progeny of interspecific crosses between S. officinarum and S. spontaneum were
found to be distinctively more robust than either parent. When S. officinarum clones
were used as the female parent, progeny tended to be larger stalked, higher in sucrose
levels, and generally more vigorous than when S. spontaneum clones were used as the
female parent. The pollination of noble cane with S. spontaneum followed by
repeated backcrosses to the noble canes has come to be called ‘nobilization’, with
selected hybrid progenies referred to as ‘nobilized’ canes (Bremer 1961). A key
event in the evolution of modern sugarcane cultivars was the production of the
nobilized cultivar, ‘POJ2878’, of Proefstation Oost, Java, in 1921 (Jeswiet 1929).
Further study of the clade that includes Saccharum may prove intriguing. This
clade also includes Miscanthus, which has biomass yields that are similarly high as
Saccharum, but is better adapted to temperate climates and therefore has stimulated
much interest for bioenergy production in the USA, Europe, and China (Heaton et al.
2008). Miscanthus species have a basic set of n = x = 19 chromosomes, versus the
x = 10 that is characteristic of many Saccharinae including Saccharum. One
attractive hypothesis to explain the transition from 10 to 19 chromosomes is that
Miscanthus, like Saccharum (Ming et al. 1998), may have experienced a polyploidization event in the 8–9 my since its divergence from sorghum—but unlike
Saccharum, which is largely autopolyploid, Miscanthus homologs have diverged
sufficiently that they no longer normally pair with one another—that is, there is now
preferential pairing of chromosomes. Genome evolution may have included a
chromosomal fusion to get from n = 20 to 19. If Miscanthus and Saccharum shared
a genome doubling event, a possibility that we are continuing to investigate, it would
be an intriguing and perhaps unprecedented case in which one lineage (Miscanthus)
adapted to the duplicated state by re-establishing disomy, while a sister lineage
continued to have the option of polysomy.
CO
RR
190
191
UN
Editor Proof
6 Ancient and Recent Polyploidy in Monocots
Layout: T1 Standard SC
Chapter No.: 6
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 100/107
A. H. Paterson et al.
234
6.5 Adapting to the Polyploid State
235
6.5.1 Centromeric Divergence and Restoration of Disomy
255
6.5.2 Karyotype Evolution
242
243
244
245
246
247
248
249
250
251
252
253
256
257
258
259
260
261
262
263
264
265
266
267
268
269
270
271
PR
OO
241
D
240
TE
239
EC
238
A polyploidy event may result in genomic instability, consequently incurring a
process of diploidization characterized by widespread DNA rearrangements often
accompanied by large-scale gene losses (Paterson et al. 2004; Van de Peer 2004;
Wang et al. 2006; Xiong et al. 2011). These DNA rearrangements may result in
chromosome number variations. Grasses range from n = 2 to 18 in their basic
chromosome sets (Soderstrom et al. 1987; Hilu 2004). In the sequenced genomes,
rice, sorghum, and Brachypodium have n = 12, 10, and 5 chromosomes, respectively. Although maize (Zea mays) experienced a WGD since divergence from
sorghum, modern-day maize retains the same chromosome number (n = 10) as
sorghum. Some lineages of Sorghum have experienced chromosome condensations
even in the absence of polyploidization (Spangler et al. 1999; Spangler 2003).
Comparison of grass genomes has shed light on chromosome number evolution and
ancestral grass karyotypes (Salse et al. 2009; Murat et al. 2010). An ancestral
karyotype of n = 5 chromosomes was inferred (Salse et al. 2009; Murat et al. 2010)
before the pan-grass polyploidization, with n = 2x = 10 chromosomes after the
duplication, then two fissions to result in n = 2x = 12 chromosomes in the common
CO
RR
237
F
254
One important challenge facing a newly formed autopolyploid may be that the
presence of sets of four homologous chromosomes may tend to hinder purging of
deleterious alleles. Transition to diploid inheritance would, in principle, allow
more rapid allele frequency changes and reduced genetic load.
Centromeric divergence may have been a mechanism by which paleopolyploid
grasses restored diploid inheritance (Bowers et al. 2005). A high concentration of
rice genes duplicated by ancient polyploidy falls near Ks 0.85, while rice gene pairs
with Ks 0.2–0.6 tend to be located in peri/centromeric regions. This suggests that
shortly after polyploidization, a substantial restructuring of centromeric regions
began that lasted until about 16 mya [based on the synonymous substitution rate
used (Lynch and Conery 2000)]. About 18 % of the rice genome shows highly
significant concentrations of matches (p \ 1 9 10-5) in the Ks range of 0.2–0.6.
The restructuring of non-syntenic regions largely involves migration of DNA
between pericentromeric regions of different chromosomes. The concentrations of
relatively recent single–locus duplications near the centromeres may, but does not
necessarily, reflect a higher duplication rate in these regions. Alternatively, recent
duplications may be preserved more frequently in pericentromeric regions, i.e.,
there may be more rapid loss of single-gene duplications in euchromatic regions
(Bowers et al. 2005).
236
UN
Editor Proof
100
Layout: T1 Standard SC
Chapter No.: 6
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 101/107
101
286
6.5.3 Gene Retention and Loss
278
279
280
281
282
283
284
287
288
289
290
291
292
293
294
295
296
297
298
299
300
301
302
303
304
305
306
307
308
309
310
311
312
PR
OO
277
D
276
Among the genes duplicated in the polyploidy events shared by the grass lineage,
the vast majority of pairs have lost at least one duplicated copy. The finding that
only a small fraction of genes show differential gene losses after the split of rice
(1.8 %) and sorghum (3.1 %) demonstrates empirically that, after the 70-mya
polyploidization, the genome of the last universal common ancestor of grasses had
already experienced most gene loss and reached a relatively stable state prior to the
divergence of the major grass lineages about 50 mya (Paterson 2008; The International Brachypodium Initiative 2010).
Gene losses have often occurred in a complementary and segmental manner,
that is, with non-random patterns of retention/loss on corresponding duplicated
DNA segments, in a process known as fractionation (Thomas et al. 2006). Genes
may be removed by a short-DNA deletion mechanism (Woodhouse et al. 2010),
and in a pair of duplicates, gene loss may be universally biased to preserve the
gene that is responsible for the majority of expression (Schnable et al. 2011). For
more than 90 % of the preserved duplicated genes, the two copies have the same
transcriptional orientations (Wang et al. 2005), and the exceptions may be a result
of local DNA inversions or differential gains/losses of new tandemly duplicated
genes in the paleo-duplicated regions.
In grasses, and indeed across angiosperms, we find three ‘fates’ of individual gene
pairs following duplication (see also Chap. 1, this volume). Most gene functional
groups show post-duplication gene preservation/loss rates that are indistinguishable
from the genome-wide average. Such ‘neutral’ loss of duplicated genes presumably
involves inactivating mutations opposed by very weak selection (Haldane 1933), as
the fate of the vast majority of duplicated genes. Population genetic models suggest
that loss of duplicated genes may happen over a few million years (Lynch and
Conery 2000). Genes in some specific functional categories duplicate and reduplicate
TE
275
EC
274
CO
RR
273
F
285
ancestor of major cereals. However, the authors noted that an ancestral karyotype of
n = 6–7 was also possible. They inferred that chromosome number variation/
reduction from the common ancestor may be attributable to non-random centric
double-strand break repair events. It was suggested that centromeric/telomeric
illegitimate recombination between non-homologous chromosomes led to nested
chromosome fusions and synteny break points, and concluded that these break
points were meiotic recombination hotspots that corresponded to high sequence
turnover loci through repeat invasion. These rules seem to explain most chromosome number changes in the grass genomes sequenced so far, especially the previously observed nested chromosome fusions in Brachypodium (The International
Brachypodium Initiative 2010). However, many details related to dynamics of
centromeres and telomeres during the rearrangements remain unclear, and the wide
range in possible ancestral karyotypes (from 12 to 24) suggests that further revision
of thinking on this subject is likely.
272
UN
Editor Proof
6 Ancient and Recent Polyploidy in Monocots
Layout: T1 Standard SC
Chapter No.: 6
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 102/107
A. H. Paterson et al.
325
6.6 The Unique Case of Rice Chromosomes 11 and 12
322
323
326
327
328
329
330
331
332
333
334
335
336
337
338
339
340
341
342
343
344
345
346
347
348
349
350
351
352
353
PR
OO
320
321
Rice chromosomes 11 and 12 (R11 and R12) are a striking exception among
chromosomes affected by the 70-mya polyploidization (Wang et al. 2011). R11
and R12 share a *3-Mb duplicated DNA segment at the termini of their short
arms, the formation of which had been dated based on synonymous substitutions to
*5–7 mya (The Rice Chromosomes 11 and 12 Sequencing Consortia 2005; Wang
et al. 2005; Yu et al. 2005). Remarkably, in the second grass genome sequence,
sorghum, the corresponding region(s) of its orthologous chromosomes (S5 and S8,
respectively) also contained such an apparently recent duplication despite having
diverged from rice about 50 mya (Paterson et al. 2009b). Physical and genetic
maps also suggest shared terminal segments of the corresponding chromosomes in
wheat (4, 5), foxtail millet (VII, VIII), and pearl millet (linkage groups 1, 4)
(Devos et al. 2000; Singh et al. 2007). It would be exceedingly unlikely for
segmental duplications to happen independently at such closely corresponding
locations in reproductively isolated lineages. A much more parsimonious
hypothesis is that the R11/12 and S5/8 regions each resulted from the pan-grass
duplication 70 mya but have an unusual evolutionary history (Paterson et al.
2009b).
Detailed analysis of R11 and R12 suggested that illegitimate recombination has
continued for millions of years after the 70-mya divergence of these homoeologs,
indeed remaining ongoing in the past 400,000 years since divergence of subspecies japonica and indica (Wang et al. 2007). Gradual and step-by-step restrictions
on recombination, starting around the time of the 70-mya polyploidization, have
resulted in ‘strata’ along the chromosome pair that differ in the degree of DNA
sequence similarity between homoeologous genes (Wang et al. 2011). Sequence
similarity between homoeologs in the strata reflects the time(s) of recombination
suppression rather than the times of their origination. Indeed, the most terminal
stratum in rice (RSA) appears \0.5 my old, while two more internal strata (RSB,
RSC) appear to have been restricted in their ability to recombine 9.4 and 39.1 mya,
D
319
TE
317
318
EC
316
CO
RR
315
F
324
(Blanc and Wolfe 2004; Seoighe and Gehring 2004; Maere et al. 2005; Chapman et al.
2006; Paterson et al. 2006; Tang et al. 2008a), and in many instances can be related to
the ‘‘gene balance’’ hypothesis, that stoichiometry among members of pathways and
networks is important to biological function (Birchler et al. 2005; Birchler and Veitia
2007; Veitia et al. 2008). Other specific genes and gene functional groups show more
extensive loss of duplicate copies than the genome-wide average, and this loss has
often been convergent following independent duplications separated by hundreds of
millions of years during the evolution of grasses, Arabidopsis, yeast, and Tetraodon
(pufferfish) (Paterson et al. 2006). Much greater knowledge of gene functions,
particularly regarding those less-explored genes that are recurrently restored to the
singleton state, may provide new insights into the ‘adaptation’ of a newly formed
polyploid to the duplicated state.
313
314
UN
Editor Proof
102
Layout: T1 Standard SC
Chapter No.: 6
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 103/107
103
PR
OO
F
Editor Proof
6 Ancient and Recent Polyploidy in Monocots
Fig. 6.2 Gene repertoire and organization in a terminal segment of homoeologous rice and
sorghum chromosomes experiencing concerted evolution. This region includes 4 rice-sorghum
quartets (Q), 3 triplets [T: losses on S5 (2), S8], 13 pairs of taxon-specific genes [P: 5 sorghum, 8
rice]; 8 taxon-specific singletons [S: 6 sorghum, 2 rice], and one lineage-specific duplication
(X: two terminal genes on S5 and S8 share common ancestor with single rice genes).
Classification letters for a gene family appear in only one member (members connected by lines).
Reprinted from Wang et al. (2011) with permission. Copyright is owned by the American Society
of Plant Biology
359
360
361
362
363
364
365
366
367
368
369
370
371
372
373
374
375
376
377
378
379
380
381
D
TE
357
358
EC
356
respectively. In sorghum, even the most terminal (of only two) strata dates to
13.4 mya, reflecting the parallel but independent evolution of this unusual chromosome segment in divergent lineages (Wang et al. 2011). The corresponding
regions in maize and Brachypodium also show prominent homoeologous recombination. However, widespread chromosomal rearrangement, especially in maize
after its lineage-specific polyploidization, makes the stratification patterns more
difficult to compare than in rice and sorghum.
Both intriguing and perplexing is that the distal chromosomal region with the
greatest DNA similarity between surviving duplicated genes also has the highest
concentration of lineage-specific gene pairs found anywhere in these genomes, and a
significantly elevated gene evolutionary rate (Wang et al. 2011). Of 33 and 23 riceand sorghum-specific gene pairs on these chromosomes, a respective 100 and 90 %
of them are in the young strata. Both members of a remarkable 50 % of the 16
duplicated RSA gene pairs are absent from sorghum, and 15 (38 %) of 39 SSA pairs
are absent from rice (Fig. 6.2). Gene losses on either one of a pair of homoeologs
experiencing concerted evolution may be commuted to the other, perhaps
explaining the more than tenfold higher rate of gene loss in the RSA and SSA
regions than the genome-wide averages of 1.8 % in rice and 3.1 % in sorghum since
their divergence about 50 mya.
The singular evolutionary history of this pair of grass chromosomes needs
further exploration. Elevated gene loss rates and elevated evolutionary rates of the
preserved genes in young strata may facilitate speciation in that the loss of
alternative copies of duplicated genes leads to reproductive isolation (Werth and
Windham 1991; Lynch and Force 2000; Scannell et al. 2006; see also Chap. 1, this
volume). The recently proposed inter-relationship between reproductive isolation
and autoimmune responses (Bomblies et al. 2007; Yin et al. 2008) draws attention
to the finding that orthologs R11 and S5 each contain *25 % of the NBS-LRR
resistance genes (Zhou et al. 2004; Paterson et al. 2009b). Second, a high level of
CO
RR
355
UN
354
Layout: T1 Standard SC
Chapter No.: 6
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 104/107
A. H. Paterson et al.
392
References
393
394
395
396
397
398
399
400
401
402
403
404
405
406
407
408
409
410
411
412
413
414
415
416
417
418
419
420
421
422
423
424
425
426
427
428
Birchler JA, Veitia RA (2007) The gene balance hypothesis: from classical genetics to modern
genomics. Plant Cell 19:395–402
Birchler JA, Riddle NC, Auger DL, Veitia RA (2005) Dosage balance in gene regulation:
biological implications. Trends Genet 21:219–226
Blanc G, Wolfe KH (2004) Functional divergence of duplicated genes formed by polyploidy
during Arabidopsis evolution. Plant Cell 16:1679–1691
Bomblies K, Lempe J, Epple P, Warthmann N, Lanz C, Dangl JL, Weigel D (2007) Autoimmune
response as a mechanism for a dobzhansky-muller-typeincompatibility syndrome in Plants.
PLOS Biology 5:1962-1972
Bowers JE, Arias MA, Asher R, Avise JA, Ball RT, Brewer GA, Buss RW, Chen AH, Edwards
TM, Estill JC, Exum HE, Goff VH, Herrick KL, Steele CLJ, Karunakaran S, Lafayette GK,
Lemke C, Marler BS, Masters SL, McMillan JM, Nelson LK, Newsome GA, Nwakanma CC,
Odeh RN, Phelps CA, Rarick EA, Rogers CJ, Ryan SP, Slaughter KA, Soderlund CA, Tang
HB, Wing RA, Paterson AH (2005) Comparative physical mapping links conservation of
microsynteny to chromosome structure and recombination in grasses. Proc Nat Acad Sci USA
102:13206–13211
Bremer G (1923) A cytological investigation of some species and species-hybrids of the genus
Saccharum. Genetica 5:273–326
Bremer G (1961) Problems in breeding and cytology of sugar cane. 4. Origin of increase of
chromosome number in species hybrids of Saccharum. Euphytica 10:325–342
Chapman BA, Bowers JE, Feltus FA, Paterson AH (2006) Buffering crucial functions by
paleologous duplicated genes may impart cyclicality to angiosperm genome duplication. Proc
Nat Acad Sci USA 103:2730–2735
Charlesworth B (2002) The evolution of chromosomal sex determination. Novartis Found Symp
244:207–219 (discussion 220–204, 253–207)
Chittenden LM, Schertz KF, Lin YR, Wing RA, Paterson AH (1994) A detailed Rflp map of
Sorghum-bicolor X S-propinquum, suitable for high-density mapping, suggests ancestral
duplication of sorghum chromosomes or chromosomal segments. Theor Appl Genet
87:925–933
Devos KM, Pittaway TS, Reynolds A, Gale MD (2000) Comparative mapping reveals a complex
relationship between the pearl millet genome and those of foxtail millet and rice. Theor Appl
Genetics 100:190–198
Feldman M, Levy AA (2005) Allopolyploidy—a shaping force in the evolution of wheat
genomes. Cytogenet Genome Res 109:250–258
Freeling M (2001) Grasses as a single genetic system: reassessment 2001. Plant Physiol
125:1191–1197
389
390
PR
OO
388
D
387
TE
385
386
EC
384
CO
RR
383
F
391
concerted evolution, associated stratification of chromosomal segments, and
extensive homoeologous gene loss are each characteristics of sex chromosomes in
organisms from divergent branches of the tree of life, including humans (Lahn and
Page 1999), chickens (Lawson Handley et al. 2006), fungi (Charlesworth 2002),
and plants (Ming and Moore 2007). Moreover, unexpectedly close proximity
between, and co-expansion of, NBS-LRR and several sex-determining gene
analogs is found, particularly on S5 (Wang et al. 2011). A hypothesis for further
study is whether genes on the various orthologs and paralogs of these chromosomes (or regions therein) could have some ‘functional coherence’ resembling that
of the human Y chromosome (Lahn and Page 1999).
382
UN
Editor Proof
104
Layout: T1 Standard SC
Chapter No.: 6
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 105/107
105
EC
TE
D
PR
OO
F
Goff SA, Ricke D, Lan TH, Presting G, Wang RL, Dunn M, Glazebrook J, Sessions A, Oeller P,
Varma H, Hadley D, Hutchinson D, Martin C, Katagiri F, Lange BM, Moughamer T, Xia Y,
Budworth P, Zhong JP, Miguel T, Paszkowski U, Zhang SP, Colbert M, Sun WL, Chen LL,
Cooper B, Park S, Wood TC, Mao L, Quail P, Wing R, Dean R, Yu YS, Zharkikh A, Shen R,
Sahasrabudhe S, Thomas A, Cannings R, Gutin A, Pruss D, Reid J, Tavtigian S, Mitchell J,
Eldredge G, Scholl T, Miller RM, Bhatnagar S, Adey N, Rubano T, Tusneem N, Robinson R,
Feldhaus J, Macalma T, Oliphant A, Briggs S (2002) A draft sequence of the rice genome
(Oryza sativa L. ssp japonica). Science 296:92–100
Haldane JBS (1933) The part played by recurrent mutation in evolution. Am Nat 67:5–19
Heaton EA, Dohleman FG, Long SP (2008) Meeting US biofuel goals with less land: the
potential of Miscanthus. Glob Change Biol 14:2000–2014
Hilu KW (2004) Phylogenetics and chromosomal evolution in the Poaceae (grasses). Aust J Bot
52:10
International Rice Genome Sequencing P (2005) The map-based sequence of the rice genome.
Nature 436:793–800
Jaillon O, Aury JM et al (2007) The grapevine genome sequence suggests ancestral
hexaploidization in major angiosperm phyla. Nature 449:463–467
Jannoo N, Grivet L, Chantret N, Garsmeur O, Glaszmann JC, Arruda P, D’Hont A (2007)
Orthologous comparison in a gene-rich region among grasses reveals stability in the
sugarcane polyploid genome. Plant J 50:574–585
Jeswiet J (1929) The development of selection and breeding of the sugarcane in Java. Int Soc
Sugar Cane Technol 3:44–57
Kim C, Tang H, Paterson AH (2009) Duplication and divergence of grass genomes: integrating
the chloridoids. Trop Plant Biol 2:51–62
Kishimoto N, Higo H, Abe K, Arai S, Saito A, Higo K (1994) Identification of the duplicated
segments in rice chromosomes 1 and 5 by linkage analysis of cDNA markers of known
functions. Theor Appl Genet 88:722–726
Lahn BT, Page DC (1999) Four evolutionary strata on the human X chromosome. Science (New
York) 286:964–967
Lawrence WJC (1931) The secondary association of chromosomes. Cytologia 2:352–384
Lawson Handley LJ, Hammond RL, Emaresi G, Reber A, Perrin N (2006) Low Y chromosome
variation in Saudi-Arabian hamadryas baboons (Papio hamadryas hamadryas). Heredity
96:298–303
Liu H, Sachidanandam R, Stein L (2001) Comparative genomics between rice and Arabidopsis
shows scant collinearity in gene order. Genome Res 11:2020–2026
Lohithaswa HC, Feltus FA, Singh HP, Bacon CD, Bailey CD, Paterson AH (2007) Leveraging
the rice genome sequence for comparative genomics in monocots. Theor Appl Genetics
115:237–243
Lynch M, Conery JS (2000) The evolutionary fate and consequences of duplicate genes. Science
290:1151–1155
Lynch M, Force AG (2000) The origin of interspecific genomic incompatibility via gene
duplication. Am Nat 156:590–605
Maere S, De Bodt S, Raes J, Casneuf T, Van Montagu M, Kuiper M, Van de Peer Y (2005)
Modeling gene and genome duplications in eukaryotes. Proc Nat Acad Sci USA
102:5454–5459
Mayer KFX, Martis M, Hedley PE, Simkova H, Liu H, Morris JA, Steuernagel B, Taudien S,
Roessner S, Gundlach H, Kubalakova M, Suchankova P, Murat F, Felder M, Nussbaumer T,
Graner A, Salse J, Endo T, Sakai H, Tanaka T, Itoh T, Sato K, Platzer M, Matsumoto T,
Scholz U, Dolezel J, Waugh R, Stein N (2011) Unlocking the barley genome by chromosomal
and comparative genomics. The Plant Cell 23:1249–1263
Ming R, Moore PH (2007) Genomics of sex chromosomes. Curr Opin Plant Biol 10:123–130
Ming R, Liu SC, Lin YR, da Silva J, Wilson W, Braga D, van Deynze A, Wenslaff TF, Wu KK,
Moore PH, Burnquist W, Sorrells ME, Irvine JE, Paterson AH (1998) Detailed alignment of
CO
RR
429
430
431
432
433
434
435
436
437
438
439
440
441
442
443
444
445
446
447
448
449
450
451
452
453
454
455
456
457
458
459
460
461
462
463
464
465
466
467
468
469
470
471
472
473
474
475
476
477
478
479
480
481
UN
Editor Proof
6 Ancient and Recent Polyploidy in Monocots
Layout: T1 Standard SC
Chapter No.: 6
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 106/107
EC
TE
D
PR
OO
F
Saccharum and sorghum chromosomes: comparative organization of closely related diploid
and polyploid genomes. Genetics 150:1663–1682
Ming R, Liu SC, Moore PH, Irvine JE, Paterson AH (2001) QTL analysis in a complex
autopolyploid: genetic control of sugar content in sugarcane. Genome Res 11:2075–2084
Ming R, Del Monte TA, Hernandez E, Moore PH, Irvine JE, Paterson AH (2002a) Comparative
analysis of QTLs affecting plant height and flowering among closely-related diploid and
polyploid genomes. Genome 45:794–803
Ming R, Wang YW, Draye X, Moore PH, Irvine JE, Paterson AH (2002b) Molecular dissection
of complex traits in autopolyploids: mapping QTLs affecting sugar yield and related traits in
sugarcane. Theor Appl Genet 105:332–345
Murat F, Xu JH, Tannier E, Abrouk M, Guilhot N, Pont C, Messing J, Salse J (2010) Ancestral
grass karyotype reconstruction unravels new mechanisms of genome shuffling as a source of
plant evolution. Genome Res 20:1545–1557
Nagamura Y, Inoue T, Antonio B, Shimano T, Kajiya H, Shomura A, Lin S, Kuboki Y,
Harushima Y, Kurata N, Minobe Y, Yano M, Sasaki T (1995) Conservation of duplicated
segments between rice chromosomes 11 and 12. Breed Sci 45:373–376
Paterson AH (2008) Paleopolyploidy and its impact on the structure and function of modern plant
genomes. Genome Dyn 4:1–12
Paterson AH, Lan TH, Reischmann KP, Chang C, Lin YR, Liu SC, Burow MD, Kowalski SP,
Katsar CS, DelMonte TA, Feldmann KA, Schertz KF, Wendel JF (1996) Toward a unified
genetic map of higher plants, transcending the monocot-dicot divergence. Nat Genet
14:380–382
Paterson A, Bowers J, Peterson D, Estill J, Chapman B (2003) Structure and evolution of cereal
genomes. Curr Opin Genet Dev 13:644–650
Paterson AH, Bowers JE, Chapman BA (2004) Ancient polyploidization predating divergence of
the cereals, and its consequences for comparative genomics. Proc Nat Acad Sci USA
101:9903–9908
Paterson AH, Chapman BA, Kissinger J, Bowers JE, Feltus FA, Estill J, Marler BS (2006)
Convergent retention or loss of gene/domain families following independent whole-genome
duplication events in Arabidopsis, Oryza, Saccharomyces, and Tetraodon. Trends Genet
22:597–602
Paterson AH, Bowers JE, Feltus FA, Tang H, Lin L, Wang X (2009a) Comparative genomics of
grasses promises a bountiful harvest. Plant Physiol 149:125–131
Paterson AH, Bowers JE, Bruggmann R, Dubchak I, Grimwood J, Gundlach H, Haberer G,
Hellsten U, Mitros T, Poliakov A, Schmutz J, Spannagl M, Tang H, Wang X, Wicker T,
Bharti AK, Chapman J, Feltus FA, Gowik U, Grigoriev IV, Lyons E, Maher CA, Martis M,
Narechania A, Otillar RP, Penning BW, Salamov AA, Wang Y, Zhang L, Carpita NC,
Freeling M, Gingle AR, Hash CT, Keller B, Klein P, Kresovich S, McCann MC, Ming R,
Peterson DG, Mehboob ur R, Ware D, Westhoff P, Mayer KF, Messing J, Rokhsar DS
(2009b) The Sorghum bicolor genome and the diversification of grasses. Nature 457:551–556
Raven PH, Evert RF, Eichhorn SE (2005) Biology of plants, 7th edn. W. H. Freeman, New York
Salse J, Abrouk M, Bolot S, Guilhot N, Courcelle E, Faraut T, Waugh R, Close TJ, Messing J,
Feuillet C (2009) Reconstruction of monocotelydoneous protochromosomes reveals faster
evolution in plants than in animals. Proceedings of the National academy of sciences of the
United States of America 106:14908–14913
Salse J, Bolot S, Throude M, Jouffe V, Piegu B, Quraishi UM, Calcagno T, Cooke R, Delseny M,
Feuillet C (2008) Identification and characterization of shared duplications between rice and
wheat provide new insight into grass genome evolution. Plant Cell 20:11–24
Scannell DR, Byrne KP, Gordon JL, Wong S, Wolfe KH (2006) Multiple rounds of speciation
associated with reciprocal gene loss in polyploid yeasts. Nature 440:341–345
Schnable PS, Ware D, Fulton RS, Stein JC, Wei FS, Pasternak S, Liang CZ, Zhang JW, Fulton L,
Graves TA, Minx P, Reily AD, Courtney L, Kruchowski SS, Tomlinson C, Strong C,
Delehaunty K, Fronick C, Courtney B, Rock SM, Belter E, Du FY, Kim K, Abbott RM,
Cotton M, Levy A, Marchetto P, Ochoa K, Jackson SM, Gillam B, Chen WZ, Yan L,
CO
RR
482
483
484
485
486
487
488
489
490
491
492
493
494
495
496
497
498
499
500
501
502
503
504
505
506
507
508
509
510
511
512
513
514
515
516
517
518
519
520
521
522
523
524
525
526
527
528
529
530
531
532
533
534
535
A. H. Paterson et al.
UN
Editor Proof
106
Layout: T1 Standard SC
Chapter No.: 6
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 107/107
107
EC
TE
D
PR
OO
F
Higginbotham J, Cardenas M, Waligorski J, Applebaum E, Phelps L, Falcone J, Kanchi K,
Thane T, Scimone A, Thane N, Henke J, Wang T, Ruppert J, Shah N, Rotter K, Hodges J,
Ingenthron E, Cordes M, Kohlberg S, Sgro J, Delgado B, Mead K, Chinwalla A, Leonard S,
Crouse K, Collura K, Kudrna D, Currie J, He RF, Angelova A, Rajasekar S, Mueller T,
Lomeli R, Scara G, Ko A, Delaney K, Wissotski M, Lopez G, Campos D, Braidotti M, Ashley
E, Golser W, Kim H, Lee S, Lin JK, Dujmic Z, Kim W, Talag J, Zuccolo A, Fan C, Sebastian
A, Kramer M, Spiegel L, Nascimento L, Zutavern T, Miller B, Ambroise C, Muller S,
Spooner W, Narechania A, Ren LY, Wei S, Kumari S, Faga B, Levy MJ, McMahan L, Van
Buren P, Vaughn MW, Ying K, Yeh CT, Emrich SJ, Jia Y, Kalyanaraman A, Hsia AP,
Barbazuk WB, Baucom RS, Brutnell TP, Carpita NC, Chaparro C, Chia JM, Deragon JM,
Estill JC, Fu Y, Jeddeloh JA, Han YJ, Lee H, Li PH, Lisch DR, Liu SZ, Liu ZJ, Nagel DH,
McCann MC, SanMiguel P, Myers AM, Nettleton D, Nguyen J, Penning BW, Ponnala L,
Schneider KL, Schwartz DC, Sharma A, Soderlund C, Springer NM, Sun Q, Wang H,
Waterman M, Westerman R, Wolfgruber TK, Yang LX, Yu Y, Zhang LF, Zhou SG, Zhu Q,
Bennetzen JL, Dawe RK, Jiang JM, Jiang N, Presting GG, Wessler SR, Aluru S, Martienssen
RA, Clifton SW, McCombie WR, Wing RA, Wilson RK (2009) The B73 maize genome:
complexity, diversity, and dynamics. Science 326:1112–1115
Schnable JC, Springer NM, Freeling M (2011) Differentiation of the maize subgenomes by
genome dominance and both ancient and ongoing gene loss. Proc Nat Acad Sci USA
108:4069–4074
Seoighe C, Gehring C (2004) Genome duplication led to highly selective expansion of the
Arabidopsis thaliana proteome. Trends Genet 20:461–464
Singh NK, Dalal V, Batra K, Singh BK, Chitra G, Singh A, Ghazi IA, Yadav M, Pandit A, Dixit
R, Singh PK, Singh H, Koundal KR, Gaikwad K, Mohapatra T, Sharma TR (2007) Singlecopy genes define a conserved order between rice and wheat for understanding differences
caused by duplication, deletion, and transposition of genes. Funct Integr Genomics 7:17–35
Soderstrom TR, Hilu KW, Campbell CS, Barkworth MA (1987) Grass systematics and evolution.
Smithsonian Institution Press, Washington
Soltis DE, Smith S, Cellinese N, Refulio-Rodriquez NF, Olmstead R, Crawley S, Black C, Diouf
D, Hilu KW, Latvis M, Wurdack K, Xi Z, Davis C, Donoghue M, Soltis PS (2011) Inferring
angiosperm phylogeny: a 17-gene analysis. Am J Bot 98:704–730
Spangler R (2003) Taxonomy of Sarga, Sorghum, and Vacoparis (Poaceae: Andropogoneae).
Aust Syst Bot 16:279–299
Spangler R, Zaitchik B, Russo E, Kellogg E (1999) Andropogoneae evolution and generic limits
in Sorghum (Poaceae) using ndhF sequences. Syst Bot 24:267–281
Tang H, Wang X, Bowers JE, Ming R, Alam M, Paterson AH (2008) Unraveling ancient
hexaploidy through multiply-aligned angiosperm gene maps. Genome Res 18:1944–1954
Tang H, Bowers JE, Wang X, Ming R, Alam M, Paterson AH (2008) Synteny and colinearity in
plant genomes. Science 320:486–488
Tang HB, Bowers JE, Wang XY, Paterson AH (2010) Angiosperm genome comparisons reveal
early polyploidy in the monocot lineage. Proc Nat Acad Sci USA 107:472–477
The International Brachypodium Initiative (2010) Genome sequencing and analysis of the model
grass Brachypodium distachyon. Nature 463:763–768
Thomas BC, Pedersen B, Freeling M (2006) Following tetraploidy in an Arabidopsis ancestor,
genes were removed preferentially from one homeolog leaving clusters enriched in dosesensitive genes. Genome Res 16:934–946
Van de Peer Y (2004) Computational approaches to unveiling ancient genome duplications. Nat
Rev Genet 5:752–763
Vandepoele K, Simillion C, Van de Peer Y (2003) Evidence that rice and other cereals are ancient
aneuploids. Plant Cell 15:2192–2202
Veitia RA, Bottani S, Birchler JA (2008) Cellular reactions to gene dosage imbalance: genomic,
transcriptomic and proteomic effects. Trends Genet 24:390–397
Wang X, Shi X, Hao B, Ge S, Luo J (2005) Duplication and DNA segmental loss in the rice
genome: implications for diploidization. New Phytol 165:937–946
CO
RR
536
537
538
539
540
541
542
543
544
545
546
547
548
549
550
551
552
553
554
555
556
557
558
559
560
561
562
563
564
565
566
567
568
569
570
571
572
573
574
575
576
577
578
579
580
581
582
583
584
585
586
587
588
589
UN
Editor Proof
6 Ancient and Recent Polyploidy in Monocots
Layout: T1 Standard SC
Chapter No.: 6
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 108/107
EC
TE
D
PR
OO
F
Wang XY, Shi XL, Li Z, Zhu QH, Kong L, Tang W, Ge S, Luo JC (2006) Statistical inference of
chromosomal homology based on gene colinearity and applications to arabidopsis and rice.
BMC Bioinformatics 7:447
Wang X, Tang H, Bowers JE, Feltus FA, Paterson AH (2007) Extensive concerted evolution of
rice paralogs and the road to regaining independence. Genetics 177:1753–1763
Wang X, Tang H, Paterson AH (2011) Seventy million years of concerted evolution of a
homoeologous chromosome pair, in parallel, in major Poaceae lineages. Plant Cell 23:27–37
Werth CR, Windham MD (1991) A model for divergent, allopatric speciation of polyploid
pteridophytes resulting from silencing of duplicate-gene expression. Am Nat 137:515–526
Wicker T, Mayer KFX, Gundlach H, Martis M, Steuernagel B, Scholz U, Simkova H,
Kubalakova M, Choulet F, Taudien S, Platzer M, Feuillet C, Fahima T, Budak H, Dolezel J,
Keller B, Stein N (2011) Frequent gene movement and pseudogene evolution is common to
the large and complex genomes of wheat, barley, and their relatives. The Plant Cell
23:1706–1718
Woodhouse MR, Schnable JC, Pedersen BS, Lyons E, Lisch D, Subramaniam S, Freeling M
(2010) Following tetraploidy in maize, a short deletion mechanism removed genes
preferentially from one of the two homologs. PLoS Biol 8:e1000409
Xiong Z, Gaeta RT, Pires JC (2011) Homoeologous shuffling and chromosome compensation
maintain genome balance in resynthesized allopolyploid Brassica napus. Proc Nat Acad Sci
USA 108:7908–7913
Yin T, Difazio SP, Gunter LE, Zhang X, Sewell MM, Woolbright SA, Allan GJ, Kelleher CT,
Douglas CJ, Wang M, Tuskan GA (2008) Genome structure and emerging evidence of an
incipient sex chromosome in Populus. Genome Res 18:422–430
Yu J, Wang J, Lin W, Li SG, Li H, Zhou J, Ni PX, Dong W, Hu SN, Zeng CQ, Zhang JG, Zhang
Y, Li RQ, Xu ZY, Li ST, Li XR, Zheng HK, Cong LJ, Lin L, Yin JN, Geng JN, Li GY, Shi JP,
Liu J, Lv H, Li J, Deng YJ, Ran LH, Shi XL, Wang XY, Wu QF, Li CF, Ren XY, Wang JQ,
Wang XL, Li DW, Liu DY, Zhang XW, Ji ZD, Zhao WM, Sun YQ, Zhang ZP, Bao JY, Han
YJ, Dong LL, Ji J, Chen P, Wu SM, Liu JS, Xiao Y, Bu DB, Tan JL, Yang L, Ye C, Zhang JF,
Xu JY, Zhou Y, Yu YP, Zhang B, Zhuang SL, Wei HB, Liu B, Lei M, Yu H, Li YZ, Xu H,
Wei SL, He XM, Fang LJ, Zhang ZJ, Zhang YZ, Huang XG, Su ZX, Tong W, Li JH, Tong
ZZ, Li SL, Ye J, Wang LS, Fang L, Lei TT, Chen C, Chen H, Xu Z, Li HH, Huang HY, Zhang
F, Xu HY, Li N, Zhao CF, Dong LJ, Huang YQ, Li L, Xi Y, Qi QH, Li WJ, Hu W, Zhang YL,
Tian XJ, Jiao YZ, Liang XH, Jin JA, Gao L, Zheng WM, Hao BL, Liu SQ, Wang W, Yuan
LP, Cao ML, McDermott J, Samudrala R, Wong GKS, Yang HM (2005) The genomes of
Oryza sativa: a history of duplications. Plos Biology 3:266–281
Zhou JH, Wang JL, Xu JC, Lei CL, Ling ZZ (2004) Identification and mapping of a rice blast
resistance gene Pi-g(t) in the cultivar Guangchangzhan. Plant Pathol. 53:191–196
Zhang Y, Xu GH, Guo XY, Fan LJ (2005) Two ancient rounds of polyploidy in rice genome.
J Zhejiang Univ Sci B 6:87–90
CO
RR
590
591
592
593
594
595
596
597
598
599
600
601
602
603
604
605
606
607
608
609
610
611
612
613
614
615
616
617
618
619
620
621
622
623
624
625
626
627
628
A. H. Paterson et al.
UN
Editor Proof
108
Metadata of the chapter that will be visualized in
SpringerLink
Book Title
Polyploidy and Genome Evolution
Series Title
Chapter Title
Genomic Plasticity in Polyploid Wheat
Copyright Year
2012
Copyright HolderName
Springer-Verlag Berlin Heidelberg
Corresponding Author
Family Name
Kashkush
Particle
Given Name
Khalil
Suffix
Author
Division
Department of Life Sciences
Organization
Ben-Gurion University
Address
84105, Beer-Sheva, Israel
Email
kashkush@bgu.ac.il
Family Name
Feldman
Particle
Given Name
Moshe
Suffix
Division
Plant Sciences Department
Organization
The Weizmann Institute of Science
Address
76100, Rehovot, Israel
Email
Author
Family Name
Levy
Particle
Given Name
Avraham
Suffix
Division
Plant Sciences Department
Organization
The Weizmann Institute of Science
Address
76100, Rehovot, Israel
Email
Author
Family Name
Chalhoub
Particle
Given Name
Boulos
Suffix
Division
UMR INRA 1165—CNRS 8114—UEVE, Organization and Evolution of
Plant Genomes (OEPG)
Organization
Unité de Recherche En Génomique Végétale (URGV)
Address
2 Rue Gaston Crémieux, 91057, Evry Cedex, France
Email
Abstract
The importance of hybridization and polyploidization in wheat speciation has been recognized for close to a
century (Sakamura 1918; Kihara 1919, 1924, 1954; Percival 1921; Sax 1927). Following these pioneering
works, it quickly became apparent that polyploid wheats are not the sum of their constituent genomes. This
is not unexpected because the nascent hybrids/polyploids are equipped with a complex set of regulatory
elements and of copy number variation that originate from two or more divergent genomes and that generate
novel types of interactions and dosage effects. Moreover, they have to adjust at the cytological level, at the
level of gene expression, and at the protein level. They also have to maintain genome stability through the
regulation of meiotic pairing and recombination, the orchestration of cell division, and the silencing of
transposons. The recent studies described here provide an impressive account with regard to the extent and
the rapid time course at which a new genetic variant was established upon hybridization and polyploidization.
We describe here the current knowledge on the changes that occurred in the wheat genome upon
allopolyploidization, starting from the early evolutionary and cytological studies to the recent genomic
analyses.
Book ISBN: 978-3-642-31441-4
Page: 109/134
Chapter 7
2
Genomic Plasticity in Polyploid Wheat
3
5
4
Moshe Feldman, Avraham Levy, Boulos Chalhoub
and Khalil Kashkush
6
Abstract
PR
OO
1
7
13
14
15
16
17
18
19
20
21
22
23
D
12
TE
10
11
The importance of hybridization and polyploidization in wheat speciation has been
recognized for close to a century (Sakamura 1918; Kihara 1919, 1924, 1954;
Percival 1921; Sax 1927). Following these pioneering works, it quickly became
apparent that polyploid wheats are not the sum of their constituent genomes. This
is not unexpected because the nascent hybrids/polyploids are equipped with a
complex set of regulatory elements and of copy number variation that originate
from two or more divergent genomes and that generate novel types of interactions
and dosage effects. Moreover, they have to adjust at the cytological level, at the
level of gene expression, and at the protein level. They also have to maintain
genome stability through the regulation of meiotic pairing and recombination, the
orchestration of cell division, and the silencing of transposons. The recent studies
described here provide an impressive account with regard to the extent and the
rapid time course at which a new genetic variant was established upon hybridization and polyploidization. We describe here the current knowledge on the
changes that occurred in the wheat genome upon allopolyploidization, starting
from the early evolutionary and cytological studies to the recent genomic analyses.
EC
9
CO
RR
8
F
Book ID: 272454_1_En
Date: 16-8-2012
M. Feldman A. Levy
Plant Sciences Department, The Weizmann Institute of Science,
76100, Rehovot, Israel
B. Chalhoub
UMR INRA 1165—CNRS 8114—UEVE,
Organization and Evolution of Plant Genomes (OEPG),
Unité de Recherche En Génomique Végétale (URGV),
2 Rue Gaston Crémieux, 91057, Evry Cedex, France
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: 7
K. Kashkush (&)
Department of Life Sciences, Ben-Gurion University, 84105,
Beer-Sheva, Israel
e-mail: kashkush@bgu.ac.il
P. S. Soltis and D. E. Soltis (eds.), Polyploidy and Genome Evolution,
DOI: 10.1007/978-3-642-31442-1_7, Springer-Verlag Berlin Heidelberg 2012
109
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 110/134
110
M. Feldman et al.
Diploids
(2n = 2x = 14)
Tetraploids
(2n = 4x = 28)
Hexaploids
(2n = 6x = 42)
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
D
28
TE
27
Spelt (hulled)
wheats
Naked (free threshing)
wheats
The discovery of wild emmer wheat, the progenitor of most domesticated wheats
(Aaronsohn and Schweinfurth 1906; Aaronsohn 1910), made it possible for Schulz
(1913) to assemble the first natural classification of the wheats. He divided the
genus Triticum into three major groups: einkorn, emmer, and dinkel. This classification was supported by the pioneering cytological study of Sakamura (1918),
who was the first to determine the correct chromosome number of the wheats.
Sakamura discovered that Schulz’s three groups of wheats differ in their chromosome number: the einkorns are diploids (2n = 14), the emmers are tetraploids
(2n = 28), and the dinkels are hexaploids (2n = 42) (Table 7.1). It then became
obvious that the species of Triticum represent a polyploid series with diploid,
tetraploid, and hexaploid species.
Since then, the species of the wheat group (Triticum and its closely related
genus Aegilops) have been subjected to extensive taxonomic, cytogenetic, genetic,
biochemical, molecular, and evolutionary study by numerous scientists (see
reviews of Kihara 1954; Mac Key 1966; Morris and Sears 1967; Kimber and Sears
1987; Feldman et al. 1995; Feldman 2001; Gupta et al. 2005; Dvorak 2009). Due
to these extensive studies the allopolyploid species of the wheat group became a
classic example of evolution through allopolyploidy. The cytogenetic studies of
Kihara (1919, 1924, 1954), Percival (1921), Sax (1927), and others on chromosome pairing in hybrids among species of different ploidy levels showed that all
the polyploid species of the group form an allopolyploid series based on x = 7.
Each allopolyploid species was identified as a product of hybridization followed
by chromosome doubling (Fig. 7.1). Von Tschermak and Bleier (1926) were the
first to identify a spontaneous chromosome doubling in the cross of wild emmer
(T. turgidum ssp. dicoccoides) with Aegilops geniculata, thus demonstrating the
possibility of species formation via allopolyploidy in the wheat group. Subsequent
studies showed that the frequency of unreduced gametes in intergeneric hybrids of
wheat could be in some hybrids as high as 50 % (Kihara and Lilienfeld 1949).
Therefore, one might assume that there is a high potential for the frequent and
EC
26
Domesticated wheats
Einkorn (one-grained T. aegilopoides T. monococcum None
wheat)
—[
Emmer (two-grained T. dicoccoides T. dicoccum — T. durum
wheat)
—[
[
T. turgidum
T. polonicum
Dinkel
None
T. spelta — [ T. compactum
T. vulgare
CO
RR
25
Wild
progenitors
7.1 The Wheat Group: Natural and Synthetic Polyploids
UN
24
Series
F
Ploidy level
PR
OO
Editor Proof
Table 7.1 Superimposition of Sakamura’s (1918) finding of the right chromosome number on
Schulz’s (1913) natural classification of the wheats
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 111/134
111
PR
OO
F
Editor Proof
7 Genomic Plasticity in Polyploid Wheat
59
60
61
62
63
64
65
66
67
68
69
70
71
72
73
74
75
76
77
TE
58
EC
56
57
recurrent formation of interspecific or intergeneric hybrids and allopolyploids in
the wheat group.
The discovery by Blakeslee (1937) that colchicine can induce chromosome
doubling opened new possibilities for the study of wheat evolution through allopolyploidization. It also provided an easy method to synthesize different wheat
allopolyploids, some of which have similar genomes to natural allopolyploids and
others having new genomic combinations (see examples in Ozkan et al. 2001).
Synthetic allopolyploids, either induced or occurring spontaneously, offer excellent tools to mimic the evolutionary speciation events that occurred in nature and
to test in a controlled manner; the new features of the hybrid/polyploid genome
compared to those of its parents.
Since the discovery that the polyploid species of wheat comprise an allopolyploid series, attempts have been made to identify the diploid donors of the different
genomes to the allopolyploids of the wheat group. Most of these attempts used the
cytogenetic approach of genome analysis, developed by Kihara (1919, 1924) and
based on the concept of genome stability, assuming that the genomes of the
allopolyploid species remain similar to those of their diploid parents. However, the
accumulating cytogenetic and molecular evidence has indicated that this is not the
case; while one genome remains relatively unchanged, the second genome(s) of
the allopolyploids have changed considerably from those of their parental diploids.
These genomes were termed modified genomes by Kihara (1954) and other wheat
cytogeneticists. Every allopolyploid species of Aegilops and Triticum contains an
unchanged genome side-by-side with a modified one whose diploid origin has
been intricate and difficult to trace (Zohary and Feldman 1962).
CO
RR
55
UN
54
D
Fig. 7.1 Evolutionary history of allotetraploid and allohexaploid wheat. Diploid wheats
(2n = 2x = 14) from the Triticum-Aegilops group diverged *4 million years ago (MYA) from
a common progenitor (Huang et al. 2002). Interspecific hybridization between the diploid
T. urartu (genome AA) as male and the donor of the B genome (an unknown species similar to
Ae. speltoides) as female, followed by chromosome doubling, gave rise, *0.5 MYA, to wild
allotetraploid wheat, Triticum turgidum ssp. dicoccoides (2n = 4x = 28, genome BBAA), an
allotetraploid considered as the direct progenitor of durum wheat. Domestication of allotetraploid
wheat took place *10,500 YA and was rapidly followed (*9,500 YA) by a second round of
intergeneric hybridization and chromosome doubling between domesticated allotetraploid wheat
and the donor of the D genome, Ae. tauschii (2n = 2x = 14, genome DD), giving rise to bread
wheat, an allohexaploid with 2n = 6x = 42 chromosomes (genome BBAADD). Adapted from
Levy and Feldman (2004)
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 112/134
M. Feldman et al.
95
7.2 Cytological Diploidization
87
88
89
90
91
92
93
96
97
98
99
100
101
102
103
104
105
106
107
108
109
110
111
112
113
114
115
116
117
118
PR
OO
85
86
D
83
84
TE
81
82
Because of the close relationship of the progenitors and the similarity of the two
different genomes of allotetraploid wheat and of the three different genomes of
allohexaploid wheat (Morris and Sears 1967), the successful establishment of the
polyploid species in nature required the acquisition during the allopolyploidization
process of molecular and genetic systems that would prevent intergenomic pairing
and recombination. By restricting pairing to fully homologous chromosomes
(intragenomic pairing), the cytological-diploidizing systems ensure exclusive
bivalent pairing at meiosis and, consequently, regular segregation of genetic
material, complete fertility, and genetic stability. Cytological diploidization has
been brought about in allopolyploid wheat by two independent systems that
complement each other: One system is based on genetic control of pairing, and the
second is based on physical divergence of chromosomes.
Historically, the first mechanism of cytological diploidization that received
much attention was a genetic system involved in sustaining the exclusive bivalent
pairing in allopolyploids. It consisted of the activity of the genetic loci Ph1 on
chromosome arm 5BL and Ph2 on chromosome arm 3DS (Sears 1976). These loci
suppress pairing of homoeologous chromosomes while allowing homologs to pair
regularly. The mechanism controlling the Ph1 mode of action is still unclear.
Mapping data suggests the involvement of a complex locus that affects cell cycle
progression through the regulation of Cyclin-dependent kinases (Griffiths et al.
2006); however, direct evidence through complementation studies is still missing.
It was suggested by several authors that Ph-like genes exist in diploid species of
the wheat group (Okamoto and Inomata 1974; Avivi 1976; Waines 1976; Maan
EC
80
CO
RR
79
F
94
Nevertheless, genome analysis studies revealed that allotetraploid wheat
(genome BBAA) originated from hybridization events involving two diploid
progenitors classified in the genera Aegilops and Triticum. Genome B, which is a
modified genome, was derived from Ae. speltoides and underwent changes on the
polyploid level or, more likely, from a closely related species to Ae. speltoides
which is extinct or extant (Feldman et al. 1995). Genome A, which has been
modified relatively little, was derived from T. urartu (Chapman et al. 1976;
Dvorak 1976). Allohexaploid wheat (genome BBAADD) originated from
hybridization between allotetraploid wheat and Ae. tauschii, the donor of the
D genome (Kihara 1944; McFadden and Sears 1944, 1946). The evolution of the
wheats is presented in Fig. 7.1 (for details, see Feldman et al. 1995; Feldman
2001). Modern classification for the Triticum group (Van Slageren 1994;
Table 7.2) recognizes two diploid species, T. monococcum L. and T. urartu Tum.
ex Gand., two tetraploid species, T. turgidum L. and T. timopheevii (Zhuk.) Zhuk.,
and two hexaploid species, T. aestivum L. and T. zhukovskyi Men. & Er. The
economically important wheats are T. aestivum (bread wheat, comprising 95 % of
the global wheat production) and T. turgidum (macaroni wheat).
78
UN
Editor Proof
112
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 113/134
7 Genomic Plasticity in Polyploid Wheat
113
AmAm
AA
Tetraploids (2n = 4x = 28) GGAA
Triticum urartu
Triticum timopheevii
ssp. armeniacum
ssp. timopheevii
Triticum turgidum
ssp. dicoccoides
ssp. dicoccon
ssp paleocochicum
ssp. parvicoccumb
ssp. durum
ssp. turgidum
ssp. polonicum
ssp. turanicum
ssp. carthlicum
Triticum zhukovskyi
D
BBAA
Triticum monococcum
ssp. aegilopoides
ssp. monococcum
EC
TE
Hexaploids (2n = 6x = 42) GGAA
AmAm
BBAADD Triticum aestivum
ssp. spelta
ssp. macha
ssp. aestivum
ssp. compactum
ssp. sphaerococcum
a
Wild einkorn
Domesticated einkorn or
small spelt
None (wild form)
F
Diploids (2n = 2x = 14)
PR
OO
Editor Proof
Table 7.2 The nomenclature of the commercially cultivated wheats and their immediate wild
relatives (after van Slageren, 1994)a
Ploidy level
Genome
Species and subspecies Common name
Wild timopheevii
Domesticated timopheevii
Wild emmer
Domesticated emmer
Georgian wheat
None
Macaroni or hard wheat
Rivet, cone or pollard wheat
Polish wheat
Khorassan wheat
Persian wheat
None
Dinkel or large spelt
None
Common or bread wheat
Club wheat
Indian dwarf or short wheat
120
121
122
123
124
125
126
127
128
129
130
131
1977) and they became more effective at the polyploid level as a result of
duplication. This dosage-dependent effect might have been selected to improve the
fertility of the allopolyploid. This genetic system superimposes itself on, takes
advantage of, and thereby reinforces the system of physical homoeologous differentiation already in existence and described below. The genetic system is very
effective in suppressing homoeologous pairing in interspecific and intergeneric F1
hybrids. However, its suppressive effect on homoeologous pairing in allopolyploid
wheat might not be essential since in plants deficient for Ph1 there is relatively
very little such pairing (Sears 1976). Interestingly, and in accord with the above,
gene(s) like Ph were not found in all the allopolyploid species of the closely
related genus Aegilops. In spite of this, these species, relying solely on the
structural homoeologous differentiation, exhibit exclusive bivalent pairing of fully
homologous chromosomes, i.e., exclusive intragenomic pairing. It might be that
UN
119
CO
RR
Taxa derived from a single spontaneous or induced mutation and are not commercially cultivated, such as diploid T. sinskajae Filat and Kurk., tetraploid T. militinae Zhuk. and Migush.,
and hexaploid T. vavilovii (Tum.) Jakubz., are not included
b
Extinct, described by Kislev 1980
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 114/134
114
M. Feldman et al.
Occur during or immediately after allopolyploidization
F
Genetic and epigenetic changes
Species-specific
Lead to cytological diploidization
Occur during the life of the
allopolyploid species
Mostly genetic changes
Population- or biotype-specific
Promote genetic diversity,
flexibility, and adaptability
Improve harmonic functioning of the divergent genomes
Stabilize the nascent allopolyploid and facilitate its
establishment as a new species in nature
PR
OO
Editor Proof
Table 7.3 Types and characteristics of genomic changes in allopolyploid wheat
Revolutionary changes (triggered by allopolyploidization) Evolutionary changes (Facilitated
by allopolyploidy)
143
7.3 Genomic Structural Changes
137
138
139
140
141
144
145
146
147
148
149
150
151
152
153
154
155
156
TE
136
EC
135
Studies with synthetic polyploids as well as genome sequencing data indicate that
a broad range of DNA rearrangements occurred during, or soon after, hybridization and polyploidization. What triggers these changes is a fascinating and still
open question, but what is clear is that these changes are extensive, including DNA
loss, transposon activation, gene duplication, and pseudogenization, and are relatively rapid. Feldman and Levy (2005) have distinguished between the revolutionary changes that occur rapidly and evolutionary changes that take place
throughout the evolution of the allopolyploid (Tables 7.3 and 7.4). Note that
evolutionary changes might also occur in an accelerated manner, thanks to the
buffering of mutations in the polyploid background (Mac Key 1954, 1958; Sears
1972; Thompson et al. 2006), leading to rapid neo- or subfunctionalization of
genes and to a process of diploidization and of divergence from the diploid progenitor genomes.
CO
RR
134
UN
133
D
142
stringent selection for fertility under domestication has favored the development of
two systems to ensure suppression of multivalent formation and to promote
bivalent pairing. Moreover, the cytological diploidization of the allopolyploid
wheats that leads to disomic inheritance prevents independent segregation of genes
from the different genomes. This mode of inheritance leads to permanent maintenance of favorable intergenomic genetic interactions. It enables fixation of heterotic interaction between genomes and sustained division of tasks (genome
asymmetry) between genomes. A series of DNA rearrangements in the allopolyploid further contributes to the physical divergence between the homoeologous
chromosomes and to the strengthening of the disomic genetic system. These
changes are described below.
132
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 115/134
7 Genomic Plasticity in Polyploid Wheat
115
Structural
• Elimination of low-copy DNA sequences
Chromatin remodeling
Chromatin modifications
Heterochromatinization
DNA methylation
Small RNAs activation or
repression
Methylation (leading to
silencing)
Demethylation (leading to
gene activation)
Release transposons from
silencing
Silencing transposons
PR
OO
• Elimination, reduction, or amplification of highcopy sequences
• Inter-genomic invasion of DNA sequences
• Elimination of rRNA and 5S RNA genes
Functional • Gene loss
•
•
•
•
•
F
Editor Proof
Table 7.4 Revolutionary changes induced by allopolyploidization in wheat
Level
Genetic
Epigenetic
•
161
162
163
164
165
166
167
168
169
170
171
172
173
174
175
176
177
178
179
180
181
TE
160
Allopolyploidization causes immediate nonrandom elimination of specific noncoding, lowcopy, and high-copy DNA sequences. These sequences are present in
all the diploid species of Aegilops and Triticum but occur in only one pair of
chromosomes (chromosome-specific sequences) or in several chromosome pairs of
one genome (genome-specific sequences) at the polyploid level (Fig. 7.2)
(Feldman et al. 1997; Liu et al. 1998a; Liu et al. 1998b; Ozkan et al. 2001; Shaked
et al. 2001; Han et al. 2003; Salina et al. 2004; Han et al. 2005). The extent of
DNA elimination was estimated by the determination of the nuclear DNA amount
in natural allopolyploids and in their diploid progenitors as well as in newly
synthesized allopolyploids and in their parental plants (Ozkan et al. 2003; Eilam
et al. 2008; Eilam et al. 2010). Natural wheat allopolyploids contain 2–10 % less
DNA than the sum of their diploid parents, and synthetic allopolyploids exhibit a
similar loss, indicating that DNA elimination occurs soon after allopolyploidization (Eilam et al. 2008; Eilam et al. 2010). Also, from the very little variation in
DNA amount that exists at the intraspecific level, it was concluded that the
reduction of DNA content occurred immediately after the formation of the
polyploids, and that after this there was almost no change in DNA amount during
the life of the allopolyploid species. In triticale (an allopolyploid between wheat
and rye, Secale cereale), Boyko et al. (1984, 1988) found that there was a great
reduction in DNA content in the course of triticale formation with about 9 % for
octoploid and 28–30 % for hexaploid triticale. The different genomes were not
affected equally in triticale; wheat genomic sequences were relatively conserved,
whereas rye genomic sequences were predominantly involved in a very high
level of variation and elimination (Ma et al. 2004; Ma and Gustafson 2005, 2006).
EC
159
CO
RR
158
7.3.1 Revolutionary changes
UN
157
D
• Rewiring of gene expression through novel inter- •
genomic interactions
• New dosage response (positive, negative, dosage •
compensation)
• Gene suppression or activation
•
• Transcriptional activation of transposons (that may
affect nearby genes in cis)
• New transpositions of transposons
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 116/134
M. Feldman et al.
D
PR
OO
F
Editor Proof
116
184
185
186
187
188
189
190
191
192
193
194
195
196
Also in hexaploid wheat the genomes were not affected equally; genome D
underwent considerable reduction in DNA amount whereas the wheat A and B
genomes did not shrink in size (Eilam et al. 2008; Eilam et al. 2010). DNA
elimination seems to be nonrandom at the intrachromosomal level as well: Liu
et al. (1997) found in allohexaploid wheat that the chromosome-specific sequences
of chromosome arm 5BL are not distributed at random along this chromosome arm
but cluster in terminal (subtelomeric), subterminal, and interstitial regions of this
arm, making these regions extremely chromosome-(homologous-)specific. Hence,
it was tempting to suggest that these homologous-specific regions play a critical
role in homology search and initiation of meiotic pairing (the classical pairing
initiation sites) (Feldman et al. 1997). In some studies, DNA elimination was not
observed in the first generations of synthetic allopolyploids (Mestiri et al. 2010;
Zhao et al., 2011). It is possible that the timing, as well as the extent of elimination
depends on the type of genomic combinations involved or on the type of sequences
analyzed.
CO
RR
183
UN
182
EC
TE
Fig. 7.2 Schematic representation of the wheat karyotype. The wheat karyotype is arranged into
genomes A, B, and D and into seven homoeologous groups. Examples of the different types of
sequences are drawn on top of the chromosomes, namely: triplicated sequences (group-specific
sequences), chromosome-specific sequences (CSSs) that are present in only one chromosome
pair, genome-specific sequences (GSSs) that can be on more than one chromosome pair but only
in one of the genomes, and dispersed repeats that are present on both homoeologous and
nonhomoeologous chromosomes. Adapted from Levy and Feldman (2004)
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 117/134
7 Genomic Plasticity in Polyploid Wheat
117
Editor Proof
Table 7.5 Evolutionary changes facilitated by allopolyploidy in wheat
Level
Type
• Chromosomal repatterning (intra- and inter-genomic translocations)
• Introgression of chromosomal segments from alien genomes and production of
recombinant genomes
Functional • Nonfunctionalization (deletion or pseudogenization)
• Subfunctionalization
• Neofunctionalization
• Copy number changes
• New allelic variation
202
203
204
205
206
207
208
209
210
211
212
213
214
215
216
217
218
219
220
221
222
223
224
225
226
227
228
229
D
201
TE
200
Several structural changes occur in the allopolyploid wheat genomes, which
generate new variants that could not take place in the diploid parental genomes and
that occur almost exclusively in an allopolyploid background (Table 7.5). This
includes events, such as intergenomic horizontal transfer of chromosomal segments, repetitive sequences, transposons, or genes among the constituent genomes.
These events may occur sporadically throughout the history of the allopolyploid
species. Intergenomic translocations that are population- or biotype-specific are
widespread in allohexaploid wheat (Maestra and Naranjo 1999). Invasion of the A
genome by sequences from the B genome—most probably transposons—was
detected in wild allotetraploid wheat using GISH (Belyayev et al. 2000). The
possibility of intergenomic transfer adds to the allopolyploid genomes’ plasticity
and enables the creation of new genetic combinations that are beyond the addition
of two genomes.
Moreover, in contrast to diploids, which are genetically isolated from each
other and have undergone divergent evolution, allopolyploids in the wheat group
exhibit convergent evolution, because they contain genetic material from two or
more different diploid genomes and can exchange genes with each other via
hybridization and introgression, resulting in the production of new genomic
combinations. Examples of such introgression between allotetraploid Aegilops
species that share one genome and differ in the other genome(s) were provided by
Zohary and Feldman (1962) and Feldman (1965a, b, c). Such hybridizations are
eased by the shared genome, which acts as a buffer and ensures some fertility in
the resulting hybrids. In such hybrids, chromosomes of two dissimilar genomes,
brought together from different parents, may pair and exchange genetic material
and recombine (Feldman 1965a). Additional evidence for the existence of introgressed genomes in allopolyploid Aegilops was obtained from C-banding analysis
(Badaeva et al. 2004). Introgression of a DNA sequence from allopolyploid wheat
to the allotetraploid Aegilops species, Ae. peregrina, was described by Weissmann
et al. (2005).
In addition to evolutionary changes that are almost unique to an allopolyploid
background, it might be that other types of mutations that can cause structural or
functional changes on an evolutionary scale (e.g., point mutations, satellite
EC
199
CO
RR
198
7.3.2 Evolutionary Changes
UN
197
PR
OO
F
Structural
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 118/134
M. Feldman et al.
240
7.4 Functional Changes
232
233
234
235
236
237
238
PR
OO
231
F
239
instability, transposition, etc.) can occur in an accelerated manner in the polyploid
background. The presence of duplication or triplication of the genetic material in
wheat allopolyploids might have relaxed constraints on gene structure and function. Thus, the accumulation of genetic variation through mutations or hybridization might be tolerated more readily in allopolyploid than in diploid species.
While there is no direct evidence for this assertion, there is indirect support from
experimental data showing a higher resistance of allohexaploid wheat to irradiation compared to the diploid progenitors (Mac Key 1954, 1958; Sears 1972). Such
increase in resistance to mutation with increased ploidy level was shown to be
correlated with increased evolvability and fitness in yeast (Thompson et al. 2006).
230
254
7.4.1 Functional Diploidization
246
247
248
249
250
251
252
255
256
257
258
259
260
261
262
263
264
265
266
TE
244
245
EC
243
CO
RR
242
D
253
Allopolyploidy affects gene or protein function through a variety of mechanisms
(Table 7.4). It has been widely suggested that following polyploidization,
individual genes follow one of many possible evolutionary fates including
nonfunctionalization (deletion or pseudogenization), neofunctionalization (evolution of novel functions among alleles or homoeoalleles), generating new phenotypes, or subfunctionalization (evolution of partitioned ancestral functions among
alleles or homoeoalleles) (Lynch and Force 2000; Prince and Pickett 2002;
Chaudhary et al. 2009). Functional changes are often regulated by genetic and
epigenetic interactions among homoeoalleles and might provide the plasticity that
is required to improve the fitness and adaptation of the newly formed allopolyploid
and to increase its competitive efficiency with its parental species as well as other
plant species, leading to its successful establishment in nature. Moreover, as
discussed above, polyploidization seems to facilitate gene evolvability.
241
Increased gene dosage may lead to redundancy or in some cases may have a
deleterious effect, for example, due to the formation of an unbalanced system
(Veitia et al. 2008). Regulating gene action in a duplicated genome might be
achieved through dosage compensation (nonlinear gene dosage response). A
regulatory process that can bring redundant or unbalanced gene systems in
polyploids toward a diploid-like mode of expression is functional diploidization
(Ohno 1970). Functional diploidization is the process whereby existing genes in
multiple doses can be eliminated, become inactive via mutations (base substitution, insertion, or deletion leading to pseudogenization) or diverted to new
functions.
Examples of functional diploidization in polyploid wheat involve mainly genes
that code for structural or storage proteins, e.g., histones, subunits of tubulins,
UN
Editor Proof
118
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 119/134
273
274
275
276
277
278
279
280
281
282
283
284
285
286
287
288
289
290
291
292
293
294
295
296
297
298
299
300
301
302
303
304
305
306
307
308
309
310
311
F
272
PR
OO
271
D
270
TE
269
119
subunits of glutenins and gliadins, and ribosomal RNA (and possibly also tRNA).
In such genes, expression of all homoeoalleles might be redundant and even
deleterious, due to overproduction and inefficiency. Also, activity of all homoeoalleles may produce intermediate phenotypes in several traits that decrease the
viability of the plants. In this case, traits controlled by genes from only one
genome may have a higher adaptive value. It is, therefore, expected that such gene
loci would have been targets for genetic diploidization. In hexaploid wheat, intergenomic suppression, as seen by the disappearance of a storage protein subunit,
was observed immediately upon formation of a wheat allohexaploid (genome
BBAADD; see Galili and Feldman 1984; Galili et al. 1986). This is a common way
to instantaneously reduce the negative effect of overproduction and inefficiency of
genes that exist in super-optimal dose. Fascinatingly, suppression was reversible:
the storage protein reappeared upon extraction of the tetraploid BBAA genomes
and disappeared when the D genome was added. Similarly, attempts to transfer a
leaf-rust resistance gene from tetraploid to hexaploid wheat failed because of a
suppressor gene that was mapped to the D genome (Kerber and Green 1980).
Intergenomic suppression of disease resistance genes is a common phenomenon as
was noticed in natural and in several newly formed allopolyploids (Anikster, Y.,
Manisterski, J., and Feldman, M., unpublished data). Comparable results were
obtained by Dhaliwal and co-workers (Aghaee-Sarbarzeh et al. 2001) who found
that in Triticum durum-Aegilops amphiploids dominant leaf rust and stripe rust
resistance genes from the Aegilops parents were suppressed by genes on the AB
genomes of the wheat parent. Another well-studied example of intergenomic
suppression is the silencing of rye ribosomal RNA genes in the presence of the
wheat genome. Cytosine methylation is involved in this silencing, as suggested by
reactivation of the rye ribosomal RNA genes upon treatment with 5-aza-cytidine
and analyzed by the use of methylation sensitive/insensitive isoschizomers
(Houchins et al. 1997).
Recent analysis of the sequences of wheat group 1 chromosomes shows significant deviations from synteny with many of the nonsyntenic genes representing
pseudogenes (Wicker et al. 2011). Part of this pseudogenization might have
occurred after polyploidization as suggested from the analysis of the Q-factor. A
recent study shows that a combination of mutations in Q genes contributed to the
domestic spike phenotype, namely nonfragile, soft glumes, and free threshing
(Zhang et al. 2011). The mutation with the most significant phenotypic effect is an
amino acid substitution in the protein coded by the 5A locus, while other mutations, such as pseudogenization of the locus on 5B or subfunctionalization of the
locus on 5D, also contributed to the domestication phenotype, but to a lesser extent
(Zhang et al. 2011). Remarkably, these mutations occurred after polyploidization.
The Hardness (Ha) gene constitutes another example of genetic diploidization,
through gene deletion, in polyploid wheat (Chantret et al. 2005).
Genetic diploidization might also be achieved through epigenetic control.
Epigenetic silencing can be brought about via cytosine methylation of DNA
sequences (Kashkush et al. 2002) or chromatin modifications or remodeling as
well as the activity of small RNA molecules (Kenan-Eichler et al. 2011). Gene
EC
268
CO
RR
267
UN
Editor Proof
7 Genomic Plasticity in Polyploid Wheat
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 120/134
314
315
316
317
318
319
320
silencing can also be achieved via novel regulatory interactions such as intergenomic suppression (Galili and Feldman 1984). Shaked et al. (2001) and Kashkush
et al. (2002) reported the occurrence of alterations in cytosine methylation in 13 %
of the loci affecting both repetitive DNA sequences and low-copy DNA in
approximately equal proportions. Changes in microRNAs, such as miR168 which
targets the Argonaute1 gene, were shown to occur in newly synthesized wheat
(Kenan-Eichler et al. 2011).
F
313
PR
OO
312
M. Feldman et al.
7.4.2 Subfunctionalization Through Partitioning
and Compensation of Duplicated Gene Expression
338
7.4.3 New Interactions Between Maintained Genes/Proteins
326
327
328
329
330
331
332
333
334
335
336
339
340
341
342
343
344
345
346
347
TE
324
325
EC
323
CO
RR
322
D
337
Functional diversification of duplicated genes (subfunctionalization), i.e., differential or partitioning of expression of homoeoalleles in different tissues and/or in
different developmental stages, is also a form of genetic diploidization (Adams
et al. 2003). Subfunctionalization, an important aspect of allopolyploidy, has been
studied relatively little in allopolyploid wheat. Koebner and co-workers (Bottley
et al. 2006) found that differential expression of homoeoalleles in different plant
tissues is common in hexaploid wheat. The activity of silenced genes could be
restored in aneuploid lines, suggesting that no mutation was involved but rather
new cis–trans interactions or reversible epigenetic alterations were responsible.
The data of Bottley et al. (2006) suggest that for leaf transcripts, there is a modest
bias toward silencing of the D genome copies, but this pattern does not extend to
root transcripts. Mochida et al. (2006) also presented evidence for differential
expression of homoeoalleles in wheat and suggested that inactivation of homoeoalleles is a nonrandom effect. The molecular basis for these cases of instant
subfunctionalization remains to be determined. Similarly, subfunctionalization of
all three homoeologs of the Q-locus in hexaploid wheat was recently described
(Zhang et al. 2011).
321
In allohexaploid wheat many gene loci exist in triplicate dose, and the extra gene
dosage per se may produce improved or even novel traits. Homoeoalleles may
differ from one another by allelic variation, and in this case, activity of all the
duplicated genes may produce desirable intergenomic interactions and heterotic
effects. These are mainly genes that code for functional proteins (enzymes). This
occurrence of enzyme diversity (isozymes) increases the biochemical potential of
the allopolyploids. In fact, in wheat allopolyploids the expression of most homoeoalleles coding for functional enzymes is retained (Mitra and Bhatia 1971;
Hart 1983a, b, 1987). Also, intergenomic gene interactions may be, in some cases,
UN
Editor Proof
120
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 121/134
355
356
357
358
359
360
361
362
363
364
365
366
367
368
369
370
371
372
373
374
375
376
377
378
379
380
381
382
383
384
385
386
387
388
389
390
391
392
F
354
PR
OO
353
D
351
352
TE
350
121
expressed in novel traits that do not exist in their parental diploids. Some of these
traits may have great adaptive value. Intergenomic gene interactions have direct
relevance also to wheat cultivation. For example, the baking quality of allohexaploid wheat (bread wheat) is due to the unique properties of its gluten—a product
derived from the combined contribution of the three genomes of hexaploid wheat,
and thus exists only at the hexaploid level. In addition, the combination of a large
number of spikelets per spike derived from T. urartu (the donor of the A genome),
with several fertile florets per spikelet originating from the donors of the B genome, facilitated the high fertility of durum and bread wheat. Likewise, merging of
Ae. tauschii (the donor of the D genome) displaying the cold hardiness phenotype
with the prolific nature of allotetraploid wheat (the donor of the A and B genomes)
enabled the expansion of wheat cultivation into colder regions. Enzyme multiplicity, derived from the activity of all homoeoalleles, increases the ability of the
allopolyploid to adapt to a wider range of environments. This might account for
the very wide distribution of wheat under cultivation—much wider than that of any
other cultivated plant.
Genome-wide reprograming in gene expression may occur in a polyploid as a
result of new interactions among regulatory factors of the parents. For example,
novel interactions between the trans factor from one species and the cis or trans
factors of the other parental species, as was shown in yeast in a cross between wide
hybrids (Tirosh et al. 2009), may account for the observed cases of gene repression
or for cases of activation via overexpression.
The consequences of allopolyploidy on gene expression have been widely
studied at the genome-wide level in several natural and synthetic allopolyploids of
Triticum. The majority of studies relied on the comparison of the expression level
in the allopolyploid to those of its parents and/or to the average of its parents,
expressed as the mid-parental value (MPV). In hexaploid wheat, Pumphrey et al.
(2009) found that approximately 16 % of the 825 analyzed genes displayed
nonadditive expression in the first generation of synthetic hexaploid wheat.
Chagué et al. (2010) analyzed 55,052 transcripts in two lines of synthetic allohexaploid wheat and found that 7 % of the genes had nonadditive expression,
while Akhunova et al. (2010) found in synthetic allohexaploid wheat that about
19 % of the studied genes showed nonadditive expression. Similar studies (He
et al. 2003) showed that the expression of a significant fraction of genes (7.7 %)
was altered in the synthetic allohexaploid T. turgidum-Ae. tauschii, and that Ae.
tauschii genes were affected much more frequently than those of T. turgidum.
Strikingly, these different studies show that deviation of gene expression in allopolyploids from their parents or average of parents appears to be a common
feature, although evaluated in various allopolyploids and using different techniques and approaches. Interestingly, silencing of the same genes was also found
in natural hexaploid wheat, i.e., in the variety Chinese Spring (He et al. 2003).
Chagué et al. (2010) suggested, based on similar gene expression patterns
observed between natural and synthetic wheat allohexaploids, that regulation of
gene expression is established immediately after allohexaploidization and maintained over generations. It is of interest to note that several genes that are silent in
EC
349
CO
RR
348
UN
Editor Proof
7 Genomic Plasticity in Polyploid Wheat
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 122/134
M. Feldman et al.
407
7.5 Genome asymmetry
400
401
402
403
404
405
408
409
410
411
412
413
414
415
416
417
418
419
420
421
422
423
424
425
426
427
428
429
430
431
PR
OO
399
D
398
The rapid processes of cytological and genetic diploidization allow for the
development and occurrence of two contrasting and highly important genetic
phenomena in allopolyploid wheat that contribute to the evolutionary success of
these polyploids: (1) build up and maintenance of enduring intergenomic favorable
genetic combinations, and (2) genome asymmetry in the control of a variety of
morphological, physiological, and molecular traits, i.e., complete or principal
control of certain traits by only one of the constituent genomes. However, while
the first phenomenon was taken for granted by plant geneticists, genomic asymmetry in interspecific hybrids and allopolyploids was mainly known in ribosomal
RNA genes (reviewed in Pikaard 2000) and only recently has also been documented in other traits (Peng et al. 2003a, b; Fahima et al. 2006; Feldman and Levy
2009; Flagel et al. 2009; Rapp et al. 2009; Flagel and Wendel 2010). The phenomena of diploidization and of dominance in gene expression lead to genome
asymmetry, which is manifested in a clear-cut division of tasks among the constituent genomes of allopolyploid wheat (Levy and Feldman 2004; Feldman and
Levy 2009; Feldman et al., in press). Genome A controls morphological traits
while genome B in allotetraploid wheat and genomes B and D in allohexaploid
wheat control the reaction to biotic and abiotic factors (Tables 7.6 and 7.7). Intergenomic pairing would lead to both disruption of the linkage of the homoeoalleles that contribute to positive intergenomic interactions and segregation of
genes that participate in the control of certain traits by a single genome. Intergenomic recombination may, therefore, result in many intermediate phenotypes
that may affect, in a negative manner, the functionality, adaptability, and stability
of the allopolyploids.
TE
396
397
EC
395
CO
RR
394
F
406
the parental species became active in the newly formed allohexaploid (He et al.
2003). Similarly, cDNA-AFLP gels also revealed several cDNAs that were
expressed only in the allopolyploids and not in the diploid progenitors (Shaked
et al. 2001; Kashkush et al. 2002).
The proportion of genes for which expression in the polyploids is different from
the average of the parents is underestimated in these studies, as most of the
technologies used do not allow separate analysis of the expression level of each of
the gene copies (homoeoalleles) and their respective contribution to overall gene
expression (Pumphrey et al. 2009; Chagué et al. 2010; Akhunova et al. 2010). It is
not possible to detect, for example, a situation where repression of one homoeoallele is compensated by the activation of the other using microarray technologies. The progress towards sequencing of wheat genomes and the promise of
next-generation technologies should allow in the future the resolution of expression at the level of individual homoeologues.
393
UN
Editor Proof
122
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 123/134
7 Genomic Plasticity in Polyploid Wheat
123
Glumes with keels
The shape of the edge of the glumes (beaked
glumes)
Hairs at the base of every spikelet
Plant habitus
Growth habit
Autogamous behavior
Many domestication genes
436
437
438
439
440
441
442
443
444
445
446
447
448
449
450
451
452
453
454
D
In the preceding sections, we have discussed structural changes and changes in
gene expression resulting from genetic or epigenetic phenomena. Transposable
elements (TEs) are discussed here separately due to their specific mode of action,
their abundance (up to 90 % of the wheat genomes; Sabot et al. 2005), and their
impact on both structure and expression of the genome.
TE
435
EC
434
7.6.1 Transcriptional Activation of Retrotransposons in Synthetic
Wheat Polyploids
CO
RR
433
Higher polymorphism of HMW glutenin
genes
Larger amount of repetitive sequences
Activity on nucleolar organizers
Larger number of rRNA genes
7.6 Response of Transposable Elements
to Allopolyploidization
It is now clear that some eukaryote retrotransposon promoters retain activity under
normal conditions and initiate either read-in transcripts of the transposon itself or
read-out transcripts into flanking host sequences (Vicient et al. 2001; Kashkush
et al. 2002; Nigumann et al. 2002; Kashkush et al. 2003; Kashkush and Khasdan
2007). Following allopolyploidization events in wheat, the steady-state level of
expression of LTR retrotransposons was massively elevated (Kashkush et al. 2002,
2003, unpublished data), similarly to what was observed in synthetic Arabidopsis
allopolyploid hybrids (Madlung et al. 2005). In addition, the transcriptional
activity of a LTR element termed Wis2-1A (Lucas et al. 1992) leads to the production of read-out transcripts toward flanking host DNA sequences, a process that
occurred in a genome-wide manner (Kashkush et al. 2003). In many cases, these
read-out transcripts were associated with the expression of adjacent genes,
depending on their orientation: knocking down or knocking out the gene product if
the read-out transcript was in the antisense orientation relative to the orientation of
UN
432
Regulation of ecological adaptation
Double the number of disease resistance
genes
Contains more stress–related genes?
Higher polymorphism of molecular markers
F
Inflorescence morphology
Free caryopsis
PR
OO
Editor Proof
Table 7.6 Genome asymmetry in the control of various traits in the wild allotetraploid wheat,
T. turgidum subsp. dicoccoides (genome BBAA)
Traits under control of Genome A
Traits under control of Genome B
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 124/134
M. Feldman et al.
Table 7.7 Genome asymmetry in the control of agronomic traits in domesticated durum (genome BBAA) and bread wheat (genome BBAADD)a
Traits
Traits under control of
br B1 on 3BS
tg2 on 2BS
PR
OO
Eg P2 on 7BL (?)
Rht B1 on 4BS; Rh4
on 2Bl;
Ga1, Ga3 on 4BS
W1 on 2BS
W1I on 2BS; W3I on
1BL
ms1 on 4BS
Ph1 on 5BL
Ne1 on 5BL; Ne2 on
2BS
EC
CO
RR
Hybrid chlorosis
Ch1 on 2A
Aluminum tolerance
Boron tolerance
Low cadmium uptake
Iron deficiency
Herbicide response
Difenzoquat insensitivity
Chlortoluron insensitivity Imi3 on 6AL
Imidazolinone resistance
Response to photoperiod
Response to vernalization Vrn-A1 on
5AL
Response to salinity
Frost resistance
Number of resistance
genes to diseases and
pests
a
Fr1 on 5AL
45
tg1 on 2DS
Rht D1 on 4DS; Rht8 on
2DL;
Rht5 on 3BS; Rht9 on
7BS; Rht13 on
7BS
Gpc B1 on 6BS
Pro1 on 5DL; Pro2 on 5Ds
Ha on 5DS
Pin D1 on 5DS
TE
Grain protein content
Grain hardness
Puroindolines and grain
softness protein
Gibberellic acid response
Waxiness
Epistatic inhibitors of
waxiness
Male sterility
Ms3 on 5AS;
ms5 on 3A
Pairing homoeologous
Hybrid necrosis
Genome D
F
Genome B
D
Genome A
Inflorescence morphology
Elongated glumes
Eg P1 on 7AL
Branched spikes
Bh on 2AS
Nonbrittle rachis
br A1 on 3AS
br A2 on 2A
Nontenacious glume (lax
glume)
Reduce plant height
Rht7 on 2A;
Rht12 on
5AL;
UN
Editor Proof
124
Bo1 on 7BL
Cdu1 on 5BL
Fe2 on 7BS
Dfg 1 on 2BL
Su1 on 6BS
Imi2 on 6BL
Ppd-B1 on 2BS
Vrn-B1 on 5BL;
Vrn-B3 on 7BS
88
Ga2 on 4DS
W2 on 2DS (?)
W2I on 2DS
Ms2 on 4DS; Ms4 on 4DS
Ph2 on 3DS
Ch2 on 3DL
Alt2 on 4DL
Fe1 on 7DL
Imi1 on 6DL
Ppd-D1 on 2DS
Vrn-D1 on 5DL; Vern-D4
on 5DL;Vern-D5 on
5DL
Kna1 on 4DL
Fr2 on 5DL
51
Data from the 2008 Wheat Gene Catalogue (http://wheat.pw.usda.gov/GG2/index.shtml)
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 125/134
461
462
463
464
465
466
467
468
469
470
471
472
473
474
475
476
477
478
479
480
481
482
483
484
485
486
487
488
489
490
491
492
493
494
F
460
PR
OO
459
7.6.2 Massive Methylation of TE-Adjacent DNA Sequences
Following Allopolyploidization
D
458
Alterations in the genomic methylation patterns following allopolyploidization
have been examined in several polyploid systems, including Arabidopsis (Madlung
et al. 2002; Belzile et al. 2009), Spartina (Salmon et al. 2005; Parisod et al. 2009),
Brassica (Lukens et al. 2006; Wang et al. 2009), and wheat (Shaked et al. 2001).
The methylation alterations are either hyper- or hypomethylation, depending on the
sequence analyzed, and are reproducible. Recent studies in wheat have investigated
in detail the methylation of CCGG sites flanking several TE families (Kraitshtein
et al. 2010; Yaakov and Kashkush 2011a, b; Zhao et al. 2011). In one study
(Kraitshtein et al. 2010), transposon methylation display (TMD) analysis was
applied (see Kashkush and Khasdan 2007) to analyze a terminal-repeat retrotransposon in miniature (TRIM), termed Veju, in Triticum turgidum ssp. durum
(genome AABB) and Aegilops tauschii (genome DD), and the first four generations
of the derived allohexaploid. It was estimated that over 50 % of the CCGG sites
flanking Veju elements showed altered TMD patterns in the first four generations of
the newly formed allohexaploid. Hypomethylation of Veju-flanking CCGG sites
was predominant in the first generation of the newly formed allohexaploid, while
hypermethylation was predominant in subsequent generations. This might indicate
reduced Veju transcriptional activity after the third generation of the synthetic
allohexaploid. A similar pattern of hypomethylation of Veju elements was also
observed in the first three generations of a synthetic allotetraploid that was derived
from a cross between Ae. sharonensis (genome SlSl) and T. monococcum ssp.
aegilopoides (genome AmAm) (Yaakov and Kashkush 2011b). However, unlike in
the synthetic allohexaploid, Veju elements remained hypomethylated up to the
fourth generation of the synthetic allotetraploid. In support of these studies, Zhao
et al. (2011) observed massive methylation changes around Veju in three different
combinations of newly formed wheat allohexaploids.
TE
457
125
the gene transcript (such as the iojap-like gene) or overexpressing the gene if the
read-out transcript was in the sense orientation (such as the puroindoline-b gene).
The mechanisms by which transcriptional activation of TEs influences the
expression of neighboring genes are poorly understood. In some cases, the correlation between the reduction of the sense expression of the gene and the production of the antisense strand that initiated from the adjacent transposon promoter
(Kashkush et al. 2003; Puig et al. 2004) might indicate that post-transcriptional
gene silencing is a major mechanism for inactivating adjacent genes. Recent
studies on tracking methylation changes around a LTR retrotransposon in the first
four generations of a newly formed wheat allopolyploid (Kraitshtein et al. 2010)
may indicate that this read-out activity is restricted to the first generations of the
nascent polyploid species.
EC
456
CO
RR
455
UN
Editor Proof
7 Genomic Plasticity in Polyploid Wheat
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 126/134
M. Feldman et al.
507
7.6.3 Changes in TE Composition Following Allopolyploidization
501
502
503
504
505
508
509
510
511
512
513
514
515
516
517
518
519
520
521
522
523
524
525
526
527
528
529
530
531
532
533
534
535
PR
OO
500
The prevalence of TEs and their inherent sequence similarity make them a prime
target for illegitimate and nonhomologous recombination. TEs have been shown to
undergo rearrangements following allopolyploidization in Spartina (Parisod et al.
2009), tobacco (Petit et al. 2010), and Triticale (Bento et al. 2008). Recent data for
synthetic allohexaploid wheat indicate that rearrangements of retrotransposoncontaining sequences occur rapidly and reproducibly (repeated in independently
newly formed allopolyploid lines) in the first generations following polyploidization. In addition, a change in the methylation status (usually hypomethylation) in
the first generation was followed by deletion of retrotransposon-containing
sequences in subsequent generations (Kraitshtein et al. 2010). These data suggest a
correlation between methylation and post-allopolyploidization rearrangements that
occur via a mechanism that has yet to be identified. One possible explanation is
that hypomethylation confers an open chromatin structure to the TE sequences,
which exposes these demethylated elements to be targeted for deletion by the host.
There is evidence that small RNAs corresponding to Veju elements might play a
pivotal role in Veju methylation in the newly formed wheat allohexaploid (KenanEichler et al. 2011).
Despite the altered methylation status and transcriptional activation of TEs
following allopolyploidization, there are very few reports on the transpositional
activity of transposons. Madlung et al. (2005) showed both methylation alterations
and limited transpositional activation of a Sunfish transposon in polyploid
Arabidopsis. Petit et al. (2010) reported an increase in the copy number of a Tnt1
retrotransposon in allotetraploid tobacco. No transposition bursts were reported in
Spartina (Parisod et al. 2009) or in wheat (Kashkush et al. 2003; Kraitshtein et al.
2010; Yaakov and Kashkush 2011a). These reports suggest that the transpositional
activity of TEs following allopolyploidization might be restricted to specific TE
families (Parisod et al. 2010). Recently, it was shown that the immense loss of
Veju sequences in the first generation of the synthetic allohexaploid is probably
D
499
TE
498
EC
497
CO
RR
496
F
506
The methylation patterns of three TEs [Balduin (belonging to the CACTA
superfamily), Apollo (belonging to the MuDR/Foldback superfamily), and Thalos
(a stowaway-like MITE belonging to the Tc1/mariner superfamily)] have also been
analyzed in allopolyploid wheat. The CCGG sites flanking the three elements
underwent massive hypermethylation in the first four generations of the synthetic
allohexaploid (Yaakov and Kashkush 2011a, b), while they underwent massive
hypomethylation in the first four generations of the synthetic allotetraploid
(Yaakov and Kashkush 2011b). The massive hypermethylation of these elements
in the synthetic allohexaploid might be connected to the lack of transpositional
activity (Yaakov and Kashkush 2011a). It is important to mention that transcriptional activation of LTR retrotransposons does not correlate with transpositional
activity (Kashkush et al. 2003).
495
UN
Editor Proof
126
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 127/134
127
552
7.7 Concluding Remarks
543
544
545
546
547
548
549
550
553
554
555
556
557
558
559
560
561
562
563
564
565
566
567
568
569
570
571
572
573
574
575
576
PR
OO
542
D
541
TE
539
540
The studies reviewed above indicate that, in the wheat group, hybridization and
chromosome doubling induce a burst of genomic alterations, some of which could
not occur at the diploid level. Some of these changes might improve the ability of
the newly formed allopolyploids to survive in nature and to compete with their
parental species, corresponding thus to phenotypic and adaptive novelty. Other
changes are probably deleterious. TEs seem to play an important role in the
various responses to hybridization and polyploidization due to their abundance and
also due to their tendency to be dysregulated as a result of genomic shocks. The
balance between the beneficial and deleterious changes associated with allopolyploidization is probably what determines the fate of the nascent species.
The formation of an allopolyploid species is accomplished rapidly via the
combined processes of hybridization and genome doubling, but its establishment
in nature as a successful species probably depends on a high degree of plasticity
that enables it to overcome potential incompatibilities and to gain new traits. The
studies reported here suggest that wheat can achieve genomic plasticity through
the induction of a series of cardinal nonadditive genomic changes. Some changes,
genetic and epigenetic, are rapid and nonMendelian, occurring during or immediately after the formation of the F1 hybrid or the allopolyploid (revolutionary
changes). Other changes occur sporadically over a long time period during the life
of the allopolyploid species (evolutionary changes). From a population point of
view, the chance of a new individual, such as a nascent hybrid/allopolyploid, to
establish itself as a new species is almost nil, unless it has some increased fitness
over its parents. This fitness advantage must be manifested within a few generations of formation or the new species will rapidly be extinct. The revolutionary
EC
538
CO
RR
537
F
551
followed by retrotransposition in subsequent generations, a process that causes
new insertions to accumulate in allohexaploids (Kraitshtein et al. 2010). The same
study also suggests that these new insertions are targeted for methylation. Methylation of the new Veju elements protects the genome from deleterious transposon
insertions. Investigating the scale of eliminated DNA sequences, including TE
sequences, by identifying the deletion breakpoints will allow a better understanding of the mechanism(s) involved and of the nature of the connection between
methylation and rearrangements.
In summary, different classes of wheat TEs appear to respond differently to the
allopolyploidization event. Table 7.8 summarizes the type of response of 12 different class I and class II elements. It can be seen clearly that epigenetic response,
mainly methylation changes, is a common factor for all studied TEs, while genetic
response that includes rearrangements and/or transpositions is restricted to specific
TE families. Transcriptional data for most studied TEs are still lacking (Table 7.8).
However, there is a good basis for suggesting a connection among methylation
changes in TEs with alteration in their expression patterns.
536
UN
Editor Proof
7 Genomic Plasticity in Polyploid Wheat
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 128/134
128
M. Feldman et al.
Editor Proof
Table 7.8 Summary of the genetic and epigenetic responses of several TE families to allopolyploidization in wheat
Reference
Family
TE
Genetic and epigenetic alterations in synthetic
allopolyploids (compared to parental lines)a
Methylation Transcription Rearrangementb Transpositionc
4
4
4
Copia
Angela
BARE1
Wis2-1
A
4
4
NA
NA
NA
9
NA
4
9
NA
NA
9
9
MuDR/
Foldback
Apollo
4
NA
4
NA
a
b
c
578
579
580
581
582
583
584
585
4
9
9
9
9
9
9
4
4
4
4
?
4
?
?
EC
NA
NA
NA
NA
Kraitshtein
et al.
(2010)
KenanEichler
et al.
(2011)
Zhao
et al.
(2011)
Unpublished
Zhao et al.
(2011)
Kashkush
et al.
(2003)
Unpublished
Yaakov and
Kashkush
(2011a)
Yaakov and
Kashkush
(2011a)
Yaakov and
Kashkush
(2011a)
Unpublished
Unpublished
Unpublished
Unpublished
4 altered, 9 no change, ? not validated, NA data not available
Include deletion and/or insertion
Typical TE transposition
changes described here may contribute to the establishment of the new species.
Instantaneous elimination of sequences from one genome in the newly formed
allopolyploids increases the divergence of the homoeologous chromosomes, and
thus leads to exclusive intragenomic pairing that improves fertility. Mechanisms
such as loss of deleterious genes (e.g. genetic incompatibilities) or positive dosage
effects or new intergenomic heterotic interactions may all rapidly increase the
fitness of the nascent species. The evolutionary changes, on the other hand, contribute to the build-up of genetic variability and thus increase adaptability, fitness,
competitiveness, and colonizing ability. It is clear that most hybridization events in
UN
577
4
4
CO
RR
Stowaway- Thalos
like
MITE
Eos
Minos
Oleus
Fortuna
9
D
Sabrina 4
Balduin 4
9
9
TE
Gypsy
CACTA
?
F
Veju
PR
OO
TRIM
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 129/134
129
610
References
611
612
613
614
615
616
617
618
619
620
621
622
623
624
625
626
627
628
629
630
Aaronsohn A (1910) Agricultural and botanical explorations in Palestine. Bull Plant Ind 180:1–63
Aaronsohn A, Schweinfurth G (1906) Die auffindung des wilden emmers (Triticum dicoccum) in
Nordpalästina. Altneuland Monatsschrift für die irtschaft. Erschliessung Palästinas
7(8):213–220
Adams KL, Cronn R, Percifield R, Wendel JF (2003) Genes duplicated by polyploidy show
unequal contributions to the transcriptome and organ-specific reciprocal silencing. Proc Natl
Acad Sci U S A 100(8):4649–4654
Aghaee-Sarbarzeh M, Dhaliwal HS, Harjit-Singh (2001) Suppression of rust resistance genes
from distantly related species in Triticum durum-Aegilops amphiploids. In: Johnson R,
Yahyaoui A, Wellings C, Saidi A, Ketata H (eds) Meeting the challenge of yellow rust in
cereal crops. Proceedings of the First Regional Conference on Yellow Rust in the Central and
West Asia and North Africa Region, Karaj, Iran. pp 8–14
Akhunova AR, Matniyazov RT, Liang H, Akhunov ED (2010) Homoeolog-specific transcriptional bias in allopolyploid wheat. BMC genomics 11:505
Avivi L (1976) The effect of genes controlling different degrees of homoeologous pairing on
quadrivalent frequency in induced autotetraploid lines of Triticum longissimum. Can J Genet
Cytol 18:357–364
Badaeva ED, Amosova AV, Samatadze TE, Zoshchuk SA, Shostak NG, Chikida NN, Zelenin
AV, Raupp WJ, Friebe BR, Gill BS (2004) Genome differentiation in Aegilops. 4. Evolution
of the U-genome cluster. Plant Syst Evol 246:45–76
592
593
594
595
596
597
598
599
600
601
602
603
604
605
606
607
608
PR
OO
591
D
590
TE
589
EC
588
CO
RR
587
F
609
nature do not lead to the formation of a new species, but remarkably, the wheat
group is equipped with a battery of molecular mechanisms that provide the
potential for phenotypic novelty and for successful speciation to occur. Future
work should enable better understanding of the role of specific genes and DNA
sequences in speciation, the mechanisms that confer robustness of the genome to
the shock of allopolyploidy and to the activation of TEs, and the mechanisms that
enable the orchestration of chromosome division and the control of bivalent
pairing during meiosis.
Altogether, the reported revolutionary and evolutionary genomic changes
emphasize the dynamic plasticity of the wheat allopolyploid genome with regard
to both structure and function. Presumably, these changes have improved the
adaptability of the newly formed allopolyploids and facilitated their rapid colonization of new ecological niches. No wonder, therefore, that cultivated allopolyploid wheats exhibit a wider range of genetic flexibility than diploid wheats
and could adapt themselves to a great variety of environments. In contrast to
Stephens (1951), who had the insight that allopolyploidy might lead to new
evolutionary opportunities, Stebbins (1971, 1980) stated that while polyploidy has
been of great importance for the origin of species it has contributed little to
progressive evolution. He assumed that polyploids evolve more slowly than their
diploid relatives. Stebbins (1971, 1980) did not take into consideration that allopolyploidization triggers a burst of genomic alterations that are not feasible at the
diploid level and that lead to new evolutionary opportunities. Allopolyploidy has
proved to be a powerful evolutionary factor that has played a decisive role in the
evolution of the wheat group.
586
UN
Editor Proof
7 Genomic Plasticity in Polyploid Wheat
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 130/134
EC
TE
D
PR
OO
F
Belyayev A, Raskina O, Korol A, Nevo E (2000) Coevolution of A and B genomes in
allotetraploid Triticum dicoccoides. Genome 43(6):1021–1026
Belzile F, Beaulieu J, Jean M (2009) The allotetraploid Arabidopsis thaliana-Arabidopsis lyrata
subsp petraea as an alternative model system for the study of polyploidy in plants. Mol Genet
Genomics 281(4):421–435
Bento M, Pereira HS, Rocheta M, Gustafson P, Viegas W, Silva M (2008) Polyploidization as a
retraction force in plant genome evolution: sequence rearrangements in Triticale. PLoS ONE
3:1402–1413
Blakeslee AF (1937) Redoublement du nombre de chromosomes chez les plantes par traitement
chimique. Compt Rend Acad Sci Paris 205:476–479
Bottley A, Xia GM, Koebner RMD (2006) Homoeologous gene silencing in hexaploid wheat.
Plant J 47(6):897–906
Boyko EV, Badaev NS, Maximov NG, Zelenin AV (1984) Does DNA content change in the
course of triticale breeding. Cereal Res Commun 12(1–2):99–100
Boyko EV, Badaev NS, Maximov NG, Zelenin AV (1988) Regularities of genome formation and
organization in cereals. I. DNA quantitative changes in the process of allopolyploidization.
Genetika 24:89–97
Chague V, Just J, Mestiri I, Balzergue S, Tanguy AM, Huneau C, Huteau V, Belcram H, Coriton
O, Jahier J, Chalhoub B (2010) Genome-wide gene expression changes in genetically stable
synthetic and natural wheat allohexaploids. New phytol 187(4):1181–1194
Chaudhary B et al (2009) Reciprocal silencing, transcriptional bias and functional divergence of
homeologs in polyploid cotton (gossypium). Genetics 182:503–517
Chantret N, Salse J, Sabot F, Rahman S, Bellec A, Laubin B, Dubois I, Dossat C, Sourdille P,
Joudrier P, Gautier MF, Cattolico L, Beckert M, Aubourg S, Weissenbach J, Caboche M,
Bernard M, Leroy P, Chalhoub B (2005) Molecular basis of evolutionary events that shaped
the hardness locus in diploid and polyploid wheat species (Triticum and Aegilops). Plant Cell
17(4):1033–1045
Chapman V, Miller TE, Riley R (1976) Equivalence of the A genome of bread wheat and that of
Triticum urattu. Genet Res 27:69–76
Dvorak J (1976) The relationship between the genome of Triticum urattu and the A and B
genomes of Triticum aestivum. Can J Genet Cytol 18:371–377
Dvorak J (2009) Triticeae genome structure and evolution. In: Feuiller C, Muehlbauer GJ (eds)
Genetics and genomics of the Triticeae, plant genetics and genomics: crops and models 7.
Springer, Berlin. pp 685–711
Eilam T, Anikster Y, Millet E, Manisterski J, Feldman M (2008) Nuclear DNA amount and
genome downsizing in natural and synthetic allopolyploids of the genera Aegilops and
Triticum. Genome 51(8):616–627
Eilam T, Anikster Y, Millet E, Manisterski J, Sagi-Assif O, Feldman M (2010) Genome size in
diploids, allopolyploids, and autopolyploids of mediterranean triticeae. doi:10.1155/2010/
341380
Fahima T, Cheng JP, Peng JH, Nevo E, Korol A (2006) Asymmetry distribution of disease
resistance genes and domestication synrome QTLs in tetraploid wheat genome. 8th
International Congress of Plant Molecular Biology, Adelaide, Australia
Feldman M (1965a) Chromosome pairing between differential genomes in hybrids of tetraploid
Aegilops species. Evolution 19:563–568
Feldman M (1965b) Fertility of interspecific F1 hybrids and hybrid derivatives involving
tetraploid species of Aegilops Section Pleionathera. Evolution 19:556–562
Feldman M (1965c) Further evidence for natural hybridization between tetraploid tetraploid
species of Aegilops Section Pleionathera. Evolution 19:162–174
Feldman M (2001) The origin of cultivated wheat. In: Bonjean A, Angus W (eds) The wheat
book. Lavoisier Tech and Doc, Paris, pp 1–56
Feldman M, Levy AA (2005) Allopolyploidy—a shaping force in the evolution of wheat
genomes. Cytogenet Genome Res 109(1–3):250–258
CO
RR
631
632
633
634
635
636
637
638
639
640
641
642
643
644
645
646
647
648
649
650
651
652
653
654
655
656
657
658
659
660
661
662
663
664
665
666
667
668
669
670
671
672
673
674
675
676
677
678
679
680
681
682
683
M. Feldman et al.
UN
Editor Proof
130
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 131/134
131
EC
TE
D
PR
OO
F
Feldman M, Levy AA (2009) Genome evolution in allopolyploid wheat—a revolutionary
reprogramming followed by gradual changes. J Genet Genomics 36(9):511–518
Feldman M, Levy AA, Fahima T, Korol A (2012) Genomic asymmetry in allopolyploid plants wheat as a model. J. Exp. Bot. (in press)
Feldman M, Liu B, Segal G, Abbo S, Levy AA, Vega JM (1997) Rapid elimination of low-copy
DNA sequences in polyploid wheat: a possible mechanism for differentiation of homoeologous chromosomes. Genetics 147(3):1381–1387
Feldman M, Lupton FGH, Miller TE (1995) Wheats. In: Smartt J, Simmonds NW (eds) Evolution
of crop plants, 2nd edn. Longman Scientific, London, pp 184–192
Flagel LE, Chen LP, Chaudhary B, Wendel JF (2009) Coordinated and fine-scale control of
homoeologous gene expression in allotetraploid cotton. J Hered 100(4):487–490
Flagel LE, Wendel JF (2010) Evolutionary rate variation, genomic dominance and duplicate gene
expression evolution during allotetraploid cotton speciation. New Phytol 186(1):184–193
Galili G, Feldman M (1984) Inter-genomic suppression of endosperm- protein genes in common
wheat. Can J Genet Cytol 26:651–656
Galili G, Levy AA, Feldman M (1986) Gene-dosage compensation of endosperm proteins in
hexaploid wheat Triticum aestivum. Proc Natl Acad Sci U S A 83:6524–6528
Griffiths S, Sharp R, Foote TN, Bertin I, Wanous M, Reader S, Colas I, Moore G (2006)
Molecular characterization of Ph1 as a major chromosome pairing locus in polyploid wheat.
Nature 439(7077):749–752
Gupta PK, Kulwal PL, Rustgi S (2005) Wheat cytogenetics in the genomics era and its relevance
to breeding. Cytogenet Genome Res 109(1–3):315–327
Han FP, Fedak G, Guo WL, Liu B (2005) Rapid and repeatable elimination of a parental genomespecific DNA repeat (pGcIR-1a) in newly synthesized wheat allopolyploids. Genetics
170(3):1239–1245
Han FP, Fedak G, Ouellet T, Liu B (2003) Rapid genomic changes in interspecific and
intergeneric hybrids and allopolyploids of Triticeae. Genome 46(4):716–723
Hart GH (1983a) Genetic and evolution of mulilocus isozymes in hexaploid wheat. In: Ratazzi
MC, Scandalios JG, Whitt GS (eds) Isozymes: current topics in biological and medical
research, vol 10., Genetics and Evolution Alan R. Liss., Inc., New York, pp 365–380
Hart GH (1983b) Hexaploid wheat (Triticum aestivum L. em Thell.). In: Tanksley SD, Orton TJ
(eds) Isozymes in plant genetics and breeding, Part. B, Elsvier Science Publishers B.V.,
Amsterdam, pp 35–56
Hart GH (1987) Genetic and biochemical studies of enzymes. In: Heyne EG (ed) Wheat and
wheat improvement, Second Ed., Amer. Soc. Agronomy, Madison, Wisconsin, USA
He P, Friebe BR, Gill BS, Zhou JM (2003) Allopolyploidy alters gene expression in the highly
stable hexaploid wheat. Plant Mol Biol 52(2):401–414
Houchins K, ODell M, Flavell RB, Gustafson JP (1997) Cytosine methylation and nucleolar
dominance in cereal hybrids. Mol Gen Genet 255(3):294–301
Huang S, Sirikhachornkit A, Su X, Faris J, Gill B, Haselkorn R, Gornicki P (2002) Genes
encoding plastid acetyl-CoA carboxylase and 3-phosphoglycerate kinase of the Triticum/
Aegilops complex and the evolutionary history of polyploid wheat. Proc Natl Acad Sci U S A
99(12):8133–8138
Kashkush K, Feldman M, Levy AA (2002) Gene loss, silencing and activation in a newly
synthesized wheat allotetraploid. Genetics 160(4):1651–1659
Kashkush K, Feldman M, Levy AA (2003) Transcriptional activation of retrotransposons alters
the expression of adjacent genes in wheat. Nature Genet 33(1):102–106
Kashkush K, Khasdan V (2007) Large-scale survey of cytosine methylation of retrotransposons,
and the impact of readout transcription from LTRs on expression of adjacent rice genes.
Genetics 177:1975–1985
Kenan-Eichler M, Leshkowitz D, Tal L, Noor E, Melamed-Bessudo C, Feldman M, Levy AA
(2011) Wheat hand polyploidization results in deregulation of small RNAs. Genetics
188:263–272
CO
RR
684
685
686
687
688
689
690
691
692
693
694
695
696
697
698
699
700
701
702
703
704
705
706
707
708
709
710
711
712
713
714
715
716
717
718
719
720
721
722
723
724
725
726
727
728
729
730
731
732
733
734
735
736
UN
Editor Proof
7 Genomic Plasticity in Polyploid Wheat
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 132/134
EC
TE
D
PR
OO
F
Kerber ER, Green GJ (1980) Suppression of stem rust resistance in hexaploid wheat cv Canthach
by chromosome 7DL. Can J Bot 58:1347–1350
Kihara H (1919) Über cytologische studien bei einigen getreidearten. I. Species-bastarde des
weizens und weizenroggen-bastarde. Bot Mag 33:17–38
Kihara H (1924) Cytologische und genetische studien bei wichtigen getreidearten mit besonderer
rücksicht ouf das verhalten der chromosomen und die sterilitat in den bastarden. Mem Cell
Sci, Kyoto Imp University, B1: 1–200
Kihara H (1944) Discovery of the DD-analyser, one of the ancestors of Triticum vulgare. Agric
Hortic 19:13–14
Kihara H (1954) Considerations on the evolution and distribution of Aegilops species based on
the analyzer-method. Cytologia 19:336–357
Kihara H, Lilienfeld F (1949) A new synthesized 6x-wheat. In: Larsson GBaR (ed) Proceedings
of Eighth International Congress of Genetics, Stockholm, Sweden, 1949. Hereditas (Suppl),
pp 307–319
Kimber G, Sears ER (1987) Evolution in the genus Triticum and the origin of cultivated wheat.
In: Heyne EG (ed) Wheat and wheat improvement. American Society of Agronomy, Madison,
pp 154–164
Kislev ME (1980) Triticum parvicoccum sp. nov., the oldest naked wheat. Isr J Bot 28:95–107
Kraitshtein Z, Yaakov B, Khasdan V, Kashkush K (2010) Genetic and epigenetic dynamics of a
retrotransposon after allopolyploidization of wheat. Genetics 186(3):U801–U889
Levy AA, Feldman M (2004) Genetic and epigenetic reprogramming of the wheat genome upon
allopolyploidization. Biol J Linn Soc 82(4):607–613
Liu B, Segal G, Vega JM, Feldman M, Abbo S (1997) Isolation and characterization of
chromosome-specific DNA sequences from a chromosome arm genomic library of common
wheat. Plant J 11(5):959–965
Liu B, Vega JM, Feldman M (1998a) Rapid genomic changes in newly synthesized amphiploids
of Triticum and Aegilops. II. Changes in low-copy coding DNA sequences. Genome
41(4):535–542
Liu B, Vega JM, Segal G, Abbo S, Rodova H, Feldman M (1998b) Rapid genomic changes in
newly synthesized amphiploids of Triticum and Aegilops. I. Changes in low-copy noncoding
DNA sequences. Genome 41(2):272–277
Lucas H, Moore G, Murphy G, Flavell RB (1992) Inverted repeats in the long-terminal repeats of
the wheat retrotransposon wis 2–1A. Mol Bio Evol 9(4):716–728
Lukens LN, Pires JC, Leon E, Vogelzang R, Oslach L, Osborn T (2006) Patterns of sequence loss
and cytosine methylation within a population of newly resynthesized Brassica napus
allopolyploids. Plant Physiol 140(1):336–348
Lynch M, Force A (2000) The probability of duplicate gene preservation by subfunctionalization.
Genetics 154:459–473
Ma XF, Fang P, Gustafson JP (2004) Polyploidization-induced genome variation in triticale.
Genome 47(5):839–848
Ma XF, Gustafson JP (2005) Genome evolution of allopolyploids: a process of cytological and
genetic diploidization. Cytogenet Genome Res 109(1–3):236–249
Ma XF, Gustafson JP (2006) Timing and rate of genome variation in triticale following
allopolyploidization. Genome 49(8):950–958
Maan SS (1977) Fertility of amphiploids in Triticinae. J Heredity 68:87–94
Mac Key J (1954) Mutation breeding in polyploid cereals. Acta Agriculturae Scandinavica
4:549–557
Mac Key J (1958) Mutagenic response in Triticum at different lrvels of ploidy. In: Jenkins CB
(ed) Proceedings 1st
Mac Key J (1966) Species relationship in Triticum. Proceedings 2nd International Wheat
Genetics Symposium, Lund 1963, Hereditas Suppl. 2, pp 237–276
Madlung A, Masuelli RW, Watson B, Reynolds SH, Davison J, Comai L (2002) Remodeling of
DNA methylation and phenotypic and transcriptional changes in synthetic Arabidopsis
allotetraploids. Plant Physiol 129(2):733–746
CO
RR
737
738
739
740
741
742
743
744
745
746
747
748
749
750
751
752
753
754
755
756
757
758
759
760
761
762
763
764
765
766
767
768
769
770
771
772
773
774
775
776
777
778
779
780
781
782
783
784
785
786
787
788
789
790
M. Feldman et al.
UN
Editor Proof
132
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 133/134
133
EC
TE
D
PR
OO
F
Madlung A, Tyagi AP, Watson B, Jiang HM, Kagochi T, Doerge RW, Martienssen R, Comai L
(2005) Genomic changes in synthetic Arabidopsis polyploids. Plant J 41(2):221–230
Maestra B, Naranjo T (1999) Structural chromosome differentiation between Triticum
timopheevii and T-turgidum and T-aestivum. Theor Appl Genet 98(5):744–750
McFadden ES, Sears ER (1944) The artificial synthesis of Triticum spelta. Records Genet Soc
Amer 13:26–27
McFadden ES, Sears ER (1946) The origin of Triticum spelta and its free-threshing hexaploid
relatives. J Heredity 37(81–89):107–116
Mestiri I, Chague V, Tanguy AM, Huneau C, Huteau V, Belcram H, Coriton O, Chalhoub B,
Jahier J (2010) Newly synthesized wheat allohexaploids display progenitor-dependent meiotic
stability and aneuploidy but structural genomic additivity. New phytol 186(1):86–101
Mitra R, Bhatia C (1971) Isoenzymes and polyploidy. 1. Qualitative and quantitative isoenzyme
studies in the Triticinae. Genet Res Camb 18:57–69
Mochida K, Kawaura K, Shimosaka E, Kawakami N, Shin-I T, Kohara Y, Yamazaki Y, Ogihara
Y (2006) Tissue expression map of a large number of expressed sequence tags and its
application to in silico screening of stress response genes in common wheat. Mol Genet
Genomics 276(3):304–312
Morris R, Sears ER (1967) The cytogenetics of wheat and its relatives. In: Quisenberry KS, Reitz
LP (eds) Wheat and wheat improvement. Madison, U.S.A., pp 19–87
Nigumann P, Redik K, Matlik K, Speek M (2002) Many human genes are transcribed from the
antisense promoter of L1 retrotransposon. Genomics 79(5):628–634
Ohno S (1970) Evolution by gene duplication. Springer, Berlin
Okamoto M, Inomata N (1974) Possibility of 5B-like effect in diploid species. Wheat Inform Serv
38:15–16
Ozkan H, Levy AA, Feldman M (2001) Allopolyploidy-Induced rapid genome evolution in the
wheat (Aegilops-Triticum) group. Plant Cell 13:1735–1747
Ozkan H, Tuna M, Arumuganathan K (2003) Nonadditive changes in genome size during
allopolyploidization in the wheat (Aegilops-Triticum) group. J Hered 94(3):260–264
Parisod C, Alix K, Just J, Petit M, Sarilar V, Mhiri C, Ainouche M, Chalhoub B, Grandbastien
MA (2010) Impact of transposable elements on the organization and function of allopolyploid
genomes. New Phytol 186(1):37–45
Parisod C, Salmon A, Zerjal T, Tenaillon M, Grandbastien MA, Ainouche M (2009) Rapid
structural and epigenetic reorganization near transposable elements in hybrid and allopolyploid genomes in Spartina. New Phytol 184(4):1003–1015
Peng I, Ronin Y, Fahima T, Röder MS, Li Y, Nevo E, Korol A (2003a) Genomic distribution of
domestication QTLs in wild emmer wheat, Triticum dicoccoides. In Proceedings 10th
International Wheat Genetics Symposium, Paestum, Italy, pp 34–37
Peng JH, Ronin Y, Fahima T, Roder MS, Li YC, Nevo E, Korol A (2003b) Domestication
quantitative trait loci in Triticum dicoccoides, the progenitor of wheat. Proc Natl Acad Sci U S
A 100(5):2489–2494
Percival J (1921) The wheat plant. E.P. Dutton and Company, New York, pp 1–463
Petit M, Guidat C, Daniel J, Denis E, Montoriol E, Bui QT, Lim KY, Kovarik A, Leitch AR,
Grandbastien MA, Mhiri C (2010) Mobilization of retrotransposons in synthetic allotetraploid
tobacco. New Phytol 186(1):135–147
Pikaard CS (2000) The epigenetics of nucleolar dominance. Trends Genet 16(11):495–500
Prince VE, Pickett FB (2002) Splitting pairs: the diverging fates of duplicated genes. Nat Rev
Genet 3:827–837
Puig M, Caceres M, Ruiz A (2004) Silencing of a gene adjacent to the breakpoint of a widespread
Drosophila inversion by a transposon-induced antisense RNA. P Natl Acad Sci U S A
101(24):9013–9018
Pumphrey M, Bai J, Laudencia-Chingcuanco D, Anderson O, Gill BS (2009) Nonadditive
expression of homoeologous genes is established upon polyploidization in hexaploid wheat.
Genetics 181(3):1147–1157
CO
RR
791
792
793
794
795
796
797
798
799
800
801
802
803
804
805
806
807
808
809
810
811
812
813
814
815
816
817
818
819
820
821
822
823
824
825
826
827
828
829
830
831
832
833
834
835
836
837
838
839
840
841
842
843
UN
Editor Proof
7 Genomic Plasticity in Polyploid Wheat
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 134/134
EC
TE
D
PR
OO
F
Rapp RA, Udall JA, Wendel JF (2009) Genomic expression dominance in allopolyploids. Bmc
Biol 7
Sabot F, Guyot R, Wicker T, Chantret N, Laubin B, Chalhoub B, Leroy P, Sourdille P, Bernard M
(2005) Updating of transposable element annotations from large wheat genomic sequences
reveals diverse activities and gene associations. Mol Genet Gen 274(2):119–130
Sakamura T (1918) Kurze mitteilung über die chromosomenzahalen und die verwandtschaftsverhältnisse der Triticum Arten. Bot Mag 32(1918):151–154
Salina EA, Numerova OM, Ozkan H, Feldman M (2004) Alterations in subtelomeric tandem
repeats during early stages of allopolyploidy in wheat. Genome 47(5):860–867
Salmon A, Ainouche ML, Wendel JF (2005) Genetic and epigenetic consequences of recent
hybridization and polyploidy in Spartina (Poaceae). Mol Ecol 14(4):1163–1175
Sax K (1927) Chromosome behavior in Triticum hybrids, Verhandlungen des V Int. Kongresses
für Vererbungswissenchaft, Berlin, 2:1267–1284
Schulz A (1913) Die geschichte der kultivierten getreide. Nebert, Halle
Sears ER (1972) The nature of mutation in hexaploid wheat. Symp Biol Hung 12:73–82
Sears ER (1976) Genetic control of chromosome pairing in wheat. Annu Rev Genet 10:31–51
Shaked H, Kashkush K, Ozkan H, Feldman M, Levy AA (2001) Sequence elimination and
cytosine methylation are rapid and reproducible responses of the genome to wide
hybridization and allopolyploidy in wheat. Plant Cell 13:1749–1759
Stebbins GLJ (1980) Polyploidy in plants: unsolved problems and prospect, in polyploidy—
biological relevance. In: Lewis WH (ed) Plenum Press, New York
Stebbins GLJ (1971) Chromosomal evolution in higher plants. Addison-Wesley, New York
Stephens SG (1951) Possible significance of duplication in evolution. Adv Genet 4:247–265
Thompson DA, Desai MM, Murray AW (2006) Ploidy controls the success of mutators and
nature of mutations during budding yeast evolution. Curr Biol: CB 16(16):1581–1590
Tirosh I, Reikhav S, Levy AA, Barkai N (2009) A yeast hybrid provides insight into the evolution
of gene expression regulation. Science 324(5927):659–662
Van Slageren MW (1994) Wild wheats: a monograph of Aegilops L. and Amblyopyrum (Jaub.
and Spach) Eig (Poaceae). Agricultural University, Wageningen, The Netherlands
Veitia RA, Bottani S, Birchler JA (2008) Cellular reactions to gene dosage imbalance: genomic,
transcriptomic and proteomic effects. Trends Genet 24(8):390–397
Vicient CM, Jaaskelainen MJ, Kalendar R, Schulman AH (2001) Active retrotransposons are a
common feature of grass genomes. Plant Physiol 125(3):1283–1292
Von Tschermak E, Bleier H (1926) Über fruchtbare Aegilops-weizenbastarde, der deutsch. Bot
Ges 44:110–132
Waines JG (1976) A model for the origin of diploidizing mechanisms in polyploid species. Amer
Natur 110:415–430
Wang JB, Xu YH, Zhong L, Wu XM, Fang XP (2009) Rapid alterations of gene expression and
cytosine methylation in newly synthesized Brassica napus allopolyploids. Planta
229(3):471–483
Weissmann S, Feldman M, Gressel J (2005) Sequence evidence for sporadic intergeneric DNA
introgression from wheat into a wild Aegilops species. Mol Biol Evol 22:2055–2062
Wicker T, Mayer KFX, Gundlach H, Martis M, Steuernagel B, Scholz U, Simkova H,
Kubalakova M, Choulet F, Taudien S, Platzer M, Feuillet C, Fahima T, Budak H, Dolezel J,
Keller B, Stein N (2011) Frequent gene movement and pseudogene evolution is common to
the large and complex genomes of wheat, barley, and their relatives. Plant Cell
23(5):1706–1718
Yaakov B, Kashkush K (2011a) Massive alterations of the methylation patterns around DNA
transposons in the first four generations of a newly formed wheat allohexaploid. Genome
54(1):42–49
Yaakov B, Kashkush K (2011b) Methylation, transcription, and rearrangements of transposable
elements in synthetic allopolyploids. Int J Plant Genomics. doi:10.1155/2011/569826
Zhang ZC, Belcram H, Gornicki P, Charles M, Just J, Huneau C, Magdelenat G, Couloux A,
Samain S, Gill BS, Rasmussen JB, Barbe V, Faris JD, Chalhoub B (2011) Duplication and
CO
RR
844
845
846
847
848
849
850
851
852
853
854
855
856
857
858
859
860
861
862
863
864
865
866
867
868
869
870
871
872
873
874
875
876
877
878
879
880
881
882
883
884
885
886
887
888
889
890
891
892
893
894
895
896
897
M. Feldman et al.
UN
Editor Proof
134
Layout: T1 Standard SC
Chapter No.: 7
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 135/134
135
EC
TE
D
PR
OO
F
partitioning in evolution and function of homoeologous Q loci governing domestication
characters in polyploid wheat. P Natl Acad Sci U S A 108(46):18737–18742
Zhao N, Zhu B, Li M, Wang L, Xu L, Zhang H, Zheng S, Qi B, Han F, Liu B (2011) Extensive
and heritable epigenetic remodeling and genetic stability accompany allohexaploidization of
wheat. Genetics. doi:10.1534/genetics.111.127688
Zohary D, Feldman M (1962) Hybridization between amphiploids and the evolution of polyploids
in the wheat (Aegilops-Triticum) group. Evolution 16:44–61
CO
RR
898
899
900
901
902
903
904
UN
Editor Proof
7 Genomic Plasticity in Polyploid Wheat
Metadata of the chapter that will be visualized in
SpringerLink
Book Title
Polyploidy and Genome Evolution
Series Title
Chapter Title
Maize (Zea Mays) as a Model for Studying the Impact of Gene and Regulatory Sequence Loss Following
Whole-Genome Duplication
Copyright Year
2012
Copyright HolderName
Springer-Verlag Berlin Heidelberg
Corresponding Author
Family Name
Freeling
Particle
Given Name
Michael
Suffix
Author
Division
Department of Plant and Microbial Biology
Organization
University of California
Address
Berkeley, USA
Email
freeling@uclink.berkeley.edu
Family Name
Schnable
Particle
Given Name
James C.
Suffix
Division
Department of Plant and Microbial Biology
Organization
University of California
Address
Berkeley, USA
Email
Abstract
Modern maize (2n = 20) is functionally diploid, and its chromosomes pair normally, forming 10 bivalents
during meiosis. Sufficient genomic rearrangement has occurred that no two maize chromosomes are
homologous across their entire lengths. Yet comparisons of genetic maps, duplicate gene sequences, and later
genome assemblies revealed maize is descended from a polyploid ancestor which lived 5–12 million years
ago. In the time since that polyploid ancestor lived 8,000–9,000 genes conserved at syntenic positions in other
grass species have been reduced to single copy in maize while 4,000–5,000 genes are still retained as
homologous gene pairs. The consequences of this polyploidy are continuing to resolve in modern maize
accessions. With a wide range of data sets generated by an active research community, maize is an unparalleled
model for the in silico study of the changes in genome structure, gene content, and gene regulation that a
successful polyploidy brings about in a plant lineage.
1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 137/144
Chapter 8
6
James C. Schnable and Michael Freeling
11
12
13
14
15
16
17
18
19
22
23
24
25
26
27
CO
RR
20
21
PR
OO
9
10
Abstract Modern maize (2n = 20) is functionally diploid, and its chromosomes
pair normally, forming 10 bivalents during meiosis. Sufficient genomic rearrangement has occurred that no two maize chromosomes are homologous across
their entire lengths. Yet comparisons of genetic maps, duplicate gene sequences,
and later genome assemblies revealed maize is descended from a polyploid
ancestor which lived 5–12 million years ago. In the time since that polyploid
ancestor lived 8,000–9,000 genes conserved at syntenic positions in other grass
species have been reduced to single copy in maize while 4,000–5,000 genes are
still retained as homologous gene pairs. The consequences of this polyploidy are
continuing to resolve in modern maize accessions. With a wide range of data sets
generated by an active research community, maize is an unparalleled model for the
in silico study of the changes in genome structure, gene content, and gene regulation that a successful polyploidy brings about in a plant lineage.
D
7
8
TE
4
EC
3
F
5
Maize (Zea Mays) as a Model for Studying
the Impact of Gene and Regulatory
Sequence Loss Following Whole-Genome
Duplication
2
8.1 Background on the Maize Polyploidy
Suspicion of a polyploid origin for maize first came from the prevalence of
duplicate mutants identified at unlinked locations throughout the genome; these
have been observed since the early days of maize genetics (Rhoades 1951). Over
time, the evidence that the maize lineage was descended from an ancient polyploid
grew to include the arrangement of duplicate genes and markers in similar orders
on multiple maize chromosomes reported in a number of genetic maps developed
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: 8
J. C. Schnable M. Freeling (&)
Department of Plant and Microbial Biology, University of California, Berkeley, USA
e-mail: freeling@uclink.berkeley.edu
P. S. Soltis and D. E. Soltis (eds.), Polyploidy and Genome Evolution,
DOI: 10.1007/978-3-642-31442-1_8, Springer-Verlag Berlin Heidelberg 2012
137
Layout: T1 Standard SC
Chapter No.: 8
Book ID: 272454_1_En
Date: 16-8-2012
J. C. Schnable and M. Freeling
42
8.2 Timing of the Maize Polyploidy
37
38
39
40
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
61
62
63
64
65
66
67
68
PR
OO
35
36
D
34
Unlike other important crops species, such as bread wheat and potato, the maize
polyploidy was not directly associated with domestication, but occurred millions
of years earlier. Analysis of divergence between duplicate genes on opposite maize
subgenomes—known as homeologs—has placed the split of the two progenitor
genomes found within modern maize at *12 million years before present (Swigoňová et al. 2004). The date of divergence between the two subgenomes of maize
does not necessarily reflect how long ago the actual event of polyploidization
occurred, as fertile allopolyploids can form between related species that have been
evolving independently for millions of years. However, based on a single case of
gene conversion it can be concluded that the two subgenomes of maize have
shared a single nucleus for at least 5 million years (Swigoňová et al. 2004).
Recent evidence has allowed the phylogenetic placement of the maize tetraploidy to be narrowed to a discrete interval within the diversification of the Andropogoneae—the tribe of grasses within which maize is placed (Mathews et al.
2002). Phylogenetic trees of the homologous genes zfl1 and zfl2 show that these
whole genome duplicates had already diverged in the common ancestor of the
genus Zea (maize and teosinte) and the sister genus Tripsacum (Bomblies and
Doebley 2005). Comparison of genetic maps in maize and sorghum prior to the
publication of complete genome sequences for these species demonstrated that
sorghum—and by extension other relatives within the ‘‘core’’ Andropogoneae—
did not share the maize polyploidy (Wei et al. 2007). Following the publication of
complete genome assembles for both maize (Schnable et al. 2009) and sorghum
(Paterson et al. 2009), a study of thousands of maize homologous gene pairs found
that both maize subgenomes appear equally diverged from sorghum (Woodhouse
et al. 2010). Given these constraints, the phylogenetic placement of the maize
tetraploidy can be inferred (Fig. 8.1).
TE
33
EC
31
32
CO
RR
30
F
41
during the 1980s (Goodman et al. 1980; Wendel et al. 1986; Helentjaris et al.
1988). With the availability of abundant sequence data for cloned duplicate genes,
it became possible to definitively classify maize as an ancient polyploid (Gaut and
Doebley 1997).
In the absence of diploid species showing higher genetic similarity to one maize
subgenome or the other, it will be impossible to conclusively prove maize is not an
ancient autopolyploid. The most closely related genus to Zea, Tripsacum, is descended from the same polyploid ancestor (Bomblies and Doebley 2005). The most
closely related species with a sequenced genome is Sorghum bicolor, which shows
an equal divergence from both subgenomes of maize (Swigoňová et al. 2004;
Woodhouse et al. 2010). Almost no molecular data are available for the most
closely related, and apparently diploid, genera to maize, Elionurus (13 sequences
in GenBank) and Coelorachis (14 sequences in GenBank). The question of autoversus allo-polyploidy may never be conclusively answered for maize.
28
29
UN
Editor Proof
138
Book ISBN: 978-3-642-31441-4
Page: 138/144
Layout: T1 Standard SC
Chapter No.: 8
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 139/144
139
PR
OO
F
Editor Proof
8 Maize (Zea Mays) as a Model for Studying the Impact of Gene
Fig. 8.1 A highly pruned tree of grass species. Branches are scaled by modal synonymous
substitutions per site among all syntenic orthologs as measured by SynMap and QuotaAlign
(Lyons et al. 2008b; Tang et al. 2011). The position of the branch between Zea and Tripsacum is
only approximated, as the genome of Tripsacum has not yet been sequenced
73
74
75
76
77
78
79
80
81
82
83
84
85
86
87
88
89
90
91
92
93
D
TE
72
While all plant genomes sequenced to date have experienced at least one ancient
polyploidization (Paterson et al. 2010), in the vast majority of cases, duplicate
regions have been too heavily rearranged to reconstruct which regions can be
grouped together as originating from the same parental genome. Such was not the
case with maize. The polyploid ancestor of maize possessed 20 chromosomes, 2
equivalent to each of the 10 chromosomes of sorghum. Researchers could assign
segments of the ten chromosomes of modern maize back to each of these ancestral
chromosomes even before the completion of either the sorghum or maize genomes
(Wei et al. 2007). With the publication of the completed maize and sorghum
genomes, the same reconstruction can now be carried out using web-based tools by
anyone with a fondness for puzzles (Fig. 8.2).
A cursory examination of Fig. 8.2 will reveal for any position in the sorghum
genome there are two syntenic orthologous regions in maize, while for any
position in the maize genome there is only one syntenic orthologous region in
sorghum. In no case are the two regions orthologous to the same region in sorghum
are present on the same maize chromosome. In 15 of the 20 inferred ancestral
chromosomes, all segments are contained within a single chromosome of modern
maize. In the remaining five cases, orthologous regions belonging to a single
inferred ancestral maize chromosome are split between two—or in one case
three—modern maize chromosomes. Fusion of chromosomes has often resulted
from the insertion of one chromosome in between the arms of another—often
linked with inversions of whole chromosome arms—as previously observed in the
reduction of the Brachypodium distachyon genome to five chromosomes from the
EC
71
CO
RR
70
8.3 Changes in Genome Arrangement Following Polyploidy
in Maize
UN
69
Layout: T1 Standard SC
Chapter No.: 8
Book ID: 272454_1_En
Date: 16-8-2012
J. C. Schnable and M. Freeling
TE
D
PR
OO
F
Editor Proof
140
Book ISBN: 978-3-642-31441-4
Page: 140/144
CO
RR
EC
Fig. 8.2 A dotplot generated using SynMap (Lyons et al. 2008b) to compare the arrangement of
orthologous regions of the sorghum (x-axis) and maize (y-axis) genome. Each dot represents a
pair of homologous genes in maize and sorghum within a syntenic region, and each dot is color
coded by synonymous substitutions per site between those two genes. In general, purple lines
represent syntenic orthologous regions between the maize and sorghum genomes whilegreen
diagonals represent homologous syntenic regions from a more ancient whole-genome duplication
shared by all grass species. An interactive version of this graphic can be regenerated using the
following link: http://genomevolution.org/r/3vpl
99
8.4 Ancient Gene Loss in Maize
95
96
97
100
101
102
UN
98
inferred ancestral number of 12 (The International Brachypodium Initiative 2010).
Researchers can search for differences across whole pairs of reconstructed maize
ancestral chromosomes, and, by using differential gene loss as a mark to distinguish chromosome copies, even compare entire subgenomes (Schnable et al.
2011).
94
Being able to compare pairs of complete ancestral chromosomes enabled a thorough investigation of an odd observation reported for Arabidopsis paleopolyploidy: the loss of genes was biased between homologous regions of the genome
Layout: T1 Standard SC
Chapter No.: 8
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 141/144
141
135
8.5 Ongoing Gene Loss in Maize
109
110
111
112
113
114
115
116
117
118
119
120
121
122
123
124
125
126
127
128
129
130
131
132
133
136
137
138
139
140
141
142
143
PR
OO
108
D
107
TE
106
EC
105
CO
RR
104
F
134
(‘‘biased fractionation’’), with one copy of each region retaining more total genes
than the other (Thomas et al. 2006). Depending on the definition of a functional
gene, between 4,000 and 5,000 homologous gene pairs are retained within the
maize genome (Schnable et al. 2009; Woodhouse et al. 2010). Another
8,000–9,000 genes conserved at syntenic location in both rice and sorghum have
fractionated back to a single copy in maize (Schnable et al. 2011). The same
fractionation bias was observed across each of the ten pairs of ancestral maize
chromosomes, with one chromosome copy retaining 70–90 % of the ancestral
grass gene content and the other chromosome copy retaining only 40–60 %. By
grouping high gene loss ancestral chromosomes together into one parental subgenome and low gene loss ancestral chromosomes into the other parental subgenome and using the wealth of maize RNA-seq data being produced by other
research projects, it was shown that genes on the subgenome with low gene loss,
referred to as maize1, tend to be expressed at higher levels than their duplicates on
the subgenome with high gene loss, or maize2 (Schnable et al. 2011). This
inequality of expression between parental subgenomes has previously been
reported in allopolyploid cotton—among other species—where it was termed
‘‘genomic dominance’’ (Flagel and Wendel 2010).
The correlation between bias in gene expression levels and bias in gene loss
rates suggests a simple explanation for the bias in gene loss rates observed in all
ancient polyploid species studied to date (Sankoff et al. 2010). The deletion of the
gene copy that contributed the majority of total gene pair expression should be
more likely to have a functional impact than the deletion of the gene copy which
contributes the minority of total gene pair expression. Therefore, even if the base
rate of gene deletion is equal on both subgenomes, null alleles of more highly
expressed gene copies would be more likely to be purged from the population by
purifying selection. Over time, deletion of redundant gene copies would tend to
cluster on the less expressed subgenome. In support of this hypothesis, it was noted
that known maize morphological mutants are significantly more likely to result
from the disruption of genes on the high-expression subgenome than expected
given the overall distribution of expressed genes between the subgenomes (Schnable and Freeling 2011).
103
In addition to ancient fixed patterns of gene loss, fractionation continues within
maize today, with genes present in some maize inbreds lost in other accessions.
Concurrent with the publication of the maize genome, the first report emerged
describing a high incidence of presence–absence variation (PAV) between different
maize inbreds. A comparison of two of the most-studied maize lines, B73 and
Mo17 (the former being the line used to generate the reference genome), determined that thousands of sequences were present in the former but entirely absent
from the latter, including at least 180 single-copy genes (Springer et al. 2009).
UN
Editor Proof
8 Maize (Zea Mays) as a Model for Studying the Impact of Gene
Layout: T1 Standard SC
Chapter No.: 8
Book ID: 272454_1_En
Date: 16-8-2012
J. C. Schnable and M. Freeling
160
8.6 Sequence Deletion in Maize
151
152
153
154
155
156
157
158
161
162
163
164
165
166
167
168
169
170
171
172
173
174
175
176
177
178
179
180
181
182
183
PR
OO
150
D
149
TE
147
148
The loss of genes is the result of the deletion of sequence from the genome. Given
the extensive observation of chromosomal mispairing, rearrangements, and partial
or complete aneuploidy in synthetic and recent natural polyploids, it might be
expected that much duplicate gene loss is the result of large deletions which
remove whole chromosomal segments from one subgenome. The fact that only a
portion of the genomes of ancient polyploid species such as rice—65.7 % of the
genes covered by duplicated blocks (Yu et al. 2005)—and Arabidopsis—89 % of
genes covered (Bowers et al. 2003)—would seem consistent with this expectation.
However, the pattern of gene loss in maize is not consistent with large deletions
following whole-genome duplication. A search for regions of the sorghum genome
represented by only one syntenic orthologous region within maize revealed only
one putative deletion of [30 genes (Schnable et al. 2012). A comparison of the
patterns of gene loss observed in duplicate regions of the maize genome to simulations assuming different lengths of sequence deletion found that the pattern
observed is consistent with [85 % of deletions removing only a single gene, or a
portion there of, with the remainder of deletions removing two, or occasionally
three, adjacent genes (Woodhouse et al. 2010). The present genome of maize is
littered with the partially deleted fragments of homologous genes (Fig. 8.3a).
The prevalence of short direct repeats flanking deletions within these fragmentary genes indicates the loss of genes in polyploids is a result of intrastrand
non-homologous recombination: short direct repeats found by chance throughout
the genome pair with each other, splicing out any intervening sequence, and
leaving a single copy of the repeat sequence (Woodhouse et al. 2010).
EC
146
CO
RR
145
F
159
A follow-up study which examined 33 inbreds and accessions of wild teosinte
identified 3,410 high-confidence genes in the genome of B73 which had been lost in
one or more of these lines—a total that represents more than 10 % of all highconfidence genes present in the maize genome (Swanson-Wagner et al. 2010).
While most PAV in maize was initially observed using comparative genomic
hybridization to microarrays, the same variation in gene content was observed in a
recent study that resequenced six inbreds using Illumina short read technology
(Lai et al. 2010).
While many genes which exhibit PAVs between B73 and other maize inbreds
possess no syntenic orthologs in other grass species and may have recently inserted
into their current locations, 4–6 % of maize genes with syntenic orthologs in both
sorghum and rice—indicating these genes have been functionally conserved for at
least 50 million years—are also involved in PAV between diverse maize lines
(Schnable et al. 2011). Among genes where both homologous are still retained in
B73, genes on the ‘high-gene-loss, low-expression’ subgenome (maize2) are more
likely to be involved in PAV (Swanson-Wagner et al. 2010; Schnable et al. 2011).
144
UN
Editor Proof
142
Book ISBN: 978-3-642-31441-4
Page: 142/144
Layout: T1 Standard SC
Chapter No.: 8
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 143/144
143
PR
OO
F
Editor Proof
8 Maize (Zea Mays) as a Model for Studying the Impact of Gene
TE
D
Fig. 8.3 a GEvo panel (Lyons et al. 2008a) illustrating an example of a gene conserved in rice,
sorghum, and one of two maize subgenomes (maize1), while on the second maize subgenome
(maize 2) only the 5’ and 3’ ends of the gene remain conserved. The colored boxes mark regions
of sequence similarity, as determined by blastn, between the orthologous genomic regions in this
figure. An interactive version of the same panel can be regenerated at the following link: http://
genomevolution.org/r/3vbu. b A second GEvo panel comparing a single sorghum gene to its two
coorthologs in maize. Purple rectangles represent functionally constrained noncoding sequences
identified by the comparison of this gene to its syntenic ortholog in rice. The red box highlights a
cluster of conserved noncoding sequences conserved upstream of one maize gene but lost from
the promoter of the other. c A comparison of the relative expression of these two maize genes in a
wide range of tissues using RNA-seq data produced by a number of maize research groups (Wang
et al. 2009; Jia et al. 2009; Li et al. 2010; Davidson et al. 2011; Waters et al. 2011)
191
8.7 Future Prospects
187
188
189
192
193
194
195
196
197
198
199
CO
RR
186
The mechanism responsible for the unequal expression of duplicate genes from
different parents in polyploid species remains unknown. With an extensive collection
of well-characterized mutants, including knockouts of a variety of epigenetic
mechanisms, maize is an excellent system for the investigation of this inexplicable
behavior. If the whole-genome duplication in maize was indeed the result of allopolyploidy, the identification of a wild species more closely related to one maize
subgenome than the other remains an exciting possibility. The wealth of RNA-seq
data being generated in maize for unrelated purposes creates an opportunity to
UN
185
EC
190
Fractionation and sequence deletion are not confined to only the protein-coding
regions of genes. Sequence deletion, presumably by the same mechanism of intrastrand non-homologous recombination, can also remove conserved noncoding regulatory sequences from one of two duplicate copies of a maize gene (Fig. 8.3b).
These genes often show different patterns of tissue-specific expression in existing
maize RNA-seq data sets, allowing researchers to develop testable hypotheses about
the function of specific regulatory sequences (Fig. 8.3c) (Freeling et al. 2012).
184
Layout: T1 Standard SC
Chapter No.: 8
Book ID: 272454_1_En
Date: 16-8-2012
J. C. Schnable and M. Freeling
210
211
212
213
Acknowledgments We thank Vincent Li, a high school intern in the Freeling lab from Project
SEED, for identifying the gene fragment shown in Fig. 8.3 and Addie M. Thompson for critical
reading of an early version of this text. Funding provided by NSF Plant Genome Research
Program grant 0701871 to MF and a Chang-Lin Tien Graduate Fellowship to JCS.
214
References
215
216
217
218
219
220
221
222
223
224
225
226
227
228
229
230
231
232
233
234
235
236
237
238
239
240
241
242
243
244
245
Bomblies K, Doebley JF (2005) Molecular evolution of FLORICAULA/LEAFY orthologs in the
Andropogoneae (Poaceae). Mol Biol Evol 22:1082–1094
Bowers JE et al (2003) Unravelling angiosperm genome evolution by phylogenetic analysis of
chromosomal duplication events. Nature 422:433–438
Davidson RM et al (2011) Utility of RNA sequencing for analysis of maize reproductive
transcriptomes. Plant Genome 4:191–203
Flagel LE, Wendel JF (2010) Evolutionary rate variation, genomic dominance and duplicate gene
expression evolution during allotetraploid cotton speciation. New Phytol 186:184–193
Freeling M et al (2012) Fractionation mutagenesis and similar consequences of mechanisms
removing dispensable or less-expressed DNA in plants. Curr Opin Plant Biol, Advance Online
Publication
Gaut BS, Doebley JF (1997) DNA sequence evidence for the segmental allotetraploid origin of
maize. Proc Natl Acad Sci USA 94:6809–6814
Goodman MM et al (1980) Linkage relationships of 19 enzyme Loci in maize. Genetics 96:697–710
Helentjaris T et al (1988) Identification of the genomic locations of duplicate nucleotide sequences
in maize by analysis of restriction fragment length polymorphisms. Genetics 118:353–363
Jia Y et al (2009) Loss of RNA–dependent RNA polymerase 2 (RDR2) function causes
widespread and unexpected changes in the expression of transposons, genes, and 24-nt small
RNAs. PLoS Genet 5:e1000737
Lai J et al (2010) Genome-wide patterns of genetic variation among elite maize inbred lines. Nat
Genet 42:1027–1030
Li P et al (2010) The developmental dynamics of the maize leaf transcriptome. Nat Genet
42:1060–1067
Lyons E et al (2008a) Finding and comparing syntenic regions among Arabidopsis and the
outgroups papaya, poplar, and grape: CoGe with Rosids. Plant Physiol 148:1772–1781
Lyons E et al (2008b) The value of non-model genomes and an example using SynMap within
CoGe to dissect the hexaploidy that predates the Rosids. Trop Plant Biol 1:181–190
Mathews S et al (2002) Phylogeny of Andropogoneae inferred from phytochrome B, GBSSI, and
ndhF. Int J Plant Sci 163:441–450
Paterson AH et al (2010) Insights from the comparison of plant genome sequences. Annu Rev
Plant Biol 61:349–372
206
207
208
PR
OO
205
D
204
TE
203
EC
202
CO
RR
201
F
209
investigate how the expression patterns of homologous gene pairs in maize have
diverged in silico. Soon, it may also be possible to study how the functions of
duplicate genes have diverged in the 5–12 million years since the maize polyploidy in
silico, as high-resolution genome-wide associate studies begin to identify the loci
responsible for variation in a wide range of maize phenotypes. In addition to a higher
likelihood of being lost entirely, has the reduced importance of less expressed genes
on the non-dominant subgenome also given these genes greater freedom for innovation, even if their new role comes at the express of their ancestral function? We
predict that higher levels of neofunctionalization will be observed on the nondominant subgenome, but this hypothesis remains untested.
200
UN
Editor Proof
144
Book ISBN: 978-3-642-31441-4
Page: 144/144
Layout: T1 Standard SC
Chapter No.: 8
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 145/144
145
EC
TE
D
PR
OO
F
Paterson AH et al (2009) The sorghum bicolor genome and the diversification of grasses. Nature
457:551–556
Rhoades MM (1951) Duplicate genes in maize. Am Nat 85:105–110
Sankoff D et al (2010) The collapse of gene complement following whole genome duplication.
BMC Genomics 11:313
Schnable JC et al (2011) Differentiation of the maize subgenomes by genome dominance and
both ancient and ongoing gene loss. Proc Natl Acad Sci USA 108:4069–4074
Schnable JC et al (2012) Genome-wide analysis of syntenic gene deletion in the grasses. Genome
Biol Evol 4:265–277
Schnable JC, Freeling M (2011) Genes identified by visible mutant phenotypes show increased
bias toward one of two subgenomes of maize. PLoS ONE 6:e17855
Schnable PS et al (2009) The B73 maize genome: complexity, diversity, and dynamics. Science
326:1112–1115
Springer NM et al (2009) Maize inbreds exhibit high levels of copy number variation (CNV) and
presence/absence variation (PAV) in genome content. PLoS Genet 5:e1000734
Swanson-Wagner RA et al (2010) Pervasive gene content variation and copy number variation in
maize and its undomesticated progenitor. Genome Res 20:1689–1699
Swigoňová Z et al (2004) Close split of sorghum and maize genome progenitors. Genome Res
14:1916–1923
Tang H et al (2011) Screening synteny blocks in pairwise genome comparisons through integer
programming. BMC Bioinf 12:102
The International Brachypodium Initiative (2010) Genome sequencing and analysis of the model
grass Brachypodium distachyon. Nature 463:763–768
Thomas BC et al (2006) Following tetraploidy in an Arabidopsis ancestor, genes were removed
preferentially from one homologue leaving clusters enriched in dose-sensitive genes. Genome
Res 16:934–946
Wang X et al (2009) Genome-wide and organ-specific landscapes of epigenetic modifications and
their relationships to mRNA and small RNA transcriptomes in maize. Plant Cell 21:1053–1069
Waters AJ et al (2011) Parent-of-origin effects on gene expression and DNA methylation in the
maize endosperm. Plant Cell 23:4221–4233
Wei F et al (2007) Physical and genetic structure of the maize genome reflects its complex
evolutionary history. PLoS Genet 3:e123
Wendel JF et al (1986) Duplicated chromosome segments in maize (Zea mays L.): further
evidence from hexokinase isozymes. Theoret Appl Genet 72:178–185
Woodhouse MR et al (2010) Following tetraploidy in maize, a short deletion mechanism
removed genes preferentially from one of the two homeologs. PLoS Biol 8:e1000409
Yu J et al (2005) The Genomes of Oryza sativa: a history of duplications. PLoS Biol 3:e38
CO
RR
246
247
248
249
250
251
252
253
254
255
256
257
258
259
260
261
262
263
264
265
266
267
268
269
270
271
272
273
274
275
276
277
278
279
280
281
282
UN
Editor Proof
8 Maize (Zea Mays) as a Model for Studying the Impact of Gene
Metadata of the chapter that will be visualized in
SpringerLink
Book Title
Polyploidy and Genome Evolution
Series Title
Chapter Title
Polyploidy in Legumes
Copyright Year
2012
Copyright HolderName
Springer-Verlag Berlin Heidelberg
Corresponding Author
Family Name
Doyle
Particle
Given Name
Jeff J.
Suffix
Abstract
Division
Department of Plant Biology
Organization
Cornell University
Address
14850, Ithaca, NY, USA
Email
jjd5@cornell.edu
Legumes are the third largest family of flowering plants, with over 700 genera and more than 19,000 species.
Genomic evidence has shown that a whole-genome duplication (WGD) occurred shortly after the origin of
the family, in an ancestor that gave rise to the papilionoids, the clade that comprises 65 % of the genera and
71 % of the species, including nearly all of the economically important crop legumes. This polyploidy event
may have been associated with the origin of nitrogen-fixing symbiosis (nodulation) in the papilionoids.
Nodulation most likely evolved independently in other legumes outside the papilionoids, hence there appears
to be no requirement for polyploidy in the evolution of this important symbiosis. More recent polyploidy, as
inferred from chromosome counts, occurs in approximately a quarter of all legume genera for which data are
available. In most cases, polyploidy is confined to individual genera, species within genera, or cytotypes
within species. An exception is the core clade of the genistoid legumes, a major papilionoid group that includes
lupines (Lupinus). This group is probably fundamentally polyploid and also has a propensity for further
polyploidy and aneuploidy in many of its genera. The frequency of polyploidy varies considerably among
clades of the family, being most common (outside the genistoids) in the largely temperate, herbaceous
Hologalegina (including pea and clover), and low in woody tropical groups such as the caesalpinioids.
Book ID: 272454_1_En
Date: 16-8-2012
Chapter 9
2
Polyploidy in Legumes
3
Jeff J. Doyle
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
PR
OO
D
8
TE
7
EC
6
Abstract Legumes are the third largest family of flowering plants, with over 700
genera and more than 19,000 species. Genomic evidence has shown that a wholegenome duplication (WGD) occurred shortly after the origin of the family, in an
ancestor that gave rise to the papilionoids, the clade that comprises 65 % of the
genera and 71 % of the species, including nearly all of the economically important
crop legumes. This polyploidy event may have been associated with the origin of
nitrogen-fixing symbiosis (nodulation) in the papilionoids. Nodulation most likely
evolved independently in other legumes outside the papilionoids, hence there
appears to be no requirement for polyploidy in the evolution of this important
symbiosis. More recent polyploidy, as inferred from chromosome counts, occurs in
approximately a quarter of all legume genera for which data are available. In most
cases, polyploidy is confined to individual genera, species within genera, or cytotypes within species. An exception is the core clade of the genistoid legumes, a
major papilionoid group that includes lupines (Lupinus). This group is probably
fundamentally polyploid and also has a propensity for further polyploidy and
aneuploidy in many of its genera. The frequency of polyploidy varies considerably
among clades of the family, being most common (outside the genistoids) in the
largely temperate, herbaceous Hologalegina (including pea and clover), and low in
woody tropical groups such as the caesalpinioids.
CO
RR
4
5
F
1
Book ISBN: 978-3-642-31441-4
Page: 147/179
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: 9
J. J. Doyle (&)
Department of Plant Biology, Cornell University, Ithaca, NY 14850, USA
e-mail: jjd5@cornell.edu
P. S. Soltis and D. E. Soltis (eds.), Polyploidy and Genome Evolution,
DOI: 10.1007/978-3-642-31442-1_9, Springer-Verlag Berlin Heidelberg 2012
147
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 148/179
24
J. J. Doyle
9.1 Introduction
38
9.2 A Brief Overview of Legume Phylogeny
31
32
33
34
35
36
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
61
62
PR
OO
30
D
29
TE
28
Along with Polygalaceae, Surianaceae, and Quillajaceae, Leguminosae form the
order Fabales, one of eight-orders in the Fabidae clade of rosid eudicots (Wang
et al. 2009). Bello et al. (2009) suggested that the Fabales are the product of a rapid
radiation, with legumes probably sister to Surianaceae plus Quillajaceae.
The Leguminosae has been the focus of considerable phylogenetic study, culminating in solid, chloroplast based, working hypotheses of generic relationships
(Fig. 9.1), notably those of Wojciechowski et al. (2004) and Bruneau et al. (2008).
The older classification of the family into three subfamilies, Caesalpinioideae,
Mimosoideae, and Papilionoideae (sometimes treated as separate families), is not
supported by molecular phylogenetic studies, in that although the Mimosoideae
(mimosas, acacias) and Papilionoideae (pea, bean, soybean, etc.) are monophyletic, the former is embedded in one clade of a paraphyletic caesalpinioid grade.
Relationships at the base of the family are uncertain and differ among the studies
of Wojciechowski et al. (2004), which focused most heavily on Papilionoideae,
and Bruneau et al. (2008), which emphasized caesalpinioids and included few
papilionoids. However, both studies identified caesalpinioids as the earliestdiverging lineages, including such taxa as the tribe Cercideae, which includes
Cercis (the redbud or Judas tree) and the large genus, Bauhinia (orchid tree).
Relationships at the bases of the two monophyletic subfamilies are also uncertain.
Fossil evidence places the origin of the family in the Paleocene, around
60 million years ago (MYA; see Lavin et al. 2005 for discussion; see also Bell
et al. 2010). Divergence times of all major groups within the family have been
estimated from fossil-calibrated molecular data (Lavin et al. 2005; Bruneau et al.
2008) and suggest rapid diversification of many clades, such that within
EC
27
CO
RR
26
F
37
The legumes (Leguminosae or, less preferably, Fabaceae, according to Lewis et al.
2005) are the third largest family of flowering plants, and are tremendously diverse
ecologically, morphologically, chemically, and cytologically (Doyle and Luckow
2003; Lewis et al. 2005). Not surprisingly, the family is also cytologically diverse.
As in other families, polyploidy is implicated as a major force at all levels of
legume evolution, from the early stages of radiation in the family to the origin and
recent diversification of modern genera, such as Glycine (soybean and allies) and
species within genera, such as the Medicago sativa complex (alfalfa and allies).
After summarizing progress in understanding the phylogeny of the family, this
review will discuss the role of paleopolyploidy during the early stages of the
radiation of the entire family and the possible connections with nodulation. The
occurrence of polyploidy in each of the major clades of the family will then be
reviewed.
25
UN
Editor Proof
148
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 149/179
Editor Proof
9 Polyploidy in Legumes
149
x = 14
} Early-diverging
papilionoid lineages
PR
OO
polyploidy
(to x= 28)
Genistoids x = 9
Dalbergioids x = 10
Baphioids x = 11
Hypocalyptus
Mirbelioids x = 9
H-Loteae x = 7, 8
H-Robinieae x = 10, 11
H-IRLC x = 8
Indigofereae x = 7, 8
Millettioids x = 10, 11
F
Cercideae x = 7 or 14
Detarieae x = 12
Duparquetia
Dialiinae x = 14
MCC clade
(incl. Mimosoideae) x = 14
(incl. Phaseoleae)
60
55
50
million years
TE
D
Fig. 9.1 Phylogeny of legumes. Caesalpinioids are shown in green, papilionoids in black.
Relationships among caesalpinoid lineages are shown as unresolved due to conflict among
published studies. Papilionoid taxa marked ‘‘H’’ are members of the Hologalegina. Divergence
dates for the origins of major clades are from Lavin et al. (2005). Base chromosome numbers are
given for groups with published counts. Minimum (solid arrow) and maximum (dashed arrow)
dates for the papilionoid polyploidy event are shown
66
9.3 Polyploidy and the Early Diversification of Legumes
68
69
70
71
72
73
74
75
CO
RR
67
To what degree has polyploidy shaped the radiation of legumes? Given the
growing understanding that polyploidy can drive phenotypic diversification (e.g.,
Freeling et al. 2006) and has played a role in the preservation of lineages during
periods of extinction (Fawcett et al. 2009), it might be expected that polyploidy
would be an important feature of evolution in a family that is ‘‘successful’’ as
judged by its sheer size and ecological dominance in some tropical biomes (e.g.,
rain forests, woody savannas, and dry forests), and in which a significant adaptive
novelty—the symbiotic association with nitrogen-fixing soil bacteria, termed
nodulation—has arisen (e.g., Doyle 2011).
UN
64
EC
65
10 million years of divergence from a common ancestor, all of the major lineages
in the family had evolved, including the two monophyletic subfamilies and all of
the major clades within the Papilionoideae.
63
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 150/179
82
83
84
85
86
87
88
89
90
91
92
93
94
95
96
97
98
99
100
101
102
103
104
105
106
107
108
109
110
111
112
113
114
115
116
117
118
F
81
PR
OO
80
D
79
Until the advent of genomic data, chromosome number was the prime source of
information available for inferring the existence of polyploidy. Goldblatt’s (1981)
review of the distribution of chromosome numbers in Leguminosae, published in
Advances in Legume Systematics, Part 2, remains the most comprehensive treatment of chromosomal variation in the family, and includes hypotheses concerning
the base numbers and ploidy levels of its constituent subfamilies and tribes. The
information from which his summary was drawn was included in the descriptions
of genera in Advances in Legume Systematics, Part 1 (Polhill and Raven 1981),
known to researchers in the family as the legume ‘‘bible.’’ The taxonomic treatments provided by this key resource were recently updated in Legumes of the
World (Lewis et al. 2005), taking into account the rapid progress in legume
phylogenetics. The phylogenetic studies that have revolutionized our understanding of relationships within the family also provide a new phylogenetic context for understanding chromosome number evolution that was not available
previously, but unfortunately the otherwise excellent Legumes of the World does
not include any cytological information.
The key contribution of objective phylogenetic data to our understanding of
cytological evolution in the family is the confirmation of caesalpinioid legumes as
a grade rather than as a natural subfamily. Chromosome numbers for the major
clades that comprise the caesalpinioid grade are relatively constant, principally
2n = 24–28 (Fig. 9.1). Standing out from these higher chromosome numbers are
Chamaecrista and Cercis. The large, mostly, pantropical genus, Chamaecrista, is
cytologically complex, with 2n = 14, 16, and 28. Goldblatt (1981) considered its
lower numbers to be the products of aneuploid reduction, and this hypothesis has
been recently supported (Torres et al. 2011, see below). Phylogenetic studies
(Wojciechowski et al. 2004; Bruneau et al. 2008) now nest Chamaecrista and
other Cassieae s.s. within the Mimosoideae-Caesalpinioideae-Cassieae (MCC)
clade (2n = 28), supporting this hypothesis.
Cercis is a small genus (10 species) with disjunct worldwide distribution and
2n = 14. It is a member of the Cercideae, all other members of which are
2n = 28, including the large pantropical genus, Bauhinia s.l. (ca. 250 species).
Phylogenetic studies show that Cercis is sister to the remaining genera (Bruneau
et al. 2008), which may be consistent with Goldblatt’s (1981) conclusion that it is
diploid and the remainder of the tribe is fundamentally polyploid. This is of some
importance given the relatively early divergence of Cercideae in some phylogenies. In the rbcL phylogeny of Kajita et al. (2001) the tribe was sister to the
remainder of the family, though with relatively weak support, and this topology
also appears in the phylogenetic summary of Lewis et al. (2005). The chloroplast
matK tree of Wojciechowski et al. (2004), which emphasized Papilionoideae,
placed Cercideae, and Detarieae (mainly 2n = 24) together as the first-diverging
legume lineage. In these topologies, it is possible that, as Goldblatt (1981) suggested, the legumes are fundamentally x = n = 7, with subsequent independent
TE
78
EC
77
9.3.1 Chromosome Number Evidence for Polypoidy in Legumes
CO
RR
76
J. J. Doyle
UN
Editor Proof
150
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 151/179
127
128
129
130
131
132
133
134
135
136
137
138
139
140
141
142
143
144
145
146
147
148
149
150
151
152
153
154
155
156
157
158
F
125
126
PR
OO
123
124
D
122
TE
121
chromosomal increase both within Cercideae and in the ancestor of all remaining
legumes.
In contrast, the concatenated chloroplast matK/trnK ? trnL-F tree of Bruneau
et al. (2008) placed Detarieae as the first branch in the legume phylogeny, sister to
a trichotomy composed of Cercideae, Duparquetia, and the remainder of the
family. In this phylogeny, then, the base number for the family would be
x = n = 12, with 2n = 14 in Cercis representing a reduction. Interestingly, the
genome size of Cercis canadensis is comparable to measurements from the several
species of Bauhinia in the Kew C-value database (http://data.kew.org/cvalues/;
Leitch and Doyle, unpublished data), supporting this reduction hypothesis.
Even a high base number for early diverging lineages, as suggested by the
Bruneau et al. (2008) topology, would not definitively suggest polyploidy at the
base of the family, given what is known of chromosome numbers from other
Fabales. No information is available for Quillaja in the Index of Plant Chromosome Numbers (IPCN; http://www.tropicos.org/Project/IPCN), but Surianaceae is
represented by a single species of Stylobasium, with a number of 2n = 30, suggesting that the common ancestor of legumes and Surianaceae could have had a
high chromosome number.
Patterns of chromosomal evolution among major groups of legumes are complex even outside of the earliest branching. The bulk of the family belongs to two
sister clades: the MCC clade and the Papilionoideae. The two tribes that comprise
the MCC clade along with Mimosoideae (Caesalpinieae and Cassieae s.s.) are both
diploid based on x = 14. Given the presence of taxa with 2n = 28 in the grade at
the base of Papilionoideae, it is likely that the common ancestor of that group and
the MCC was diploid based on x = 14 as well. The majority of papilionoids,
however, have lower base chromosome numbers, ranging from x = 7–11,
depending on the tribe. These presumably represent reductions in chromosome
number, as discussed below; they certainly give no evidence for polyploidy.
EC
120
151
CO
RR
119
9.3.2 Genetic and Genomic Evidence for Polyploidy in the Early
Evolution of Legumes
Genomic studies, starting with linkage maps and continuing through studies of
expressed sequence tags (ESTs) and genome sequencing, have revolutionized
understanding of polyploidy in seed plants. Although it was long known that
diploidization can erase chromosomal evidence of polyploidy over time, it is now
clear that plant genomes comprise nested sets of WGD. The common ancestor of
all seed plants underwent a polyploid duplication, with a later WGD in the
ancestor of all angiosperms (Jiao et al. 2011) and numerous lineage-specific
duplications in various groups of flowering plants (Soltis et al. 2009).
It has been known for some time that cryptic polyploidy occurs in legume
genomes. For example, Shoemaker et al. (1996) used linkage map information to
UN
Editor Proof
9 Polyploidy in Legumes
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 152/179
165
166
167
168
169
170
171
172
173
174
175
176
177
178
179
180
181
182
183
184
185
186
187
188
189
190
191
192
193
194
195
196
197
198
199
200
201
202
203
F
164
PR
OO
163
D
162
TE
161
hypothesize that the soybean genome shows evidence of a more ancient duplication than the one that is responsible for its high chromosome number relative to
allied phaseoloid genera (millettioid clade, Fig. 9.1). In 2004, two different groups
mined the extensive EST collections of soybean and the diploid model legume,
Medicago truncatula (2n = 14; a member of the Hologalegina IRLC clade), to
search for the genomic signature of ancient polyploidy events (Blanc and Wolfe
2004; Schleueter et al. 2004). This signature is produced when all genes in the
genome are duplicated by autopolyploidy or when homoeologous loci are brought
together by allopolyploidy. It is observed by plotting the frequency distribution of
pairwise Ks (synonymous substitutions per synonymous site—a stand-in for time)
values for hundreds to thousands of paralogous gene pairs. Simple gene duplication is an ongoing phenomenon in all eukaryotes, but most duplicates are purged
from the genome rapidly, producing a characteristic distribution with many recent
duplicates with low Ks values and relatively few older pairs with high Ks (Lynch
and Conery 2003). Polyploid duplications appear as additional components
(‘‘peaks’’) against this background; the mode of such a Ks peak is taken as an
estimate for the age of the polyploid, though it is generally an overestimate of that
age (Doyle and Egan 2010).
Both Blanc and Wolfe (2004) and Schleueter et al. (2004) identified two Ks
peaks in soybean, as expected (Fig. 9.2); both reported similar Ks modes for these
peaks but because they used different substitution rates for plant nuclear genes, this
led to different estimates of the age of polyploidy (or homoeologue divergence).
The Schleueter et al. (2004) estimates are more in keeping with divergence dates
for papilionoid legume taxa (Lavin et al. 2005) and are preferred for that reason
(Shoemaker et al. 2006); in addition, the rate used by Schleueter et al. (2004) is
much closer to the rate recently estimated for Arabidopsis (Ossowski et al. 2010).
Of great interest was the finding, by both groups, of two Ks peaks in the M.
truncatula EST collection (Fig. 9.2). The younger of the two peaks is recent
enough that if due to polyploidy, it would most likely have left chromosomal
evidence, and has yet to be explained (Young et al. 2011). The older Medicago
peak, on the other hand, was estimated by Schleueter et al. (2004) to be around
54.6 MYA, very close to the 54 MY age estimated by Lavin et al. (2005) for the
divergence of the soybean (millettioid) and Medicago (Hologalegina) lineages,
and also similar to the age estimated for the older soybean peak (41.6 MYA). This
raised the possibility that the two species shared an ancient WGD.
This hypothesis was tested by Pfeil et al. (2005) using a phylogenomic
approach with 39 gene pairs chosen from among those used by Schleueter et al.
(2004) to identify the Ks peak in soybean. Topologies of gene trees overwhelmingly favored the hypothesis that their common ancestor was polyploid. Comparisons of linkage relationships between Medicago and the other legume model
species, Lotus japonicus (in the Hologalegina Loteae clade, Fig. 9.1), provided
further support for this hypothesis, and also showed that the duplication was not
found in poplar. Using the Lavin et al. (2005) date for the divergence of the
millettioid (Glycine) and Hologalegina (Lotus, Medicago) clades, the WGD event
had taken place by around 54 MYA. Thus, the common ancestor of the two major
EC
160
CO
RR
159
J. J. Doyle
UN
Editor Proof
152
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 153/179
153
CO
RR
EC
TE
D
PR
OO
F
Editor Proof
9 Polyploidy in Legumes
UN
Fig. 9.2 Evidence for polyploidy in the genomes of: a Glycine max and b Medicago truncatula.
The graphs plot the number of pairs of paralogous sequences (‘‘density’’ or ‘‘percent of pairs’’)
versus binned Ks (‘‘synonymous distances’’) classes. Pairs with very low divergence (produced by
ongoing recent duplications) were not plotted. Curves were fit to the binned divergence data and
are interpreted as groups of genes duplicated simultaneously in large-scale genomic events such
as polyploidy; modes of peaks provide a maximum age for allopolyploid events (Doyle and Egan
2010). Divergence time was estimated from synonymous distances using standard clock methods;
note the different estimated ages (modes of curves) for the older event in the two species. Data are
from Schleueter et al. (2004), who used expressed sequence tags (ESTs). Figure courtesy of
Jessica Schlueter (UNC-Charlotte)
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 154/179
J. J. Doyle
237
9.3.3 Polyploidy and Nodulation in Legumes
212
213
214
215
216
217
218
219
220
221
222
223
224
225
226
227
228
229
230
231
232
233
234
235
238
239
240
241
242
243
PR
OO
210
211
D
209
TE
207
208
EC
206
CO
RR
205
F
236
sister clades that comprise nearly 9,000 species––around 45 % of all legumes and
63 % of papilionoids––was polyploid.
Subsequently, Bertioli et al. (2009) studied Arachis (peanut) and showed that its
genome also shows evidence of the [54 MYA event, indicating that the large
dalbergioid clade is also fundamentally polyploid. Unpublished information
reported by McClean (personal communication) at the 2009 International Conference of Legume Genetics and Genomics showed that Lupinus, and hence the
genistoid clade, also shares this WGD. Thus, all of the lineages of the main
radiation of the papilionoids share a polyploid ancestor.
The possibility that all legumes share this polyploidy event remained tenable
until transcriptomic data from the caesalpinioid genus, Chamaecrista, became
available (Singer et al. 2009). Cannon et al. (2010) produced and analyzed over
1,200 gene phylogenies from these data, which overwhelmingly supported the
conclusion that the Chamaecrista genome shows no evidence of any polyploidy
event subsequent to the prerosid triplication, and notably lacks the WGD found in
core papilionoids (all but the early diverging lineages in Fig. 9.1). Chamaecrista
belongs to the MCC clade, which is sister to the papilionoid clade. Therefore, the
absence of the WGD in Chamaecrista indicates that the common ancestor of the
MCC clade and papilionoids was not polyploid, placing the WGD within papilionoids (Fig. 9.1). Whether the WGD took place in the papilionoid common
ancestor is still unknown, because the lineages that comprise the paraphyletic
grade lacking the putative molecular synapomorphy for the major papilionoid
radiation (chloroplast genome 50 kb inversion) remain to be sampled.
Thus, it is possible that this core papilionoid WGD facilitated the radiation of
the most species-rich lineage of legumes, comprising 69 % of the species (13,390/
19,327) and 59 % (438/741) of the genera of the third largest family of flowering
plants. This is the group that is more uniformly characterized by the eponymous
legume fruit, by the bilaterally symmetric papilionoid flower, and by the ability to
nodulate. The early diverging grade of papilionoids does contain some genera with
papilionoid flowers and legume fruits, but many lineages in this part of the tree are
characterized by unusual, nonpapilionaceous corollas and drupaceous or samaroid
fruits (Pennington et al. 2000); this grade also contains nearly all of the papilionoid
genera that do not nodulate (Doyle 2011).
204
The correspondence between nodulation and polyploidy in the family is interesting. Core papilionoids nearly all nodulate, but this is also true of Mimosoideae,
and Chamaecrista is among a handful of caesalpinioids known to be able to form a
nodulation symbiosis (Sprent 2009). It remains unclear whether there was a single
origin of nodulation in the common ancestor of the papilionoid and MCC clades,
followed by many losses of nodulation, or whether there were multiple origins of
UN
Editor Proof
154
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 155/179
155
264
9.3.4 Harmonizing Chromosomal and Genomic Evolution
252
253
254
255
256
257
258
259
260
261
262
265
266
267
268
269
270
271
272
273
274
275
276
277
278
279
280
281
282
283
284
PR
OO
250
251
D
249
TE
248
EC
246
247
Whatever the original basic chromosome number of the family, the earliest radiation from the common ancestor does not seem to have involved polyploidy,
despite Goldblatt’s (1981, p. 457) conclusion that the ‘‘… initial phase of polyploidy is probably very ancient and may have taken place in the late Cretaceous,
when major groups of Leguminosae began differentiating and were probably
evolving rapidly into new habitats.’’ Evidence against Goldblatt’s view of polyploidy in the ancestor of the entire family is the absence of any trace of polyploidy
in gene families of Chamaecrista other than the prerosid whole-genome triplication
(WGT; Jaillon et al. 2007). This indicates that the ancestor of the MCC and older
ancestors back to the prerosid WGT did not experience polyploid duplications. The
uniformity of base chromosome numbers in the major radiations of the family—
Cercideae, detarioids, MCC, and probably papilionoids (see above) suggest that
relatively high numbers (2n = 24–28) are plesiomorphic in the family.
The most parsimonious hypothesis for papilionoids is that the earliest papilionoid ancestor was also 2n = 28. Shortly after the divergence of this ancestor
from the MCC ancestor, the papilionoids radiated rapidly, and polyploidy occurred
nearly simultaneously, no later than the divergence of the first major lineage to
diverge in the core clade (genistoids, e.g., Lupinus). This WGD did not leave
evidence in higher chromosome numbers; to the contrary, polyploidy is associated
with chromosome number reduction in core papilionoids (Fig. 9.1). Goldblatt
CO
RR
245
F
263
nodulation in the MCC clade and an independent origin in the ancestor of the core
papilionoids (Doyle 2011).
The demonstration that Chamaecrista not only lacks the core papilionoid
WGD, but also does not show any genomic evidence of other polyploidy events,
indicates that polyploidy is not a prerequisite for nodulation in legumes as a whole
(Cannon et al. 2010). Thus, in a model of a single origin of nodulation in the
family, polyploidy would have played no role (Fig. 9.3). At the other extreme,
polyploidy could not have been involved in an origin of nodulation unique to
Chamaecrista, nor would it have been essential for the origin of nodulation in a
model where the symbiosis evolved in a common ancestor of Chamaecrista and
other members of the MCC clade (e.g., Mimosoideae).
This is not to say, of course, that polyploidy was not important in the origin or
evolution of nodulation in core papilionoids or in other nodulating taxa whose
genomes have yet to be explored, such as mimosoids. In the case of papilionoids, it
is possible that nodulation and the WGD will be found to coincide, either in the
ancestor of the main radiation of core papilionoids or at the first papilionoid
ancestor, but better phylogenetic resolution is required before this can be tested
(Pennington et al. 2001). In either case, refinement of the nodulation symbiosis in
taxa such as Medicago may well have been facilitated by the availability of homoeologues produced in the core papilionoid WGD (Young et al. 2011).
244
UN
Editor Proof
9 Polyploidy in Legumes
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 156/179
PR
OO
F
Cercideae
Detarieae
Amherstieae
Dialiinae
Umtiza clade
Batesia
Chamaecrista
Melanoxylon
Cassia
Senna
Caesalpinia clade
Moldenhawera
Erythrophleum
Calpocalyx
Pentaclethra
Parkia
core Mimosoideae
Campsiandra
Dimorphandra
Burkea
Dinizia
Arapatiella
Jacqueshuberia
Sclerolobium
Tachigali
Peltophorum clade
Swartzia clade
Non-nod. papil.
Mimosoideae-Caesalpinieae-Cassieae (MCC) clade
J. J. Doyle
Editor Proof
156
TE
D
N
60
50
40
EC
P
20
30
million years
10
“core papilionoids”
0
285
286
287
UN
CO
RR
Fig. 9.3 Relationship between polyploidy and nodulation in Leguminosae. Genera known to
nodulate are shown in bold face; Pentaclethra includes both nodulating species and species that
apparently cannot nodulate. Possible origins of nodulation are indicated with symbols and colors.
A single origin of nodulation for the entire family could have occurred in the common ancestor of
the papilionoids and the MCC clade (indicated by ‘‘N’’); this would have required many
independent losses in the course of legume evolution. Independent origins within the MCC clade
and Papilionoideae would require fewer losses. In the MCC clade, a single origin could have
occurred in the common ancestor of all genera known to nodulate (triangle), or once in each
major lineage of nodulating taxa (circles), or additional times within some clades (squares).
Similarly, a single origin could be hypothesized for papilionoids (green circle), or twice
(squares). The placement of the papilionoid polyploidy event (red ‘‘P’’) is indicated as in
Fig. 9.1. Polyploidy is associated with nodulation only in the papilionoids and might not be
directly associated there, given the uncertainty about the placement of both the polyploidy event
and the origin(s) of nodulation. Figure adapted from Doyle (2011)
(1981, p. 457) notes that ‘‘the cytological history of legumes seems to involve
some descending aneuploidy in every major evolutionary line but is most pronounced in Papilionoideae, in which most predominantly herbaceous tribes or
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 157/179
157
299
9.4 Polyploidy in Tribes and Genera of Legumes
295
296
297
300
301
302
303
304
305
306
307
308
309
310
311
312
313
314
315
316
317
318
319
320
321
322
323
324
325
326
327
328
PR
OO
294
Genomic and phylogenomic information currently is limited to the few genera
discussed above. More information can be expected with the advent of new
sequencing technologies, notably (at this writing) Illumina and 454. Twenty-five
legume genera, representing all three subfamilies, are included in the 1,000 Transcriptomes project (1 kp, see http://www.onekp.com/angiosperms.html), but results
from these species have not yet been analyzed (Steven Cannon, personal communication). Chromosome numbers provide a better guide at this level, but clearly
should be interpreted with the caveat that high numbers are likely to indicate
polyploidy, but low numbers cannot be assumed to be fundamentally diploid, given
the potential for cryptic polyploidy, a phenomenon discussed further below.
Three sources of data were used for the following survey of polyploidy in
legume clades: Goldblatt (1981); chromosome numbers provided for each genus
listed in the tribal treatments in Advances in Legume Systematics, Part 1 (Polhill
and Raven 1981); and the on-line Index of Plant Chromosome Numbers (IPCN;
http://www.tropicos.org/projectwebportal.aspx?pagename=Home&projectid=9).
IPCN was searched for all genera in Lewis et al. (2005) for which no data were
available in Goldblatt (1981) or Polhill and Raven (1981), and for genera with
polyploid counts, to identify infrageneric patterns of polyploidy, particularly when
published phylogenies were available. For genera with evidence of polyploidy,
BIOSIS was searched using the genus name in conjunction with either \polyploid*[and/or\phylogeny[. For some larger genera, IPCN was also consulted to
search for polyploid counts published since 1981. There remain many gaps in our
knowledge of legume chromosome numbers. Overall, only around 54 % of legume
genera have counts reported (Table 9.1), and the percentage is far lower for
tropical woody groups such as the caesalpinioid tribe Detarieae (30 %). This is
perhaps not too surprising, given the size of the family, the large number of small
genera, and the tropical distributions of many groups. It is also no doubt a commentary on how counting chromosomes has fallen from favor in this age of highthroughput science.
D
292
293
TE
291
EC
290
CO
RR
289
F
298
genera have achieved relatively low base numbers.’’ The details of this process
await elucidation of phylogenetic relationships of the early diverging papilionoid
lineages, and sampling of these taxa for the presence of polyploidy. What is clear
is that, at some point early in the history of Papilionoideae, chromosome numbers
fell from as high as n = 28 to n = 7–11. This process could have begun in, or
prior to, the common ancestor of the core papilionoids (Fig. 9.1). In any case, the
initial reduction was rapid, taking place within 10 MY after the polyploid event,
and perhaps within only 5 MY of the WGD. Detailed studies of synteny in the
various core papilionoid lineages should help elucidate whether initial reorganization occurred in a common ancestor or was completely independent in the longdiverged major clades of the core papilionoids.
288
UN
Editor Proof
9 Polyploidy in Legumes
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 158/179
158
J. J. Doyle
171 (2251)
82 (3271)
40 (415)
83 (2354)
53 (1514)
7 (58)
32 (763)
34 (414)
62
39
15
59
32
6
22
23
4
10
2
24
11
1
5
9
6
26
13
41
34
17
23
39
54 (4351)
7 (768)
40
2
15
1
38
50
56 (1104)
22
1
5
112 (2064)
731
71
393
12
95
17
24
a
F
Percent genera
with polyploidy
(%)
PR
OO
Caesalpinioidsb
Mimosoideae
Swartzieae ? Sophoreae
Genistoids
Dalbergioids
Baphioids
Mirbelioids
Hologalegina:
Robinioids ? Loteae
Hologalegina: IRLC
Millettioids:
indigofereae
Millettioids: core
Millettioids
Millettioids: phaseoloids
Total
based on chromosome numbers
Genera with
Number of
chromosome
genera with
counts
polyploidy
D
Editor Proof
Table 9.1 Polyploidy in legume clades,
Clade
Number of
genera
(species)a
Source: Lewis et al. (2005)
Summation of the following monophyletic groups: Cercideae, 12 genera total, 2 of 6 genera
with chromosome counts have reports of polyploidy, 33 % (12, 2/6, 33 %); Dialiinae (17, 0/7,
0 %); Detarieae (82, 2/25, 8 %); MCC clade minus mimosoids (59, 0/24, 0 %)
TE
b
342
9.4.1 Caesalpinioids and Mimosoideae
332
333
334
335
336
337
338
339
340
343
344
345
CO
RR
331
UN
330
EC
341
Goldblatt (1981) made a distinction among genera that are exclusively polyploid relative to genera in the same tribe, and genera that include species with both
‘‘diploid’’ and ‘‘polyploid’’ chromosome numbers. In light of our current understanding of polyploidy, this is an artificial distinction, but exclusively polyploid
genera are perhaps worth noting, because they represent lineages where polyploids
may have replaced their diploid progenitors entirely. Such lineages would constitute evidence of polyploid ‘‘success’’ (e.g., Mayrose et al. 2011) in measures of
diversity a million years from now.
The results of this survey show that polyploidy, as inferred solely from chromosome numbers, occurs in nearly a quarter of all legume genera, but varies
widely in frequency among different lineages (Table 9.1). As Goldblatt (1981)
noted, polyploidy is rare in woody, tropical groups such as caesalpinioids and early
diverging papilionoid lineages.
329
9.4.1.1 Clades of the Caesalpinioid Grade, Excluding the MCC Clade
Goldblatt (1981) stated that polyploidy (assuming a basic diploid number of
2n = 28) is uncommon in caesalpinioid groups. As noted above, unpublished
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 159/179
159
360
9.4.1.2 The MCC Clade
355
356
357
358
361
362
363
364
365
366
367
368
369
370
371
372
373
374
375
376
377
378
379
380
381
382
383
384
385
386
387
PR
OO
353
354
D
351
352
Chromosome numbers of the caesalpinioid members of the MCC clade range from
2n = 20–28, with the exception of Chamaecrista, in which numbers of 2n = 14,
16, and 28 occur. Goldblatt (1981) raised the possibility that low chromosome
numbers in this genus are not ancestral, but instead represent aneuploid reduction
from ancestral x = 14 species. This makes good sense given the prevalence of
2n = 28 counts throughout much of the MCC clade. A recent molecular phylogenetic analysis of Chamaecrista (Torres et al. 2011) supported the hypothesis that
the 2n = 14 species of sect. Xerocalyx form a monophyletic group nested within
species having higher chromosome numbers. Chamaecrista is a genus of considerable interest as a potential model for nonpapilionoid legumes (Singer et al.
2009), making it an attractive system for exploring chromosome evolution in the
caesalpinioids.
Mimosoideae comprise a monophyletic group embedded in the MCC clade, and
the subfamily is dominated by taxa with n = 13. Goldblatt (1981) listed only three
genera with base numbers higher than n = 14: Schleinitzia (2n = 52, 54), Leucaena (2n = 52, 56, 104, 112), and Dichrostachys (2n = 50, 56). These genera are
all members of tribe Mimoseae and are relatively closely related within that tribe
(Lewis et al. 2005), with the former two being members of the same clade and
Dichrostachys being part of a sister clade (Luckow et al. 2003). However,
Schleinitzia and Leucaena are not sisters within their clade (Hughes et al. 2003;
Luckow et al. 2005), suggesting that polyploidy has originated independently in
each case.
Leucaena itself has been fertile ground for systematic investigation. Boff and
Schifino-Wittmann (2003) concluded that its species are segmental paleopolyploids. A series of studies has built a strong foundation for understanding the
complex history of hybridization and polyploidy in the genus, and the impact of
these phenomena on characters such as nrDNA ITS pseudogene evolution, and
TE
350
EC
349
CO
RR
348
F
359
genome size data do not support the hypothesis that Bauhinia (commonly
2n = 28) is tetraploid relative to Cercis (2n = 14). However, this is negative
evidence, and given the prevalence of genome downsizing (Leitch and Bennett
2004) in polyploids, it remains possible that Bauhinia (along with Adenolobus and
Griffonia) is polyploid. More recent polyploidy occurs in the group, with Tylosema
having 2n = 52; Tylosema is nested within Bauhinia s. l. (Sinou et al. 2009). The
large Detarieae s.l. clade is overwhelmingly 2n = 24. The two detarioid genera
having higher chromosome numbers (Hardwickia, 2n = 34; Colophospermum,
2n = 36) are strongly supported as sisters within the Prioria clade of Bruneau
et al. (2008); they could represent independent aneuploid reduction from a polyploid ancestor. Goldblatt (1981) mentioned polyploidy in Anthonotha (2n = 24,
28, 72), but only 2n = 24 and 28 are listed by Cowan and Polhill (1981), and no
counts are listed in IPCN. All known counts from the Dialiinae clade are 2n = 24
or 28.
346
347
UN
Editor Proof
9 Polyploidy in Legumes
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 160/179
J. J. Doyle
418
9.4.2 Papilionoideae
395
396
397
398
399
400
401
402
403
404
405
406
407
408
409
410
411
412
413
414
415
416
419
420
421
422
423
424
425
426
PR
OO
394
D
393
TE
392
EC
391
CO
RR
390
F
417
their role in domestication (Hughes et al. 2002, 2007; Govindarajulu et al. 2011a,
b). Govindarajulu et al. (2011b) concluded that ‘‘… a comprehensive picture of the
complex evolutionary dynamics of polyploidy in Leucaena is emerging. This
includes paleotetraploidization, diploidization of the last common ancestor to
Leucaena, allopatric divergence among diploids, and recent allopolyploid origins
for tetraploid species likely associated with human translocation of seed.’’
Acacia (sens. lat.) is reported to have 2n = 26, 52, 76, and 104, though the
majority of its species are diploid (Gallagher et al. 2011). Polyploidy has not been
a focus of recent phylogenetic analyses (e.g., Brown et al. 2010; Murphy et al.
2010). Similarly, phylogenetic studies of Prosopis (2n = 28, 52, 56) or Prosopidastrum (2n = 28, 56) do not discuss polyploidy (e.g., Bessega et al. 2006;
Catalano et al. 2008). No phylogenetic studies appear to have addressed polyploidy in Neptunia (2n = 28, 36, 54, 56, 72, 78), though Pandit et al. (2006) note
that N. plena, an invasive species in Singapore, is a polyploid (2n = 72). No
studies appear to exist on polyploidy and phylogeny of Inga (2n = 26, 52; the
latter reported by Hanson 1995), Albizia (2n = 26, c. 78), or Calliandra (2n = 16,
22, 32, 44).
Mimosa (2n = 24, 26, 28, 40, 52) is a genus of around 500 species; a second
genus whose species vary in ploidy, Schrankia (2n = 16, 22, 24, 26, 52), is deeply
nested within Mimosa (Simon et al. 2011). Dahmer et al. (2011) concluded that the
phylogenetic pattern ‘‘… suggests that duplication of chromosome numbers
evolved several times in the genus and that polyploidy is not restricted to any
particular clade within Mimosa. On the contrary, it seems that polyploids arose
independently from ancestors with lower ploidy levels and are present in divergent
lineages in the genus.’’ Seijo and Fernandez (2001) reported chromosome numbers
from the southern extreme of the range and discovered polyploidy within M.
balansae. Chromosomal and morphological studies by Morales et al. (2010)
clarified relationships in the M. debilis-M. nuda complex, demonstrating that
hybridization and polyploidy are responsible for taxonomic complexity in the
group.
388
389
Relatively few papilionoid genera appear to be exclusively polyploid based on
chromosome number. As noted above, the early diverging papilionoid lineages
have relatively high numbers, like the caesalpinioid and mimosoid groups. Ateleia
(Swartzieae) is 2n = 40, presumably representing a second polyploidy event
followed by aneuploid reduction. Goldblatt listed Dipteryx (Dipterygeae) as being
a polyploid genus but with a questionable count of 2n = 32; this number was not
reported in the treatment of the tribe by Polhill (1981a), nor is a count for the
genus listed in IPCN.
UN
Editor Proof
160
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 161/179
434
435
436
437
438
439
440
441
442
443
444
445
446
447
448
449
450
451
452
453
454
455
456
457
458
459
460
461
462
463
464
465
466
467
468
469
470
F
432
433
PR
OO
431
D
430
The genistoid clade, which is weakly supported as sister to the remaining core
papilionoids (Fig. 9.1), is one of the most complex groups with respect to polyploidy, and detailed discussion is beyond the scope of this review. The frequency
of polyploidy in the genistoid clade is the highest for any well-sampled group of
legumes (Table 9.1). Cusma-Velari and Feoli-Chiapella (2009) discussed cytology
in ‘‘so-called ‘primitive’ genera of Genisteae’’ in light of molecular phylogenetic
data. Genistoids clearly have a base number of x = 9 (Goldblatt 1981), with
2n = 18 being common in most of its tribes.
Sophora s.l. has been divided into several segregates that vary in chromosome
number. Sophora s.s. is part of the genistoid clade and is x = 9. Boatwright and
van Wyk (2011) reported on the relationships of several of these based on nrDNA
ITS sequences. They focused on the placement of the South African species, S.
ihambanensis, which is polyploid (2n = 36); in their tree it is sister to S. tomentosa, a diploid, but they do not discuss origins of the polyploid. A count of
2n = 18 is common in Sophora s.l., and several additional species are polyploids
with 2n = 36 (S. alopecuroides, S. pachycarpa, and S. songarica); S. leachiana is
listed by ICPN as having 2n = 36 and 2n = 54 cytotypes. The small segregate,
Calia, has 2n = 18 and may be sister to the entire genistoid clade. The remainder
of Sophora s.l. comprises 2n = 28 species transferred to Styphnolobium in the
early diverging papilionoid grade.
In Thermopsideae, Thermopsis has both diploid and polyploid species
(2n = 18, 36). In Podalyrieae, two genera are exclusively polyploid: Virgilia
(2n = 54) and Cyclopia (2n = 36); they are not supported as sisters in Boatwright
et al. (2008). Crotalarieae are sister to Genisteae and include the exclusively
polyploid Buchenroedera (2n = 28; Van Wyk and Schutte 1988), as well as
polyploids within Crotalaria (2n = 14, 16, 32) and Lotononis (2n = 18, 28, 36).
Genisteae are by far the most cytologically complex group in the entire Leguminosae. Even genera with low numbers may be polyploid, such as Anarthrophyllum (2n = 24; Goldblatt 1981) and Dichilus (2n = 28). These genera, along
with the polyploid Polhillia (2n = 32) and complex Melolobium (2n = 18, 32),
were once placed in Crotalarieae. It is in the core Genisteae that polyploidy and
aneuploidy have run rampant. The group includes Argyrolobium (2n = 24, 26, 30,
32, 48), Adenocarpus (2n = 26, 46, 48, 52, 54), Laburnum (2n = 48, 50), Cytisophyllum (2n = 50, 52), Petteria (2n = 52), Argyrocytisus (2n = 50), Chamaecytisus (2n = 48, 96), Cytisus (2n = 22, 24, 46, 48, 92, 96), Calicotome
(2n = 24, 48, 50), Erinacea (2n = 52), Spartium (2n = 48, 52, 54, 56), Retama
(2n = 48), Genista (2n = 18, 22, 24, 26, 28, 30, 32, 36, 40, 42, 44, 46, 48, 50, 52,
56, 72, 80, 96), Echinospartium (2n = 44, 52), Stauracanthus (2n = 28, 48, ca.
128), Ulex (2n = 32, 64, 80, 96), and Lupinus (2n = 24, 30, 32, 34, 36, 38, 40, 42,
48, 50, 52, 96).
Bisby (1981) considered the plethora of chromosome numbers attributed to
individual genera to be partly a real phenomenon, but also due to difficulties in
obtaining reliable counts given the small size and high numbers of chromosomes,
TE
429
9.4.2.1 Genistoids
EC
428
161
CO
RR
427
UN
Editor Proof
9 Polyploidy in Legumes
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 162/179
J. J. Doyle
497
9.4.2.2 Dalbergioids
479
480
481
482
483
484
485
486
487
488
489
490
491
492
493
494
495
498
499
500
501
502
503
504
505
506
507
508
509
510
511
512
PR
OO
478
D
477
TE
475
476
EC
473
474
CO
RR
472
F
496
combined with the taxonomic complexity of the groups. Some of the taxonomic
complexity is being resolved by molecular phylogenetic studies focused on Cytisus
and Genista (Cubas et al. 2002; Pardo et al. 2004), but these studies do not discuss
polyploidy per se. For Lupinus, three studies have documented the rapid radiation
of the genus in its New World center of diversity (Hughes and Eastwood 2006;
Drummond 2008; Drummond et al. 2012), and another phylogenetic study focused
on the Old World species (Ainouche et al. 2004). Drummond (2008) noted that,
‘‘While a complex history of aneuploidy (2n = 32, 34, 36, 38, 40, 42, 50, 52) in
the Old World and eastern New World … implies that allopolyploidy may have
provided an additional mechanism for reproductive isolation and evolutionary
divergence, chromosomal numbers in the western New World species (2n = 48
with occasional autopolyploids of 2n = 96) are relatively stable.’’ It is this western
group that has radiated explosively, presumably driven by ecology and not due to
polyploidy per se (Drummond 2008). Conterato and Schifino-Wittmann (2006)
described chromosome numbers and meiotic behavior in diploid and polyploid
American lupines, and noted consistencies with phylogenetic relationships in the
genus.
The placement of polyploid former Crotalarieae in the same clade with core
Genisteae may lend support to the idea that polyploidy arose early in the entire
clade, as suggested by Goldblatt (1981), Lavin et al. (2005) dated the common
ancestor of Crotalarieae and Genisteae at around 41 MYA. On the other hand,
Goldblatt also noted (1981, p. 452) that ‘‘basic numbers for these genera (Genista,
Ulex, Cytisus) are however in the diploid range and a basic number of x = 12 for
the group as a whole and for several genera has been suggested …’’. Genomic data
for members of Genisteae should eventually allow the determination of the number
and relative timing of polyploid events in the group.
471
The dalbergioid s.l. clade is split into two major subclades, Amorpheae and a
second clade comprising Adesmieae, Aeschynomeneae, and many members of the
polyphyletic Dalbergieae (Lewis et al. 2005). The entire dalbergioid clade is
dominated by 2n = 20 species. Within Amorpheae, polyploidy occurs in genera
from each of the major subclades described by McMahon (2005). In the daleoid
clade, Dalea has 2n = 14, 16, 28, and 42. Spellenberg (1981) hypothesized that
tetraploids and hexaploids of D. formosa (2n = 28, 42) were autopolyploids
derived from the diploid (2n = 14) cytotype. In the amorphoid clade, Amorpha
includes both diploids and polyploids (2n = 20, 40), all native to the New World.
The widespread A. fruticosa is exclusively polyploid and morphologically complex (Wilbur 1975). It has become an invasive weed in Europe (e.g., Hulina 2010),
illustrating a common feature of polyploidy (e.g., Pandit et al. 2011; te Beest et al.
2011). Its relationships to other members of the genus appear to be complex,
sharing chloroplast haplotypes with different sympatric diploids across its range
(Straub and Doyle, unpublished data). Studies of the A. georgiana complex
UN
Editor Proof
162
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 163/179
163
547
9.4.2.3 Baphioids
521
522
523
524
525
526
527
528
529
530
531
532
533
534
535
536
537
538
539
540
541
542
543
544
545
548
549
550
551
552
PR
OO
520
D
518
519
TE
517
EC
515
516
CO
RR
514
F
546
identified mixed populations of diploids and polyploids in what had been assumed
to be an exclusively diploid species; these included an apparent allopolyploid
between A. georgiana and A. herbacea (Straub and Doyle 2009). Straub
(unpublished data) has identified additional polyploid species in the genus and
hypothesized their origins.
The core dalbergioid clade is split into the small Adesmia clade (six genera) and
the much larger clade comprising the Dalbergia and Pterocarpus sister clades
(Lewis et al. 2005). Polyploidy occurs in all three clades. In the Adesmia clade,
Adesmia includes both diploids and polyploids (2n = 20, 40), and Amicia is
exclusively polyploid (2n = 38).
In the Dalbergia clade, Smithia (2n = 38) is exclusively polyploid and is
considered closely related to Kotschya (Lewis et al. 2005), a genus that includes
species with chromosome numbers indicative of polyploidy and aneuploidy
(2n = 28, 30, 36, 40). These two genera are grouped with Aeschynomene species
(Lavin et al. 2001), a genus that also includes diploids and polyploids (2n = 18,
20, 40). Another dalbergioid-clade genus, Ormocarpum (2n = 24, 26), was not
listed in Goldblatt’s discussion of polyploid genera, but could potentially be a
cryptic polyploid with aneuploid reduction. Information on chromosome numbers
of other members of the Ormocarpum group (Thulin and Lavin 2001) would be
useful in addressing this issue.
The Pterocarpus clade also includes several genera with both diploids
(including presumed aneuploids) and polyploids scattered among its subclades:
Platymiscium (2n = 16, 18, 20, 32), Pterocarpus (2n = 22, 24, 44), Geoffroea
(2n = 20, 60), and Arachis (2n = 20, 40). Arachis includes the tetraploid peanut
or groundnut (A. hypogaea), as well as three other tetraploid species, one of which
(A. glabrata) is a tropical forage crop. Peanut is hypothesized to be an allopolyploid derived from A- and B-genome species (e.g., Burow et al. 2009). Seijo et al.
(2007) provide a useful summary of hypotheses concerning the origin (or origins)
of peanut; controversy exists concerning such issues as the exact progenitor species of both homoeologous genomes, mode and number of origins, and whether
there was subsequent introgression from wild species into the cultigen. They
identified likely diploid progenitors of A. hypogaea using GISH and studied
meiotic behavior of two other tetraploids (Ortiz et al. 2011) and a spontaneous
autotriploid of A. pintoi (Lavia et al. 2011).
513
This small group appears to be interesting from the standpoint of polyploidy.
Chromosome numbers are known from six of its seven genera. Of these, four are
listed as 2n = 22, one (Dalhousiea) is 2n = 44, and Baphia was reported in
Polhill and Raven (1981) to have both numbers, though no reports for any of the
47 species of the genus exists in IPCN.
UN
Editor Proof
9 Polyploidy in Legumes
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 164/179
553
J. J. Doyle
9.4.2.4 Mirbelioids
576
9.4.2.5 Hologalegina
562
563
564
565
566
567
568
569
570
571
572
573
574
577
578
579
580
581
582
583
584
585
586
587
588
589
590
591
592
593
PR
OO
561
D
559
560
TE
557
558
EC
556
The Hologalegina clade includes robinioids and the Inverted Repeat Loss Clade
(IRLC; named for the absence of a prominent feature of the chloroplast genome).
Goldblatt (1981) concluded that ‘‘Species polyploidy is overwhelmingly concentrated in temperate to cool Eurasia’’ so it is not surprising that this largest clade of
legumes, which includes many temperate genera, has a higher frequency of
polyploidy than the family average, nearly 40 % in both of its major subclades
(Table 9.1).
The robinioids comprise two clades: one with Sesbania plus Loteae (including
Coronilleae), the other being Robinieae (s.s.). Diploid chromosome numbers vary
considerably within Robinieae s.s., and Goldblatt (1981) suggested several possible base numbers, the most likely being x = 10 or 11. There is one apparently
exclusively polyploid genus, Poissonia, which Goldblatt (1989) counted as
(2n = ca. 32). Although only numbers of 2n = 10 and 11 were given for Robinia
in Polhill and Sousa (1981), more recent counts of 2n = 30 for R. hispida suggest
polyploidy within this small genus.
Even with the limited sampling of Loteae in Wojciechowski et al. (2004), it is
clear that numerous problems exist with the genera as circumscribed, notably that
CO
RR
555
F
575
Polyploidy in the mirbeliod clade (23 %) is close to the average for the whole
family (Table 9.1). Isotropis (2n = 16, 18, 32) is sister to the large ‘‘Pultenaea s.l.
group’’ in Orthia et al. (2005), which includes Oxylobium (2n = 16). Chandler et al.
(2001) sank both Brachysema (2n = 16, 32) and the monotypic Jansonia
(2n = 32) in Gastrolobium, previously a genus with only 2n = 16 species.
Chandler et al. (2001) placed the single sampled species of Jansonia sister to
Brachysema celsianum (not listed in IPCN), in a clade that also included B.
praemorsum; that species is a diploid at 2n = 16, as is at least one species in the
sister clade, Nemcia coriacea. Thus, it is likely that polyploidy has arisen more
than once just within Gastrolobium s.l., and another time in Isotropis. Among
several ‘‘strongly paraphyletic’’ Pultenaea s.l. genera, nearly all of which are
2n = 16, are two genera with polyploidy: Pultenaea s.s. (2n = 8, 12, 14, 16, 18,
27, 32) and Chorizema (2n = 16, 32). Sorting out how the various chromosome
numbers in Pultenaea s.s. are related will be of considerable interest but will
require more complete phylogenies than appear to be available at present. Of the
10 Chorizema species (out of 27 in the genus) listed in IPCN, polyploidy is only
reported from C. aciculare, which has both diploid and tetraploid cytotypes.
Smaller genera in Pultenaea s.l. with known polyploidy are Eutaxia (2n = 16,
32) and Dillwynia (2n = 14, 21, 28), which are in the same weakly supported
clade in Orthia et al. (2005). One of the two Dillwynia species (D. phylicoides)
included in the Orthia et al. (2005) tree has both diploid and tetraploid cytotypes
listed in IPCN.
554
UN
Editor Proof
164
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 165/179
603
604
605
606
607
608
609
610
611
612
613
614
615
616
617
618
619
620
621
622
623
624
625
626
627
628
629
630
631
632
633
634
635
636
637
638
F
PR
OO
601
602
D
600
TE
598
599
Anthyllis (2n = 10, 12, 14, 16, 28) and Ornithopus (2n = 14) are nested within
Lotus (2n = 10, 12, 14, 24, 28). Other genera with known polyploidy are Coronilla (2n = 12, 20, 24); Hippocrepis (2n = 14, 28); Dorycnium (2n = 14, 28;
both cytotypes in D. axilliflorum), and Scorpiurus (S. muricatus has 2n = 14, 16,
28; other species are 2n = 14 or 28). Degtjareva et al. (2006) provided phylogenetic hypotheses for Lotus, with sampling of other genera, but did not discuss
polyploidy. Rosello and Castro (2008) discussed polyploidy in the flora of the
Balearic Isles, among which are species of Anthyllis and Coronilla.
The genus Lotus includes the genomic model legume, L. japonicus (Sato et al.
2008), which is part of the L. corniculatus (birdsfoot trefoil) polyploid complex.
Grant and Small (1996) summarized many studies of this complex and concluded
that it was a fertile topic for further study, particularly to identify the diploid
progenitors of L. corniculatus itself, which they considered to be an allopolyploid.
Gauthier et al. (1998a, b) discussed evolutionary patterns in the L. corniculatus/L.
alpinus polyploid complex in the Alps of Europe; they described morphological
and genetic consequences of autopolyploidy in L. alpinus and suggested introgression at the tetraploid level between it and L. corniculatus.
The majority of genera and species in the IRLC clade belong to two sister
clades in Wojciechowski et al. (2004): one includes the Astragalean clade
(Astragalus and allies) and Hedysareae, and the second includes the Vicioid clade.
The remainder of the IRLC phylogeny, moving successively further from these
clades, consists of a clade with Wisteria (2n = 16) and one species of Callerya,
followed by a clade with Glycyrrhiza (2n = 16) and a second species of Callerya.
The Astragalean clade has extensive polyploidy. Perhaps most striking is the
clade that includes the New Zealand endemic tribe Carmichaelieae plus the
Australian Swainsona and an additional New Zealand genus, Montigera, all of
which are polyploid (Wagstaff et al. 1999). The only exclusively polyploid genera
that Goldblatt (1981) listed for the IRLC clade belong to this clade: Swainsona
(2n = 32); Clianthus (2n = 32); Carmichaelia (2n = 32, ca. 96), Chordospartium
(2n = 32); and Corallospartium (2n = 32), with the latter two subsumed in
Carmichaelia in Lewis et al. (2005). Wagstaff et al. (1999) concluded that the New
Zealand radiation was recent, involved an already polyploid colonizer, and may
have been associated with orogeny and glaciation.
Elsewhere in the Astragalean clade are two large genera with extensive polyploidy and aneuploidy, Oxytropis (300–400 species: 2n = 16, 32, 48, 64, 96) and
Astragalus (ca. 2500 species: 2n = 16, 22, 24, 26, 28, 32, 44, 48, 64). In
Astragalus, polyploidy appears to be more common among Old World than among
New World species. Wojciechowski (2005) summarized phylogenetic results for
this huge genus, showing that aneuploid species form a clade. According to Gohil
and Ashraf (2008), polyploidy occurs in around only 17 % of Astragalus species.
However, Astragalus is one of the largest genera of plants with as many as 2500
species (Lewis et al. 2005), so if this percentage is correct, then there are over 400
polyploid species in the genus. There does not seem to be a comprehensive
phylogeny that discusses origins of polyploidy in Oxytropis. However, a series of
papers describe autopolyploidy, including multiple autopolyploid origins, in
EC
596
597
165
CO
RR
594
595
UN
Editor Proof
9 Polyploidy in Legumes
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 166/179
646
647
648
649
650
651
652
653
654
655
656
657
658
659
660
661
662
663
664
665
666
667
668
669
670
671
672
673
674
675
676
677
678
679
680
681
682
683
F
PR
OO
644
645
D
642
643
TE
641
Oxytropis chankaensis (e.g., Artyukova et al. 2011). Jorgensen et al. (2003) suggested ‘‘a scenario of multiple formations of polyploids, possibly including
hybridization among diverged Alaskan Oxytropis populations.’’
Within the Hedysareae, phylogenetic studies of Caragana (2n = 16, 24, 32, 48)
suggest that polyploidy is confined to a single group of species, and that triploids,
tetraploids, and hexaploids may all be autopolyploid in origin (Zhang et al. 2009).
Neither Hedysarum (2n = 14, 16, 48) nor Onobrychis (2n = 14, 16 28, 32)
appears to be monophyletic on the basis of nrDNA ITS phylogenies (Ahangarian
et al. 2007). Hejazi et al. (2010) discussed karyotypic evolution in diploid and
polyploid species but did not provide a phylogenetic context or identify origins of
polyploids. Based on IPCN listings, most polyploidy reported for Hedysarum in
IPCN appears to involve multiple cytotypes within a single species (e.g., H.
arcticum and H. hedysaroides, both 2n = 14, 28; H. dasycarpum and H. mackenziei, both 2n = 16, 32; H. gmelinii, 2n = 16, 28, 56), but some species are
exclusively polyploid (e.g., H. inundatum, 2n = 28). Similarly, in Onobrychis
there is variation within species (e.g., O. aequidentata, 2n = 14, 16, 28; O. arenaria, O. bobrovii, 2n = 14, 28; O. crista-galli, 2n = 16, 32), with other species
being exclusively polyploid (e.g., O. biebersteinii, O. cyri, O. dielsii, all 2n = 28).
There appear to be no phylogenies or evolutionary studies of polyploidy in Alhagi
(2n = 16, 28; the latter number is not listed in IPCN).
The majority of the vicioid clade forms two sister clades, one with Fabeae
(Vicieae) plus Trifolium, and a second comprising the remaining Trifolieae genera;
polyploidy occurs in both clades. Successive sisters to this clade (Cicer, Galega,
and Parochetus) are all 2n = 16.
Vicia includes both diploids and tetraploids (2n = 10, 12, 14, 24, 28), but
polyploidy was considered rare in the genus by Kupicha (1981). Indeed, Vicia is
best known for its extensive non-polyploid variation in genome size (Chooi 1971;
Neumann et al. 2006), which shows only weak correlation with ploidy: diploid
(2n = 14) V. peregrina has a genome size of 9.48 pg/1C, double that of tetraploid
(2n = 24) V. tenuifolia (4.73 pg/1C). Endo et al. (2008) did not discuss either
issue in their phylogenetic study of New Wold Vicia. Travnicek et al. (2010)
studied the history of polyploidy in V. cracca, determining the ploidy of over
6,500 individuals at more than 250 localities in Europe and mapping the distributions of diploids, triploids, and tetraploids; they noted the rarity of triploids,
suggesting strong reproductive barriers between diploids and tetraploids.
Polyploidy is also noted to be rare in Lathyrus (2n = 14, 28, 42; Kupicha
1981). Gutierrez et al. (1994) hypothesized autopolyploid origins of L. pratensis
and L. palustris from conspecific diploids, but an allopolyploid origin of L.
venosus from two diploid species (L. ochroleucus and L. palustris). Only 2n = 14
counts are listed for the closely related Pisum in IPCN, for which Kupicha (1981)
listed polyploidy as ‘‘rare.’’
Turini et al. (2010) reconstructed nrDNA ITS and chloroplast phylogenies for
69 of the 86 species of Ononis (2n = 16, 20, 30, 32, 60, 64) and identified several
well-supported clades. They concluded that, ‘‘Unfortunately, only limited information is available … on chromosome numbers to test support for these groups’’.
EC
640
CO
RR
639
J. J. Doyle
UN
Editor Proof
166
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 167/179
691
692
693
694
695
696
697
698
699
700
701
702
703
704
705
706
707
708
709
710
711
712
713
714
715
716
717
718
719
720
721
722
723
724
725
726
727
728
F
690
PR
OO
689
D
687
688
TE
686
However, chromosome counts are available for nearly half of the species in their
phylogeny in IPCN, and some conclusions can be drawn. For example, the clade
that is strongly supported as sister to the remainder of the genus in their nrDNA
ITS tree includes only polyploids (O. tridentata and O. fruticosa are both 2n = 30;
O. rotundifolia is 2n = 32), suggesting that the genus as a whole could be polyploid. Only three species have low, potentially non-polyploid numbers in IPCN.
These occur in different clades, and in two cases species with low numbers have
higher numbers as well (O. variegata, 2n = 16, 30; O. ornithopodioides, 2n = 20,
32), raising the possibility that they are independent reductions from typical
polyploid numbers. The exception, O. adenotricha, is only reported as 2n = 16; its
position varies between the nrDNA ITS and trnL-F trees of Turini et al. (2010),
being sister to the tridentata clade in the trnL-F tree; however, this entire group is
not resolved as sister to the remainder of the genus in that tree. Elsewhere in the
genus, O. spinosa has multiple cytotypes (2n = 30, 32, 60), whereas O. pendula is
only known at 2n = 64. Kloda et al. (2008) studied patterns of genetic diversity in
several diploid and polyploid species in England and concluded that gene flow was
occurring within ploidy levels, but not between diploids and tetraploids.
Medicago includes the genomic model legume, M. truncatula (Young et al.
2011). Steele et al. (2010) provided a phylogeny for Medicago (2n = 14, 16, 32,
48), including multispecies sampling of its sister clade, which comprises the
interdigitated species of the two paraphyletic genera Melilotus (2n = 16, 24, 32;
though tetraploids are not reported in IPCN) and Trigonella (2n = 16, 28, 32, 44).
Aneuploid change from 2n = 16 to 2n = 14 has occurred several times in
Medicago (Steele et al. 2010). Polyploidy is concentrated in a clade that comprises
most species of sect. Medicago, along with M. arborea (sect. Dendrotelis); an
additional polyploid species, M. scutellata, occurs in the clade sister to this sect.
Medicago clade. Rosato et al. (2008) used fluorescence in situ hybridization
(FISH) to study relationships between polyploids and diploids in sect. Dendrotelis.
Some Medicago species are exclusively polyploid whereas others possess
multiple cytotypes. The M. sativa complex, which includes cultivated autotetraploid alfalfa (M. sativa ssp. sativa) as well as other diploid (2n = 16) and autopolyploid (2n = 32) species and their hybrids, has been the focus of several recent
studies (Sakiroglu et al. 2010; Havananda et al. 2010, 2011, and unpublished data).
Two major autopolyploid pairs in the complex are: (1) M. s. caerulea and M. s.
sativa, both with blue flowers and coiled pods, distinguishable by the larger size of
the tetraploid (M. s. sativa) for several characters; and (2) M. s. falcata, a yellowflowered taxon with falcate pods whose diploid and polyploid cytotypes are
indistinguishable morphologically. Interestingly, although M. s. sativa and M. s.
caerulea are undifferentiated for chloroplast haplotypes, the two cytotypes of M. s.
falcata possess nearly mutually exclusive sets of haplotypes, with haplotypes in
the tetraploid most likely derived by introgression from M. prostrata, a species
from outside the complex (Havananda et al. 2011). Jenczewski et al. (1999)
reported gene flow between wild and cultivated M. sativa populations; however,
based on chloroplast data, there does not appear to be significant gene flow
between blue- and yellow-flowered taxa in the complex either at the diploid or
EC
685
167
CO
RR
684
UN
Editor Proof
9 Polyploidy in Legumes
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 168/179
J. J. Doyle
743
9.4.2.6 Indigofereae
736
737
738
739
740
741
744
745
746
747
748
749
750
751
752
753
754
755
756
757
758
759
760
761
762
763
764
765
766
767
768
769
770
PR
OO
735
D
733
734
Schrire et al. (2009) provided a detailed phylogeny for this tribe, a monophyletic
group that is sister to the millettioid clade. Schrire et al. (2009) did not comment
on chromosomal variation or polyploidy, but numerous records are readily
available in IPCN, and mapping these onto the phylogeny provides some insights
into cytological evolution of the group.
The tribe is dominated by the very large genus Indigofera (ca. 700 species),
which Goldblatt (1981) and Polhill (1981b) listed as having 2n = 14, 16, 32, 48.
The higher numbers thus would be interpreted as representing tetraploids and
hexaploids. However, Frahm-Leliveld (1966), summarizing the cytotaxonomy of
the tribe, cited two x = 6 species, I. macrocalyx (2n = 12) and I. emarginella
(2n = 24), and concluded that ‘‘… the 48-chromosome Himalayan and EastAsiatic shrubby Indigoferas may not be hexaploids with base number x = 8, but
octoploids in an x = 6 range.’’ None of the species is listed in IPCN, but Reddy
and Revathi (1993) reported 2n = 12 for I. anil, confirming the presence of x = 6
in the genus.
The Schrire et al. (2009) phylogeny does not support the Frahm–Leliveld
(1966) hypothesis. One of the two 2n = 12 species, I. macrocalyx, is placed in the
large Palaeotropical clade of Schrire et al. (2009) and is sister to a group of species
that includes I. pulchra (2n = 16). All three sampled species with 2n = 48 are in
the Palaeotropical clade, but are placed nowhere near I. macrocalyx. In the Pantropical clade, I. rhynchocarpa (2n = 16) is sister to the clade that includes I.
emarginella, which is on a long branch sister to several other species; the only
other species counted from this subclade is also 2n = 16. Thus, there is no evidence that 2n = 48 species are derived from x = 12 species.
Tetraploids based on x = 8 are scattered throughout the phylogeny (Schrire
et al. 2009), supporting the observation of Frahm-Leliveld (1966) that 2n = 32 is
common in the genus. In the Palaeotropical clade, I. atriceps (2n = 32) is in a
TE
732
EC
731
CO
RR
730
F
742
tetraploid levels, despite the existence of morphologically intermediate hybrid
subspecies (Havananda et al. 2011, and unpublished data). Much is known about
the genetics of polyploidy in alfalfa, where unreduced gametes have received
considerable study as a breeding tool (e.g., Bingham 1972; Veronesi et al. 1986;
Tondini et al. 1993; Calderini and Mariani 1997).
Ellison et al. (2006) constructed a phylogeny of Trifolium (2n = 10, 14, 16, 28,
32) that included 218 of its ca. 255 species, as well as species from 11 genera of
the vicioid clade. They showed that the genus is monophyletic; incongruence
within the genus between nuclear and chloroplast markers suggests considerable
hybridization. They also hypothesized a minimum of 19 shifts to aneuploidy and
22 instances of polyploidy from a base number of 2n = 16. They identified the
progenitors of two important species, both shown to be allopolyploids: the
widespread weed, T. dubium, and the most commonly cultivated clover species, T.
repens (Ellison et al. 2006).
729
UN
Editor Proof
168
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 169/179
169
794
9.4.2.7 Millettioids
780
781
782
783
784
785
786
787
788
789
790
791
792
795
796
797
798
799
800
801
802
803
804
805
806
807
808
809
810
811
812
PR
OO
778
779
D
776
777
TE
775
EC
773
774
With the recognition that Wisteria and Callerya are part of the IRLC, and that
Cyclolobium and Poecilanthe belong in the Brongniartieae, chromosome numbers
in the Millettieae (Tephrosieae in Polhill and Raven (1981)) are mostly 2n = 20 or
22, with 2n = 24 in Xeroderris, though many genera have no reported counts in
IPCN. Interestingly, Xeroderris is placed as sister to the remainder of the entire
millettioid clade in Wojciechowski et al. (2004), suggesting that 2n = 20 or 22
may be synapomorphic for the remainder of the millettioid clade (including
phaseoloids; see below). Millettieae comprises the bulk of one of the two major
millettioid clades (core millettioids), along with Abrus (Abreae), and much of the
subtribe Diocleinae of tribe Phaseoleae. Both diploid and tetraploid cytotypes
(2n = 22, 44) have been reported in three species of the ca. 40 IPCN records for
the large (ca. 350 spp.) genus Tephrosia (e.g., Srivastav and Raina 1986). The low
frequency of polyploidy in the core millettioid clade (5 %) is nearly identical to
the frequency in caesalpinioids, both of which are largely woody, tropical groups.
The other large clade (phaseoloids) contains most of the tribe Phaseoleae as
well as the tribes Desmodieae and Psoraleeae and is dominated by 2n = 20 or 22
counts. Polyploidy is more frequent in the phaseoloid clade (17 %), but still less
than half as common as in Hologalegina (Table 9.1). Within the phaseoloid clade,
CO
RR
772
F
793
subclade that also includes diploids. In another subclade, I. mysorensis includes
both 2n = 16 and 32 cytotypes; other two members of its subclade are diploid.
Indigofera microcalyx, in yet another subclade, is 2n = 32; no other members of
its subclade has been counted, but the only counts from its sister clade are diploid.
In the pantropical clade, a small subclade in Schrire et al. (2009) includes I.
koreana (2n = 32), I. grandiflora (2n = 32, 48), I. decora (2n = 48 in Choi and
Kim (1997) but not listed in IPCN), as well as I. venulosa (no count available) and
I. kirilowii (2n = 16). Topologies differ between Schrire et al. (2009) and Choi
and Kim (1997), who focused on this group of mostly Korean endemics. Choi and
Kim (1997) listed I. grandiflora as 2n = 16, and given this count their topology
could suggest independent derivation of polyploidy in I. koreana (from I. grandiflora) and I. decora (from I. venulosa if it is diploid). An alternative explanation
is a single derivation of polyploidy within this clade. Elsewhere in the pantropical
clade, I. heterantha (2n = 48) is sister to I. hebepetala (2n = 16); the clade sister
to these two species includes I. amblyantha (2n = 48) and I. cassioides (2n = 16).
In a different subclade, I. suffruticosa is reported to have both 2n = 16 and 32
cytotypes; the closest reported species to it is diploid. In the Tethyan clade, I.
sessiliflora (2n = 32) is the only member of its subclade with a count in IPCN.
Indigofera hochstetteri has both 2n = 16 and 32 counts; its sister species, I.
arabica, is diploid. Indigofera angulosa is 2n = 32; no other species in its clade
has counts in IPCN.
Thus, there appear to be no large clades composed exclusively of polyploids in
Indigofera. Instead, as in other large legume genera, polyploidy is sporadic.
771
UN
Editor Proof
9 Polyploidy in Legumes
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 170/179
820
821
822
823
824
825
826
827
828
829
830
831
832
833
834
835
836
837
838
839
840
841
842
843
844
845
846
847
848
849
850
851
852
853
854
855
856
F
PR
OO
818
819
D
816
817
TE
815
chromosome numbers suggest that several genera are polyploid. The best known
of these is Glycine, with around 30 species whose lowest chromosome numbers are
2n = 38 and 40, in contrast to most of its phylogenetic neighbors (e.g., Doyle et al.
2003; Stefanovic et al. 2009) which are typical millettioids with 2n = 20 or 22. As
noted above, genomic data confirm the presence of two cycles of polyploidy in G.
max (soybean) since the origin of the legumes. The more recent of these has
resulted in homoeologous gene pairs that diverged around 10 MYA (Shoemaker
et al. 2006; Egan and Doyle 2010), setting a maximum date for the polyploidy
event, with the minimum date set by the earliest divergence of the various Glycine
species around 5 MYA (Innes et al. 2008; Doyle and Egan 2010). Phylogenetic
evidence is consistent either with autopolyploidy or with allopolyploidy from
extinct species more closely related to one another than to any extant genera
outside of Glycine (Straub et al. 2006). The presence of two classes of centromeric
heterochromatin repeats suggests that Glycine could be an allopolyploid, with the
two repeat types each derived from one of the diploid progenitor species (Gill et al.
2009). Such a hypothesis is difficult to test due to the extensive rearrangement of
homoeologous segments in the soybean genome (Schmutz et al. 2010) and also
requires complex patterns of concerted evolution among repeats on different
chromosomes.
At least three other phaseoloid genera are likely to be exclusively polyploid
based on chromosome number alone (Lackey 1981). In Erythrina (coral bean), all
sampled species are 2n = 42. The single count in IPCN for the small genus
Cologania is 2n = 44, and the monotypic Teyleria is also 2n = 44 (Kumar and
Hymowitz 1989). Goldblatt (1981) considered Calopogonium, with counts of
2n = 36 and ca. 37 in C. mucunoides, to be a polyploid, presumably with aneuploid reduction from a base of x = 10; however, Gill and Husaini (1986) reported
a count of 2n = 24, which could suggest a more recent derivation of polyploidy
within the genus. Similarly, counts of 2n = 28—considered polyploid in T. mollis
by Kumari and Bir (1990)—predominate in Teramnus species, though T. labialis is
variously listed as 2n = 20, 22, and 28. Strongylodon is also 2n = 28.
Polyploidy appears to be rare within Phaseoleae genera. Even relatively large
and well-studied genera such as Rhynchosia (ca. 230 species), Phaseolus (60–65
species), and Vigna (ca. 100 species) were reported in Polhill and Raven (1981) as
being exclusively diploid, though Sen and Bhattacharya (1988) later reported a
count of 2n = 44 in V. glabrescens. Polyploidy has also been reported within
species of Amphicarpaea and Neonotonia by Kumar and Hymowitz (1989; both
2n = 22, 44). Apios americana includes both diploid and triploid cytotypes
(2n = 22, 33); Joly and Bruneau (2004) reported multiple origins of autotriploidy
and high heterozygosity in this species. Glycine not only is a relatively recent
polyploid at the generic level (see above) but also includes several recently formed
allopolyploid species whose genomic relationships to extant diploids have been
worked out using molecular phylogenies (reviewed by Doyle et al. 2004), and
which are the focus of physiological and transcriptomic studies (Coate and Doyle
2010; Ilut et al. (in press)).
EC
814
CO
RR
813
J. J. Doyle
UN
Editor Proof
170
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 171/179
865
866
867
868
869
870
871
872
873
874
875
876
877
878
879
880
881
882
883
884
885
886
887
888
889
890
891
892
893
894
895
896
897
F
PR
OO
864
9.5 Searching for Cryptic Polyploidy in the Phaseoloid
Legumes
Clearly, all papilionoids are fundamentally polyploid, even those with low chromosome numbers. The tempo and mechanism(s) of chromosomal diploidization
are unknown (Doyle et al. 2008; Soltis et al. 2010), and without that information it
is difficult to estimate the prevalence of cryptic polyploidy. As noted above,
consideration of the divergence times for major lineages suggests that the rate of
chromosomal diploidization is rapid—likely 10 MY or less.
On the other hand, ‘‘polyploid’’ chromosome numbers have persisted for at
least 5–10 MY in Glycine (Fig. 9.2). A cryptic polyploid papilionoid legume is a
taxon that has experienced an additional polyploidy event subsequent to the ca. 50
MYA papilionoid WGD but has a low chromosome number typical of its clade.
Thus, in the phaseoloid clade, cryptic polyploids would have chromosome numbers of 2n = 20 or 22. Polyploids on the way to diploidization would have
chromosome numbers between these numbers and 2n = 40-44. As noted above,
Calopogonium and Teramnus are candidates for this class; one perennial Glycine
species with 2n = 38 is likely at the first stages of this process.
We know from the paralog Ks profile of Glycine (Schleueter et al. 2004) that no
additional polyploidy events took place between the two WGD episodes detectable
in its genome. Therefore, we can infer that all of the ancestors of Glycine experienced only the papilionoid WGD; this includes the ancestors that form the backbone of the phaseoloid clade (Fig. 9.4), as well as the common ancestor of
phaseoloids and Indigofereae, and also its common ancestor with the IRLC clade.
Given these conditions, candidates for cryptic polyploidy include lineages that are
connected to the phaseoloid backbone by branches longer than 5–10 MY. This
includes several major groups, such as subtribes Phaseolinae and Cajaninae
(Fig. 9.4). Initial sampling of one species of each lineage would provide information on another set of ancestors, and subsequent searches can then be focused on
lineages connected to these ancestors by suitably long branches. This has now been
done for Cajanus cajan (pigeonpea), which Varshney et al. (2011) have shown has
no history of recent polyploidy. We are sampling other phaseoloids using 454
D
862
863
TE
861
Polyploidy occurs within at least one genus of Desmodieae, Lespedeza, which
was listed in Polhill and Raven (1981) as 2n = 18, 20, 22, 36. However, IPCN
gives higher numbers, for example L. bicolor with both 2n = 22 and 42 cytotypes,
as well as L. daurica and L. potaninii, both exclusively 2n = 42. Triploidy occurs
in Campylotropis polyantha var. leiocarpa (2n = 22, 33) and possibly in the genus
Pseudarthria, listed as 2n = 22, 26, 34. Only diploid counts (2n = 20, 22) were
reported by Ohashi et al. (1981) from the large (ca. 275 spp.) genus, Desmodium.
However, additional counts are found in IPCN, both at the diploid (2n = 24, 26)
and tetraploid levels, the latter in D. styracifolium (2n = 42) and D. incanum
(2n = 22, 44).
EC
859
860
171
CO
RR
857
858
UN
Editor Proof
9 Polyploidy in Legumes
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 172/179
Bituminaria
Cullen
Glycine 2n = 38, 40
Teramnus 2n = 28
Amphicarpaea 2n = 22, 44
Pueraria lobata
Neonotonia 2n = 22, 44
Pachyrhizus
Pueraria phaseoloides
Pseudovigna
Dumasia
Cologania 2n = 44
Macroptilium
Vigna
Macrotyloma
Psophocarpus
Erythrina 2n = 42
Cajanus
Bolusafra
Hardenbergia
Kennedia
Lespedeza
Campylotropis
Desmodium pauciflorum
Desmodium barbatum
Mucuna
Shuteria
Apios 2n = 22, 33
20
10
million years
TE
D
PR
OO
}
polyploidy
F
J. J. Doyle
Editor Proof
172
CO
RR
EC
Fig. 9.4 Phylogeny and polyploidy in the phaseoloid clade. Topology and dates of the
chronogram are taken from Stefanovic et al. (2009). Chromosome numbers of the species used in
that study are color coded as follows: Green, 2n = 20; blue, 2n = 22; black, 2n = 18; red,
known or possible polyploid numbers, with the numbers shown following the taxon name.
Species known to have multiple cytotypes are indicated by multiple colors corresponding to
chromosome numbers listed above. The two numbers shown for Glycine are from different
species, only one of which (G. max, 2n = 40) was used in the Stefanovic et al. (2009) study. The
range of dates for the polyploidy event in Glycine is indicated. Yellow dots indicate ancestral
nodes that lacked any polyploidy event subsequent to the papilionoid WGD
903
9.6 Conclusions
899
900
901
904
905
906
UN
902
transcriptome sequencing, including taxa with both low and high chromosome
numbers; for the latter we wish to estimate maximum ages of polyploidy. Thus far
we have not found examples of cryptic polyploidy, but have determined that
polyploidy in Erythrina probably took place on roughly the same time scale as in
Glycine (\10 MYA; Egan and Doyle, unpublished data).
898
Among the most persistent questions concerning polyploidy in plants are how
successful the phenomenon is as an evolutionary mechanism. Is polyploidy a ticket
to innovation, adaptation, invasiveness, survival in the face of global catastrophes,
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 173/179
173
942
943
944
945
946
947
948
Acknowledgments I thank many colleagues and lab members for discussions of polyploidy, and
Jane Doyle for her support and encouragement. I also thank Jane Doyle, Sue Sherman-Broyles,
Iben Sorensen, and Toby Pennington for critical reading of the manuscript, and Melissa Luckow
for help with mimosoid systematics. I am grateful for many years of funding from the National
Science Foundation for work on polyploidy, most recently grants DEB-0948800, IOS-0939423,
IOS-0822258, and IOS-0744306. I thank Doug Soltis for helpful suggestions in review of the
manuscript.
913
914
915
916
917
918
919
920
921
922
923
924
925
926
927
928
929
930
931
932
933
934
935
936
937
938
939
940
PR
OO
912
D
911
TE
910
EC
909
CO
RR
908
F
941
or is it an evolutionary dead end … or both? Based on rates of polyploid formation
and extinction in the phylogenetic record, Mayrose et al. (2011) conclude that
‘‘polyploidy is most often an evolutionary dead end, but the possibility remains
that the expanded genomic potential of those polyploids that do persist drives longterm evolutionary success.’’
Legumes may illustrate both of these points. The most diverse and species-rich
clade of this third largest family of flowering plants, the core papilionoids, is
ancestrally polyploid. Clearly, the ancestor of this group of around 450 genera and
13,000 species, like the ancestor of seed plants and the ancestor of flowering plants
(Jiao et al. 2011), was most emphatically not a ‘‘dead end.’’ It remains to be
determined whether there is a perfect correlation between the papilionoid polyploidy event and the origin of nodulation in core papilionoids, and it will take
much more work to demonstrate that the two are causally related (Doyle 2011;
Young et al. 2011). It is also clear that nodulation is not sufficient to explain the
explosive radiation of papilionoid legumes, because other nodulating groups both
in legumes and elsewhere in the rosids have not proliferated to the same extent as
papilionoids (Doyle 2011).
Despite the obvious success of the core papilionoid lineage, the pattern of
evolution within the core papilionoids suggests that polyploidy has not been a
major feature in establishing new lineages, similar to the conclusion of Mayrose
et al. (2011) for angiosperms generally. It is not that polyploidy is rare within the
family—indeed, around a quarter of all legumes for which chromosome data are
available have one or more species that are polyploid (Table 9.1). However, much
of the polyploidy in the family occurs as single polyploid genera embedded within
diploids, as scattered species within genera, or as multiple cytotypes within species. Two significant exceptions are the Genisteae, which may be entirely polyploid and within which nearly all genera show a propensity for polyploidy and
aneuploidy, and the lineage that includes the IRLC tribe Carmichaelieae. The
largest papilionoid genera, including Astragalus, are not fundamentally polyploid.
‘‘Success’’ is a very ambiguous term and can be measured in many ways.
Species with short evolutionary histories that have not been involved in subsequent
speciation, yet have invaded extensive new territories and had major impact on the
environment, certainly could be considered ‘‘successful.’’ Many plant polyploids,
including genera and species of legumes, fit this description. So does Homo
sapiens.
907
UN
Editor Proof
9 Polyploidy in Legumes
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 174/179
TE
D
PR
OO
F
Ahangarian S, Osaloo SK, Maassoumi AA (2007) Molecular phylogeny of the tribe Hedysareae
with special reference to Onobrychis (Fabaceae) as inferred from nrDNA ITS sequences. Iran
J Bot 13:64–74
Ainouche A, Bayer RJ, Misset M- (2004) Molecular phylogeny, diversification and character
evolution in Lupinus (Fabaceae) with special attention to Mediterranean and African lupines.
Plant Syst Evol 246:211–222
Artyukova EV, Kozyrenko MM, Kholina AB, Zhuravlev YN (2011) High chloroplast haplotype
diversity in the endemic legume Oxytropis chankaensis may result from independent
polyploidization events. Genetica 139:221–232
Bell CD, Soltis DE, Soltis PS (2010) The age and diversification of the angiosperms re-revisited.
Am J Bot 97:296–313
Bello MA, Bruneau A, Forest F, Hawkins JA (2009) Elusive relationships within order Fabales:
phylogenetic analyses using matK and rbcL sequence data. Syst Bot 34:102–114
Bertioli D, Moretzsohn M, Madsen L, Sandal N, Leal-Bertioli S, Guimaraes P, Hougaard B,
Fredslund J, Schauser L, Nielsen A, Sato S, Tabata S, Cannon S, Stougaard J (2009) An
analysis of synteny of Arachis with Lotus and Medicago sheds new light on the structure,
stability and evolution of legume genomes. BMC Genomics 10:45
Bessega C, Vilardi JC, Saidman BO (2006) Genetic relationships among American species of the
genus Prosopis (Mimosoideae, Leguminosae) inferred from ITS sequences: evidence for longdistance dispersal. J Biogeogr 33:1905–1915
Bingham ET (1972) Sexual poly ploidy in Medicago-Sativa-D. Genetics 71:S5
Bisby FA (1981) Genisteae. In: Polhill RM, Raven PH (eds) Advances in legume systematics,
Part 1. Royal Botanic Gardens, Kew, pp 409–425
Blanc G, Wolfe KH (2004) Widespread paleopolyploidy in model plant species inferred from age
distributions of duplicate genes. Plant Cell 16:1667–1678
Boatwright JS, Van Wyk B- (2011) The systematic position of Sophora inhambanensis
(Fabaceae: Sophoreae). S Afr J Bot 77:249–250
Boatwright JS, Savolainen V, Van Wyk B, Schutte-Vlok AL, Forest F, van der Bank M (2008)
Systematic position of the anomalous genus Cadia and the phylogeny of the tribe Podalyrieae
(Fabaceae). Syst Bot 33:133–147
Boff T, Schifino-Wittmann MT (2003) Segmental allopolyploidy and paleopolyploidy in species
of Leucaena Benth: evidence from meiotic behaviour analysis. Hereditas (Lund) 138:27–35
Brown GK, Clowes C, Murphy DJ, Ladiges PY (2010) Phylogenetic analysis based on nuclear
DNA and morphology defines a clade of eastern Australian species of Acacia s.s. (section
Juliflorae): the ‘Acacia longifolia group’. Aust Syst Bot 23:162–172
Bruneau A, Mercure M, Lewis GP, Herendeen PS (2008) Phylogenetic patterns and
diversification in the caesalpinioid legumes. Botany-Botanique 86:697–718
Burow MD, Simpson CE, Faries MW, Starr JL, Paterson AH (2009) Molecular biogeographic
study of recently described B- and A-genome Arachis species, also providing new insights
into the origins of cultivated peanut. Genome 52:107–119
Calderini O, Mariani A (1997) Increasing 2n gamete production in diploid alfalfa by cycles of
phenotypic recurrent selection. Euphytica 93:113–118
Cannon SB, Ilut D, Farmer AD, Maki SL, May GD, Singer SR, Doyle JJ (2010) Polyploidy did
not predate the evolution of nodulation in all legumes. PLoS ONE 5:e11630
Catalano SA, Vilardi JC, Tosto D, Saidman BO (2008) Molecular phylogeny and diversification
history of Prosopis (Fabaceae: Mimosoideae). Biol J Linn Soc 93:621–640
Chandler GT, Bayer RJ, Crisp MD (2001) A molecular phylogeny of the endemic Australian
genus Gastrolobium (Fabaceae: Mirbelieae) and allied genera using chloroplast and nuclear
markers. Am J Bot 88:1675–1687
Choi B, Kim J (1997) ITS sequences and speciation on far eastern Indigofera (Leguminosae).
J Plant Res 110:339–346
EC
950
951
952
953
954
955
956
957
958
959
960
961
962
963
964
965
966
967
968
969
970
971
972
973
974
975
976
977
978
979
980
981
982
983
984
985
986
987
988
989
990
991
992
993
994
995
996
997
998
999
1000
References
CO
RR
949
J. J. Doyle
UN
Editor Proof
174
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 175/179
175
EC
TE
D
PR
OO
F
Chooi WY (1971) Variation in nuclear DNA content in the genus Vicia-D. Genetics 68:195–211
Coate JE, Doyle JJ (2010) Quantifying whole transcriptome size, a prerequisite for understanding
transcriptome evolution across species: an example from a plant allopolyploid. Genome Biol
Evol 2:534–546
Conterato IF, Schifino-Wittmann MT (2006) New chromosome numbers, meiotic behaviour and
pollen fertility in American taxa of Lupinus (Leguminosae): contributions to taxonomic and
evolutionary studies. Bot J Linn Soc 150:229–240
Cowan RS, Polhill RM (1981) Amherstieae. In: Polhill RM, Raven PH (eds) Advances in legume
systematics, Part 1. Royal Botanic Gardens, Kew, pp 135–142
Cubas P, Pardo C, Tahiri H (2002) Molecular approach to the phylogeny and systematics of
cytisus (Leguminosae) and related genera based on nucleotide sequences of nrDNA (ITS
region) and cpDNA (trnL-trnF intergenic spacer). Plant Syst Evol 233:223–242
Cusma-Velari T, Feoli-Chiapella L (2009) The so-called primitive genera of Genisteae
(Fabaceae): systematic and phyletic considerations based on karyological data. Bot J Linn
Soc 160:232–248
Dahmer N, Simon MF, Schifino-Wittmann MT, Hughes CE, Sfoggia Miotto ST, Giuliani JC
(2011) Chromosome numbers in the genus Mimosa L.: cytotaxonomic and evolutionary
implications. Plant Syst Evol 291:211–220
Degtjareva GV, Kramina TE, Sokoloff DD, Samigullin TH, Valiejo-Roman CM, Antonov AS
(2006) Phylogeny of the genus Lotus (Leguminosae, Loteae): evidence from nrITS sequences
and morphology. Can J Bot 84:813–830
Doyle JJ (2011) Phylogenetic perspectives on the origins of nodulation. Molec Plant Microbe
Interact 24:1289–1295
Doyle JL, Rauscher JT, Brown AHD (2004) Diploid and polyploid reticulate evolution
throughout the history of the perennial soybeans (Glycine …. New Phytologist)
Doyle JJ, Egan AN (2010) Dating the origins of polyploidy events. New Phytol 186:73–85
Doyle JJ, Flagel LE, Paterson AH, Rapp RA, Soltis DE, Soltis PS, Wendel JF (2008)
Evolutionary genetics of genome merger and doubling in plants. Annu Rev Genet 42:443–461
Doyle JJ, Doyle JL, Harbison C (2003) Chloroplast-expressed glutamine synthetase in Glycine
and related Leguminosae: phylogeny, gene duplication, and ancient polyploidy. Syst Bot
28:567–577
Doyle JJ, Luckow MA (2003) The rest of the iceberg. Legume diversity and evolution in a
phylogenetic context. Plant Physiol (Rockville) 131:900–910
Drummond CS (2008) Diversification of Lupinus (Leguminosae) in the western new world:
derived evolution of perennial life history and colonization of montane habitats. Mol
Phylogenet Evol 48:408–421
Drummond CS, Eastwood RJ, Miotto STS, Hughes CE (2012) Multiple continental radiations
and correlates of diversification in lupinus (Leguminosae): testing for key innovation with
incomplete taxon sampling. Syst Biol 61:443–460
Egan AN, Doyle J (2010) A comparison of global, gene-specific, and relaxed clock methods in a
comparative genomics framework: dating the polyploid history of soybean (Glycine max).
Syst Biol 59:534–547
Ellison NW, Liston A, Steiner JJ, Williams WM, Taylor NL (2006) Molecular phylogenetics of
the clover genus (Trifolium––Leguminosae). Mol Phylogenet Evol 39:688–705
Endo Y, Choi B, Ohashi H, Delgado-Salinas A (2008) Phylogenetic relationships of New World
Vicia (Leguminosae) inferred from nrDNA internal transcribed spacer sequences and floral
characters. Syst Bot 33:356–363
Fawcett JA, Maere S, Van de Peer Y (2009) Plants with double genomes might have had a better
chance to survive the cretaceous-tertiary extinction event. Proc Natl Acad Sci U S A
106:5737–5742
Frahm-Leliveld JA (1966) Cytotaxonomic notes on the genera Indigofera L. and Cyamopsis DC.
[Leguminosae]. Genetica 37:403–426
Freeling M, Thomas BC (2006) Gene-balanced duplications, like tetraploidy, provide predictable
drive to increase morphological complexity. Genome Res 16:805–814
CO
RR
1001
1002
1003
1004
1005
1006
1007
1008
1009
1010
1011
1012
1013
1014
1015
1016
1017
1018
1019
1020
1021
1022
1023
1024
1025
1026
1027
1028
1029
1030
1031
1032
1033
1034
1035
1036
1037
1038
1039
1040
1041
1042
1043
1044
1045
1046
1047
1048
1049
1050
1051
1052
1053
1054
UN
Editor Proof
9 Polyploidy in Legumes
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 176/179
EC
TE
D
PR
OO
F
Gallagher RV, Leishman MR, Miller JT, Hui C, Richardson DM, Suda J, Travnicek P (2011)
Invasiveness in introduced Australian Acacias: the role of species traits and geneome size.
Divers Distrib 17:884–897
Gauthier P, Lumaret R, Bedecarrats A (1998a) Genetic variation and gene flow in Alpine diploid
and tetraploid populations of Lotus (L. alpinus (D.C.) Schleicher/L. corniculatus L.).
I. Insights from morphological and allozyme markers. Heredity 80:683–693
Gauthier P, Lumaret R, Bedecarrats A (1998b) Genetic variation and gene flow in Alpine diploid
and tetraploid populations of Lotus (L. alpinus (D.C.) Schleicher/L. corniculatus L.). II.
Insights from RFLP of chloroplast DNA. Heredity 80:694–701
Gill LS, Husaini (1986) Cytological observations in Leguminosae from southern Nigeria.
Willdenowia 15:521–527
Gill N, Findley S, Walling JG, Hans C, Ma J, Doyle J, Stacey G, Jackson SA (2009) Molecular
and chromosomal evidence for allopolyploidy in soybean. Plant Physiol 151:1167–1174
Gohil RN, Ashraf M (2008) Cytological parameters viz a viz probable modes of evolution in
Astragalus L. Proc Nat Acad Sci India Sect B-Biol Sci 78:281–287
Goldblatt P (1989) Miscellaneous chromosome counts in Asteraceae Bignoniaceae Proteaceae
and Fabaceae. Ann Mo Bot Gard 76:1186–1188
Goldblatt P (1981) Cytology and the phylogeny of leguminosae. In: Polhill RM, Raven PH (eds)
Advances in legume systematics, Part 2. Royal Botanic Gardens, Kew, pp 427–464
Govindarajulu R, Hughes CE, Bailey D (2011a) Phylogenetic and population genetic analyses of
diploid Leucaena (Leguminosae-Mimosoideae) reveal cryptic species diversity and patterns of
divergent allopatric speciation. Am J Bot 98:2049–2063
Govindarajulu R, Hughes CE, Alexander P, Bailey D (2011b) The complex dynamics of ancient
and recent polyploidy in Leucaena (Leguminosae). Am J Bot 98:2064–2076
Grant WF, Small E (1996) The origin of the Lotus corniculatus (Fabaceae) complex: a synthesis
of diverse evidence. Can J Bot 74:975–989
Gutierrez JF, Vaquero F, Vences FJ (1994) Allopolyploid vs. autopolyploid origins in the genus
Lathyrus (Leguminosae). Heredity 73:29–40
Hanson L (1995) Some new chromosome counts in the genus Inga (Leguminosae: Mimosoideae).
Kew Bull 50:801–804
Havananda T, Brummer EC, Maureira-Butler IJ, Doyle JJ (2010) Relationships among diploid
members of the Medicago sativa (Fabaceae) species complex based on chloroplast and
mitochondrial DNA sequences. Syst Bot 35:140–150
Havananda T, Brummer EC, Doyle JJ (2011) Complex patterns of autopolyploid evolution in
alfalfa and allies (Medicago sativa: Leguminosae). Am J Bot 98:1633-1646
Hejazi H, Mohsen S, Nasab MZ (2010) Cytotaxonomy of some Onobrychis (Fabaceae) species
and populations in Iran. Caryologia 63:18–31
Hughes CE, Eastwood R (2006) Island radiation on a continental scale: exceptional rates of plant
diversification after uplift of the Andes. Proc Natl Acad Sci U S A 103:10334–10339
Hughes CE, Govindarajulu R, Robertson A, Filer DL, Harris SA, Bailey CD (2007) Serendipitous
backyard hybridization and the origin of crops. Proc Natl Acad Sci U S A 104:14389–14394
Hughes CE, Bailey CD, Harris SA (2002) Divergent and reticulate species relationships in
Leucaena (Fabaceae) inferred from multiple data sources: insights into polyploid origins and
nrDNA polymorphism. Am J Bot 89:1057–1073
Hughes CE, Bailey CD, Krosnick S, Luckow MA (2003) Relationships among genera of the
informal Dichrostachys and Leucaena groups (Mimosoideae) inferred from nuclear ribosomal
ITS sequences. In: Klitgaard B, Bruneau A (eds) Advances in legume systematics, Part 1.0.
Royal Botanic Gardens, Kew, pp 221–238
Hulina N (2010) ‘‘Planta hortifuga’’ in flora of the continental part of Croatia. Agriculturae
Conspectus Scientificus 75:57–65
Ilut DC, Coate JE, Luciano AK, Owens TG, May GD, Farmer A, Doyle JJ (2012) A comparative
transcriptomic study of an allotetraploid and its diploid progenitors illustrates the unique
advantages and challenges of RNA-Seq in plant species. Am J Bot 99:383–396
CO
RR
1055
1056
1057
1058
1059
1060
1061
1062
1063
1064
1065
1066
1067
1068
1069
1070
1071
1072
1073
1074
1075
1076
1077
1078
1079
1080
1081
1082
1083
1084
1085
1086
1087
1088
1089
1090
1091
1092
1093
1094
1095
1096
1097
1098
1099
1100
1101
1102
1103
1104
1105
1106
1107
J. J. Doyle
UN
Editor Proof
176
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 177/179
177
EC
TE
D
PR
OO
F
Innes RW, Ameline-Torregrosa C, Ashfield T, Cannon E, Cannon SB, Chacko B, Chen NWG,
Couloux A, Dalwani A, Denny R, Deshpande S, Egan AN, Glover N, Hans CS, Howell S, Ilut
D, Jackson S, Lai H, Mammadov J, del Campo SM, Metcalf M, Nguyen A, O’Bleness M,
Pfeil BE, Podicheti R, Ratnaparkhe MB, Samain S, Sanders I, Segurens B, Sevignac M,
Sherman-Broyles S, Thareau V, Tucker DM, Walling J, Wawrzynski A, Yi J, Doyle JJ,
Geffroy V, Roe BA, Maroof MAS, Young ND (2008) Differential accumulation of
retroelements and diversification of NB-LRR disease resistance genes in duplicated regions
following polyploidy in the ancestor of soybean. Plant Physiol (Rockville) 148:1740–1759
Jaillon O, Aury J, Noel B, Policriti A, Clepet C, Casagrande A, Choisne N, Aubourg S, Vitulo N,
Jubin C, Vezzi A, Legeai F, Hugueney P, Dasilva C, Horner D, Mica E, Jublot D, Poulain J,
Bruyere C, Billault A, Segurens B, Gouyvenoux M, Ugarte E, Cattonaro F, Anthouard V,
Vico V, Del Fabbro C, Alaux M, Di Gaspero G, Dumas V, Felice N, Paillard S, Juman I,
Moroldo M, Scalabrin S, Canaguier A, Le Clainche I, Malacrida G, Durand E, Pesole G,
Laucou V, Chatelet P, Merdinoglu D, Delledonne M, Pezzotti M, Lecharny A, Scarpelli C,
Artiguenave F, Pe ME, Valle G, Morgante M, Caboche M, Adam-Blondon A, Weissenbach J,
Quetier F, Wincker P, French-Italian Public (2007) The grapevine genome sequence suggests
ancestral hexaploidization in major angiosperm phyla. Nature (London) 449:463
Jenczewski E, Prosperi J, Ronfort J (1999) Evidence for gene flow between wild and cultivated
Medicago sativa (Leguminosae) based on allozyme markers and quantitative traits. Am J Bot
86:677–687
Jiao Y, Wickett NJ, Ayyampalayam S, Chanderbali AS, Landherr L, Ralph PE, Tomsho LP, Hu
Y, Liang H, Soltis PS, Soltis DE, Clifton SW, Schlarbaum SE, Schuster SC, Ma H, LeebensMack J, dePamphilis CW (2011) Ancestral polyploidy in seed plants and angiosperms. Nature
473:97–100
Joly S, Bruneau A (2004) Evolution of triploidy in Apios americana (Leguminosae) revealed by
genealogical analysis of the histone H3-D gene. Evolution 58:284–295
Jorgensen JL, Stehlik I, Brochmann C, Conti E (2003) Implications of ITS sequences and RAPD
markers for the taxonomy and biogeography of the Oxytropis campestris and O. arctica
(Fabaceae) complexes in Alaska. Am J Bot 90:1470–1480
Kajita T, Ohashi H, Tateishi Y, Bailey CD, Doyle JJ (2001) rbcL and legume phylogeny, with
particular reference to phaseoleae, millettieae, and allies. Syst Bot 26:515–536
Kloda JM, Dean PDG, Maddren C, MacDonald DW, Mayes S (2008) Using principle component
analysis to compare genetic diversity across polyploidy levels within plant complexes: an
example from British Restharrows (Ononis spinosa and Ononis repens). Heredity
100:253–260
Kumar PS, Hymowitz T (1989) Where are the diploid 2n equals 2x equals 20 genome donors of
glycine willd. Leguminosae Papilionoideae. Euphytica 40:221–226
Kumari S, Bir SS (1990) Karyomorphological evolution in Papilionaceae. J Cytol Genet
25:173–219
Kupicha FK (1981) Vicieae. In: Polhill RM, Raven PH (eds) Advances in legume systematics,
Part 1. Royal Botanic Gardens, Kew, pp 377–381
Lackey JA (1981) Phaseoleae. In: Polhill RM, Raven PH (eds) Advances in legume systematics,
Part 1. Royal Botanic Gardens, Kew, pp 301–328
Lavia GI, Ortiz MA, Robledo G, Fernandez A, Seijo G (2011) Origin of triploid Arachis pintoi
(Leguminosae) by autopolyploidy evidenced by FISH and meiotic behaviour. Ann Bot
(London) 108:103–111
Lavin M, Herendeen PS, Wojciechowski MF (2005) Evolutionary rates analysis of Leguminosae
implicates a rapid diversification of lineages during the tertiary. Syst Biol 54:575–594
Lavin M, Pennington RT, Klitgaard BB, Sprent JI, de Lima HC, Gasson PE (2001) The
dalbergioid legumes (Fabaceae): delimitation of a pantropical monophyletic clade. Am J Bot
88:503–533
Leitch IJ, Bennett MD (2004) Genome downsizing in polyploid plants. Biol J Linn Soc
82:651–663
CO
RR
1108
1109
1110
1111
1112
1113
1114
1115
1116
1117
1118
1119
1120
1121
1122
1123
1124
1125
1126
1127
1128
1129
1130
1131
1132
1133
1134
1135
1136
1137
1138
1139
1140
1141
1142
1143
1144
1145
1146
1147
1148
1149
1150
1151
1152
1153
1154
1155
1156
1157
1158
1159
1160
UN
Editor Proof
9 Polyploidy in Legumes
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 178/179
EC
TE
D
PR
OO
F
Lewis GP, Schrire B, MacKinder B, Lock M (2005) Legumes of the world. Royal Botanic
Gardens, Kew
Luckow M, Fortunato RH, Sede S, Livshultz T (2005) The phylogenetic affinities of two
mysterious monotypic mimosoids from southern South America. Syst Bot 30:585–602
Luckow M, Miller JT, Murphy DJ, Livshultz T (2003) A phylogenetic analysis of the
Mimosoideae (Leguminosae) based on chloroplast DNA sequence data. In: Klitgaard B,
Bruneau A (eds) Advances in legume systematics, Part 1.0. Royal Botanic Gardens, Kew,
pp 197–220
Lynch M, Conery JS (2003) The origins of genome complexity. Science 302:1401–1404
Mayrose I, Zhan SH, Rothfels CJ, Magnuson-Ford K, Barker MS, Rieseberg LH, Otto SP (2011)
Recently formed polyploid plants diversify at lower rates. Science (Washington DC)
333:1257
McMahon MM (2005) Phylogenetic relationships and floral evolution in the papilionoid legume
clade Amorpheae. Brittonia 57:397–411
Morales M, Wulff AF, Fortunato RH, Poggio L (2010) Chromosome and morphological studies
in the Mimosa debilis complex (Mimosoideae, Leguminosae) from southern South America.
Aust J Bot 58:12–22
Murphy DJ, Brown GK, Miller JT, Ladiges PY (2010) Molecular phylogeny of Acacia Mill.
(Mimosoideae: Leguminosae): evidence for major clades and informal classification. Taxon
59:7–19
Neumann P, Koblizkova A, Navratilova A, Macas J (2006) Significant expansion of Vicia
pannonica genome size mediated by amplification of a single type of giant retroelement.
Genetics 173:1047–1056
Ohashi H, Polhill RM, Schubert BG (1981) Desmodieae. In: Polhill RM, Raven PH (eds)
Advances in legume systematics, Part 1. Royal Botanic Gardens, Kew, pp 292–300
Orthia LA, Cook LG, Crisp MD (2005) Generic delimitation and phylogenetic uncertainty: an
example from a group that has undergone an explosive radiation. Aust Syst Bot 18:41–47
Ortiz MA, Guillermo Seijo J, Fernandez A, Lavia GI (2011) Meiotic behavior and pollen viability
of tetraploid Arachis glabrata and A. nitida species (section Rhizomatosae, Leguminosae):
implications concerning their polyploid nature and seed set production. Plant Syst Evol
292:73–83
Ossowski S, Schneeberger K, Lucas-Lledo JI, Warthmann N, Clark RM, Shaw RG, Weigel D,
Lynch M (2010) The rate and molecular spectrum of spontaneous mutations in Arabidopsis
thaliana RID E-2139-2011 RID C-1418-2008. Science 327:92–94
Pandit MK, Tan HTW, Bisht MS (2006) Polyploidy in invasive plant species of Singapore. Bot J
Linn Soc 151:395–403
Pandit MK, Pocock MJO, Kunin WE (2011) Ploidy influences rarity and invasiveness in plants.
J Ecol 99: 1108-1115
Pardo C, Cubas P, Tahiri H (2004) Molecular phylogeny and systematics of Genista
(Leguminosae) and related genera based on nucleotide sequences of nrDNA (ITS region)
and cpDNA (trnL-trnF intergenic spacer). Plant Syst Evol 244:93–119
Pennington, RT, Klitgaard BB, Ireland H, Lavin M (2000) New insights into floral evolution of
basal Papilionoideae from molecular phylogenies. In: Herendeen PS Bruneau A (eds.)
Advances in legume systematics Part 9. Royal Botanic Gardens, Kew p 233–248
Pennington, RT, Lavin M, Ireland H, Klitgaard BB, Preston J, Hu J-M (2001) Phylogenetic
relationships of basal papilionoid legumes based upon sequences of the chloroplast trnL
intron. Syst Bot 26:537–556
Pfeil BE, Schlueter JA, Shoemaker RC, Doyle JJ (2005) Placing paleopolyploidy in relation to
taxon divergence: a phylogenetic analysis in legumes using 39 gene families. Syst Biol
54:441–454
Polhill RM (1981a) Dipteryxeae. In: Polhill RM, Raven PH (eds) Advances in legume
systematics, Part 1. Royal Botanic Gardens, Kew, pp 231–232
Polhill RM (1981b) Indigofereae. In: Polhill RM, Raven PH (eds) Advances in legume
systematics, Part 1. Royal Botanic Gardens, Kew, pp 289–291
CO
RR
1161
1162
1163
1164
1165
1166
1167
1168
1169
1170
1171
1172
1173
1174
1175
1176
1177
1178
1179
1180
1181
1182
1183
1184
1185
1186
1187
1188
1189
1190
1191
1192
1193
1194
1195
1196
1197
1198
1199
1200
1201
1202
1203
1204
1205
1206
1207
1208
1209
1210
1211
1212
1213
1214
J. J. Doyle
UN
Editor Proof
178
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 179/179
179
EC
TE
D
PR
OO
F
Polhill RM, Raven PH (1981) Advances in legume systematics, Part 1. Royal Botanic Gardens,
Kew
Polhill RM, Sousa M (1981) Robinieae. In: Polhill RM, Raven PH (eds) Advances in legume
systematics, Part 1. Royal Botanic Gardens, Kew, pp 283–288
Reddy VRK, Revathi R (1993) Chemotaxonomic studies in the genus Indigofera Linn. J Econ
Taxon Bot 17:115–120
Rosato M, Castro M, Rossello JA (2008) Relationships of the woody Medicago species (section
Dendrotelis) assessed by molecular cytogenetic analyses. Ann Bot (London) 102:15–22
Rossello JA, Castro M (2008) Karyological evolution of the angiosperm endemic flora of the
Balearic Islands. Taxon 57:259–273
Sakiroglu M, Doyle JJ, Brummer EC (2010) Inferring population structure and genetic diversity
of broad range of wild diploid alfalfa (Medicago sativa L.) accessions using SSR markers.
Theor Appl Genet 121:403–415
Sato S, Nakamura Y, Kaneko T, Asamizu E, Kato T, Nakao M, Sasamoto S, Watanabe A, Ono A,
Kawashima K, Fujishiro T, Katoh M, Kohara M, Kishida Y, Minami C, Nakayama S,
Nakazaki N, Shimizu Y, Shinpo S, Takahashi C, Wada T, Yamada M, Ohmido N, Hayashi M,
Fukui K, Baba T, Nakamichi T, Mori H, Tabata S (2008) Genome structure of the legume,
Lotus japonicus. DNA Res 15:227–239
Schleueter J, Dixon P, Granger C, Grant D, Clark L, Doyle JJ, Shoemaker RC (2004) Mining
EST databases to resolve evolutionary events in major crop species. Genome 47:868–876
Schmutz J, Cannon SB, Schlueter J, Ma J, Mitros T, Nelson W, Hyten DL, Song Q, Thelen JJ,
Cheng J, Xu D, Hellsten U, May GD, Yu Y, Sakurai T, Umezawa T, Bhattacharyya MK,
Sandhu D, Valliyodan B, Lindquist E, Peto M, Grant D, Shu S, Goodstein D, Barry K, FutrellGriggs M, Du J, Tian Z, Zhu L, Gill N, Joshi T, Libault M, Sethuraman A, Zhang XC,
Shinozaki K, Nguyen HT, Wing RA, Cregan P, Specht J, Grimwood J, Rokhsar D, Stacey G,
Shoemaker RC, Jackson SA (2010) Genome sequence of the paleopolyploid soybean. Nature
463:178–183
Schrire BD, Lavin M, Barker NP, Forest F (2009) Phylogeny of the Tribe Indigofereae
(Leguminosae-Papilionoideae): geographically structured more in succulent-rich and
temperate settings than in grass-rich environments. Am J Bot 96:816–843, 844–852
Seijo G, Lavia GI, Fernandez A, Krapovickas A, Ducasse DA, Bertioli DJ, Moscone EA (2007)
Genomic relationships between the cultivated peanut (Arachis hypogaea, Leguminosae) and
its close relatives revealed by double GISH. Am J Bot 94:1963–1971
Seijo G, Fernandez A (2001) Chromosome numbers of some southernmost species of Mimosa L.
(Leguminosae). Cytologia 66:19–23
Sen O, Bhattacharya S (1988) Cytomixis in vigna-glabrescens Ttk-1 wild. Cytologia 53:437–440
Shoemaker RC, Polzin K, Labate J, Specht J, Brummer EC, Olson T, Young N, Concibido V,
Wilcox J, Tamulonis JP, Kochert G, Boerma HR (1996) Genome duplication in soybean
(Glycine subgenus soja). Genetics 144:329–338
Shoemaker RC, Schlueter J, Doyle JJ (2006) Paleopolyploidy and gene duplication in soybean
and other legumes. Curr Opin Plant Biol 9:104–109
Simon MF, Grether R, de Queiroz LP, Saerkinen TE, Dutra VF, Hughes CE (2011) The
evolutionary history of Mimosa (Leguminosae): toward a phylogeny of the sensitive plants.
Am J Bot 98:1201–1221
Singer SR, Maki SL, Farmer AD, Ilut D, May GD, Cannon SB, Doyle JJ (2009) Venturing
beyond beans and peas: what can we learn from chamaecrista? Plant Physiol 151:1041–1047
Sinou C, Forest F, Lewis GP, Bruneau A (2009) The genus Bauhinia s.l. (Leguminosae): a
phylogeny based on the plastid trnL-trnF region. Botany-Botanique 87:947–960
Soltis DE, Buggs RJA, Doyle JJ, Soltis PS (2010) What we still don’t know about polyploidy.
Taxon 59:1387–1403
Soltis DE, Albert VA, Leebens-Mack J, Bell CD, Paterson AH, Zheng C, Sankoff D, dePamphilis
CW, Wall PK, Soltis PS (2009) Polyploidy and angiosperm diversification. Am J Bot
96:336–348
CO
RR
1215
1216
1217
1218
1219
1220
1221
1222
1223
1224
1225
1226
1227
1228
1229
1230
1231
1232
1233
1234
1235
1236
1237
1238
1239
1240
1241
1242
1243
1244
1245
1246
1247
1248
1249
1250
1251
1252
1253
1254
1255
1256
1257
1258
1259
1260
1261
1262
1263
1264
1265
1266
1267
UN
Editor Proof
9 Polyploidy in Legumes
Layout: T1 Standard SC
Chapter No.: 9
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 180/179
EC
TE
D
PR
OO
F
Spellenberg R (1981) Poly ploidy in Dalea-Formosa Fabaceae on the Chihuahuan desert.
Brittonia 33:309–324
Sprent JI (2009) Legume nodulation: a global perspective. Wiley-Blackwell, Ames
Srivastav PK, Raina SN (1986) Cytogenetics of Tephrosia Vi. Meiotic systems in some taxa.
Cytologia 51:359–374
Steele KP, Ickert-Bond SM, Zarre S, Wojciechowski MF (2010) Phylogeny and character
evolution in Medicago (Leguminosae): evidence from analyses of plastid Trnk/matk and
nuclear Ga3ox1 sequences. Am J Bot 97:1142–1155
Stefanovic S, Pfeil BE, Palmer JD, Doyle JJ (2009) Relationships among phaseoloid legumes
based on sequences from eight chloroplast regions. Syst Bot 34:115–128
Straub SCK, Doyle JJ (2009) Conservation genetics of Amorpha georgiana (Fabaceae), an
endangered legume of the Southeastern United States. Mol Ecol 18:4349–4365
Straub SCK, Pfeil BE, Doyle JJ (2006) Testing the polyploid past of soybean using a low-copy
nuclear gene––is Glycine (Fabaceae: Papilionoideae) an auto- or allopolyploid? Mol
Phylogenet Evol 39:580–584
te Beest M, Le Roux JJ, Richardson DM, Brysting AK, Suda J, Kubešová M, Pyšek P (2011) The
more the better? The role of polyploidy in facilitating plant invasions. Ann Bot 109:19–45
Thulin M, Lavin M (2001) Phylogeny and biogeography of the Ormocarpum group (Fabaceae): a
new genus Zygocarpum from the horn of Africa region. Syst Bot 26:299–317
Tondini F, Tavoletti S, Mariani A, Veronesi F (1993) A statistical approach to estimate the
frequency of n, 2n and 4n pollen grains in diploid alfalfa. Euphytica 69:109–114
Torres DC, Matos Santos Lima JP, Fernandes AG, Nunes EP, Grangeiro TB (2011) Phylogenetic
relationships within chamaecrista sect. Xerocalyx (Leguminosae, Caesalpinioideae) inferred
from the cpDNA trnE- trnT intergenic spacer and nrDNA ITS sequences. Genet Mol Biol
34:244–251
Travnicek P, Eliasova A, Suda J (2010) The distribution of cytotypes of Vicia cracca in Central
Europe: the changes that have occurred over the last four decades. Preslia (Prague)
82:149–163
Turini FG, Braeuchler C, Heubl G (2010) Phylogenetic relationships and evolution of
morphological characters in Ononis L. (Fabaceae). Taxon 59:1077–1090
van Wyk BE, Schutte AL (1988) Chromosome numbers in Lotononis and Buchenroedera
(Fabaceae-Crotalarieae). Ann Missouri Bot Gard 75:1603–1607
Varshney RK, Chen W, Li Y, Bharti AK, Saxena RK, Schlueter JA et al (2011) Draft genome
sequence of Pigeonpea (Cajanus cajan), an orphan legume crop of resource-poor farmers. Nat
Biotech. doi:10.1038/nbt.2022
Veronesi F, Mariani A, Bingham ET (1986) Unreduced gametes in diploid medicago and their
importance in alfalfa breeding. Theor Appl Genet 72:37–41
Wagstaff S, Heenan P, Sanderson M (1999) Classification, origins, and patterns of diversification
in New Zealand Carmichaelinae (Fabaceae). Am J Bot 86:1346–1356
Wang H, Moore MJ, Soltis PS, Bell CD, Brockington SF, Alexandre R, Davis CC, Latvis M,
Manchester SR, Soltis DE (2009) Rosid radiation and the rapid rise of angiosperm-dominated
forests. Proc Nat Acad Sci 106:3853–3858
Wilbur RL (1975) A revision of the North American genus Amorpha Leguminosae Psoraleae.
Rhodora 77:337–409
Wojciechowski MF, Lavin M, Sanderson MJ (2004) A phylogeny of legumes (Legumenosae)
based on analyses of the plastid matK gene resolves many well-supported subclades within
the family. Am J Bot 91:1846–1862
Wojciechowski M (2005) Astragalus (Fabaceae): a molecular phylogenetic perspective.
Brittonia 57:382–396
Young N, Debellé F, Oldroyd G, Geurts R, Cannon SB et al (2011) The medicago genome
provides insight into the evolution of rhizobial symbioses. Nature. doi:10.1038/nature10625
Zhang M, Fritsch PW, Cruz BC (2009) Phylogeny of Caragana (Fabaceae) based on DNA
sequence data from rbcL, trnS-trnG, and ITS. Mol Phylogenet Evol 50:547–559
CO
RR
1268
1269
1270
1271
1272
1273
1274
1275
1276
1277
1278
1279
1280
1281
1282
1283
1284
1285
1286
1287
1288
1289
1290
1291
1292
1293
1294
1295
1296
1297
1298
1299
1300
1301
1302
1303
1304
1305
1306
1307
1308
1309
1310
1311
1312
1313
1314
1315
1316
1317
1318
1319
1320
J. J. Doyle
UN
Editor Proof
180
Metadata of the chapter that will be visualized in
SpringerLink
Book Title
Polyploidy and Genome Evolution
Series Title
Chapter Title
Jeans, Genes, and Genomes: Cotton as a Model for Studying Polyploidy
Copyright Year
2012
Copyright HolderName
Springer-Verlag Berlin Heidelberg
Corresponding Author
Family Name
Wendel
Particle
Given Name
Jonathan F.
Suffix
Author
Division
Department of Ecology, Evolution, and Organismal Biology
Organization
Iowa State University
Address
50011, Ames, IA, USA
Email
jfw@iastate.edu
Family Name
Flagel
Particle
Given Name
Lex E.
Suffix
Author
Division
Department of Biology
Organization
Duke University
Address
90338, 27708, Durham, NC, USA
Email
lex.flagel@duke.edu
Family Name
Adams
Particle
Given Name
Keith L.
Suffix
Abstract
Division
Department of Botany, and UBC Botanical Garden and Centre for Plant
Research
Organization
University of British Columbia
Address
6268 University Blvd, V6T 1Z4, Vancouver, BC, Canada
Email
keitha@mail.ubc.ca
We present an overview of the cotton genus (Gossypium) as a model for the study of polyploidy. A synopsis
of the origin and evolution of polyploid cotton is provided, offering an organismal framework and
phylogenetic perspective that is critical for understanding modes and mechanisms of gene and genome
evolution. Sequence data from thousands of genes implicate a mid-Pleistocene (1–2 mya) origin of polyploid
cotton, following trans-oceanic dispersal of an Old World, A-genome diploid to the New World and
subsequent hybridization with an indigenous D-genome diploid. This chance biological reunion, occurring
after 5–10 million years of diploid evolution in isolation, has led to an array of molecular genetic interactions
in the newly formed allopolyploid lineage, including nonreciprocal homoeologous recombination and perhaps
other forms of interlocus concerted evolution, differential rates of genomic evolution, intergenomic spread
of transposable elements, and myriad forms of alterations in duplicate expression relative to that experienced
in the ancestral diploids. The latter include developmental, organ-, tissue-, and cell-specific biases in
homoeologous gene expression, which can be sensitive to various forms of environmental perturbation and
stress. The allopolyploid Gossypium transcriptome is exceptionally dynamic, with homoeolog expression
ratios being subject to change even during development of the single-celled cotton fiber. Expression evolution
is temporally partitioned into changes accompanying genome merger (hybridization) at the diploid level,
polyploidization, and longer term evolution at the allopolyploid level. Evidence indicates that allopolyploidy
facilitated colonization of a new ecological niche for the genus and led to an enhanced capacity for developing
agronomically superior cotton varieties. The myriad mechanisms that underlie genomic and regulatory
evolution are suggested to have contributed to both ecological success and agronomic potential.
Book ISBN: 978-3-642-31441-4
Page: 181/206
Chapter 10
4
Jonathan F. Wendel, Lex E. Flagel and Keith L. Adams
9
10
11
12
13
14
15
16
17
18
19
20
21
22
D
8
TE
7
Abstract We present an overview of the cotton genus (Gossypium) as a model for
the study of polyploidy. A synopsis of the origin and evolution of polyploid cotton
is provided, offering an organismal framework and phylogenetic perspective that is
critical for understanding modes and mechanisms of gene and genome evolution.
Sequence data from thousands of genes implicate a mid-Pleistocene (1–2 mya)
origin of polyploid cotton, following trans-oceanic dispersal of an Old World,
A-genome diploid to the New World and subsequent hybridization with an indigenous D-genome diploid. This chance biological reunion, occurring after 5–10
million years of diploid evolution in isolation, has led to an array of molecular
genetic interactions in the newly formed allopolyploid lineage, including nonreciprocal homoeologous recombination and perhaps other forms of interlocus
concerted evolution, differential rates of genomic evolution, intergenomic spread
of transposable elements, and myriad forms of alterations in duplicate expression
relative to that experienced in the ancestral diploids. The latter include developmental, organ-, tissue-, and cell-specific biases in homoeologous gene expression,
which can be sensitive to various forms of environmental perturbation and stress.
The allopolyploid Gossypium transcriptome is exceptionally dynamic, with
homoeolog expression ratios being subject to change even during development of the
EC
6
CO
RR
5
PR
OO
3
Jeans, Genes, and Genomes: Cotton
as a Model for Studying Polyploidy
2
F
1
Book ID: 272454_1_En
Date: 16-8-2012
J. F. Wendel (&)
Department of Ecology, Evolution, and Organismal Biology,
Iowa State University, Ames, IA 50011, USA
e-mail: jfw@iastate.edu
L. E. Flagel
Department of Biology, Duke University, 90338 Durham, NC 27708, USA
e-mail: lex.flagel@duke.edu
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: 10
K. L. Adams
Department of Botany, and UBC Botanical Garden and Centre for Plant Research,
University of British Columbia, 6268 University Blvd, Vancouver, BC V6T 1Z4, Canada
e-mail: keitha@mail.ubc.ca
P. S. Soltis and D. E. Soltis (eds.), Polyploidy and Genome Evolution,
DOI: 10.1007/978-3-642-31442-1_10, Springer-Verlag Berlin Heidelberg 2012
181
Layout: T1 Standard SC
Chapter No.: 10
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 182/206
25
26
27
28
29
30
single-celled cotton fiber. Expression evolution is temporally partitioned into
changes accompanying genome merger (hybridization) at the diploid level, polyploidization, and longer term evolution at the allopolyploid level. Evidence indicates
that allopolyploidy facilitated colonization of a new ecological niche for the genus
and led to an enhanced capacity for developing agronomically superior cotton
varieties. The myriad mechanisms that underlie genomic and regulatory evolution
are suggested to have contributed to both ecological success and agronomic
potential.
F
24
PR
OO
23
J. F. Wendel et al.
31
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
61
62
63
D
36
TE
35
Because of its economic importance, the cotton genus (Gossypium L.) has long
attracted the attention of agricultural scientists, taxonomists, and biologists in
multiple disciplines. Accordingly, a great deal is understood about the origin and
diversification of the genus, its basic plant biology, and its properties as a crop
plant (Paterson 2009; Stewart et al. 2010; Wendel et al. 2009). One of the most
salient features of the genus is that its history is so encompassing in scope,
involving a global (mostly austral) phylogenetic diversification and a repeated
history of trans-oceanic dispersal. Superimposed on this natural diversification has
been a many-thousand-year history of human manipulation, tracing to ancient
human cultures on several continents who independently domesticated four species, two from the Americas, G. hirsutum and G. barbadense, and two from
Africa-Asia, G. arboreum and G. herbaceum. In each of these four cases,
aboriginal peoples discovered that the unique properties of cotton ‘‘fibers’’, which
are unicellular epidermal seed trichomes, made them useful for ropes, textiles, and
other applications. Each of these crop species has its own history of domestication,
diversification, and current utilization (Brubaker et al. 1999a; Hutchinson 1951,
1954; Hutchinson et al. 1947; Percy and Wendel 1990; Wendel et al. 2009).
This rich history of scientific study has made Gossypium one of the best systems
for studies of polyploidy in plants. As testified by the many papers in this volume,
plant evolution has been characterized by repeated rounds of whole-genome doubling. Evolutionary footprints of paleopolyploidy in ‘‘diploid’’ (n = 13) cotton
have long been evidenced using both classic and more modern techniques,
including chromosome banding, comparative genome mapping, and analysis of
synonymous substitution rate curves (Muravenko et al. 1998; Brubaker et al. 1999b;
Reinisch et al. 1994; Paterson 2009; Lin and Paterson 2009; Lin et al. 2011). These
more ancient events, which must have profoundly impacted morphological, ecological, and physiological diversification, predate a more recent allopolyploidy
event that traces to the mid-Pleistocene (Wendel 1989; Wendel et al. 2009; Wendel
and Cronn 2003). The most experimentally tractable polyploid event is this most
recent one, resulting from the merger that led to the evolution of the five extant
species of New World allopolyploid cotton (2n = 4x = 52), including the two
EC
33
34
10.1 Introduction
CO
RR
32
UN
Editor Proof
182
Layout: T1 Standard SC
Chapter No.: 10
Book ISBN: 978-3-642-31441-4
Page: 183/206
Jeans, Genes, and Genomes: Cotton as a Model for Studying Polyploidy
183
88
10.1.1 Origin and Diversification of the Diploid Cottons
72
73
74
75
76
77
78
79
80
81
82
83
84
85
86
89
90
91
92
93
94
95
96
97
98
99
100
101
102
103
104
PR
OO
71
D
70
TE
68
69
EC
67
The cotton genus belongs to a small tribe, the Gossypieae, that includes only eight
genera and *120 species (Fryxell 1968, 1979). Four of these genera are either
monotypic or contain only several species with restricted geographic distributions,
Lebronnecia (Marquesas Islands), Cephalohibiscus (New Guinea, Solomon Islands),
Gossypioides (east Africa, Madagascar), and Kokia (Hawaii). The tribe also includes
four moderately sized genera with broader ranges: Hampea, with 21 neotropical
species; Cienfuegosia, with 25 species from the neotropics and parts of Africa;
Thespesia, with 17 tropical species; and last but not least, Gossypium, the largest and
most widely distributed genus in the tribe with more than 50 species (Fryxell 1992).
Gossypium species collectively have achieved a nearly worldwide distribution,
with several primary centers of diversity in the arid or seasonally arid tropics and
subtropics. Species-rich regions include Australia, especially the Kimberley region
in NW Australia, the Horn of Africa and southern Arabian Peninsula, and the
western part of central and southern Mexico. Recognition of these groups of
species reflects decades of accumulated understanding that emerged from basic
plant exploration and taxonomic analysis (Fryxell 1979, 1992; Hutchinson et al.
CO
RR
66
F
87
cultivated species G. hirsutum (Upland cotton) and G. barbadense (Pima cotton,
Sea Island cotton). Gossypium hirsutum presently is responsible for over 90 % of
the cotton crop internationally, having spread from its original home in Mesoamerica to over 50 countries in both eastern and western hemispheres. Accumulating evidence indicates that polyploidy per se has played a critical role in enabling
the development of modern, agronomically elite varieties. Thus, it is of interest to
explore what cotton has taught us about the evolutionary consequences of genome
doubling, and in turn, what the process of genome doubling might reveal about
phenotypic diversification and the goals of agronomic improvement.
We begin by reviewing our current understanding of the diversity of the genus
and the origin of the Gossypium clade, which includes one of the classic examples
of polyploidy. This phylogenetic and temporal perspective provides the organismal
framework that serves as the foundation for all analyses of the consequences of
polyploid evolution, including those focused on genomic, epigenomic, and phenotypic levels. We then provide a synopsis of the myriad genomic consequences
that were set in motion by the evolutionary processes of genome merger, chromosome doubling, and subsequent evolutionary diversification which collectively
gave rise to modern allopolyploid cottons, drawing attention to the relationships
among evolutionary processes and temporal scale of divergence. Thus, we distinguish phenomena and processes that might characterize the earliest stages of
polyploid formation from those that are responsible for longer term genomic and
phenotypic changes. Finally, we summarize evidence that polyploidy enables the
evolution of transgressive or novel phenotypes in cotton, as exemplified by the
differences between modern cultivated diploid versus allopolyploid cotton.
64
65
UN
Editor Proof
10
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 10
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 184/206
113
114
115
116
117
118
119
120
121
122
123
124
125
126
127
128
129
130
131
132
133
134
135
136
137
138
139
140
141
142
143
144
145
146
147
148
149
F
111
112
PR
OO
109
110
D
108
TE
107
1947; Saunders 1961; Watt 1907; Wendel et al. 2009). The genus is extraordinarily
diverse; species morphologies range from fire-adapted, herbaceous perennials in
NW Australia to trees in SW Mexico that escape the dry season by dropping their
leaves. Corolla colors span a rainbow of blue to purple (G. triphyllum), mauves
and pinks (‘‘Sturt’s Desert Rose’’, G. sturtianum, is the official floral emblem of
the Northern Territory, Australia), whites and pale yellows (NW Australia,
Mexico, Africa-Arabia), and even a deep sulfur-yellow (G. tomentosum from
Hawaii). Seed coverings range from nearly glabrous (e.g., G. klotzschianum and
G. davidsonii), to short stiff, dense, brown hairs that aid in wind-dispersal
(G. australe, G. nelsonii), to the long, fine white fibers that characterize highly
improved forms of the four cultivated species (Fig. 10.1). There are even seeds
that produce fat bodies to facilitate ant-dispersal (Seelanan et al. 1999). At the
other end of the ant coevolution spectrum is G. tomentosum from the Hawaiian
Islands, which lost the foliar and extra-floral nectaries that are common in other
Gossypium species, presumably in response to the absence of native ants.
The evolution of this morphological and geographic diversity was accompanied
by a parallel diversification at the chromosomal level (Lin and Paterson 2009).
Although all diploid species share the same chromosome number (n = 13), there
is more than 3-fold variation in DNA content per genome (Hendrix and Stewart
2005). Chromosome morphology is similar among closely related species, as
reflected in the ability of related species to form hybrids that display normal
meiotic pairing and sometimes high F1 fertility. In contrast, crosses among more
distant relatives may be difficult to achieve, and those that are successful are
characterized by meiotic abnormalities. The collective observations of pairing
behavior, chromosome size, and relative fertility in interspecific hybrids led to the
designation of single-letter genome symbols (Beasley 1941) for related clusters of
species. Presently, eight diploid genome groups (A through G, plus K) are recognized (Endrizzi et al. 1985; Stewart 1995).
A genealogical framework for the genus is provided from analyses of multiple
molecular phylogenetic investigations (reviewed in Wendel and Cronn 2003).
A key phylogenetic conclusion has been the demonstration that the group of species
recognized as belonging to Gossypium do, in fact, constitute a single natural
lineage, despite their exceptionally broad geographic distribution and extraordinary
morphological and cytogenetic diversity. A second important result is that the
closest relative of Gossypium is the sister clade that includes the AfricanMadagascan genus Gossypioides and the Hawaiian endemic genus Kokia; these
latter genera may thus be used as phylogenetic outgroups for studying evolutionary
patterns and processes within Gossypium. A third phylogenetic conclusion is that
each of the classically recognized genome groups comprises a monophyletic group.
This information is summarized in a depiction of our present understanding of
relationships (Fig. 10.2), which shows four major lineages of diploid species
corresponding to three continents: Australia (C-, G-, K-genomes), the Americas
(D-genome), and Africa/Arabia (two lineages: one comprising the A-, B-, and
F-genomes, and a second containing the E-genome species). Embedded in this
result is the observation that the earliest divergence in the genus separated the New
EC
106
CO
RR
105
J. F. Wendel et al.
UN
Editor Proof
184
Layout: T1 Standard SC
Chapter No.: 10
Book ISBN: 978-3-642-31441-4
Page: 185/206
Jeans, Genes, and Genomes: Cotton as a Model for Studying Polyploidy
185
TE
D
PR
OO
F
Editor Proof
10
Book ID: 272454_1_En
Date: 16-8-2012
151
152
153
154
155
156
157
158
159
World D-genome lineage from the ancestor of all Old World taxa, and thus, that
New World and Old World diploids are phylogenetic sister groups. Following this
split in the genus, cottons comprising the Old World lineage divided into three
groups, namely, the Australian cottons (C-, G-, and K-genome species), the
African-Arabian E-genome species, and the African A-, B-, and F-genome cottons.
The African F-genome clade, which consists of the sole species G. longicalyx, is
diagnosed as sister to the A-genome species, an important realization in that this
relationship identifies the wild forms most closely related to the clade (the
A-genome) that ‘‘invented’’ long, or spinable, fiber since these two clades (A- and
F-) shared a common ancestor.
UN
150
CO
RR
EC
Fig. 10.1 Representative seed and trichome diversity in Gossypium. Seed and trichome size and
morphology are exceedingly variable in the genus. Most wild species have relatively small seeds
(\5 mm in any dimension) with equally short fibers. Long (spinnable) fiber evolved only once, in
the ancestor of modern A-genome cottons, which subsequently donated this capacity to modern
tetraploid species, including the commercially important G. hirsutum and G. barbadense, at the
time of allopolyploid formation. Key to species: Cult. AD1 = G. hirsutum TM1; Wild AD1 = G.
hirsutum Tx2094 from the Yucatan Peninsula; AD3 = G. tomentosum WT936 from Hawaii;
C1 = G. sturtianum C1-4 from Australia; Cult. A2 = G. arboreum AKA8401; Wild A1 = G.
herbaceum subsp. africanum from Botswana; D5 = G. raimondii from Peru; D3 = G. davidsonii
D3d-32 from Baja California; F1 = G. longicalyx F1-3 from Tanzania; B1 = G. anomalum B1-1
from Africa
Layout: T1 Standard SC
Chapter No.: 10
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 186/206
J. F. Wendel et al.
TE
D
PR
OO
F
Editor Proof
186
161
162
163
164
165
166
167
168
169
170
A temporal framework for the origin of Gossypium and its diversification is
provided by sequence divergence data, which may serve as a proxy for time
(Senchina et al. 2003). These analyses indicate that Gossypium diverged from its
closest relatives during the Miocene, perhaps 10–15 mya, subsequently spreading
around the world via trans-oceanic dispersal to acquire its modern geographic
range. These early estimates of divergence, and hence times, have recently been
supported by an enormous amount of new sequence data derived from a global
assembly of ESTs (Flagel et al. 2012). Consideration of the phylogeny of Fig. 10.2
in a temporal context and in light of plate tectonic history leads to the inference
that the history of Gossypium has entailed multiple episodes of trans-oceanic
dispersal. These include at least one dispersal between Australia and Africa,
UN
160
CO
RR
EC
Fig. 10.2 Evolutionary history of Gossypium, as inferred from multiple molecular phylogenetic
data sets. The closest relative of Gossypium is a lineage containing the African-Madagascan
genus Gossypioides and the Hawaiian endemic genus Kokia. Following its likely origin 5–10
mya, Gossypium split into three major diploid lineages: the New World clade (D-genome); the
African-Asian clade (A-, B-, E-, and F-genomes); and the Australian clade (C-, G-, and
K-genomes). This global radiation involved several trans-oceanic dispersal events and was
accompanied by morphological, ecological, and chromosomal differentiation (2C genome sizes
shown in white ellipses). Allopolyploid cottons formed following trans-oceanic dispersal of an
A-genome diploid to the Americas, where the immigrant underwent hybridization, as female,
with a native D-genome diploid similar to modern G. raimondii. Polyploid cotton probably
originated during the Pleistocene (1–2 mya), with the five modern species comprising the
descendants of an early and rapid colonization of the New World tropics and subtropics
Layout: T1 Standard SC
Chapter No.: 10
Book ISBN: 978-3-642-31441-4
Page: 187/206
Jeans, Genes, and Genomes: Cotton as a Model for Studying Polyploidy
187
174
10.1.2 Origin and Diversification of the Polyploid Cottons
179
180
181
182
183
184
185
186
187
188
189
190
191
192
193
194
195
196
197
198
199
200
201
202
203
204
205
206
207
208
209
210
211
PR
OO
178
D
177
A rich body of cytogenetic and experimental evidence has demonstrated that
the tetraploid species, which are entirely New World in their distribution, are
allopolyploids containing two genomes, an A-genome from Africa or Asia, and a
D-genome similar to those found in the American diploids (Endrizzi et al. 1985;
Wendel et al. 2009; Wendel and Cronn 2003). The hemisphere-scale allopatry of
these two diploid genome groups led to more than 50 years of mystery surrounding
the timing of formation and parentage of the New World allopolyploids. With
respect to the question of ‘‘when’’, gene sequence data convincingly demonstrate
that allopolyploid Gossypium originated prior to the evolution of modern humans
but relatively recently in geological terms, perhaps 1–2 mya, or in the midPleistocene (Senchina et al. 2003; Wendel 1989; Flagel et al. 2012). With respect to
the second part of the question, that of polyploid parentage, it is now clear that both
extant A-genome species (G. arboreum, G. herbaceum) are equally divergent from
the A-genome of allopolyploid cottons and that the closest living relative of the
progenitor D-genome donor is G. raimondii (Endrizzi et al. 1985; Wendel et al.
2009; Wendel and Cronn 2003). Studies using nuclear (biparentally inherited)
genes led to the same conclusion. Additionally, all allopolyploids contain an
A-genome cytoplasm, as evidenced from analysis of both mitochondrial and plastid
genomes (Galau and Wilkins 1989; Small and Wendel 1999; Wendel 1989).
Finally, the studies mentioned above, and additional, extensive DNA sequence data
from ongoing studies (Grover et al. unpublished), support a single origin for
allopolyploid cotton.
Given a Pleistocene origin for allopolyploid cotton species, one may infer that
their morphological diversification and spread must have been relatively rapid
following polyploidization. At present, five allopolyploid species are widely recognized, although a sixth species (G. ekmanianum) was recently proposed
(Krapovickas and Seijo 2008). Gossypium darwinii is native to the Galapagos
Islands, where it may form large populations in some areas (Percy and Wendel
1990). Gossypium tomentosum, from the Hawaiian Islands, has a more diffuse
population structure, occurring mostly as scattered individuals and small populations on several islands (DeJoode and Wendel 1992). A third allopolyploid,
G. mustelinum, is restricted to a small region of northeast Brazil (Wendel et al.
1994). In addition to these three truly wild species, there are two cultivated species
(G. barbadense and G. hirsutum), each of which has a large indigenous range,
collectively encompassing a wealth of morphological forms that span the wildto-domesticated continuum (Brubaker and Wendel 1993, 1994, 2001; Fryxell 1979;
Hutchinson 1951; Percy and Wendel 1990). Gossypium hirsutum is widely
TE
176
EC
175
CO
RR
172
F
173
another to the Americas leading to the evolution of the D-genome diploids, and a
second, much later colonization of the New World by the A-genome ancestor of
the AD-genome allopolyploids (see below).
171
UN
Editor Proof
10
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 10
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 188/206
J. F. Wendel et al.
232
10.2 Evolution Following Genome Duplication
233
10.2.1 Chromosomal Stabilization and Structural Stasis
220
221
222
223
224
225
226
227
228
229
230
234
235
236
237
238
239
240
241
242
243
244
245
246
247
248
249
250
251
PR
OO
219
D
218
TE
216
217
EC
214
215
Classical cytogenetic evidence indicates that chromosomes of the A- and D-genomes
of allopolyploid Gossypium are less able to pair with one another than are chromosomes of the living descendants of their diploid progenitors (reviewed in Endrizzi
et al. 1985). For example, allopolyploid-derived haploids form an average of less
than one bivalent per cell at meiotic metaphase, whereas chromosomes in hybrids
between extant A- and D- genome diploids average 5.8 and 7.8 bivalents (reviewed in
Endrizzi et al. 1985). These and similar observations indicate that natural selection
has favored the evolution of mechanisms that promote exclusive bivalent formation
in the allopolyploid. Neither the pace at which such mechanisms operate nor their
nature are understood, but it seems rational to postulate that selection would be most
intense in the first generations following allopolyploid formation, where the fitness
cost of unbalanced gametes would be the greatest.
One hypothesis for this apparent lack of bivalent formation is that genome
stabilization following polyploidization involved genomic reorganization of the
two resident genomes such that they no longer are capable of homoeologous
pairing. To evaluate the extent of structural change, genetic maps were generated
and compared among interspecific F2 progenies for diploid (both A- and D-) and
allopolyploid cottons (Brubaker et al. 1999b; Rong et al. 2004). Comparisons of
CO
RR
213
F
231
distributed in Central and northern South America, the Caribbean, and even reaches
distant islands in the Pacific (eg., Solomon Islands, Marquesas). Gossypium
hirsutum is thought to have a more northerly distribution than G. barbadense, with
wild populations occurring as far north as Tampa Bay, Florida (27̊38’N) (Stewart,
personal observation). Gossypium barbadense has a more southerly indigenous
range, centered in the northern third of South America but with a large region of
range overlap with G. hirsutum in the Caribbean.
Consideration of the distribution of the allopolyploid species suggests that
polyploidy led to the invasion of a new ecological niche. Fryxell (1965, 1979)
noted that in contrast to the majority of diploid species, allopolyploid species
typically occur in coastal habitats, at least those forms that arguably are truly wild.
Two species, both island endemics (G. darwinii and G. tomentosum), are restricted
to near coastlines, and for two others (G. barbadense and G. hirsutum), wild forms
occur in littoral habitats ringing the Gulf of Mexico, northwest South America, and
distant Pacific Islands. Fryxell speculated that following initial formation, adaptation of the newly evolved allopolyploid to littoral habitats enabled it to exploit
the fluctuating sea levels that characterized the Pleistocene. This ecological
innovation is envisioned to have facilitated initial establishment of the new
polyploid lineage and also may have provided a means for the rapid dispersal of
the salt-water-tolerant seeds.
212
UN
Editor Proof
188
Layout: T1 Standard SC
Chapter No.: 10
258
259
260
261
262
263
264
265
266
267
268
269
270
271
272
273
274
275
276
277
278
279
280
281
282
283
284
285
286
287
288
289
290
291
292
F
257
PR
OO
256
D
255
10.2.2 Genome Sizes, Transposable Element Mobilization,
and Genomic Downsizing
TE
254
189
gene order and synteny among the A- and D-genome maps, as well as those
for both genomes of the allopolyploid (A vs. AT and D vs. DT), demonstrate
that relatively few structural rearrangements have arisen in the 1–2 my since
allopolyploid formation; conservation of collinear linkage groups is the rule rather
than the exception. Thus, allopolyploidy in Gossypium has not been accompanied
by extensive chromosomal rearrangement. This implies that structural rearrangement has not been a significant aspect of the process of polyploid genome
stabilization in cotton. Additional support for this idea emerges from experiments
involving synthetic allopolyploids; for example, Liu et al. (2001) used AFLP
analysis to demonstrate almost exclusive fragment additivity for 22,000 genomic
loci in nine sets of newly synthesized allotetraploid and allohexaploid Gossypium.
Thus, and in contrast to some other plant models described in this volume (e.g.,
wheat, Tragopogon), the polyploid Gossypium genome appears to be relatively
quiescent, at least with respect to the phenomenon of rapid genome change.
A corollary is that the evolutionary enforcement of homologous pairing discussed
above originated through means other than structural rearrangements.
As shown in Fig. 10.2, genome sizes vary widely among diploid cotton species,
from *900 Mb in the D-genome diploids to *2,600 Mb in the Australian diploids
(Hendrix and Stewart 2005), reflecting primarily the differential and punctuated
proliferation of various families of copia and gypsy transposable elements (TEs), as
well as lineage-specific differences in the rate of deletions (Hawkins et al. 2008,
2006, 2009). The two progenitor genomes of allopolyploid cotton differ 2-fold in
size, and moreover, they differ in their complement of resident TEs. Thus, allopolyploidization entailed the merger of two different complements of TEs, creating
the potential for activation of TEs due to the generalized disruption of epigenetic
suppression of TE activity following the merger of two diverged regulatory systems, a process commonly referred to as ‘‘genomic shock’’. To evaluate the possibility that polyploidization in Gossypium was accompanied by a transpositional
burst, as in some other species (Kashkush et al. 2002; Shan et al. 2005; Ungerer
et al. 2006), Hu et al. (2010) used phylogenetic and quantitative methods to identify
changes in TE populations. These data showed that the major LTR retrotransposon
classes in the AD genome phylogenetically clustered with either their A- or Dgenome antecedent elements in a genome-specific fashion, with no evidence of an
impressive, recent, TE burst. Thus, hybridization and polyploidy do not appear to
have stimulated a massive TE proliferation in Gossypium.
Notwithstanding the relative TE quiescence indicated by these studies, evidence
using FISH (Hanson et al. 1999, 1998) implicates at least a modest level of TE
activity in allopolyploid cotton. These data show that a family of copia-like
retrotransposable elements ‘‘horizontally’’ transferred across genomes following
EC
253
CO
RR
252
Book ISBN: 978-3-642-31441-4
Page: 189/206
Jeans, Genes, and Genomes: Cotton as a Model for Studying Polyploidy
UN
Editor Proof
10
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 10
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 190/206
J. F. Wendel et al.
324
10.2.3 Genic Evolution in Diploid Versus Allopolyploid Cotton
299
300
301
302
303
304
305
306
307
308
309
310
311
312
313
314
315
316
317
318
319
320
321
322
325
326
327
328
329
330
331
332
333
PR
OO
298
D
297
TE
296
EC
295
CO
RR
294
F
323
allopolyploid formation. This result highlights the phenomenon of TE spread
across previously separated genomes following polyploid formation, raising the
possibility that this process has played a role in diversification and adaptation via
novel TE insertions.
An attractive feature of the Gossypium model system is that species representing both progenitor genomes remain extant, and that they vary so dramatically
(nearly 2-fold) in genome size yet retain collinearity with their orthologs in the
allopolyploid genome. Although the Gossypium genome has yet to be sequenced,
it is likely that a high-quality D-genome sequence (from G. raimondii) will be
published by the time this volume is published, with perhaps the A-genome
sequence not far behind. It is an exciting prospect to contemplate the availability
of both diploid sequences as well as one or more from allopolyploid Gossypium.
These data will offer a veritable gold mine for generating insight into the pace,
patterns, and dynamics of genome evolution that accompany diploid divergence,
allopolyploid formation, and subsequent evolution at the polyploid level.
A glimpse of this promise is provided by current work focused on comparative
sequencing of bacterial artificial chromosomes (BACs) from Gossypium in polyploid and diploid species. Two comparative BAC sequencing studies have been
published, a region surrounding the CesA1 gene (Grover et al. 2004) from both
homoeologous genomes of G. hirsutum, and a region surrounding the AdhA gene
(Grover et al. 2007) from G. hirsutum and diploids representing models of the two
progenitor genomes, i.e., G. arboreum and G. raimondii. Data generated to date
indicate that small deletions are more prevalent in the polyploid genomes (AT and
DT) than in either diploid genome, illustrative of the general phenomenon of
genomic downsizing in polyploid genomes (Bennett and Leitch 2005), while
providing a glimpse into an underlying mechanism (illegitimate recombination,
which is biased in the allopolyploid toward deletion). Extensions of this work into
20 regions and in more genomes, including a phylogenetic outgroup are underway
(Grover, Wendel, and Paterson, unpubl.), offering an opportunity to learn more
about features of genome evolution that distinguish polyploid cotton from its
diploid progenitors.
293
The most immediate and important genomic consequence of allopolyploid formation in Gossypium was simultaneous duplication of all nuclear genes. From a
phylogenetic perspective, the various fates of gene duplication may partially be
modeled as shown in Fig. 10.3. The null hypothesis for sequence evolution in
allopolyploids derives from the organismal history; if both duplicated genes evolve
independently following allopolyploid formation, then each homoeolog should be
phylogenetically sister to its ortholog from its progenitor diploid, rather than to the
other homoeolog. Similarly, if rates of sequence evolution are maintained between
the diploid and allopolyploid level, branch lengths for the two A-genome sequences
UN
Editor Proof
190
Layout: T1 Standard SC
Chapter No.: 10
Book ISBN: 978-3-642-31441-4
Page: 191/206
Recombination
TE
D
Null hypothesis
PR
OO
Gene loss
Unequal rates
191
F
Jeans, Genes, and Genomes: Cotton as a Model for Studying Polyploidy
Editor Proof
10
Book ID: 272454_1_En
Date: 16-8-2012
Novel expression
Intergenomic
transfer
334
335
336
337
UN
CO
RR
EC
Fig. 10.3 A model of various possibilities for duplicate gene evolution after allopolyploidy in
Gossypium. The null expectation (center) derives from the organismal history (left): if
homoeologs evolve independently following allopolyploid formation, then each should be
phylogenetically sister to its ortholog from the donor diploid, rather than to the other homoeolog.
Similarly, if rates of sequence evolution are similar at the diploid and allopolyploid level, branch
lengths for the two A-genome sequences (one from the diploid, ‘‘A’’, and the other from the
allopolyploid, ‘‘AT’’) should be similar, as they should for the two D-genome sequences (‘‘D’’
and ‘‘DT’’). The utility of this null hypothesis lies in its falsification; if homoeologous sequences
interact via concerted evolutionary forces or nonreciprocal homoeologous recombination, for
example, a different tree may be recovered (‘‘Recombination’’, top center), or if there is strong
directional selection or pseudogenization, rate inequalities may become evident (‘‘Unequal
rates’’, bottom center). Additional possibilities include loss of one of the homeologs (‘‘Gene
loss’’, top right), replicative transfer of sequences from one genome to the other (‘‘Intergenomic
transfer’’, bottom right), and evolutionary divergence in duplicate gene expression domains or
amounts (‘‘Novel expression’’, right middle). This latter category, novel expression, encompasses
multiple phenomena, including developmentally or environmentally regulated biases in
homoeolog expression ratios, organ- or tissue- or cell-specific homoeolog silencing, novel
expression domains, and transgressive (higher or lower than either progenitor diploid) expression
amounts
(one from the diploid, the other from the allopolyploid) should be similar, as they
should for the two D-genome sequences. The utility of the null hypothesis lies in its
falsification; if homoeologous sequences interact, for example, a different tree may
be recovered, or if there is strong directional selection or pseudogenization, rate
Layout: T1 Standard SC
Chapter No.: 10
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 192/206
J. F. Wendel et al.
342
10.2.3.1 Molecular Evolution of Homoeologs and Orthologs
347
348
349
350
351
352
353
354
355
356
357
358
359
360
361
362
363
364
365
366
367
368
369
370
371
372
373
374
375
376
377
378
PR
OO
346
D
345
Early work (Cronn et al. 1999; Senchina et al. 2003) demonstrated that duplicated
genes typically are both retained even in natural (1–2 mya -ld) allopolyploids, and
that they evolve essentially as modeled in the central panel of Fig. 10.3, i.e.,
equally, additively, and at equivalent rates. A more recent analysis (Flagel et al.
2012), based on a global assembly of 5 million Sanger and 454 ESTs supplemented by *150 million 82-bp Illumina reads, has provided a vastly expanded
and detailed view of genic evolution in diploid and allopolyploid cotton, perhaps
the most extensive yet for any plant genus. These data, representing analysis of
*10,000 genes in each comparison, show that rates of synonymous substitution
(Ks) between A- and D-genome orthologs (0.036) are nearly identical to that
experienced by their homoeologous descendants in the allopolyploid genome
(0.037), with non-synonymous substitution rates being identical in the two contrasts (0.009). These results demonstrate unequivocally that allopolyploidy has not,
in general, been accompanied by an enhanced rate of nucleotide substitution in
coding regions, as might be expected from an assumption of rapid decay of
‘‘redundant’’ duplicated copies.
An additional perspective provided by Flagel et al. (2012) is that the data also
provide an extraordinarily accurate depiction of the ancestry of polyploid cotton.
By using thousands of genic alignments, the relative distances of the A- and
D-genome diploids to their counterparts in the allopolyploid (AT, DT) can be
calculated and effectively translated into relative divergence times because of the
demonstration of rate homogeneity discussed above. The results show that the A
genome diploid has a mean Ks of 0.009 from the AT-genome, whereas the
comparable figure for the D genome comparison is 0.015. From this we infer that
modern A-genome diploids are a better model (by about 50 %) of the actual
genome donor of allopolyploid cotton than are extant D-genome diploids,
consistent with previous suggestions based on diverse data sources (Wendel et al.
2009; Wendel and Cronn 2003; Senchina et al. 2003). These new data provide
deep insight into the cotton model system, furthering its value for comparative
analyses.
A general expectation of molecular evolution in allopolyploids is that mutations
occur randomly among homoeologs, and hence that evolutionary rates will be
equivalent for homoeologs (Fig. 10.3). A corollary expectation is that duplicated
gene copies will accumulate diversity, within and among populations, at equivalent rates. Small et al. (1999) tested this hypothesis of rate equivalence among
homoeologs using the gene AdhA, for which both homoeologs were sequenced for
TE
343
344
EC
340
CO
RR
339
F
341
inequalities may become evident. Additional possibilities include silencing or loss
of one of the duplicated copies, ‘‘horizontal transfer’’ of sequences from one
genome to the other, and novel expression, the latter encompassing a variety of
phenomena discussed under Duplicate gene expression (Sect. 10.4).
338
UN
Editor Proof
192
Layout: T1 Standard SC
Chapter No.: 10
Book ISBN: 978-3-642-31441-4
Page: 193/206
Jeans, Genes, and Genomes: Cotton as a Model for Studying Polyploidy
193
407
10.2.3.2 Interaction Among Homoeologs at the DNA Level
387
388
389
390
391
392
393
394
395
396
397
398
399
400
401
402
403
404
405
408
409
410
411
412
413
414
415
416
417
418
419
PR
OO
386
D
385
TE
384
EC
382
383
CO
RR
381
F
406
22 accessions (44 alleles per genome) of G. hirsutum and for five accessions (10
alleles per genome) of G. barbadense. In both allopolyploids, estimates of
nucleotide diversity were higher for AdhA from the D-genome than from the Agenome. In a follow-up study (Small and Wendel 2002) using a second ADH gene
(AdhC), this conclusion was even more strongly supported; here 24 different
alleles were detected for the D-genome homoeolog versus only 7 for the
A-genome homoeolog, with a similar increase in allelic diversity for the
D-homoeolog in G. barbadense. These observations indicate that at least for some
genes, homoeologs may accumulate synonymous substitutions at vastly different
rates. At present, the responsible forces and underlying molecular mechanisms are
obscure, but a logical suggestion is that they are causally connected to the nearly 2fold difference in genome size between the co-resident genomes in the allopolyploid nucleus (the AT genome is approximately twice the size of the DT genome).
At present, little information exists for Gossypium that enables a thorough analysis of the relative rates of pseudogenization between diploids and allopolyploids, or
between the two homoeologous genomes of allopolyploid cotton. The ongoing
extensions of the comparative BAC sequencing approach taken by Grover and
colleagues (Grover et al. 2004, 2007) are promising in this respect, as are the ongoing
genome sequencing projects. Similarly, relatively little information exists on rates of
gene loss in cotton, though evidence to date based on comparative BAC sequencing
(Grover et al. 2004, 2007), comparative mapping analyses (Rong et al. 2004;
Brubaker et al. 1999b), EST collections (Flagel et al. 2012), and AFLP studies (Liu
et al. 2001) suggests that rates of gene loss are neither high nor particularly biased
with respect to genomic origin. A recent study using Southern hybridization,
however, detected three losses (of 27 genes studied) of D-homoeologs from
allopolyploid cotton and no losses of the A-homoeolog, suggesting a possible bias
(Rong et al. 2010). Again, key data likely will be generated soon that will permit these
speculations to be evaluated.
379
380
One of the first indications that duplicated genes could behave in an evolutionarily
dependent fashion was the study of Wendel et al. (1995), who demonstrated
interaction among the 18S–26S ribosomal genes that exist at multiple loci in the
A- and D-genomes. Instead of evolving independently, as expected if homoeologous repeats did not interact, repeats at the different arrays in allopolyploid cotton
have been ‘‘homogenized’’ to the same sequence (either ‘‘A-like’’ or ‘‘D-like’’) by
one or more processes of concerted evolution (reviewed by Elder and Turner
1995). In four of the five allopolyploid species, interlocus homogenization has
created exclusively D-genome like rDNAs, whereas in G. mustelinum nearly all
rDNA repeats have been homogenized to an A-like form. This example showed
that since polyploid formation 1–2 mya, some 3,800 repeats, each approximately
10 kb in length, were ‘‘overwritten’’ with the alternative form originating from the
UN
Editor Proof
10
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 10
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 194/206
427
428
429
430
431
432
433
434
435
436
437
438
439
440
441
442
443
444
445
446
447
448
449
450
451
452
453
454
455
456
457
458
459
460
461
462
463
F
426
PR
OO
425
D
423
424
TE
422
other parental genome, probably through unequal crossing over or gene conversion, and that this phenomenon operated bidirectionally, in different directions in
different allopolyploid lineages. Interlocus concerted evolution of rDNA repeats
has since been documented in many other plant polyploids, including Nicotiana
and Tragopogon (see this Chaps. 10 and 14, this volume). From a mechanistic
perspective, it seems likely that the homogenization has resulted from unequal
crossing over within and among arrays and nonreciprocal homoeologous recombination events, with homoeolog exchanges being facilitated by the sub-telomeric
location of rDNAs in many plant lineages, including Gossypium (Wendel 2000).
This demonstration that some repeated sequences could interact across genomes
in the allopolyploid nucleus led to additional investigations of the scope of the
phenomenon. In an analogous study, Cronn et al. (1996) showed that in contrast to
18S–26S arrays, 5S rDNA genes are not homogenized by concerted evolutionary
forces in the allopolyploid. Similarly, and as noted above, early studies demonstrated apparent independent evolution of homoeologs (Cronn et al. 1999;
Senchina et al. 2003), consistent with the null hypothesis of Fig. 10.3. Thus, until
recently it was thought that most nuclear genes duplicated by allopolyploidy largely evolve independently of one another in the polyploid nucleus, a proposition
that seemed entirely reasonable given the absence of evident cytogenetic interactions among homoeologs.
This presumption of homoeolog independence turns out to be incorrect; however, and for a surprisingly large percentage of duplicated genes. Using ESTs
generated from both A- and D-genome diploids and AD-genome allopolyploids,
Salmon et al. (2010) sought evidence of small ‘‘gene conversion’’ or nonreciprocal
exchanges among homoeologs, based on the presence in allopolyploid cotton of
genes that have genome-diagnostic (A or D) SNPs (single nucleotide polymorphisms) that occurred in patterns suggestive of genic interactions (e.g., a single
expressed sequence containing diagnostic SNPs in the following order, AADDAA,
implicating conversion of the middle section of an AT homoeolog by the DT
homoeolog). These bioinformatic inferences were validated by de novo PCR and
sequencing. Results were convincing and compelling, demonstrating that about
2 % of contigs in G. hirsutum have experienced nonreciprocal homoeologous
exchanges since the origin of polyploid cotton 1–2 mya. Moreover, when a
sampling of these homoeologous interactions was studied throughout the polyploid
clade using a phylogenetic approach, nonreciprocal homoeologous exchanges
were shown to have occurred throughout polyploid divergence and speciation, as
opposed to being concentrated at the root of the polyploid tree. Among six
homoeologous interactions, five occurred in only one species, with the sixth event
being shared. This result refutes a logical a priori prediction, namely that interactions among homoeologs should be most frequent early in allopolyploid evolution (and hence shared among species), prior to reinforcement of apparently
strict bivalent pairing. Finally, some genomic regions showed multiple patterns of
homoeologous recombination among species, suggesting that some regions or
genes may be ‘‘hot-spots’’ for nonreciprocal homoeologous exchanges.
EC
421
CO
RR
420
J. F. Wendel et al.
UN
Editor Proof
194
Layout: T1 Standard SC
Chapter No.: 10
Book ISBN: 978-3-642-31441-4
Page: 195/206
Jeans, Genes, and Genomes: Cotton as a Model for Studying Polyploidy
195
486
10.3 Duplicate Gene Expression
470
471
472
473
474
475
476
477
478
479
480
481
482
483
484
487
488
489
490
491
492
493
494
495
496
497
498
499
500
501
502
503
PR
OO
469
D
468
TE
467
EC
466
In addition to evolutionary changes in gene and genome structure, a key component of polyploid evolution concerns the consequences of genome doubling on
gene expression. The biological reunion in the allopolyploid nucleus of two regulatory systems that had evolved in isolation for 5–10 my (A- and D-genome), in
only one of the two parental cytoplasms (A-genome), might be expected to lead to
violations of the equal and additive expression modeled in Fig. 10.3. In principle,
both genomes in an allotetraploid could contribute equally to the transcriptome, for
any pair of homoeologs or overall, or alternatively, there may be preferential
transcription of one genome due to intergenomic interactions that could bias the
transcription machinery. Many aspects of this problem are of interest, including
the scope and scale of preferential homoeolog expression, its tissue and organ
specificity, the level of genomic bias, the mechanistic underpinnings that result in
regulatory responses, the temporal scale at which alterations in homoeolog expression evolve, and ultimately, its physiological and evolutionary relevance.
Steps toward answering these questions for Gossypium have been taken using
many different experimental approaches over the last decade. We summarize these
results below, focusing on lessons regarding (1) developmental and environmental
CO
RR
465
F
485
This demonstration of non-independent evolution of homoeologs has recently
been confirmed and extended to tens of thousands of genes using an expanded EST
set (Flagel et al. 2012). This analysis resulted in an even higher percentage (5 %)
of contigs in the assembly that show evidence of homoeolog contact in at least one
of the two polyploid species studied (G. hirsutum and G. barbadense), as well as
confirm the suggestion of hot-spots for homoeologous recombination. This high
frequency of homoeologous contact is an astonishing result given the absence of
prior cytogenetic or other observations that would have suggested this possibility,
and the apparently complete bivalent formation at meiosis. Flagel et al. (2012)
also detected 50 % more homoeologous exchanges in G. hirsutum than in
G. barbadense, suggesting that following polyploid formation, rates of nonreciprocal homoeologous recombination may diverge, even among closely related
species that share nearly all evident ecological and life-history characteristics.
These results are fascinating and lead to a number of questions. For example, what
is the genomic distribution of genes subject to nonreciprocal recombination events,
and does this information suggest a causal mechanism? Are nonreciprocal
recombination events random with respect to outcome (A vs D), or alternatively, is
the nuclear genome of allopolyploid cotton slowly becoming more ‘‘A-like’’ or
D-like’’? Do any of the detected ‘‘gene conversion’’ events have physiological
consequences and hence possible adaptive significance? Insight into these and
related questions in cotton and in other allopolyploids is likely to emerge in the next
several years.
464
UN
Editor Proof
10
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 10
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 196/206
510
511
512
513
514
515
516
517
518
519
520
521
522
523
524
525
526
527
528
529
530
531
532
533
534
535
536
537
538
539
540
541
542
543
F
PR
OO
508
509
The first indication that polyploidy in Gossypium is accompanied by extensive
organ-specific changes in duplicate gene expression emerged nearly a decade ago in
a study of 40 homoeologous gene pairs in different organs of G. hirsutum using
SSCP-cDNA (Adams et al. 2003). Almost one-third of the genes revealed bias
toward one homoeolog or the other, or only expression of one homoeolog, in at least
one organ. Transcript levels for the two members of each gene pair varied by gene
and, unexpectedly, by organ type. Especially noteworthy were genes that showed
organ-specific, reciprocal silencing of alternative homoeologs; that is, one member
of a duplicated gene pair displayed minimal to no transcription in some organs,
whereas a reciprocal pattern was exhibited by the alternative homoeolog in other
organ(s). In particular, floral organs showed dramatic expression patterns in this
regard, with major differences among petals, stamens, and stigmas/styles (Adams
et al. 2003, 2004). Organ-specific expression of homoeologs was assayed more
extensively by Chaudhary et al. (2009), who employed a novel high-resolution
mass-spectrometry technology (Sequenom) to investigate relative expression levels
of each homoeolog for 63 gene pairs in 24 tissues. Results from over 2,000 assays
demonstrated that 40 % of homoeologs are transcriptionally biased in at least one
stage of cotton development, that genome merger per se has a large effect on
relative expression of homeologs (see section on temporal partitioning, below), and
that the majority of alterations are caused by cis-regulatory divergence between
diploid progenitors. The study also revealed 15 cases of probable regulatory
neofunctionalization among 8 tissues, perhaps the first such demonstration in
allopolyploid plants.
These surprising indications of partitioning of duplicate gene expression during
development have since been found in many other allopolyploid plants (see also
Chap. 14, this volume, and, e.g., Buggs et al. 2010; Bottley and Koebner 2008;
Bottley et al. 2006; Wang et al. 2006), opening the experimental floodgates on the
study of duplicate gene expression. Because differences in duplicate gene
expression were found between different organ types, the question arose as to
patterns of duplicate gene expression during development of any single organ. Liu
and Adams (2007) examined the expression ratio of homoeologous AdhA genes in
eight developmental stages of hypocotyls, cotyledons, and roots, and 11 developmental stages of ovary walls and ovules. They showed that expression ratios of
the two homoeologs changed considerably when comparing some stages of organ
development, indicating that differential expression of homoeologous genes is
developmentally regulated and that determining the extent of homoeologous gene
D
507
10.3.1 Developmental and Environmental Effects on Duplicate
Gene Expression
TE
506
effects on gene expression; (2) global analyses of genomic interactions; and (3) the
temporal scale at which expression evolution arises.
EC
505
CO
RR
504
J. F. Wendel et al.
UN
Editor Proof
196
Layout: T1 Standard SC
Chapter No.: 10
Book ISBN: 978-3-642-31441-4
Page: 197/206
Jeans, Genes, and Genomes: Cotton as a Model for Studying Polyploidy
197
577
10.3.2 Global Biases in Duplicate Gene Expression
550
551
552
553
554
555
556
557
558
559
560
561
562
563
564
565
566
567
568
569
570
571
572
573
574
575
578
579
580
581
582
583
584
PR
OO
549
D
548
TE
547
EC
546
CO
RR
545
F
576
expression changes requires analysis of multiple developmental stages and organs.
An important extension of this line of analysis was to ask whether biased
expression of homoeologs can be cell type-specific. In this respect, cotton offers an
excellent opportunity because the single-celled fibers are easily accessible. To
address this question, Hovav et al. (2008a, b) used homoeolog-specific microarrays
to assay homoeologous gene expression during fiber development. In these
remarkable studies, discussed further below, a full range of duplicate gene
expression was observed among the approximately 1,400 genes, from balanced
expression of homoeologs, to varying homoeolog ratios that shifted during
development. Notably, four genes showed complete reciprocal silencing of alternative homoeologs during fiber development. Collectively, the foregoing studies
show that variation in homoeologous gene expression during development is the
rule rather than the exception, and that it occurs even during development of a
single cell type.
An additional twist on duplicate gene expression has been the demonstration that
homoeolog expression patterns are not simply developmentally fixed, but instead
are responsive to environmental conditions. Liu and Adams (2007) examined
expression of AdhA homoeologs in response to abiotic stress conditions, with a
focus on cold and water submersion (flooding simulation), at different developmental stages. They found that some stress treatments significantly altered AT to DT
expression ratios, including a case of reciprocal silencing of homoeologs in
response to abiotic stress, an unprecedented finding. In a more recent study, Dong
and Adams (2011) extended their analysis to 30 gene pairs using three organs
(leaves, roots, and cotyledons) and five abiotic stress treatments (heat, cold,
drought, high salt, and water submersion). Over 70 % of the genes showed stressinduced changes in the relative expression levels of the duplicates under one or
more stress treatments, and 12 sets of homoeologs showed opposite changes in
expression levels in response to different abiotic stress treatments. These results
indicate that abiotic stress conditions can have considerable effects on duplicate
gene expression in a polyploid, with the effects varying by gene, stress, and organ
type. Together with the profound developmental partitioning noted above, differential expression in response to environmental stresses or cues may all be factors
that contribute to the preservation of duplicated genes in polyploids.
544
To provide a more global perspective on homologous gene expression in
allopolyploids, Udall et al. (2006) developed homoeolog-specific microarrays that
utilized genome-diagnostic SNPs from ESTs generated from the two genomes of
allopolyploid cotton and the diploids G. arboreum (A-genome) and G. raimondii
(D-genome). Using leaf RNA, they found that 199 of 461 gene pairs (43 %)
deviated from equal expression, thereby providing an initial quantitative
perspective on the scale of biased homoeolog expression. This microarray
UN
Editor Proof
10
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 10
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 198/206
592
593
594
595
596
597
598
599
600
601
602
603
604
605
606
607
608
609
610
611
612
613
614
615
616
617
618
619
620
621
622
623
624
625
626
627
628
629
F
591
PR
OO
590
D
589
TE
588
EC
587
approach was refined and extended to a larger set of gene pairs (n = 1383) by
Flagel et al. (2008), who reported that 70 % of the genes in petals have biased
homoeolog expression ratios, and that more of these genes are D-genome (39.5 %)
than A-genome (30.5 %) biased. In addition, they found that the D-genome copies
of 69 genes and the A-genome copies of 46 genes were silenced, collectively
representing about 8 % of all genes. Similarly, Hovav et al. (2008a, b) used the
same SNP-specific microarray technology on RNAs extracted from trichomes
harvested from three developmental time points in wild and modern accessions of
two independently domesticated cotton species, G. hirsutum and G. barbadense.
Among these species 25–37 % of genes were significantly biased toward one of
the two genomes at each developmental stage, but these biases were not random
with respect to genome-of-origin; instead, duplicate gene expression was biased
toward the D-genome at all three time points studied, accounting for 63–76 % of
the biased genes. Finally, and to place homoeolog expression evolution in a
phylogenetic context, Flagel and Wendel (2010) extended their analysis of
duplicate gene expression in petals to all five extant allopolyploid species. Several
aspects of this study are notable, including the demonstration that all five species
display an overall preference for D-genome expression (D-genome bias accounting for 54–60 % of genes with biased homoeolog expression), that the percentage
of duplicate genes that are biased varies widely among species (from 48 to 88 %),
and that the overall magnitude of bias (as opposed to simply whether or not a gene
exhibits bias) similarly varies widely among species.
The foregoing synopsis shows that gene expression is massively altered in
polyploid cotton relative to its diploid progenitors. Unequal expression of one of
the two homoeologs likely is the rule rather than the exception, when integrated
across organs and tissues, with gene silencing representing only the endpoint in a
continuum of biased homoeolog expression. A global bias in gene expression from
one of two co-resident genomes (the D-genome; see, however, Yang et al. 2006) is
evident in allopolyploid cotton. At present, little evidence connects these observations to physiology or metabolism, and so connections to function and
ecological relevance remain obscure (see, however, section on fiber below).
Recent data (Hu et al. 2011) have extended the concept of bias to the proteomic
level, where allopolyploid cotton seeds have been shown to preferentially
accumulate proteins from the D-genome diploid parent, suggesting the possibility
of a functional connection. A promising direction for future research is targeted,
functional studies of specific gene families, protein complexes, and metabolic
pathways and networks, aiming to place homoeolog expression into a context that
permits insight into the biological significance of expression bias.
An emerging phenomenon in the study of polyploid gene expression on a large
scale is the concept of genomic dominance in expression. This phenomenon,
originally discovered and elaborated in cotton (Flagel and Wendel 2010; Rapp
et al. 2009), and just recently detected in Coffea allopolyploids (Bardil et al. 2011),
is defined as the state where total expression of a homoeolog pair mimics the
expression level of one of two diploid parents of an allopolyploid. That is, if parent
A is upregulated relative to parent B, the allopolyploid would exhibit the
CO
RR
585
586
J. F. Wendel et al.
UN
Editor Proof
198
Layout: T1 Standard SC
Chapter No.: 10
636
637
638
639
640
641
642
643
644
645
646
647
648
649
650
651
652
653
654
655
656
657
658
659
660
661
662
663
664
665
666
667
668
669
670
F
635
PR
OO
634
10.3.3 Temporal Dynamics of Duplicate Gene Expression
Evolution
One of the early (Adams et al. 2003) and subsequently further characterized
findings about duplicate gene expression in cotton is that expression evolution has
a temporal dimension; phenomena that typify genome merger and doubling need
not be the same, qualitatively or quantitatively, as those that characterize natural
allopolyploids that are 1–2 million years distant from polyploidization (Fig. 10.4).
In some cases, such as organ-specific silencing of particular homoeologs, it has
been shown that this can arise immediately upon allopolyploid formation (Adams
et al. 2003, 2004; Chaudhary et al. 2009), leading to the notion (Adams et al. 2003;
Rapp and Wendel 2005) that homoeolog silencing may be significant in duplicate
gene retention and that it may actually represent a form of instantaneous
subfunctionalization. These studies of individual genes have demonstrated that
some of the changes detected in synthetic allopolyploids are mirrored in natural
allopolyploids, whereas others are distinct between the synthetic and natural
allopolyploids (see examples in Adams et al. 2004; Chaudhary et al. 2009).
One possibility raised by these observations is that in some cases there is an
immediate epigenetic response to genome merger and doubling that becomes
either epigenetically stable or genetically fixed, for 1–2 million years.
With respect to the phenomena of genomic bias, the evolution of novel quantitative (transgressive) expression states, and genomic dominance, the studies of
Flagel et al. (2008, 2010) have been particularly illuminating with respect to this
temporal dimension to expression evolution. For example, whereas only about
one-third of genes exhibit biased homoeolog expression in petals of AxD-genome,
diploid hybrids, half or more do in petals of natural allopolyploids. Moreover, many
of the genes that display bias in the initial F1 retain or even magnify the degree of
bias after 1–2 million years. Also, whereas allopolyploid formation appears to
result in relatively few genes that exhibit quantitatively transgressive expression
levels, relative to their diploid parents, this number rises sharply in natural allopolyploids, suggesting novel opportunities for exploring gene expression space.
Finally, a comparison of genomic dominance in all five natural allopolyploids
(Flagel and Wendel 2010) to that observed in synthetic allopolyploids (Rapp et al.
2009) demonstrated that the magnitude of genomic dominance remains high after
D
633
TE
632
199
expression state of parent A; in the case where parent A is downregulated relative
to parent B, so is the allopolyploid. In this example, the A parent is genomically
dominant with respect to gene expression. Using two different genomic combinations in synthetic allopolyploid Gossypium, Rapp et al. (2009) demonstrated that
there exists a quantitative bias in genomic dominance toward one of the two
parents. In the case of the synthetic AD allopolyploid, more than 10,000 genes
display this dominant, quantitative expression phenotype. At present, the functional significance of genomic dominance is unknown.
EC
631
CO
RR
630
Book ISBN: 978-3-642-31441-4
Page: 199/206
Jeans, Genes, and Genomes: Cotton as a Model for Studying Polyploidy
UN
Editor Proof
10
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 10
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 200/206
J. F. Wendel et al.
Editor Proof
200
G. hirsutum
AD
AADD
PR
OO
G. barbadense
G. hirsutum
F
G. tomentosum
G. darwinii
G. barbadense
G. mustelinum
“Gene conversion”
Gene loss
Genomic dominance
Concerted evolution
Genetic sub- and neofunctionalization
Intergenomic transfer
Biased homoeolog expression
Transgressive expression
Homoeologous expression change
Biased genomic dominance
Epigenetic subfunctionalization
673
674
675
676
677
678
679
680
681
682
683
684
685
686
687
688
1–2 my of evolution, but that the bias in its direction dissipates (shifting from
predominantly D-genome dominance in the synthetics to similar levels of A- and
D-genome dominance in the natural allopolyploids). Thus, it appears that natural
allopolyploids have adjusted to more equally utilize the transcriptomes of the two
co-resident genomes, notwithstanding the residual D-genome homoeolog bias
described above. An intriguing possibility is that this D-genome homoeolog bias is
connected causally to the biased D-genome expression dominance that arises following genome merger; to the extent that it is, it leads to the suggestion that
homoeolog bias in other allopolyploids may be predicted by the initial conditions
established by genomic merger in the distant past, and which may be experimentally mirrored in many systems through the use of synthetic hybrids and
allopolyploids.
The partitioning of gene expression evolution into its temporal components
leads to the suggestion that these different components may entail different or at
least complementary mechanisms. The first, involving rapid or instantaneous gene
expression alteration as a consequence of genome merger and doubling, reflects
the myriad novel interactions accompanying a biological reunion of two differentiated genomes into a common nucleus. The precise nature of these interactions
CO
RR
672
UN
671
EC
TE
D
Fig. 10.4 A temporal depiction of phenomena that characterize polyploid evolution in
Gossypium. Shown are the formation and diversification of the allopolyploid clade, several
accompanying phenomena (dispersal, chromosomal stabilization), and the parallel domestication
of G. hirsutum and G. barbadense. Images to the right represent wild and domesticated states,
with close-ups of single seeds and the transformations in morphology accompanying domestication and crop improvement. Genetic and epigenetic phenomena associated with polyploidy in
cotton are shown below, with a timeline (bottom), illustrating the temporal context for each
phenomenon and whether there are increases or decreases (where known) in magnitude over time
(blue to purple gradation indicates increase, the reverse indicates decrease)
Layout: T1 Standard SC
Chapter No.: 10
Book ISBN: 978-3-642-31441-4
Page: 201/206
Jeans, Genes, and Genomes: Cotton as a Model for Studying Polyploidy
201
699
10.4 Polyploidy and Ecological Novelty
695
696
697
700
701
702
703
704
705
706
707
708
709
710
711
712
713
714
715
716
717
718
719
720
721
722
723
724
725
726
727
728
729
PR
OO
694
The foregoing discussion of the genomic and genetic attributes of allopolyploid
cotton demonstrates that polyploid formation has led to a diverse array of genetic
and genomic responses, including non-Mendelian transmission and various forms
of non-additive gene expression. The question naturally arises as to whether
allopolyploidy has stimulated novel adaptation or physiological capability.
A voluminous literature in plants documents the frequency of polyploids in various
habitats, their morphological and physiological attributes, and their ecological
success relative to diploids (Grant 1981; Stebbins 1947, 1950; Soltis and Soltis
2000). One generalization that has emerged is that polyploidy often is associated
with broader ecological amplitude and novel evolutionary opportunity, often
suggested to be mediated by the increased ‘‘buffering’’ capacity afforded by
duplicated genes and the enhanced vigor resulting from the ‘‘fixed heterozygosity’’
of their duplicated genomes. We might now rephrase or expand these suggested
mechanistic explanations to encompass a network perspective, and the vastly
increased combinatorial possibilities for regulation and evolution enabled by a
suddenly duplicated complement of genes and merged regulatory systems.
With respect to Gossypium, allopolyploidy led to the apparent invasion of a new
ecological niche. In considering the Pleistocene origin of allopolyploid cotton,
Fryxell (1965, 1979) noted that in contrast to the majority of diploid species, which
occur mostly inland in various arid to seasonally arid environments, allopolyploid
species typically occur in coastal habitats, at least those forms that arguably are
truly wild (see also Brubaker and Wendel 1994). Thus, among the five allopolyploid species, two are completely restricted to near coastlines, in that they
are island endemics (G. darwinii and G. tomentosum), and for two others
(G. barbadense and G. hirsutum), wild forms occur disparately in littoral habitats
ringing the Gulf of Mexico, northwest South America, and distant Pacific Islands.
The capacity for oceanic dispersal in Gossypium (Fryxell 1965, 1979; Stephens
1958, 1966) was associated at the allopolyploid level with specialization for
establishment in coastal communities. Fryxell (1965, 1979) forwarded the tantalizing suggestion that following initial formation, adaptation of the newly evolved
D
693
TE
692
EC
691
CO
RR
690
F
698
is not known, but probably includes disruptions in gene dosage balance, stoichiometric changes resulting from differences in competition for transcription factors,
differences in microRNA expression, and a host of novel cis and trans interactions
(Birchler et al. 2005; Birchler and Veitia 2010; Chen 2007; Osborn et al. 2003;
Veitia 2005) (see also Chaps. 1, 2, and 4, this volume). Superimposed on these
rapid evolutionary responses to polyploidy are those that arise more slowly during
the stabilization of the new polyploid genome and during evolution and speciation
over much longer periods of time. Thus, the presence of duplicated genomes
would seem to provide evolutionary opportunity and consequences immediately
on polyploid formation and for millions of years thereafter.
689
UN
Editor Proof
10
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 10
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 202/206
J. F. Wendel et al.
737
10.5 Polyploidy and Fiber
738
739
740
741
742
743
744
745
746
747
748
749
750
751
752
753
754
755
756
757
758
759
760
761
762
763
764
765
766
767
768
769
770
PR
OO
735
Finally, we would be remiss if we failed to consider the consequences of polyploidy on the development of agronomically advanced cultivars of allopolyploid
cotton. Although four separate species of Gossypium were independently
domesticated for their seed hairs, the characteristic that attracted the attention of
the earliest domesticators, the seed ‘‘lint’’ itself, evolved only once in the progenitor of the A-genome diploids (Hovav et al. 2008c; Wendel et al. 2009).
Applequist et al. (2001) generated growth curves for trichomes from cultivated and
wild diploid and allopolyploid species and demonstrated that the evolution of an
extended primary wall elongation occurred in the ancestor of wild A-genome
cotton prior to domestication and in Africa. Follow-up comparative expression
profiling experiments (Hovav et al. 2008a, b; Rapp et al. 2010) further identify
some of the metabolic pathways that were modified to enable this evolution in
fiber properties. These results led to the fascinating implication that domestication
of the New World allopolyploid cottons (which contain an A-genome, in addition
to a D-genome) that presently dominate cotton agriculture worldwide was first
precipitated by developmental and physiological transformations that occurred
hundreds of thousands of years ago in a different hemisphere.
Because fibers from all D-genome diploids are short and non-spinnable, it is
particularly interesting that fiber from the cultivated (New World) allopolyploids is
agronomically superior to that of the cultivated A-genome diploids; in this sense,
the cultivated allotetraploid fiber morphology is ‘‘non-additive’’, or perhaps
‘‘heterotic’’. A number of studies have noted this point (Jiang et al. 1998; Paterson
2005; Wright et al. 1998), suggesting that allopolyploidization provided novel
opportunities for agronomic improvement. A recent meta-analysis (Rong et al.
2007) of a large number of QTL studies in allopolyploid cotton leads to a general
picture consistent with this interpretation; more loci affecting fiber yield and
quality traits are found in the DT (n = 221) than the AT (n = 184) genome,
possibly explaining the superiority of the lint of the allopolyploids relative to the
A-genome diploids. Support for this speculation that ‘‘recruitment’’ of D-genome
genes has been important in enabling the development of advanced allotetraploid
cultivars is also emerging from comparative expression profiling studies (Hovav
et al. 2008a, b; Rapp et al. 2010), which reveal in exquisite detail the thousands of
gene expression differences that distinguish wild from domesticated cotton fiber
D
734
TE
733
EC
732
CO
RR
731
F
736
allopolyploid to littoral habitats enabled it to exploit the fluctuating sea levels
that characterized the Pleistocene. This ecological innovation not only is
envisioned to have permitted the initial establishment of the nascent polyploid
lineage, but is also suggested to have provided a means for the rapid dispersal of
the salt-water-tolerant seeds. By this means, perhaps, the mobile shorelines of the
Pleistocene facilitated exploitation of a new ecological niche, and hence colonization of the New World tropics.
730
UN
Editor Proof
202
Layout: T1 Standard SC
Chapter No.: 10
Book ISBN: 978-3-642-31441-4
Page: 203/206
Jeans, Genes, and Genomes: Cotton as a Model for Studying Polyploidy
203
788
789
790
791
792
Acknowledgments Research in the Wendel lab has largely been funded by the NSF Plant
Genome Program, with additional support from other NSF programs, the USDA NRI, and Cotton
Incorporated. Research in the Adams lab has been supported by the Natural Science and Engineering Research Council of Canada and by the USDA NRI program. We gratefully acknowledge
all of these sources of support.
793
References
794
795
796
797
798
799
800
801
802
803
804
805
806
807
808
809
810
811
812
813
Adams KL, Cronn R, Percifield R, Wendel JF (2003) Genes duplicated by polyploidy show
unequal contributions to the transcriptome and organ-specific reciprocal silencing. Proc Nat
Acad Sci USA 100:4649–4654
Adams KL, Percifield R, Wendel JF (2004) Organ-specific silencing of duplicated genes in a
newly synthesized cotton allotetraploid. Genetics 168:2217–2226
Applequist WL, Cronn RC, Wendel JF (2001) Comparative development of fiber in wild and
cultivated cotton. Evol Dev 3:3–17
Bardil A, Dantas de Almeida J, Combes MC, Lashermes P, Bertrand B (2011) Genomic
expression dominance in the natural allopolyploid Coffea arabica is massively affected by
growth temperature. New Phytol (in press)
Beasley JO (1941) Hybridization, cytology, and polyploidy of Gossypium. Chronica Bot
6:394–395
Bennett MD, Leitch IJ (2005) Genome size evolution in plants. In: Gregory TR (ed) The
evolution of the genome. Elsevier, San Diego, pp 89–162
Birchler JA, Riddle NC, Auger DL, Veitia RA (2005) Dosage balance in gene regulation:
biological implications. Trends Genet 21:219–226
Birchler JA, Veitia RA (2010) The gene balance hypothesis: implications for gene regulation,
quantitative traits and evolution. New Phytol 186:54–62
Bottley A, Koebner RM (2008) Variation for homoeologous gene silencing in hexaploid wheat.
Plant J 56:297–302
778
779
780
781
782
783
784
785
786
PR
OO
777
D
775
776
TE
774
EC
773
CO
RR
772
F
787
development, as well as a bias toward preferential expression of D-genome
homoeologs. Finally, similar implications emanate from genetic mapping experiments, where it has been shown that for 535 genes implicated in cotton fiber
development, more transcription factors were from DT than AT genome, whereas
the reverse was true for fiber development genes (Xu et al. 2010). These data are
interpreted to suggest that the D-genome ancestor provided key transcription
factors that regulate the expression of fiber genes donated by the ancestral
A-genome parent. Taken together, these studies may provide actual genetic evidence for a speculation forwarded 75 years ago by Harland (1936), who stated If
as a consequence of polyploidy a large number of genes become duplicated, and
the characters governed by such genes are of importance to the species, one of the
members may mutate, leaving the character unimpaired, with the further possibility that the mutation may be of benefit to the species. An exciting prospect is that
in the near future we will develop a deeper understanding of the nature of these
genes, the molecular genetic meaning of Harland’s invocation of the word
‘‘mutation’’, and their effects on the developmental networks that underlie altered
morphology and agronomic improvement.
771
UN
Editor Proof
10
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 10
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 204/206
EC
TE
D
PR
OO
F
Bottley A, Xia G, Koebner R (2006) Homoeologous gene silencing in hexaploid wheat. Plant J
47:897–906
Brubaker CL, Bourland FM, Wendel JF (1999a) The origin and domestication of cotton. In:
Smith CW, Cothren JT (eds) Cotton; origin, history, technology and production. Wiley,
New York, pp 3–31
Brubaker CL, Paterson AH, Wendel JF (1999b) Comparative genetic mapping of allotetraploid
cotton and its diploid progenitors. Genome 42:184–203
Brubaker CL, Wendel JF (1993) On the specific status of Gossypium lanceolatum Todaro. Genet
Res Crop Evol 40:165–170
Brubaker CL, Wendel JF (1994) Reevaluating the origin of domesticated cotton (Gossypium
hirsutum; Malvaceae) using nuclear restriction fragment length polymorphisms (RFLPs).
Am J Bot 81:1309–1326
Brubaker CL, Wendel JF (2001) RFLP diversity in cotton. In: Jenkins JN, Saha S (eds) Genetic
improvement of cotton: emerging technologies. Science Publishers, Inc., Enfield, pp 81–102
Buggs RJA, Elliott NM, Zhang L, Koh J, Viccini LF, Soltis DE, Soltis PS (2010) Tissue-specific
silencing of homoeologs in natural populations of the recent allopolyploid Tragopogon mirus.
New Phytol 186:175–183
Chaudhary B, Flagel L, Stupar RM, Udall JA, Verma N, Springer NM, Wendel JF (2009)
Reciprocal silencing, transcriptional bias and functional divergence of homeologs in
polyploid cotton (Gossypium). Genetics 182:503–517
Chen ZJ (2007) Genetic and epigenetic mechanisms for gene expression and phenotypic variation
in plant polyploids. Annu Rev Plant Biol 58:377–406
Cronn R, Small RL, Wendel JF (1999) Duplicated genes evolve independently following
polyploid formation in cotton. Proc Nat Acad Sci USA 96:14406–14411
Cronn RC, Zhao X, Paterson AH, Wendel JF (1996) Polymorphism and concerted evolution in a
tandemly repeated gene family: 5S ribosomal DNA in diploid and allopolyploid cottons.
J Mol Evol 42(6):685–705
DeJoode DR, Wendel JF (1992) Genetic diversity and origin of the Hawaiian Islands cotton,
Gossypium tomentosum. Amer J Bot 79:1311–1319
Dong S, Adams KL (2011) Differential contributions to the transcriptome of duplicated genes in
response to abiotic stresses in natural and synthetic polyploids. New Phytol 190(4):1045–1057
Elder JF, Turner BJ (1995) Concerted evolution of repetitive DNA sequences in eukaryotes.
Quart Rev Biol 70:297–320
Endrizzi JE, Turcotte EL, Kohel RJ (1985) Genetics, cytology, and evolution of Gossypium. Adv
Genet 23:271–375
Flagel L, Udall J, Nettleton D, Wendel J (2008) Duplicate gene expression in allopolyploid
Gossypium reveals two temporally distinct phases of expression evolution. BMC Biol 6:11
Flagel LE, Wendel JF (2010) Evolutionary rate variation, genomic dominance and duplicate gene
expression evolution during allotetraploid cotton speciation. New Phytol 186:184–193
Flagel LE, Wendel JF, Udall JA (2012) Duplicate gene evolution, homoeologous recombination,
and transcriptome characterization in allopolyploid cotton. BMC Genomics (in press)
Fryxell PA (1965) Stages in the evolution of Gossypium. Adv Frontiers Plant Sci 10:31–56
Fryxell PA (1968) A redefinition of the tribe Gossypieae. Bot Gaz 129:296–308
Fryxell PA (1979) The natural history of the cotton tribe Texas. A&M University Press College
Station, College Station
Fryxell PA (1992) A revised taxonomic interpretation of Gossypium L. (Malvaceae). Rheedea
2:108–165
Galau GA, Wilkins TA (1989) Alloplasmic male sterility in AD allotetraploid Gossypium
hirsutum upon replacement of its resident a cytoplasm with that of D species G. harknessii.
Theor Appl Genet 78:23–30
Grant V (1981) Plant speciation. Columbia University Press, New York
Grover CE, Kim H, Wing RA, Paterson AH, Wendel JF (2004) Incongruent patterns of local and
global genome size evolution in cotton. Genome Res 14:1474–1482
CO
RR
814
815
816
817
818
819
820
821
822
823
824
825
826
827
828
829
830
831
832
833
834
835
836
837
838
839
840
841
842
843
844
845
846
847
848
849
850
851
852
853
854
855
856
857
858
859
860
861
862
863
864
865
866
J. F. Wendel et al.
UN
Editor Proof
204
Layout: T1 Standard SC
Chapter No.: 10
205
EC
TE
D
PR
OO
F
Grover CE, Kim H, Wing RA, Paterson AH, Wendel JF (2007) Microcolinearity and genome
evolution in the AdhA region of diploid and polyploid cotton (Gossypium). Plant J 50:
995–1006
Hanson RE, Islam-Faridi MN, Crane CF, Zwick MS, Czeschin DG, Wendel JF, Mcknight TD,
Price HJ, Stelly DM (1999) Ty1-copia-retrotransposon behavior in a polyploid cotton.
Chromosome Res 8:73–76
Hanson RE, Zhao X-P, Islam-Faridi MN, Paterson AH, Zwick MS, Crane CF, McKnight TD,
Stelly DM, Price HJ (1998) Evolution of interspersed repetitive elements in Gossypium
(Malvaceae). Am J Bot 85:1364–1368
Harland SC (1936) The genetical conception of the species. Cambridge Philos Soc Biol Rev
11:83–112
Hawkins JS, Hu G, Rapp RA, Grafenberg JL, Wendel JF (2008) Phylogenetic determination of
the pace of transposable element proliferation in plants: copia and LINE-like elements in
Gossypium. Genome 51:11–18
Hawkins JS, Kim H, Nason JD, Wing RA, Wendel JF (2006) Differential lineage-specific
amplification of transposable elements is responsible for genome size variation in Gossypium.
Genome Res 16(10):1252–1261
Hawkins JS, Proulx SR, Rapp RA, Wendel JF (2009) Rapid DNA loss as a counterbalance to
genome expansion through retrotransposon proliferation in plants. Proc Nat Acad Sci USA
106(42):17811–17816
Hendrix B, Stewart JM (2005) Estimation of the nuclear DNA content of Gossypium species. Ann
Bot 95:789–797
Hovav R, Chaudhary B, Udall JA, Flagel L, Wendel JF (2008a) Parallel domestication,
convergent evolution and duplicated gene recruitment in allopolyploid cotton. Genetics
179(3):1725–1733
Hovav R, Udall J, Chaudhary B, Flagel L, Rapp R, Wendel J (2008b) Partitioned expression of
duplicated genes during development and evolution of a single cell in a polyploid plant. Proc
Nat Acad Sci USA 105:6191
Hovav R, Udall JA, Chaudhary B, Hovav E, Flagel L, Hu G, Wendel JF (2008c) The evolution of
spinnable cotton fiber entailed prolonged development and a novel metabolism. PLoS Genet
4:e25
Hu G, Hawkins JS, Grover CE, Wendel JF (2010) The history and disposition of transposable
elements in polyploid Gossypium. Genome 53:599–607
Hu G, Houston NL, Pathak D, Schmidt L, Thelan JJ, Wendel JF (2011) Genomically biased
accumulation of seed storage proteins in allopolyploid cotton. Genetics 189:1103–1115
Hutchinson JB (1951) Intra-specific differentiation in Gossypium hirsutum. Heredity 5:161–n193
Hutchinson JB (1954) New evidence on the origin of the old world cottons. Heredity 8:
225–241
Hutchinson JB, Silow RA, Stephens SG (1947) The evolution of Gossypium and the
differentiation of the cultivated cottons. Oxford University Press, London
Jiang C, Wright R, El-Zik K, Paterson A (1998) Polyploid formation created unique avenues for
response to selection in Gossypium (cotton). Proc Nat Acad Sci USA 95:4419–4424
Kashkush K, Feldman M, Levy AA (2002) Gene loss, silencing, and activation in a newly
synthesized wheat allotetraploid. Genetics 160:1651–1659
Krapovickas A, Seijo G (2008) Gossypium ekmanianum (Malvaceae), algodon silvestre de la
Republica Dominicana. Bonplandia 17:55–63
Lin L, Paterson AH (2009) Physical composition and organization of the Gossypium genomes. In:
Paterson AH (ed) Genomics of cotton, plant genetics and genomics, crops and models 3.
Springer, New York, pp 141–156
Lin LF, Tang HB, Compton RO, Lemke C, Rainville LK, Wang XY, Rong JK, Rana MK,
Paterson AH (2011) Comparative analysis of Gossypium and Vitis genomes indicates genome
duplication specific to the Gossypium lineage. Genomics 97:313–320
Liu B, Brubaker CL, Mergeai G, Cronn RC, Wendel JF (2001) Polyploid formation in cotton is
not accompanied by rapid genomic changes. Genome 44:321–330
CO
RR
867
868
869
870
871
872
873
874
875
876
877
878
879
880
881
882
883
884
885
886
887
888
889
890
891
892
893
894
895
896
897
898
899
900
901
902
903
904
905
906
907
908
909
910
911
912
913
914
915
916
917
918
919
920
Book ISBN: 978-3-642-31441-4
Page: 205/206
Jeans, Genes, and Genomes: Cotton as a Model for Studying Polyploidy
UN
Editor Proof
10
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 10
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 206/206
EC
TE
D
PR
OO
F
Liu Z, Adams KL (2007) Expression partitioning between genes duplicated by polyploidy under
abiotic stress and during organ development. Curr Biol 17:1669–1674
Muravenko O, Fedotov AR, Punina EO, Federova LI, Grif VG, Zelenin AV (1998) Comparison
of chromosome BrdU-hoechst-giemsa banding patterns of the A1 and (AD)2 genomes of
cotton. Genome 41:616–625
Osborn TC, Chris Pires J, Birchler JA, Auger DL, Jeffery Chen Z, Lee H-S, Comai L, Madlung A,
Doerge RW, Colot V, Martienssen RA (2003) Understanding mechanisms of novel gene
expression in polyploids. Trends Genet 19(3):141–147
Paterson AH (2005) Polyploidy, evolutionary opportunity, and crop adaptation. Genetica 123:191
Paterson AH (2009) Genomics of cotton, plant genetics and genomics, crops and models 3.
Springer, New York
Percy RG, Wendel JF (1990) Allozyme evidence for the origin and diversification of Gossypium
barbadense L. Theor Appl Genet 79:529–542
Rapp R, Haigler C, Flagel L, Hovav R, Udall J, Wendel J (2010) Gene expression in developing
fibres of Upland cotton (Gossypium hirsutum L.) was massively altered by domestication.
BMC Biol 8:139
Rapp R, Wendel J (2005) Epigenetics and plant evolution. New Phytol 168:81
Rapp RA, Udall JA, Wendel JF (2009) Genomic expression dominance in allopolyploids. BMC
Biol 7:18
Reinisch AJ, Dong J, Brubaker CL, Stelly DM, Wendel JF, Paterson AH (1994) A detailed RFLP
map of cotton, Gossypium hirsutum x G. barbadense: chromosome organization and evolution
in a disomic polyploid genome. Genetics 138:829–847
Rong J, Abbey C, Bowers JE, Brubaker CL, Chang C, Chee PW, Delmonte TA, Ding X, Garza JJ,
Marler BS, Park C, Pierce GJ, Rainey KM, Rastogi VK, Schulze SR, Trolinder NL, Wendel JF,
Wilkins TA, Williams-Coplin TD, Wing RA, Wright RJ, Zhao X, Zhu L, Paterson AH (2004)
A 3347-locus genetic recombination map of sequence-tagged sites reveals features of genome
organization, transmission and evolution of cotton (Gossypium). Genetics 166:389–417
Rong J, Feltus EA, Waghmare VN, Pierce GJ, Chee PW, Draye X, Saranga Y, Wright RJ,
Wilkins TA, May OL, Smith CW, Gannaway JR, Wendel JR, Paterson AH (2007) Metaanalysis of polyploid cotton QTL shows unequal contributions of subgenomes to a complex
network of genes and gene clusters implicated in lint fiber development. Genetics
176(4):2577–2588
Rong J, Feltus FA, Liu L, Lin L, Paterson AH (2010) Gene copy number evolution during
tetraploid cotton radiation. Heredity 105(5):463–472
Salmon A, Flagel L, Ying B, Udall JA, Wendel JF (2010) Homoeologous nonreciprocal
recombination in polyploid cotton. New Phytol 186:123–134
Saunders JH (1961) The wild species of Gossypium and their evolutionary history. Oxford
University Press, London
Seelanan T, Brubaker CL, Stewart JM, Craven LA, Wendel JF (1999) Molecular systematics of
Australian Gossypium section grandicalyx (Malvaceae). Syst Bot 24:183–208
Senchina DS, Alvarez I, Cronn RC, Liu B, Rong JK, Noyes RD, Paterson AH, Wing RA, Wilkins TA,
Wendel JF (2003) Rate variation among nuclear genes and the age of polyploidy in Gossypium.
Mol Biol Evol 20:633–643
Shan X, Liu Z, Dong Z, Wang Y, Chen Y, Lin X, Long L, Han F, Dong Y, Liu B (2005)
Mobilization of the active MITE transposons mPing and Pong in rice by introgression from
wild rice (Zizania latifolia Griseb.). Mol Biol Evol 22:976–990
Small RL, Ryburn JA, Wendel JF (1999) Low levels of nucleotide diversity at homoeologous Adh
loci in allotetraploid cotton (Gossypium L.). Mol Biol Evol 16:491–501
Small RL, Wendel JF (1999) The mitochondrial genome of allotetraploid cotton (Gossypium L.).
J Hered 90:251–253
Small RL, Wendel JF (2002) Differential evolutionary dynamics of duplicated paralogous Adh
loci in allotetraploid cotton (Gossypium). Mol Biol Evol 19:597–607
Soltis PS, Soltis DE (2000) The role of genetic and genomic attributes in the success of
polyploids. Proc Nat Acad Sci USA 97:7051–7057
CO
RR
921
922
923
924
925
926
927
928
929
930
931
932
933
934
935
936
937
938
939
940
941
942
943
944
945
946
947
948
949
950
951
952
953
954
955
956
957
958
959
960
961
962
963
964
965
966
967
968
969
970
971
972
973
974
J. F. Wendel et al.
UN
Editor Proof
206
Layout: T1 Standard SC
Chapter No.: 10
207
EC
TE
D
PR
OO
F
Stebbins GL (1947) Types of polyploids: their classification and significance. Adv Genet 1:
403–429
Stebbins GL (1950) Variation and evolution in plants. Columbia University Press, New York
Stephens SG (1958) Salt water tolerance of seeds of Gossypium species as a possible factor in
seed dispersal. Amer Nat 92:83–92
Stephens SG (1966) The potential for long range oceanic dispersal of cotton seeds. Amer Nat
100:199–210
Stewart JM (1995) Potential for crop improvement with exotic germplasm and genetic
engineering. In: Constable GA, Forrester NW (eds) Challenging the future: proceedings of the
world cotton research, CSIRO, Melbourne, pp 313–327
Stewart JM, Oosterhuis D, Heithholt JJ, Mauney JR (2010) Physiology of cotton. Springer, The
Netherlands
Udall JA, Swanson JM, Nettleton D, Percifield RJ, Wendel JF (2006) A novel approach for
characterizing expression levels of genes duplicated by polyploidy. Genetics 173(3):
1823–1827
Ungerer MC, Strakosh SC, Zhen Y (2006) Genome expansion in three hybrid sunflower species is
associated with retrotransposon proliferation. Curr Biol 16R:872–873
Veitia RA (2005) Gene dosage balance: deletions, duplications and dominance. Trends Genet
21:33
Wang JL, Tian L, Lee HS, Wei NE, Jiang HM, Watson B, Madlung A, Osborn TC, Doerge RW,
Comai L, Chen ZJ (2006) Genomewide nonadditive gene regulation in Arabidopsis
allotetraploids. Genetics 172:507–517
Watt G (1907) The wild and cultivated cotton plants of the world. Longmans, Green and Co,
London
Wendel JF (1989) New world tetraploid cottons contain old world cytoplasm. Proc Nat Acad Sci
USA 86:4132–4136
Wendel JF (2000) Genome evolution in polyploids. Plant Mol Biol 42:225–249
Wendel JF, Brubaker CL, Alvarez JP, Cronn RC, Stewart JM (2009) Evolution and natural
history of the cotton genus. In: Paterson AH (ed) Genomics of cotton, plant genetics and
genomics, crops and models 3. Springer, New York, pp 3–22
Wendel JF, Cronn RC (2003) Polyploidy and the evolutionary history of cotton. Adv Agron
78:139–186
Wendel JF, Rowley R, Stewart JM (1994) Genetic diversity in and phylogenetic relationships of
the Brazilian endemic cotton, Gossypium mustelinum (Malvaceae). Pl Syst Evol 192:49–59
Wendel JF, Schnabel A, Seelanan T (1995) Bidirectional interlocus concerted evolution
following allopolyploid speciation in cotton (Gossypium). Proc Nat Acad Sci USA
92:280–284
Wright RJ, Thaxton PM, El-Zik KM, Paterson AH (1998) D-subgenome bias of Xcm resistance
genes in tetraploid Gossypium (cotton) suggests that polyploid formation has created novel
avenues for evolution. Genetics 149:1987–1996
Xu Z, Yu JZ, Cho J, Yu J, Kohel RJ, Percy RG (2010) Polyploidization altered gene functions in
cotton (Gossypium spp.). PLoS ONE 5:e14351
Yang SS, Cheung F, Lee JJ, Ha M, Wei NE, Sze SH, Stelly DM, Thaxton P, Triplett B, Town CD,
Chen ZJ (2006) Accumulation of genome-specific transcripts, transcription factors and
phytohormonal regulators during early stages of fiber cell development in allotetraploid cotton.
Plant J 47:761–775
CO
RR
975
976
977
978
979
980
981
982
983
984
985
986
987
988
989
990
991
992
993
994
995
996
997
998
999
1000
1001
1002
1003
1004
1005
1006
1007
1008
1009
1010
1011
1012
1013
1014
1015
1016
1017
1018
1019
1020
Book ISBN: 978-3-642-31441-4
Page: 207/206
Jeans, Genes, and Genomes: Cotton as a Model for Studying Polyploidy
UN
Editor Proof
10
Book ID: 272454_1_En
Date: 16-8-2012
Metadata of the chapter that will be visualized in
SpringerLink
Book Title
Polyploidy and Genome Evolution
Series Title
Chapter Title
Evolutionary Implications of Genome and Karyotype Restructuring in Nicotiana tabacum L
Copyright Year
2012
Copyright HolderName
Springer-Verlag Berlin Heidelberg
Corresponding Author
Family Name
Leitch
Particle
Given Name
Andrew
Suffix
Author
Division
School of Biological and Chemical Sciences
Organization
Queen Mary University of London
Address
London, UK
Email
a.r.leitch@qmul.ac.uk
Family Name
Kovarik
Particle
Given Name
Ales
Suffix
Division
Organization
Institute of Biophysics
Address
Brno, Czech Republic
Email
Author
Family Name
Renny-Byfield
Particle
Given Name
Simon
Suffix
Division
School of Biological and Chemical Sciences
Organization
Queen Mary University of London
Address
London, UK
Email
Author
Family Name
Grandbastien
Particle
Given Name
Marie-Angèle
Suffix
Division
Organization
Institute Jean-Pierre Bourgin
Address
Versailles, France
Email
Abstract
Nicotiana tabacum is an allopolyploid that formed within the last 200,000 years from relatives of the extant
diploids N. sylvestris and N. tomentosiformis, the donors of the S- and T-genomes, respectively. Here we
review progress in our understanding of the divergence of N. tabacum subsequent to its formation, by
comparing the N. tabacum genome with those of its diploid progenitors. We also review the data from
synthetic N. tabacum, where there is evidence for much genetic change in early generations, including various
chromosomal translocations, allopolyploid-induced retroelement mobility and loss, and reductions in the copy
numbers of some tandem repeats. These observations are similar to patterns found in natural N. tabacum,
suggesting that rapid genetic divergence is induced by allopolyploidy. The T-genome of N. tabacum shows
the greatest number of genetic changes and appears to be less stable than the S-genome. We describe possible
mechanisms that may have stimulated these genetic changes and propose that these can lead to enhanced
fertility, more regular chromosome pairing, and the evolution of disomic inheritance.
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
F
PR
OO
5
Ales Kovarik, Simon Renny-Byfield, Marie-Angèle Grandbastien
and Andrew Leitch
Abstract Nicotiana tabacum is an allopolyploid that formed within the last
200,000 years from relatives of the extant diploids N. sylvestris and N. tomentosiformis, the donors of the S- and T-genomes, respectively. Here we review progress in our understanding of the divergence of N. tabacum subsequent to its
formation, by comparing the N. tabacum genome with those of its diploid progenitors. We also review the data from synthetic N. tabacum, where there is
evidence for much genetic change in early generations, including various chromosomal translocations, allopolyploid-induced retroelement mobility and loss, and
reductions in the copy numbers of some tandem repeats. These observations are
similar to patterns found in natural N. tabacum, suggesting that rapid genetic
divergence is induced by allopolyploidy. The T-genome of N. tabacum shows the
greatest number of genetic changes and appears to be less stable than the
S-genome. We describe possible mechanisms that may have stimulated these
genetic changes and propose that these can lead to enhanced fertility, more regular
chromosome pairing, and the evolution of disomic inheritance.
D
4
Evolutionary Implications of Genome
and Karyotype Restructuring in Nicotiana
tabacum L.
TE
3
Chapter 11
EC
2
Book ISBN: 978-3-642-31441-4
Page: 209/224
CO
RR
1
Book ID: 272454_1_En
Date: 16-8-2012
A. Kovarik
Institute of Biophysics, Brno, Czech Republic
S. Renny-Byfield A. Leitch (&)
School of Biological and Chemical Sciences, Queen Mary University of London,
London, UK
e-mail: a.r.leitch@qmul.ac.uk
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: 11
M.-A. Grandbastien
Institute Jean-Pierre Bourgin, Versailles, France
P. S. Soltis and D. E. Soltis (eds.), Polyploidy and Genome Evolution,
DOI: 10.1007/978-3-642-31442-1_11, Springer-Verlag Berlin Heidelberg 2012
209
Layout: T1 Standard SC
Chapter No.: 11
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 210/224
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
61
62
63
64
65
F
PR
OO
28
D
26
27
The genus Nicotiana is the sixth largest genus in the family Solanaceae and
includes 76 species found naturally in the Americas, Australia and surrounding
islands, and with a single species in Namibia (Goodspeed 1954; Knapp et al. 2004).
However, several species, including N. tabacum (tobacco), now occur more widely.
The genus has received much recent attention because of evidence for recurrent
polyploidy, resulting in nearly half the species in the genus being chromosomally
polyploid. It is likely there have been six independent polyploidy events within the
genus: at about 10 million years ago (mya) (section Suaveolentes, c. 24 species), 5
mya (section Repandae, 4 species), 1–2 mya (section Polydicleae, 2 species), and
within the last 200,000 years (N. rustica, N. arentsii, and N. tabacum) (Lim et al.
2004a). There is also reticulation at the diploid level, with at least three species
likely to be of homoploid hybrid origin (N. glauca, N. linearis, and N. spegazzinii)
(Goodspeed 1954; Kelly et al. 2010).
N. tabacum is an autotetraploid (2n = 4x = 48) derived from diploid species that
most closely resemble N. sylvestris (2n = 2x = 24), the maternal S-genome donor,
and N. tomentosiformis (2n = 2x = 24, section Tomentosae), the paternal T-genome
donor. Nicotiana sylvestris is the only species in section Sylvestres, and detailed
genetic analysis of multiple accessions indicates that there is little genetic diversity in
this species (4.2 % polymorphisms in AFLPs—Amplified Fragment Length
Polymorphisms, Petit et al. 2007). However, there is considerably more genetic
diversity among accessions of N. tomentosiformis (31.7 % AFLP polymorphism),
which form two distinct groups, one of which most closely resembles the T-genome of
N. tabacum (Murad et al. 2002; Petit et al. 2007). These data indicate that N. tabacum
formed subsequent to the divergence of the two groups of N. tomentosiformis.
Molecular clock estimates of plastid and internally transcribed spacer sequences (ITS) of nuclear ribosomal DNA (rDNA) suggest that N. tabacum formed less
than 200,000 years ago. These estimates were calibrated using likely maximum
ages of some endemic species occurring on islands of known geological ages
(Clarkson et al. 2005; Leitch et al. 2008). However, given that only feral populations of N. tabacum have been found and no truly ‘‘wild’’ population exists
(Knapp personal communication), it is quite possible that N. tabacum was born
subsequent to the origin of human agriculture, within the last 10,000 years.
The purpose of this paper is to review current understanding of the genetic
consequences of polyploidy in N. tabacum. We show that genetic change can occur
rapidly within a few generations, perhaps as a consequence of the ‘‘genomic shock’’
of allopolyploidy (McClintock 1984), a process that generates variants from which
selection favors those with enhanced fertility. This unstable phase is presumably
transient; however, there remains evidence for genome dynamism over time scales
of thousands of years, including the replacement of multiple copies of rDNA
sequences, the turnover of retroelements, and the loss of sequences targeted at the
T-genome. We discuss how such changes may promote fertility and lead to the
fixation of rearranged karyotypes during polyploid species establishment.
TE
25
EC
24
11.1 Introduction to Nicotiana
CO
RR
23
A. Kovarik et al.
UN
Editor Proof
210
Layout: T1 Standard SC
Chapter No.: 11
72
73
74
75
76
77
78
79
80
81
82
83
84
85
86
87
88
89
90
91
92
93
94
95
96
97
98
99
100
101
102
103
104
105
106
107
108
F
PR
OO
71
D
69
70
Kenton et al. (1993) were the first to use genomic in situ hybridization (GISH) to
study the chromosomes of N. tabacum. The fluorescent probes derived from total
genomic DNA of N. sylvestris and N. tomentosiformis hybridized to a separate
subset of 24 chromosomes, corresponding to the chromosomes of the S- and
T-genomes, respectively. However, the efficacy of the T-genome labeling was
inferior (Lim et al. 2000b), likely because of a loss of T-genome sequences
(Renny-Byfield et al. 2011). GISH also revealed in all N. tabacum accessions 4–9
intergenomic translocations (Kenton et al. 1993; Lim et al. 2004a; Moscone et al.
1996). Similar translocations (up to 3) were also observed in some synthetic
N. tabacum lines that were only a few generations old (Skalicka et al. 2005).
Perhaps these translocations have arisen as a consequence of multivalent formation
(see Chap. 7), or they may represent hotspots of recombination. While some
translocations are fixed (i.e. occur in all varieties), others are specific for particular
accessions. There are more translocations of S-genome origin chromatin to
T-genome chromosomes (T/s chromosome) than the reverse (S/t chromosome).
Kenton et al. (1993) hypothesized that this could be caused by selection against
S/t chromosomes. Alternatively, some of the ‘‘translocations’’ may actually be
S-genome subtelomeric satellite sequences that now occur on T-genome chromosomes, perhaps arising in their new location via recombination-based homogenization processes (Koukalova et al. 2010). Next-generation sequencing (NGS)
may enable us to distinguish among these alternative hypotheses.
As with most other plant species (Heslop-Harrison and Schwarzacher 2011), a
large fraction of Nicotiana genomes is composed of various types of retroelements. Genome sampling using NGS revealed that the major component of the
genome of N. tabacum and its diploid relatives comprises long terminal repeat
(LTR)-retrotransposons, with at least 17.1–22.5 % of the genome being Ty3/gypsy
elements and 2.2–3.4 % being Ty1/copia elements. DNA transposons comprise
around 1.7 % of the genome of these species (Renny-Byfield et al. 2011).
In addition, there are several unrelated satellite repeat families that have been
characterized (the distributions of some are shown Fig. 11.1, Lim et al. 2000b). These
repeats include: (1) The HRS60 family, which is the best-characterized tandem repeat
family in Nicotiana and includes an interstitial repeat in N. tomentosiformis and
N. tabacum (called GRS, Gazdova et al. 1995) and predominantly subtelomeric repeats
in N. sylvestris and N. tabacum (called HRS60 and NSYL2, Koukalova et al. 2010;
Koukalova et al. 1989); (2) NTS9 in N. sylvestris and N. tabacum (Jakowitsch et al.
1998); (3) NTRS in N. tomentosiformis and N. tabacum (Fig. 11.1, Matyasek et al.
1997), and (4) NicCL3, a long 2.2 kb tandem repeat comprising *2 % of the
N. tomentosiformis genome and in lower abundance in N. tabacum (Fig. 11.2,
Renny-Byfield et al. 2011 and Renny-Byfield et al. 2012). There are also a number of
satellite repeat families of known origin; these are: (5) A1/A2 satellite repeats derived
from the intergenic spacer (IGS) of rDNA, which have transposed and amplified to
multiple locations across the genome (Lim et al. 2004b); (6) tandem repeats of
TE
68
EC
67
211
11.2 Nicotiana tabacum Genome Structure
CO
RR
66
Book ISBN: 978-3-642-31441-4
Page: 211/224
Evolutionary Implications of Genome
UN
Editor Proof
11
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 11
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 212/224
A. Kovarik et al.
TE
D
PR
OO
F
Editor Proof
212
111
112
113
114
115
116
117
118
119
120
geminivirus-related DNA (GRD), occurring in two distinct families in some accessions of N. tomentosiformis and inherited to N. tabacum (Murad et al. 2004; Murad
et al. 2002), the first endogenous viruses discovered in plants (Bejarano et al. 1996); (7)
sequences of pararetroviral origin (Matzke et al. 2004), including distinct variants
found in N. tomentosiformis (NtoEPRV) and N. sylvestris (NsEPRV), both of which
have been inherited in N. tabacum.
CO
RR
110
11.3 Retroelement Response to Allopolyploid ‘‘Genomic
Shock’’
UN
109
EC
Fig. 11.1 Ideograms showing the distribution of tandem repeat sequences in N. tabacum and its
diploid progenitors, N. sylvestris and N. tomentosiformis. Taken with permission from Murad
et al. (2002). See text for a description of the repeats shown
Nicotiana hybrids were among the first in which chromosomal changes following
hybridization were demonstrated. Gerstel and Burns (1967) reported that
N. otophora 9N. tabacum hybrids have genetic instabilities of two kinds. Firstly, the
heterochromatin from N. otophora undergoes breakage causing chromatin loss; this
Layout: T1 Standard SC
Chapter No.: 11
Book ISBN: 978-3-642-31441-4
Page: 213/224
Evolutionary Implications of Genome
213
TE
D
PR
OO
F
Editor Proof
11
Book ID: 272454_1_En
Date: 16-8-2012
121
122
123
124
125
UN
CO
RR
EC
Fig. 11.2 Reduction in copy number of T-genome repeats in N. tabacum from expectation given
their abundance in N. tomentosiformis. a Fluorescence in situ hybridization (FISH) showing the
distribution of dispersed A1/A2 repeats (green fluorescence). The 18-5.8-26S rDNA locus on
chromosome 3 (N. tomentosiformis) and T3 (N. tabacum) is labeled with yellow fluorescence due
to the overlap of red and green rDNA fluorescence. Note that there is much reduced A1/A2 signal
in N. tabacum (for further information see Lim et al. 2004b). b FISH showing the distribution of
NicCL3 in both species (green fluorescence). Note the reduced abundance of this repeat in
N. tabacum. The 18-5.8-26S rDNA locus on chromosome 3 (N. tomentosiformis) and T3
(N. tabacum) is labeled with red fluorescence. For further information see Renny-Byfield et al.
(2012). c, d. NGS analysis of repeat clusters in N. tabacum and its diploid progenitors. The graphs
show the genome proportions (arcsine transformed) of repeats uniquely inherited in N. tabacum
from c N. tomentosiformis and d N. sylvestris. If N. tabacum had faithfully inherited the repeats as
found in the diploids, then all repeats would fall on the blue lines. However, in N. tomentosiformis
c most repeats at higher genome proportions are underrepresented in N. tabacum. This is not the
case for N. sylvestris d, where repeats are more or less in their expected abundance. This trend is
reflected in the abundance of repeats that are biparentally inherited, i.e., the more abundant they
are in N. tomentosiformis, the more likely they will be underrepresented in N. tabacum. For
further information see Renny-Byfield et al. (2011)
can be observed cytologically and also phenotypically by the leaf variegation it
causes. Secondly, in some cells heterochromatic blocks from N. otophora proliferate
enormously to many times their normal length, forming ‘‘megachromosomes’’. This
pioneering work revealed that interspecific hybridization can induce chromosomal
rearrangements and rapid sequence losses and gains.
Layout: T1 Standard SC
Chapter No.: 11
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 214/224
A. Kovarik et al.
157
11.4 Loss of DNA: Targeting the T-Genome
133
134
135
136
137
138
139
140
141
142
143
144
145
146
147
148
149
150
151
152
153
154
155
158
159
160
161
162
163
164
165
PR
OO
132
D
131
TE
130
EC
128
129
CO
RR
127
F
156
Melayah et al. (2001) were the first to show, using N. tabacum, that environmental stresses can induce retrotransposition. There is also evidence that allopolyploidy can stimulate transposition in N. tabacum and synthetic mimics of
N. tabacum made from the diploid progenitors (Petit et al. 2007, 2010). These
authors analyzed four populations of Ty1/copia LTR-retrotransposons (Tnt1-ol16,
Tnt1-ol13, Tnt2, and Tto1) using sequence-specific amplification polymorphism
(SSAP). They showed that the parental diploid species share essentially similar
classes of retroelements. In N. tabacum there is evidence of retrotransposon
diversification subsequent to allopolyploidy, with sequence losses concomitant
with gains. Losses of retroelements were more frequent to the T-genome, while
novel insertions of some populations, such as Tnt2, were shown to preferentially
locate to the S-genome. However, each retrotransposon population seems to
behave differently, with some populations undergoing rapid turnover, while others
display relative stasis. There was little or no diversification of retrotransposons
among individual accessions of natural N. tabacum, indicating that retroelement
diversification occurred before the accessions analyzed had diverged, perhaps
early in the species’evolution and arising as a consequence of allopolyploidy and
induced ‘‘genomic shock’’ (McClintock 1984).
To test the hypothesis that allopolyploidy can induce retroelement diversification,
Petit et al. (2010) compared insertion patterns of the Tnt1 family in the synthetic
N. tabacum Th37 (Burk 1973) with its diploid progenitors. In some Th37 individuals
in the fourth synthetic generation (S4), there was evidence of new Tnt1 insertions.
Newly, transposed copies were amplified from elements located on the N. sylvestrisderived genome and were highly similar to the Tnt1A tobacco copies amplified in
response to microbial factors (Grandbastien et al. 2005). Furthermore, a high proportion of parental SSAP bands was lost in Th37, particularly those from the
N. tomentosiformis-derived genome, again as observed in natural N. tabacum.
Together, these data indicate that retrotransposon amplification and molecular
restructuring in, or around, insertion sites occur rapidly in response to allopolyploidy,
and that similar sequences have responded in the same way to allopolyploidy in
natural and synthetic N. tabacum.
126
As with many other allopolyploid species (Leitch and Bennett 2004), N. tabacum has
a lower genome size than is expected from the sum of parental genomes (Leitch et al.
2008, N. tabacum genome size, 1C = 5110 Mpb; N. sylvestris, 1C = 2636 Mbp;
N. tomentosiformis, 1C = 2683 Mbp (http://data.kew.org/cvalues/). Recently,
(Renny-Byfield et al. 2011) used NGS analysis of N. tabacum and its two diploid
progenitors and showed that repeats derived from N. tomentosiformis was underrepresented in N. tabacum, a trend that was not observed for repeats from N. sylvestris
(Fig. 11.2). These observations lead to the conclusion that the T-genome of
UN
Editor Proof
214
Layout: T1 Standard SC
Chapter No.: 11
Book ISBN: 978-3-642-31441-4
Page: 215/224
Evolutionary Implications of Genome
215
189
11.5 Ribosomal DNA Homogenization is Rapid and Ongoing
174
175
176
177
178
179
180
181
182
183
184
185
186
187
190
191
192
193
194
195
196
197
198
199
200
201
202
203
204
205
206
PR
OO
173
D
171
172
TE
169
170
EC
168
In most eukaryotes, 5S and 18–5.8–26S rDNA units occur in tandem arrays at one
or several loci. Each large rDNA unit contains the 18S, 5.8S, and 26S ribosomal
RNA (rRNA) genes, ITS, and IGS sequences. Whilst these sequences are vital for
cell functioning, they can also be highly recombinogenic and labile sequences
influencing genome stability (Kobayashi 2011). The genes themselves are highly
conserved; however, their spacers diverge at suitable rates for resolving species
relationships within most genera, including Nicotiana (Chase et al. 2003). Early
studies revealed that the restriction fragment length polymorphism (RFLP) patterns
of the N. tabacum IGS is not additive of the diploid progenitors (Kovarik et al.
1996), but that N. tabacum has evolved its own distinct rRNA gene family(ies).
Sequence analysis revealed that the tobacco-specific units arose by reorganization
of N. tomentosiformis-inherited units followed by their subsequent amplification
(Volkov et al. 1999). The sequence changes mainly involved amplification and
reduction of subrepeats upstream and downstream of the transcription start site
within the IGS. However, the newly evolved units still occur at the four rDNA loci
that N. tabacum inherited from its parents (see Fig. 11.1). Thus, it is likely that
the parental units of S-genome origin were overwritten by the newly amplified
CO
RR
167
F
188
N. tabacum has undergone extensive sequence losses, and that genetic changes have
occurred more rapidly to the paternally derived T-genome than the S-genome.
N. tabacum harbors two families of endogenous pararetrovirus DNA, one from
each progenitor (Jakowitsch et al. 1999). These are NsEPRV from N. sylvestris,
which in N. tabacum is at the expected abundance, and NtoEPRV from
N. tomentosiformis, which occurs in N. tabacum with reduced copy number.
Likewise, the A1/A2 repeats (Lim et al. 2004b) and NicCL3 (Renny-Byfield et al.
2012), which are found in multiple locations around the genome in N. tomentosiformis (Fig. 11.2), are also underrepresented in N. tabacum. Similarly, NGS data
show that all families of Ty3/gypsy elements have undergone sequence loss in
N. tabacum (Renny-Byfield et al. 2011), and SSAP analysis of Tnt retroelements
(Ty1/copia) reveals more deletion events influencing loci from the T-genome than
the S-genome of N. tabacum (Petit et al. 2007). There is evidence of rapid and
targetted loss of repeats from the T-genome even from the earliest generations,
since in synthetic N. tabacum lines, many individuals already have reduced copy
numbers of NtoEPRV, NTRS repeats (Skalicka et al. 2005), and NicCL3
(Renny-Byfield et al. 2012), whilst most losses of Tnt1 retroelements were from
the T-genome (Petit et al. 2010). All these data suggest rapid, targetted changes
early in allopolyploid evolution. The targeting of repeat losses from the T-genome
may support the nuclear-cytoplasmic interaction hypothesis, whereby the paternally derived genome functions in a maternal cytoplasm (Gill 1991; Leitch et al.
2006), which in allopolyploids may lead to incompatibilities that preferentially
destabilize the paternally derived genome.
166
UN
Editor Proof
11
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 11
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 216/224
A. Kovarik et al.
211
N.tabacum-specific units (Lim et al. 2000a), although a few units of S genomeorigin remain intact, perhaps because they are methylated and inactive (Kovarik
et al. 2008). In contrast, the 5S rDNA follows a different evolutionary pattern in
N. tabacum, with no evidence for intergenomic homogenization, although shifts in
copy numbers of parental gene families have been observed (Fulnecek et al. 2002).
212
11.6 The Fate of Duplicated Genes
207
208
209
218
219
220
221
222
223
224
225
226
227
228
229
230
231
232
233
234
235
236
237
238
239
240
241
242
243
244
245
246
247
D
216
217
TE
215
Although there has been substantial restructuring of the repetitive fraction of the
N. tabacum genome since its formation, most genic sequences analyzed have
remained in duplicate copies, e.g., genes for drug resistance (Schenke et al. 2003),
putrescine N-methytransferases (Riechers and Timko 1999), a family of small
GTP-binding proteins (Takumi et al. 2002), lignin forming peroxidase (Matassi
et al. 1991), nitrite reductase (Kronenberger et al. 1993), nitrate reductase
(Vaucheret et al. 1989), glutamine synthetase (Clarkson et al. 2010), phytochrome
A (Intrieri et al. 2008), the developmental gene LEAFY (McCarthy 2010), and
families of DNA methyltransferases (Fulnecek et al. 2009). The only current
exception is a family of N. tabacum glucan endo-1,3-beta-glucosidase genes that
appeared to be recombinants of both ancestral sequences (Sperisen et al. 1991).
Locus additivity does not always seem to be a rule even in recently formed
synthetic allopolyploids. For example, frequent deletions of homoeologs were
found among 70 protein-coding loci in Tragopogon miscellus that formed within
the last 80 years (Buggs et al. 2009, 2012; see also Chap. 14, this volume). These
changes may not be random, as Buggs et al. (2012) showed that clusters of genes
are repeatedly lost or retained and the likelihood of retention reflects gene
ontology categories or their predicted levels of dosage sensitivity. Similarly,
synthetic lines of Brassica napus seem to eliminate much of the parental DNA,
although it is currently unknown whether coding or noncoding sequences are
preferentially targeted (Song et al. 1995; Szadkowski et al. 2011). The occurrence
of both homoeologs in N. tabacum may reflect limited sampling, or high levels of
retention. Additivity is observed in synthetic lines of Gossypium (Liu et al. 2002;
see also Chap. 10, this volume) and in synthetic wheat allopolyploids, although in
the latter case there remains some controversy (Mestiri et al. 2010; Feldman and
Levy 2009; see also Chap. 7, this volume). Additivity has also been demonstrated
at the expression level. Alleles of phytochrome A, lignin forming peroxidases,
nitrite reductase, and DNA methyltransferases are expressed from both parental
homeologs. There are also reports of epigenetic silencing of one subset of the
parental alleles, including rRNA genes that have escaped homogenization
(Kovarik et al. 2008) and the CYP82E4 locus (Chakrabarti et al. 2007) involved in
the alkaloid biosynthesis pathway.
Duplicate copies arising through polyploidy can be retained through a number
of mechanisms (Doyle et al. 2008): (1) one copy can evolve a new function
(neofunctionalization) that can become fixed through a selective advantage
EC
214
CO
RR
213
PR
OO
F
210
UN
Editor Proof
216
Layout: T1 Standard SC
Chapter No.: 11
Book ISBN: 978-3-642-31441-4
Page: 217/224
Evolutionary Implications of Genome
217
259
11.7 Genome Revolution and Allopolyploid Establishment
255
256
257
260
261
262
263
264
265
266
267
268
269
270
271
272
273
274
275
276
277
278
279
280
281
282
283
284
285
286
287
288
PR
OO
254
Until the early 1990s, it was commonly thought that chromosomal changes
establish isolation barriers among populations because of reduced fertility in
heterozygotes. Rearranged karyotypes may, then, become fixed through inbreeding
or meiotic drive, particularly in small populations (King 1993). However, it is
difficult to distinguish between karyotype changes that drive speciation events and
those arising by genetic drift after speciation. Certainly, many rearrangements are
deleterious and can cause embryo lethality or hybrid breakdown prior to reproduction (Rieseberg 1997). Only neutral or advantageous recombinants will pass
the bottleneck of selection during the early phase of speciation. Furthermore,
evolutionary geneticists pointed out that genetic mutations at the DNA level are
more common than karyotype change and are therefore likely to be more
important in speciation processes (Butlin 1993). However, recent evidence from a
number of sources, e.g., Anopheles gambiae (Turner and Hahn 2010) and
Helianthus homoploid hybrids (Strasburg et al. 2009), indicates that chromosomal
inversions can, indeed, have a role in establishing barriers between populations
and drive speciation. Strasburg et al. (2009) suggested that inversions may become
foci for the accumulation of adaptive genes, particularly at the junctions between
collinear and inverted regions, where gene flow is likely to be most impeded.
Likewise, computer modeling has shown that when recombination is eliminated at
an inverted region, and in the presence of strong selection and minimal or no gene
flow, then species isolation can be driven by such a rearrangement (Feder and
Nosil 2009).
In the context of early polyploid evolution, karyotype rearrangement may drive
enhanced fertility and potentially be fundamental to allopolyploid species establishment. Certainly, high levels of chromosomal change have been observed in
young polyploids, e.g., karyotype variability (dysploidy and intergenomic translocations) within and among populations of Tragopogon polyploids that formed
within the last 80 years (Chester et al. 2010, 2012; Lim et al. 2008), and largescale chromosomal deletions occurring in individual plants of early generation
D
253
TE
252
EC
251
CO
RR
250
F
258
(Lynch et al. 2001); (2) duplicate copies can diverge through complementary loss
of function at a particular point in the development or in particular tissues (subfunctionalization), a process that can occur through the action of drift alone; and
(3) selection can occur against gene loss because that loss would compromise
appropriate levels of gene product in relation to another gene(s) (gene balance
hypothesis, cf. Birchler and Veitia 2010). In N. tabacum, a mechanism termed
‘‘nonfunctionalization’’ has been proposed which considers a combination of
degenerative mutation and epigenetic silencing (Chakrabarti et al. 2007). Here,
‘‘non-functionalization’’ has altered alkaloid metabolism through reduced conversion of nicotine to nornicotine. Perhaps this mutation has been important to the
‘‘success’’ of N. tabacum as a recreational drug.
248
249
UN
Editor Proof
11
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 11
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 218/224
296
297
298
299
300
301
302
303
304
305
306
307
308
309
310
311
312
313
314
315
316
317
318
319
320
321
322
323
324
325
326
327
328
329
330
331
332
F
295
• Reduced chiasma frequency. Selection may favor the formation of fewer and/or
focussed chiasma at recombination hotspots. This would result in quadrivalents
formed in prophase I falling into two bivalents in metaphase I and then segregating normally (Fig. 11.3a). Computer modeling has shown that reduced
chiasma frequency results in the evolution of disomic inheritance (Le Comber
et al. 2010), although we are unaware of empirical evidence to support this
assertion.
• Structural and epigenetic divergence of homeologues. Allopolyploidy can
trigger extensive karyotype and molecular restructuring in early generations
(Gaeta et al. 2007; Renny-Byfield et al. 2011). This may result in genetic
divergence of the parental genomes, e.g., through the loss or rearrangement of
sequences shared by homeologues and recognized by the homolog recognition
machinery in meiosis (Fig. 11.3b). Such sequences are unlikely to be those that
are highly repeated across the genome since they are not chromosome specific.
Nevertheless, genetic changes may introduce regions among homeologues that
do not recombine, forming ‘‘islands of divergence’’, as in the establishment of
homoploid hybrid species (Strasburg et al. 2009). Without recombination, the
local regions around these islands will continue to diverge, leading to regular
homolog pairing, and bivalent formation. Similarly, allopolyploidy can trigger
epigenetic changes across the genome (Parisod et al. 2009). Epigenetic changes,
which are thought to alter patterns of ectopic recombination (Colot et al. 1995),
may also influence patterns of meiotic recombination and initiate islands of
divergence. In addition, newly formed epi-alleles may establish tissue or temporal patterns of gene expression (subfunctionalization, Adams and Wendel
2005). Subfunctionalization is likely to favor the evolution of regular disomic
inheritance and to select against multivalent formation. This is because multivalents result in the inappropriate segregation of homeoalleles and the
generation of individuals with reduced fitness, because they have aberrant
patterns of gene expression (Le Comber et al. 2010).
• Intergenomic translocations. In the context of a diploid, translocations among
heterologues can promote quadrivalent formation, leading to chromosome loss,
and reduced fertility. Similarly, in the context of polyploids, translocations and
aneuploidy are associated with low fertility, as observed in synthetic B. napus
lines (Xiong et al. 2011). However, it is possible that a more complex dynamic
can occur (Fig. 11.3c). If homeologues already form multivalents, then it is
conceivable that the gain of genetic material from a heterologous chromosome
may promote bivalent formation, disomic inheritance, and increased fertility.
Alternatively, complex rearrangements can result through cascades of induced
PR
OO
294
D
293
TE
292
EC
291
synthetic allopolyploids of B. napus (Gaeta et al. 2007). Typically, a major hurdle
that a newly formed polyploid must overcome is reduced fertility, often arising
through multivalent formation, where chromosomes can segregate aberrantly
leading to aneuploidy and reduction in fertility. We can envisage several allopolyploid-induced processes that may act to reduce multivalent formation and be
favored by selection, these are:
CO
RR
289
290
A. Kovarik et al.
UN
Editor Proof
218
Layout: T1 Standard SC
Chapter No.: 11
Editor Proof
11
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 219/224
Evolutionary Implications of Genome
219
(a) Reduced chiasma frequency
X
Focused / fewer chiasma may
result in elevated bivalent
formation -enhancing fertility
X
F
X
X
PR
OO
(b) Structural divergence of homologues
X
X
X
X
X
X
X
X
(c) Intergenomic translocations
X
X
X
X
X
X
X
X
X
X
Rapid homeologue
divergence may
drive bivalent
formation,
enhancing fertility
Promotes
homologue pairing,
disomic inheritance
and homologue
divergence
D
X
TE
X
X
X
X
X
X
EC
X
Homeologous translocations
promote multivalent
formation, generating further
translocations. Meiotic
abnormalities stimulate
unequal reciprocal
translocations, inversions,
end-to-end chromosome
fusions, resulting in dysploidy
.
334
335
336
337
338
340
339
341
342
343
344
pairing problems resulting from recurrent multivalent formation (unequal reciprocal translocations, inversions, end-to-end chromosome fusions), potentially
generating novel chromosomes with segments from multiple ancestral chromosomes. Such a process may be associated with a reduction in chromosome
number to a diploid-like level, as reported in the divergence of Arabidopsis
karyotypes (Mandakova et al. 2010).
UN
333
CO
RR
Fig. 11. 3 Fast molecular and chromosome evolution may enhance bivalent formation, driving
disomic inheritance, and fertility in young polyploids. Homologs are shown in the same color
(pink or pale green). Homeologues diverge over time, this is reflected in their colors becoming
more distinct (red and green). A translocation from a heterologue is shown in blue. Crossovers
among chromosomes are shown with crosses
From Nicotiana allopolyploids, we have evidence for several of the proposed
modes of chromosome evolution. First, in synthetic N. tabacum, we have observed
plants that were homozygous for an intergenomic translocation that is similar to
translocations observed in natural N. tabacum (Lim et al. 2004a). Perhaps this
translocation is selected because it enhances fertility, and that in the synthetic
Layout: T1 Standard SC
Chapter No.: 11
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 220/224
347
348
349
350
351
352
353
354
355
356
357
358
N. tabacum evolution is repeating itself. Second, the allotetraploids of sections
Polydicliae and Repandae, which formed c. 1 mya and 5 mya, respectively, have
chromosome numbers that are additive of the progenitor diploids (2n = 4x = 48).
However, the sequence organization along their chromosomes has diverged considerably, particularly in the latter. Indeed, Lim et al. (2007) suggested that in
some species of section Repandae there has been near complete genome turnover.
Perhaps these changes contributed to the establishment of disomic inheritance.
Finally, in section Suaveolentes, several species show evidence of chromosome
number reduction from expectation (chromosome numbers range from
2n = 4x = 32 to 46, depending on species, Goodspeed 1954; Knapp et al. 2004).
We anticipate that this reduction is associated with karyotype restructuring, as in
the polyploids of Arabidopsis (Mandakova et al. 2010).
F
346
PR
OO
345
A. Kovarik et al.
11.8 Advantages of the Nicotiana System and Future
Perspectives
369
Acknowledgments We thank NERC and the Czech Science Foundation for support.
370
References
371
372
373
374
375
376
377
378
379
380
Adams KL, Wendel JF (2005) Allele-specific, bidirectional silencing of an alcohol dehydrogenase gene in different organs of interspecific diploid cotton hybrids. Genetics 171:2139–2142
Bejarano ER, Khashoggi A, Witty M, Lichtenstein C (1996) Integration of multiple repeats of
geminiviral DNA into the nuclear genome of tobacco during evolution. Proc Nat Acad Sci
USA 93:759–764
Birchler JA, Veitia RA (2010) The gene balance hypothesis: implications for gene regulation,
quantitative traits and evolution. New Phytol 186:54–62
Buggs RJ, Doust AN, Tate JA, Koh J, Soltis K, Feltus FA, Paterson AH, Soltis PS, Soltis DE
(2009) Gene loss and silencing in Tragopogon miscellus (Asteraceae): comparison of natural
and synthetic allotetraploids. Heredity 103:73–81
364
365
366
367
TE
363
EC
361
362
CO
RR
360
D
368
Nicotiana provides many opportunities for studying allopolyploid genome divergence and addressing key questions facing polyploidy researchers [outlined in
Soltis et al. (2010)]. It is important, even vital for the future of humankind, to
know how allopolyploid genomes interact together. This is because many of our
most important crop species are recognizably polyploid based on their chromosome numbers, and recent evidence is emerging that all seed-bearing plants have
undergone at least one round of polyploidy in their ancestry (Jiao et al. 2011). The
allopolyploid species of Nicotiana and the synthetic allopolyploids provide a
unique opportunity to study snap shots of polyploid divergence over 10 million
years of evolution.
359
UN
Editor Proof
220
Layout: T1 Standard SC
Chapter No.: 11
221
EC
TE
D
PR
OO
F
Buggs RJA, Chamala S, Wu W, Tate JA, Schnable PS, Soltis DE, Soltis PS, Barbazuk WB (2012)
Rapid, repeated, and clustered loss of duplicate genes in allopolyploid plant populations of
independent origin. Curr Biol 22:248–252
Burk LG (1973) Partial self-fertility in a theoretical amphiploid progenitor of N. tabacum.
J Hered 64:348–350
Butlin RK (1993) Species evolution—the role of chromosome change—King, M. Nature 366:27
Chakrabarti M, Meekins KM, Gavilano LB, Siminszky B (2007) Inactivation of the cytochrome
P450 gene CYP82E2 by degenerative mutations was a key event in the evolution of the
alkaloid profile of modern tobacco. New Phytol 175:565–574
Chase MW, Knapp S, Cox AV, Clarkson JJ, Butsko Y, Joseph J, Savolainen V, Parokonny AS
(2003) Molecular systematics, GISH and the origin of hybrid taxa in Nicotiana (Solanaceae).
Ann Bot 92:107–127
Chester M, Gallagher JP, Symonds VV, Cruz da Silva AV, Mavrodiev EV, Leitch AR, Soltis PS
and Soltis DE (2012) Extensive chromosomal variation in a recently formed natural
allopolyploid species, Tragopogon miscellus (Asteraceae). Proceedings of the National
Academy of Sciences of the United States of America, 109:1176-1181
Chester M, Sykorova E, Fajkus J, Leitch AR (2010) Single integration and spread of a Copia-like
sequence nested in rDNA intergenic spacers of Allium cernuum (Alliaceae). Cytogenet
Genome Res 129:35–46
Clarkson JJ, Kelly LJ, Leitch AR, Knapp S, Chase MW (2010) Nuclear glutamine synthetase
evolution in Nicotiana: phylogenetics and the origins of allotetraploid and homoploid
(diploid) hybrids. Mol Phylogenet Evol 55:99–112
Clarkson JJ, Lim KY, Kovarik A, Chase MW, Knapp S, Leitch AR (2005) Long-term genome
diploidization in allopolyploid Nicotiana section Repandae (Solanaceae). New Phytol
168:241–252
Colot V, Goyon C, Faugeron G, Rossignol JL (1995) Methylation of repeated DNA sequences
and genome stability in Ascobolus immersus. Can J Bot-Rev Canadienne De Botanique
73:S221–S225
Doyle JJ, Flagel LE, Paterson AH, Rapp RA, Soltis DE, Soltis PS, Wendel JF (2008)
Evolutionary genetics of genome merger and doubling in plants. Ann Rev Genet 443–461
Feder JL, Nosil P (2009) Chromosomal inversions and species differences: when are geens
affecting adaptive divergence and reproductive isolation expected to reside within inversions?
Evolution 63:3061–3075
Feldman M, Levy AA (2009) Genome evolution in allopolyploid wheat-a revolutionary
reprogramming followed by gradual changes. J Genet Genomics 36:511–518
Fulnecek J, Lim KY, Leitch AR, Kovarik A, Matyasek R (2002) Evolution and structure of 5S
rDNA loci in allotetraploid Nicotiana tabacum and its putative parental species. Heredity
88:19–25
Fulnecek J, Matyasek R, Kovarik A (2009) Faithful inheritance of cytosine methylation patterns
in repeated sequences of the allotetraploid tobacco correlates with the expression of DNA
methyltransferase gene families from both parental genomes. Mol Genet Genomics
281:407–420
Gaeta RT, Pires JC, Iniguez-Luy F, Leon E, Osborn TC (2007) Genomic changes in
resynthesized Brassica napus and their effect on gene expression and phenotype. Plant Cell
19:3403–3417
Gazdova B, Siroky J, Fajkus J, Brzobohaty B, Kenton A, Parokonny A, Heslop-Harrison JS,
Palme K, Bezdek M (1995) Characterization of a new family of tobacco highly repetitive
DNA, GRS, specific for the Nicotiana tomentosiformis genomic component. Chromosome
Res 3:245–254
Gerstel DU, Burns JA (1967) Phenotypic and chromosomal abnormalities associated with the
introduction of heterochromatin from Nicotiana otophora into N. tabacum. Genetics
56:483–502
Gill BS (1991) Nucleocytoplasmic interaction (NCI) hypothesis of genome evolution and
speciation in polyploid plants. In: Sasakuma T, KinoshitaT (ed.) Proceedings of the Kihara
CO
RR
381
382
383
384
385
386
387
388
389
390
391
392
393
394
395
396
397
398
399
400
401
402
403
404
405
406
407
408
409
410
411
412
413
414
415
416
417
418
419
420
421
422
423
424
425
426
427
428
429
430
431
432
433
434
Book ISBN: 978-3-642-31441-4
Page: 221/224
Evolutionary Implications of Genome
UN
Editor Proof
11
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 11
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 222/224
EC
TE
D
PR
OO
F
memorial international symposium on cytoplasmic engineering in wheat, Kihara Memorial
Foundation, Yokohama, Japan, pp 48–53
Goodspeed TH (1954) The genus Nicotiana Massachusetts. Chronica Botanica Company, USA
Grandbastien M, Audeon C, Bonnivard E, Casacuberta JM, Chalhoub B, Costa APP, Le QH,
Melayah D, Petit M, Poncet C, Tam SM, Van Sluys MA, Mhiri C (2005) Stress activation and
genomic impact of Tnt1 retrotransposons in Solanaceae. Cytogenetic Genome Res
110:229–241
Heslop-Harrison JS, Schwarzacher T (2011) Organisation of the plant genome in chromosomes.
Plant J 66:18–33
Intrieri MC, Muleo R, Buiatti M (2008) Phytochrome A as a functional marker of phyletic
relationships in Nicotiana genus. Biol Plant 52:36–41
Jakowitsch J, Mette MF, van der Winden J, Matzke MA, Matzke AJM (1999) Integrated
pararetroviral sequences define a unique class of dispersed repetitive DNA in plants. Proc Nat
Acad Sci USA 96:13241–13246
Jakowitsch J, Papp I, Matzke MA, Matzke AJM (1998) Identification of a new family of highly
repetitive DNA, NTS9, that is located predominantly on the S9 chromosome of tobacco.
Chromosome Res 6:649–659
Jiao YN, Wickett NJ, Ayyampalayam S, Chanderbali AS, Landherr L, Ralph PE, Tomsho LP, Hu
Y, Liang HY, Soltis PS, Soltis DE, Clifton SW, Schlarbaum SE, Schuster SC, Ma H, LeebensMack J, dePamphilis CW (2011) Ancestral polyploidy in seed plants and angiosperms. Nature
473:97–113
Kelly LJ, Leitch AR, Clarkson JJ, Hunter RB, Knapp S, Chase MW (2010) Intragenic
recombination events and evidence for hybrid speciation in Nicotiana (Solanaceae). Mol Biol
Evol 27:781–799
Kenton A, Parokonny AS, Gleba YY, Bennett MD (1993) Characterization of the Nicotiana
tabacum L. genome by molecular cytogenetics. Mol Gen Genet 240:159–169
King M (1993) Species evolution the role of chromosome change. Cambridge University Press,
Cambridge
Knapp S, Chase MW, Clarkson JJ (2004) Nomenclatural changes and a new sectional
classification in Nicotiana (Solanaceae). Taxon 53:73–82
Kobayashi T (2011) Regulation of ribosomal RNA gene copy number and its role in modulating
genome integrity and evolutionary adaptability in yeast. Cell Mol Life Sci 68:1395–1403
Koukalova B, Moraes AP, Renny-Byfield S, Matyasek R, Leitch AR, Kovarik A (2010) Fall and
rise of satellite repeats in allopolyploids of Nicotiana over c. 5 million years. New Phytol
186:148–160
Koukalova B, Reich J, Matyasek R, Kuhrova V, Bezdek M (1989) A BamHI family of highly
repeated DNA sequecnes of Nicotiana tabacum. Theor Appl Genet 78:77–80
Kovarik A, Dadejova M, Lim K, Chase M, JJ C, Knapp S, Leitch A (2008) Evolution of rDNA in
Nicotiana allopolyploids: a potential link between rDNA homogenization and epigentics. Ann
Bot 101:815–823
Kovarik A, Fajkus J, Koukalova B, Bezdek M (1996) Species-specific evolution of telomeric and
rDNA repeats in the tobacco composite genome. Theor Appl Genet 92:1108–1111
Kronenberger J, Lepingle A, Caboche M, Vaucheret H (1993) Cloning and expession of distinct
nitrite reductases in tobacco-leaves and roots. Mol Gen Genet 236:203–208
Le Comber SC, Ainouche ML, Kovarik A, Leitch AR (2010) Making a functional diploid: from
polysomic to disomic inheritance. New Phytol 186:113–122
Leitch AR, Lim KY, Skalicka K, Kovarik A (2006) Nuclear cytoplasmic interaction hypothesis
and the role of translocations in Nicotiana allopolyploids. In: Cigna AAD, M. Yerevan (eds.)
Radiation risk estimates in normal and emergency situations book series: NATO security
through science series B: Physics and Biophysics (ARMENIA) pp 319–326
Leitch IJ, Bennett MD (2004) Genome downsizing in polyploid plants. Biol J Linn Soc
82:651–663
CO
RR
435
436
437
438
439
440
441
442
443
444
445
446
447
448
449
450
451
452
453
454
455
456
457
458
459
460
461
462
463
464
465
466
467
468
469
470
471
472
473
474
475
476
477
478
479
480
481
482
483
484
485
486
A. Kovarik et al.
UN
Editor Proof
222
Layout: T1 Standard SC
Chapter No.: 11
223
EC
TE
D
PR
OO
F
Leitch IJ, Hanson L, Lim KY, Kovarik A, Chase MW, Clarkson JJ, Leitch AR (2008) The ups
and downs of genome size evolution in polyploid species of Nicotiana (Solanaceae). Ann Bot
101:805–814
Lim KY, Kovarik A, Matyasek R, Bezdek M, Lichtenstein CP, Leitch AR (2000a) Gene
conversion of ribosomal DNA in Nicotiana tabacum is associated with undermethylated,
decondensed and probably active gene units. Chromosoma 109:161–172
Lim KY, Kovarik A, Matyasek R, Chase MW, Clarkson JJ, Grandbastien MA, Leitch AR (2007)
Sequence of events leading to near-complete genome turnover in allopolyploid Nicotiana
within five million years. New Phytol 175:756–763
Lim KY, Matyasek R, Kovarik A, Leitch AR (2004a) Genome evolution in allotetraploid
Nicotiana. Biol J Linn Soc 82:599–606
Lim KY, Matyasek R, Lichtenstein CP, Leitch AR (2000b) Molecular cytogenetic analyses and
phylogenetic studies in the Nicotiana section Tomentosae. Chromosoma 109:245–258
Lim KY, Skalicka K, Koukalova B, Volkov RA, Matyasek R, Hemleben V, Leitch AR, Kovarik
A (2004b) Dynamic changes in the distribution of a satellite homologous to intergenic 26-18S
rDNA spacer in the evolution of Nicotiana. Genetics 166:1935–1946
Lim KY, Soltis DE, Soltis PS, Tate J, Matyasek R, Srubarova H, Kovarik A, Pires JC, Xiong ZY,
Leitch AR (2008) Rapid chromosome evolution in recently formed polyploids in Tragopogon
(Asteraceae). Plos One, 3
Liu B, Brubaker CL, Mergeai G, Cronn RC, Wendel JF (2002) Polyploid formation in cotton is
not accompanied by rapid genomic changes. Genome 44:321–330
Lynch M, O’Hely M, Walsh B, Force A (2001) The probability of preservation of a newly arisen
gene duplicate. Genetics 159:1789–1804
Mandakova T, Joly S, Krzywinski M, Mummenhoff K, Lysak MA (2010) Fast diploidization in
close mesopolyploid relatives of Arabidopsis. Plant Cell 22:2277–2290
Matassi G, Melis R, Macaya G, Bernardi G (1991) Compositional bimodality of the nuclear
genome of tobacco. Nucleic Acids Res 19:5561–5567
Matyasek R, Gazdova B, Fajkus J, Bezdek M (1997) NTRS, a new family of highly repetitive
DNAs specific for the T1 chromosome of tobacco. Chromosoma 106:369–379
Matzke M, Gregor W, Mette MF, Aufsatz W, Kanno T, Jakowitsch J, Matzke AJM (2004)
Endogenous paragretroviruses of polyploid Nicotiana tabacum and its diploid progenitors,
N. sylvestris and N. tomentosiformis. Biol J Linn Soc 82:627–638
McCarthy EW (2010) The effect of molecular and spectral reflectance evolution on Nicotiana
polyploids of different ages. In School of Biological and Chemical Sciences. London: Ph.D.
thesis from Queen Mary University of London, p 259
McClintock B (1984) The significance of responses of the genome to challenge. Science
226:792–801
Melayah D, Bonnivard E, Chalhoub B, Audeon C, Grandbastien MA (2001) The mobility of the
tobacco Tnt1 retrotransposon correlates with its transcriptional activation by fungal factors.
Plant Journal, 28:159-168
Mestiri I, Chague V, Tanguy AM, Huneau C, Huteau V, Belcram H, Coriton O, Chalhoub B,
Jahier J (2010) Newly synthesized wheat allohexaploids display progenitor-dependent meiotic
stability and aneuploidy but structural genomic additivity. New Phytol 186:101–186
Moscone EA, Matzke MA, Matzke AJM (1996) The use of combined FISH/GISH in conjunction
with DAPI counterstaining to identify chromosomes containing transgene inserts in
amphidiploid tobacco. Chromosoma 105:231–236
Murad L, Bielawski JP, Matyasek R, Kovarik A, Nichols RA, Leitch AR, Lichtenstein CP (2004)
The origin and evolution of Geminivirus-related DNA sequences in Nicotiana. Heredity
92:352–358
Murad L, Lim KY, Christopodulou V, Matyasek R, Lichtenstein CP, Kovarik A, Leitch AR
(2002) The origin of tobacco’s T genome is traced to a particular lineage within Nicotiana
tomentosiformis (Solanaceae). Am J Bot 89:921–928
CO
RR
487
488
489
490
491
492
493
494
495
496
497
498
499
500
501
502
503
504
505
506
507
508
509
510
511
512
513
514
515
516
517
518
519
520
521
522
523
524
525
526
527
528
529
530
531
532
533
534
535
536
537
538
Book ISBN: 978-3-642-31441-4
Page: 223/224
Evolutionary Implications of Genome
UN
Editor Proof
11
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 11
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 224/224
EC
TE
D
PR
OO
F
Parisod C, Salmon A, Zerjal T, Tenaillon M, Grandbastien MA, Ainouche M (2009) Rapid
structural and epigenetic reorganization near transposable elements in hybrid and allopolyploid genomes in Spartina. New Phytol 184:1003–1015
Petit M, Guidat C, Daniel J, Denis E, Montoriol E, Bui QT, Lim KY, Kovarik A, Leitch AR,
Grandbastien MA, Mhiri C (2010) Mobilization of retrotransposons in synthetic allotetraploid
tobacco. New Phytol 186:135–147
Petit M, Lim KY, Julio E, Poncet C, de Borne FD, Kovarik A, Leitch AR, Grandbastien MA,
Mhiri C (2007) Differential impact of retrotransposon populations on the genome of
allotetraploid tobacco (Nicotiana tabacum). Mol Genet Genomics 278:1–15
Renny-Byfield S, Chester M, Kovarik A, Le Comber S, Grandbastien M, Deloger M, Nichols R,
Macas J, Novak P, Chase M, Leitch A (2011) Next generation sequencing reveals genome
downsizing in allotetraploid Nicotiana tabacum, predominantly through the elimination of
paternally derived repetitive DNAs Molecular Biology and Evolution 28:2843–2854
Renny-Byfield S, Kovarik A, Chester M, Nichols RA, Macas J, Novak P and Leitch AR (2012)
Independent, rapid and targeted loss of highly repetitive DNA in natural and synthetic
allopolyploids of Nicotiana tabacum. Plos One, 7:e36963
Riechers DE, Timko MP (1999) Structure and expression of the gene family encoding putrescine
N-methyltransferase in Nicotiana tabacum: new clues to the evolutionary origin of cultivated
tobacco. Plant Mol Biol 41:387–401
Rieseberg LH (1997) Hybrid origins of plant species. Annu Rev Ecol Syst 28:359–389
Schenke D, Sasabe M, Toyoda K, Inagaki Y, Shiraishi T, Ichinose Y (2003) Genomic structure of
the NtPDR1 gene, harboring the two miniature inverted-repeat transposable elements,
NtToya1 and NtStowaway101. Genes Genet Syst 78:409–418
Skalicka K, Lim KY, Matyasek R, Matzke M, Leitch AR, Kovarik A (2005) Preferential
elimination of repeated DNA sequences from the paternal, Nicotiana tomentosiformis genome
donor of a synthetic, allotetraploid tobacco. New Phytol 166:291–303
Soltis DE, Buggs RJA, Doyle JJ, Soltis PS (2010) What we still don’t know about polyploidy.
Taxon 59:1387–1403
Song KM, Lu P, Tang KL and Osborn TC (1995) Rapid genome change in synthetic polyploids
of Brassica and its implications for polyploid evolution. Proceedings of the National
Academy of Sciences of the United States of America, 92:7719-7723
Sperisen C, Ryals J, Meins F (1991) Comparison of cloned genes provides evidence for
intergenomic exchange of DNA in the evolution of a tobacco glucan endo-1,3 betaglucosidase gene family. Proc Nat Acad Sci USA 88:1820–1824
Strasburg JL, Scotti-Saintagne C, Scotti I, Lai Z, Rieseberg LH (2009) Genomic patterns of
adaptive divergence between chromosomally differentiated sunflower species. Mol Biol Evol
26:1341–1355
Szadkowski E, Eber F, Huteau V, Lode M, Coriton O, Jenczewski E and Chevre AM (2011)
Polyploid formation pathways have an impact on genetic rearrangements in resynthesized
Brassica napus. New Phytologist, 191:884-894
Takumi S, Ida M, Haisa Y, Ando S, Nakamura C (2002) Genomic structure and homoeologous
relationship of the two alpha-subunit genes of a heterotrimeric GTP-binding protein in
tobacco. Genome 45:626–633
Turner TL, Hahn MW (2010) Genomic islands of speciation or genomic islands and speciation?
Mol Ecol 19:848–850
Vaucheret H, Vincentz M, Kronenberger J, Caboche M, Rouze P (1989) Molecular cloning and
characterization of the 2 homologous genes coding for nitrate reductase in tobacco. Mol Gen
Genet 216:10–15
Volkov RA, Borisjuk NV, Panchuk II, Schweizer D, Hemleben V (1999) Elimination and
rearrangement of parental rDNA in the allotetraploid Nicotiana tabacum. Mol Biol Evol
16:311–320
Xiong ZY, Gaeta RT, Pires JC (2011) Homoeologous shuffling and chromosome compensation
maintain genome balance in resynthesized allopolyploid Brassica napus. Proc Nat Acad Sci
USA 108:7908–7913
CO
RR
539
540
541
542
543
544
545
546
547
548
549
550
551
552
553
554
555
556
557
558
559
560
561
562
563
564
565
566
567
568
569
570
571
572
573
574
575
576
577
578
579
580
581
582
583
584
585
586
587
588
589
590
591
592
A. Kovarik et al.
UN
Editor Proof
224
Metadata of the chapter that will be visualized in
SpringerLink
Book Title
Polyploidy and Genome Evolution
Series Title
Chapter Title
Polyploid Evolution in Spartina: Dealing with Highly Redundant Hybrid Genomes
Copyright Year
2012
Copyright HolderName
Springer-Verlag Berlin Heidelberg
Corresponding Author
Family Name
Ainouche
Particle
Given Name
M.
Suffix
Author
Division
University of Rennes 1
Organization
UMR CNRS 6553 Ecobio
Address
Bât. 14A, Campus Scientifique de Beaulieu, 35042, Rennes Cedex, France
Email
malika.ainouche@univ-rennes1.fr
Family Name
Chelaifa
Particle
Given Name
H.
Suffix
Division
University of Rennes 1
Organization
UMR CNRS 6553 Ecobio
Address
Bât. 14A, Campus Scientifique de Beaulieu, 35042, Rennes Cedex, France
Email
Author
Family Name
Ferreira
Particle
Given Name
J.
Suffix
Division
University of Rennes 1
Organization
UMR CNRS 6553 Ecobio
Address
Bât. 14A, Campus Scientifique de Beaulieu, 35042, Rennes Cedex, France
Email
Author
Family Name
Bellot
Particle
Given Name
S.
Suffix
Division
University of Rennes 1
Organization
UMR CNRS 6553 Ecobio
Address
Bât. 14A, Campus Scientifique de Beaulieu, 35042, Rennes Cedex, France
Email
Author
Family Name
Ainouche
Particle
Given Name
A.
Suffix
Division
University of Rennes 1
Organization
UMR CNRS 6553 Ecobio
Address
Bât. 14A, Campus Scientifique de Beaulieu, 35042, Rennes Cedex, France
Email
Author
Family Name
Salmon
Particle
Given Name
A.
Suffix
Division
University of Rennes 1
Organization
UMR CNRS 6553 Ecobio
Address
Bât. 14A, Campus Scientifique de Beaulieu, 35042, Rennes Cedex, France
Email
Abstract
Polyploidy and recurrent interspecific hybridization represent major features of Spartina evolution, resulting
in several superimposed divergent genomes that coexist in the currently living species. This chapter
summarizes what we presently know about Spartina history, emphasizing the recent hybridization and
polyploidization events that have important ecological and evolutionary consequences. Particular attention
is devoted to the recent formation of the allododecaploid invasive Spartina anglica, a salt-marsh “ecosystem
engineer” that resulted from hybridization between the hexaploid S. alterniflora (introduced from North
America) and tetraploid S. maritima (a European native) and subsequent genome duplication of the F1 hybrid
S. x townsendii during the nineteenth century in Western Europe. Allopolyploidy was not accompanied by
substantial restructuring of the parental genomes, as observed in some other allopolyploid systems. The major
evolutionary events affect the regulatory systems controlling gene expression (including epigenetic
regulation), which appear to have been profoundly altered by the merger of different genomes.
Methodological challenges in exploring non-model, highly redundant genomes resulting from superimposed
events of polyploidization (such as those encountered in Spartina) and the contribution of the new massive
parallel sequencing technologies are discussed.
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
F
M. Ainouche, H. Chelaifa, J. Ferreira, S. Bellot, A. Ainouche
and A. Salmon
PR
OO
5
Abstract Polyploidy and recurrent interspecific hybridization represent major
features of Spartina evolution, resulting in several superimposed divergent genomes that coexist in the currently living species. This chapter summarizes what we
presently know about Spartina history, emphasizing the recent hybridization and
polyploidization events that have important ecological and evolutionary consequences. Particular attention is devoted to the recent formation of the allododecaploid invasive Spartina anglica, a salt-marsh ‘‘ecosystem engineer’’ that resulted
from hybridization between the hexaploid S. alterniflora (introduced from North
America) and tetraploid S. maritima (a European native) and subsequent genome
duplication of the F1 hybrid S. x townsendii during the nineteenth century in
Western Europe. Allopolyploidy was not accompanied by substantial restructuring
of the parental genomes, as observed in some other allopolyploid systems. The
major evolutionary events affect the regulatory systems controlling gene expression (including epigenetic regulation), which appear to have been profoundly
altered by the merger of different genomes. Methodological challenges in
exploring non-model, highly redundant genomes resulting from superimposed
events of polyploidization (such as those encountered in Spartina) and the contribution of the new massive parallel sequencing technologies are discussed.
D
4
Polyploid Evolution in Spartina: Dealing
with Highly Redundant Hybrid Genomes
TE
3
Chapter 12
EC
2
Book ISBN: 978-3-642-31441-4
Page: 225/242
CO
RR
1
Book ID: 272454_1_En
Date: 16-8-2012
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: 12
M. Ainouche (&) H. Chelaifa J. Ferreira S. Bellot A. Ainouche A. Salmon
University of Rennes 1, UMR CNRS 6553 Ecobio, Bât. 14A, Campus Scientifique de
Beaulieu, 35042, Rennes Cedex, France
e-mail: malika.ainouche@univ-rennes1.fr
P. S. Soltis and D. E. Soltis (eds.), Polyploidy and Genome Evolution,
DOI: 10.1007/978-3-642-31442-1_12, Springer-Verlag Berlin Heidelberg 2012
225
Layout: T1 Standard SC
Chapter No.: 12
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 226/242
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
61
62
63
F
30
PR
OO
29
D
28
Polyploidy and recurrent interspecific hybridization represent major features of
Spartina evolution, resulting in several superimposed divergent genomes that
coexist in the currently living species. Particularly fascinating is the rapid range
expansion of the recently formed allododecaploid species S. anglica Hubbard that
formed in Western Europe during the end of the nineteenth century. The ecological
impact and genetic determinants of the spectacular propagation of this invasive
species that is now (deliberately or accidentally) introduced on several continents
generated an abundant literature (reviewed in Gray et al. 1990, Ainouche et al.
2004a, Triplet and Gallicé 2008, Ainouche et al. 2009) and much interest, as
illustrated by the successive International Invasive Spartina Conferences or Forums held in 1990 (Seattle WA, USA), 1997 (Olympia, WA, USA), 2004 (San
Francisco, CA, USA), and 2011 (Oakland, CA, USA). In the field of evolutionary
biology, Spartina has long represented a textbook example of recent allopolyploid
speciation in which the historical context of species formation is particularly well
documented (Huskin 1930; Stebbins 1950).
In this chapter, we summarize the current state of knowledge about Spartina
evolutionary history, including recent insights from evolutionary genetics and
genomic approaches. We also examine how genome evolution following natural
interspecific hybridization and polyploidization has contributed to diversification
and adaptation. Methodological challenges in exploring highly redundant genomes
resulting from superimposed events of polyploidization will be discussed.
TE
27
EC
26
12.1 Introduction
12.2 Recurrent Reticulate Evolution and Polyploidy
in Spartina
CO
RR
25
M. Ainouche et al.
The grass genus Spartina (‘‘cordgrasses’’) belongs to the Chloridoideae subfamily
(Fig. 12.1), one of the most poorly understood lineages of the Poaceae. Divergence
between Spartina and various grass model species is currently estimated as 35–40
MYA with Sorghum—maize–sugar cane (subfamily Panicoideae) and 50 MYA
with rice (subfamily Erhartoideae) (Christin et al. 2008, but see Prasad et al. 2011).
Phylogenetic relationships among genera of Chloridoideae are not fully resolved
and still are under debate (Hilu and Alice 2001, GPWG 2001). In recent molecular
phylogenies, Spartina appears closely related to Sporobolus, Calamovilfa and
Zoysia (Columbus et al. 2007, Fortune et al. 2007). Species from these genera
share a C4-type photosynthetic system that evolved in the Chloridoideae 25–32
MYA (Christin et al. 2008). C4 photosynthesis is generally considered an adaptation conferring higher productivity under warm temperatures. However, species
of Spartina exhibit geographic distributions that cover a range of climatic conditions, from temperate to tropical–subtropical regions; the species exhibit salt
and/or drought tolerance on coastal or inland marshes or sand dunes.
UN
Editor Proof
226
Layout: T1 Standard SC
Chapter No.: 12
(a)
Book ISBN: 978-3-642-31441-4
Page: 227/242
Polyploid Evolution in Spartina
227
(c)
Panicoideae
Cynodon dactylon
S. argentinensis
Hybrids (9x)
6x
S. foliosa
CHLORIDOIDEAE
Hybrids (6x)
4x
Cynodonteae
(b)
Zoy sieae
Pooideae
S. pectinata
6x , 8x
S. cynusoroides
Hybrids
S. densiflora (7x)
PR
OO
Aristidoideae
Bambusoideae
S. anglica
(12x)
S. alterniflora
Danthonioideae
Ehrhartoideae
S. x townsendii
(6x)
S. maritima
Arundinoideae
+ Micrairoideae
S. x neyrautii
(6x)
F
Editor Proof
12
Book ID: 272454_1_En
Date: 16-8-2012
S. x caespitosa
SPARTINA
S. bakeri
Calamovilfa
Sporobolus
Zoysia
S patens
Eragrostideae
S. arundinacea
Hybrids
Fig. 12.1 Phylogeny of Spartina (see references in the text) a position within the grass family
b position within the Chloridoideae subfamily c phylogenetic relationships among Spartina
species, recurrent hybridization and polyploidy
66
67
68
69
70
71
72
73
74
Marchant (1968a)
Marchant (1968a)
Marchant (1968a)
Marchant (1968a)
Marchant. (1968a)
Marchant (1968a)
Kim et al. (2010)
Ayres et al. (2008), Fortune et al. . (2008)
Marchant. (1968b)
Ayres et al. (2008)
Marchant (1968b)
Marchant (1968b)
Marchant (1977)
Marchant. (1968b)
TE
densiflora Brongn.
maritima (Curtis) Fern.
foliosa Trin.
alterniflora Loisel.
x townsendii H & J Groves
x neyrautii Foucaud
anglica C.E. Hubbard
EC
S.
S.
S.
S.
S.
S.
S.
40
40
40
40
40
40
40, 60, 80
70
60
62
62
62
62
120, 122, 124
CO
RR
65
patens (Aiton) Muhl.
cynusoroides (L.) Roth
backeri Merr.
gracilis Trin.
arundinacea (Thouars) Carmich.
pectinata Link
Spartina is composed of about 15 perennial species that have mostly diversified
in the New World (Mobberley 1956). Accidental or deliberate introduction of
species outside their native range over the past 150 years has accelerated
diversification by facilitating hybridization with native species, introgression or
speciation. The basic (haploid) chromosome number in this lineage is considered
to be x = 10 (Marchant 1968a), and all Spartina species recorded to date are
polyploid, ranging from tetraploids to dodecaploids (Table 12.1).
Molecular phylogenies based on nuclear and chloroplast DNA sequences
indicate that Spartina comprises two main lineages that include tetraploid and
hexaploid species, respectively (Baumel et al. 2002a). The tetraploid lineage is
composed of species native to the New World; these species colonize coastal or
UN
64
S.
S.
S.
S.
S.
S.
D
Table 12.1 Chromosome numbers reported for Spartina. Additional chromosome numbers
resulting from interspecific hybridization and/or backcrosses are presented in the text
Taxa
2n
References
Layout: T1 Standard SC
Chapter No.: 12
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 228/242
82
83
84
85
86
87
88
89
90
91
92
93
94
95
96
97
98
99
100
101
102
103
104
105
106
107
108
109
110
111
112
113
114
115
116
117
118
F
PR
OO
81
D
80
TE
78
79
EC
77
inland salt marshes from either North America (S. patens, S. bakeri, S. gracilis,
S. cynusoroides, S. pectinata) or South America (S. ciliata, S. arundinacea).
S. argentinensis (syn. S. spartinae), which has a disjunct distribution in Central
America and South America, is sister to the hexaploid lineage (Fig. 12.1c). This
hexaploid clade is composed of S. maritima, S. alterniflora, and S. foliosa, all
colonizing low-marsh zones. S. maritima, native to the Atlantic coasts of Western
Europe and Africa was, until the nineteenth century, the only Old World native
species, if we exclude recent taxa of hybrid origin. S. foliosa, a species limited to
the Pacific coast of North America (California and Mexico), is a weakly supported
sister species to S. alterniflora (Baumel et al. 2002a; Ainouche et al. 2004b) and is
distributed along the east coast of North and South America from Canada to
Argentina. This species has now one of the largest geographic distributions in the
genus, as it has been introduced to several continents (Europe, Asia).
The nature of polyploidy (auto- vs. allopolyploidy) and the origin of the
hexaploid clade are not fully understood. Up to three different duplicated (homoeologous) genes were distinguished in hexaploids for the low-copy nuclear
gene Waxy, with substitution rates ranging from 2.18 to 4.79 % among homoeologs (Fortune et al. 2007). The presence of three different homoeologous copies of
Waxy could support a hybrid (allopolyploid) origin of this lineage (Fortune et al.
2007). These hexaploid species exhibit regular bivalent pairing (Marchant 1968b).
Although allopolyploidy is the currently favored hypothesis for the origin of the
hexaploid clade, and is supported by the propensity for interspecific hybridization
in this genus, an autopolyploid origin cannot be ruled out. It has been shown that
genetic diploidization via disomic inheritance may occur rapidly following autopolyploid formation (Le Comber et al. 2010), leading to a progressive divergence
of genes duplicated by polyploidy, and to a gene topology that becomes similar to
what would be expected for allopolyploids (Staub et al. 2003).
Dating the origin of a polyploid species from (biparentally inherited) nuclear gene
divergence data is difficult in that (selective and neutral) population-level evolutionary processes and differential evolution of genes duplicated by polyploidy may
give different estimates of the date of origin at different loci. Moreover, in allopolyploids, the divergence of homoeologs actually reflects the divergence between the
parental species, thus leading to an overestimated age of the polyploid (Doyle and
Negan 2009). We recently attempted to estimate the divergence times between
different Spartina lineages from maternally inherited chloroplast sequences. Using
reconstruction of the Spartina chloroplast genome from 454 Roche GS-FLX (Titanium) pyrosequencing (Bellot S. and Ainouche M., unpublished), we were able to
analyze three coding (matK, rbcL, ndhF) and eight non-coding (intergenic spacers or
introns) chloroplast sequences. Using a Bayesian phylogenetic analysis calibrated
with known divergence times in the grass family (Wolfe et al. 1989; Gaut 2002;
Prasad et al. 2005; Chalupska et al. 2008; Christin et al. 2008), we estimated the
divergence between the tetraploid and hexaploid Spartina maternal genomes at 5
MYA, and the divergence time between the two hexaploid S. alterniflora and S.
maritima chloroplast genomes at approximately 3 MYA.
CO
RR
75
76
M. Ainouche et al.
UN
Editor Proof
228
Layout: T1 Standard SC
Chapter No.: 12
126
127
128
129
130
131
132
133
134
135
136
137
138
139
140
141
142
143
144
145
146
147
148
149
150
151
152
153
154
155
156
157
158
159
160
161
162
F
PR
OO
125
D
123
124
TE
122
EC
121
229
Several additional recent hybridization events between Spartina species have
led to the formation of homoploid hybrids or new allopolyploids. These events not
only have involved parental species of the same ploidal level within the tetraploid
or the hexaploid clades, but also involved crosses between tetraploid and hexaploid species (Fig. 12.1c). In the tetraploid lineage, early morphological analyses
suggested that S. x caespitosa was a hybrid between S. patens (Eastern North
America) and S. pectinata, which has a wide distribution across much of North
America (Mobberley 1956). Moreover, a recent survey of S. pectinata populations
across North America has revealed hexaploid and octoploid cytotypes that are
morphologically indistinguishable from the tetraploids (Kim et al. 2010). The
origin of these octoploids is unknown, and the current hypothesis that the
hexaploids derive from backcrosses between octoploids and tetraploids has to be
verified. Genetic and genomic resources are currently being developed for this
species, which is considered a good candidate for bioenergy due to its high biomass production (Gedye et al. 2010).
The South American species S. densiflora illustrates well the genetic exchanges
occurring between the tetraploid and hexaploid lineages. The history of this species was only recently elucidated: this vigorous, high marsh species originated
from southeastern coast of South America, but has been introduced to Chile,
California and Spain (Bortolus 2006). Genetic analyses and molecular phylogenies
(Baumel et al. 2002a; Ayres et al. 2008; Fortune et al. 2008) have revealed that this
species is heptaploid (2n = 70), with a hybrid origin from a tetraploid maternal
species closely related to S. arundinacea (with which it shares high chloroplast
sequence similarity), which occurs in Southern Hemisphere islands (Amsterdam,
St Paul and Tristan de Cunha islands), and a hexaploid paternal species related to
S. alterniflora that also occurs in Argentina. Interestingly, S. densiflora appears to
have hybridized with each species from the hexaploid clade in both its native area
and in more recently colonized regions where it came in contact with native
species: (1) At the mouth of the Rio de la Plata in Argentina and Uruguay,
morphologically intermediate plants named Spartina longispica were early recognized as hybrids between S. densiflora and S. alterniflora (Saint-Yves 1932;
Mobberley 1956). (2) S. densiflora was accidentally introduced to California,
probably from Chile, as suggested by geographic, historical, and molecular data
(Bortolus 2006; Fortune et al. 2008). In the San Francisco Bay, Ayres et al. (2008)
have reported hybrids between S. densiflora and the native S. foliosa (2n = 62).
Different chromosome numbers (2n = 66, 94–96) suggest that the F1 hybrids
deriving from a heptaploid and a hexaploid parent might also have backcrossed
with the parental species. (3) On the Atlantic coast of the Iberian Peninsula,
Castillo et al. (2010) have recently detected hybrids between introduced highmarsh S. densiflora and the native low-marsh hexaploid S. maritima (2n = 60). As
in California, different chromosome numbers (2n = 64–66, 2n = ca. 94) were
recorded for the hybrids, with either S. densiflora or S. maritima as maternal
genome donors. No genome doubling (allopolyploid speciation) is recorded to date
in these perennial (apparently sterile) hybrids, as occurred for S. anglica.
CO
RR
119
120
Book ISBN: 978-3-642-31441-4
Page: 229/242
Polyploid Evolution in Spartina
UN
Editor Proof
12
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 12
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 230/242
170
171
172
173
174
175
176
177
178
179
180
181
182
183
184
185
186
187
188
189
190
191
192
193
194
195
196
197
198
199
200
201
202
203
204
205
206
207
F
168
169
PR
OO
167
D
166
TE
165
Within the hexaploid clade, the outcomes of interspecific hybridization are
consistent with the phylogenetic relationships and genetic divergence among
species (Baumel et al. 2002a; Ainouche et al. 2004b): production of fertile invasive
hybrids between closely related taxa on the one hand, and formation of sterile
hybrids (followed by genome doubling) between more divergent species on the
other (Fig. 12.1c). These hybridizations resulted from recent introductions (nineteenth and twentieth centuries) of S. alterniflora from the eastern coast of the
Americas to the Pacific coast of North America and to the Atlantic coast of
Western Europe.
In California, plants of S. alterniflora spread rapidly (Daehler and Strong 1997;
Civille et al. 2005). Hybridization with its sister species S. foliosa (native to
California) has been shown to occur in both directions (Ayres et al. 1999). The
greater male fitness of S. alterniflora and recurrent backcrosses has resulted in
hybrid swarms that progressively replace original S. foliosa plants (Antilla et al.
2000; Ayres et al. 2007). Adding an additional layer of complexity, some of these
plants might also be involved in the formation of new hybrid genotypes with the
introduced S. densiflora (Ayres et al. 2008).
In Europe, S. alterniflora was accidentally introduced by ship ballast in southern
England and western France, where it hybridized with S. maritima. In England,
hybridization recorded in 1870 gave rise to S. x townsendii, a perennial, sterile
hybrid (Groves and Groves 1880). Another sterile hybrid between S. alterniflora
and S. maritima was discovered in 1892 in southwestern France in the Bidassoa
Estuary (Foucaud 1897) and named Spartina x neyrautii (Jovet 1941). Because of
their different morphology, some authors suggested that S. x neyrautii and S. x
townsendii might result from reciprocal crosses; however, molecular data revealed
that both hybrids share the same chloroplast genome of S. alterniflora, identifying it
as the maternal parent of both hybrids (Baumel et al. 2003).
After 1890 in England, fertile plants were recorded that appeared to have
resulted from chromosome doubling in S. x townsendii (Marchant 1963), thus
leading to the formation of a new allododecaploid species named S. anglica
(Hubbard 1968), with chromosome numbers 2n = 120, 122, or 124 (Marchant
et al. 1968b) suggesting aneuploidy. The vigorous plants (Fig. 12.2) have rapidly
colonized Western European salt marshes (Raybould et al. 1991a; Thompson
1991; Genegou and Levasseur 1993). Robust shoots, rhizomes and root systems
enable this new species to accumulate large volumes of tidal sediments. For this
reason, S. anglica was deliberately introduced in several parts of the world
(northern Europe, Australia, New Zealand, China, North America) for land reclamation and marsh restoration purposes. S. anglica has rapidly expanded in its
introduced range and now has a worldwide distribution (Ainouche et al. 2009).
The rapid spread of the introduced populations has led to various attempts to
control or eradicate the species; in fact, it is now listed among the 100 ‘‘world’s
worst’’ invaders (IUCN 2000). A recent survey of the ploidal levels in the original
population (Southampton area, UK) where S. anglica and S. x townsendii still
coexist, has revealed that the perennial, sterile F1 hybrids represent more than
90 % of the population (Renny-Byfield et al. 2010). Using genomic in situ
EC
164
CO
RR
163
M. Ainouche et al.
UN
Editor Proof
230
Layout: T1 Standard SC
Chapter No.: 12
Book ISBN: 978-3-642-31441-4
Page: 231/242
Polyploid Evolution in Spartina
231
TE
D
PR
OO
F
Editor Proof
12
Book ID: 272454_1_En
Date: 16-8-2012
Fig. 12.2 Spartina anglica in the Baie des Veys (Cotentin), first colonized site (1906) in France
(Corbière 1926)
211
212
213
214
215
216
217
218
219
220
221
222
223
224
EC
210
hybridization (GISH), this study also confirmed the existence of nonaploid plants
(2n = ca. 90), most likely resulting from backcrosses between S. anglica and its
maternal parent S. alterniflora.
The recurrent and continuing hybridization and genome duplication in Spartina
(Fig. 12.1) make it a particularly useful model with which to explore the consequences of these processes at various evolutionary time scales. Recent events
allow ecological, phenotypic, and genetic comparisons between the newly formed
hybrids or polyploids and their parents that are still extant in natural populations, a
situation that is only met in a few biological systems (see also Chaps. 13 and 14,
this volume).
CO
RR
209
12.3 Ecological and Adaptive Consequences of Hybridization
and Polyploidy in Spartina
UN
208
Spartina species play an important ecological role in the sedimentary dynamics of
salt marshes, where the plants are considered to be ‘‘ecosystem engineers’’(Crooks
2002). The ecological range of S. anglica along the shore is larger than either of its
parents. Spartina anglica tolerates several hours of immersion at high tides and
thus is able to occupy a vacant niche as a pioneer species in the low-tide zone. This
Layout: T1 Standard SC
Chapter No.: 12
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 232/242
232
233
234
235
236
237
238
239
240
241
242
243
244
245
246
247
248
249
250
251
252
253
254
255
256
257
258
259
260
261
262
263
264
265
266
267
268
269
F
231
PR
OO
230
D
228
229
TE
227
species may accrete large volumes of tidal sediments, making the habitat more
terrestrial, and allowing colonization by other salt marsh plant species, which
modifies the physical structure of intertidal coastal zones.
Recently formed Spartina hybrids and allopolyploids display hybrid vigor and
rapid expansion in their native range and invasive abilities when introduced, with
important implications for ecosystem management (Lambrinos 2008). S. alterniflora x foliosa hybrids have rapidly invaded the San Francisco Bay (Ayres et al.
2007). Several generations of introgressive hybridization make it difficult to differentiate invasive ‘‘cryptic hybrids’’ from ‘‘pure’’ native S. foliosa plants, and this
complicates eradication plans (The Invasive Spartina Project, www.spartina.org).
The heptaploid South American S. densiflora has also rapidly colonized Californian marshes where it increased fivefold in distribution during the last 25 years
following its introduction (Ayres et al. 2004). In Spain, the hybrid S. densiflora
x maritima exhibits greater ecological amplitude than either parental species:
hybrids are able to survive both in lower elevations in the intertidal zone where
S. maritima naturally grows and also in high marshes where S. densiflora invades
(Castillo et al. 2010).
Hybridization and polyploidy in Spartina are accompanied by various
biological changes that have influenced important adaptive traits such as breeding
system, physiology, and morphology. The Californian S. densiflora x foliosa
hybrids, derived from self-incompatible, outcrossing parents, have evolved selffertility that has contributed substantially to their rapid spread (Sloop et al. 2009).
This breakdown in self-incompatibility, also observed following allopolyploidy in
Brassica and Arabidopsis, is most likely triggered by epigenetic mechanisms
(Nasrallah et al. 2007). Several transgressive traits in height and biomass, vegetative growth rates, intertidal amplitude, and salinity tolerance are also reported in
the Spartina hybrids from California (Ayres et al. 2007).
Hybridization between S. alterniflora and S. maritima had very different morphological consequences in the two independent events that occurred in England
(S. x townsendii) and France (S. x neyrautii), even though these hybridization
events involved crosses in the same direction (S. alterniflora being the maternal
genome donor) and similar parental genotypes (Baumel et al. 2003; Yannic et al.
2004; Salmon et al. 2005). S. x neyrautii has shorter spikelets and is distinctly
more slender than S. x townsendii, which has longer fleshy leaves, resembling
more closely the maternal parent S. alterniflora, whereas S. x townsendii has
intermediate morphological features between S. maritima and S. alterniflora
(Mobberley 1956). The phenotypic differences between these two F1 hybrids of
similar genetic origin are puzzling and most likely represent differential effects of the
‘‘genomic shock’’ resulting from the merger of divergent genomes. S. x townsendii is
almost indistinguishable from its allopolyploid derivative S. anglica; moreover, the
latter species exhibits larger phenotypic plasticity (Thompson et al. 1991).
Physiological and anatomic adaptations are important components of Spartina
ecology and distribution (Maricle et al. 2006, 2009). As observed in many
polyploids (Otto 2007), stomatal cell size increases with ploidy level in Spartina
(Marchant 1967; Kim et al. 2010), which may affect photosynthetic rates (Warner
EC
226
CO
RR
225
M. Ainouche et al.
UN
Editor Proof
232
Layout: T1 Standard SC
Chapter No.: 12
277
278
279
280
281
282
283
284
285
286
287
288
289
290
291
292
293
294
295
296
297
298
299
300
301
302
303
304
305
306
307
308
309
310
F
276
PR
OO
275
D
274
TE
273
and Edwards 1993). The larger ecological amplitude of the allopolyploid
S. anglica compared to its parents has to be related to increased tolerance to highly
reducing and sulfidic sediment conditions. This increased tolerance may explain
the ability of S. anglica to colonize low-marsh zones (Maricle et al. 2006).
Survival of S. anglica in anoxic sediments likely is facilitated by its particular
ability to develop aerenchyma systems that supply the submerged plants with
atmospheric oxygen and efficiently transport oxygen to the roots (Maricle and Lee
2002). S. anglica displays enhanced mechanisms to transport O2 and exhibits five
times greater H2S removal than its progenitor species S. alterniflora (the other
parental species, S. maritima, was not investigated; Lee 2003).
An interesting function seems to have accompanied the formation of the
hexaploid lineage of Spartina: the ability to produce dimethylsulfoniopropionate
(DMSP), an osmoprotectant and anti-stress molecule (Larher et al. 1977). DMSP is
environmentally important as the main biogenic precursor of atmospheric dimethyl
sulfide (DMS), which has roles in the biogeochemical sulfur cycle, in cloud formation and in acid precipitation (Kocsis et al. 1998). DMSP is commonly produced by many marine algae, but this capacity is rare in angiosperms, where it has
been found only in three genera, one in the Asteraceae (Wollastonia) and two in
the Poaceae (Saccharum and Spartina) (Otte et al. 2004). Asteraceae and Poaceae
have independently developed different metabolic pathways to achieve this synthesis (Kocsis et al. 1998; Kocsis and Hanson 2000).
High DMSP concentrations are found in leaves of the hexaploids S. alterniflora,
S. foliosa and S. maritima (Otte et al. 2004), providing these plants with a characteristic ‘‘unpleasant sulphurous odor’’ noticed by early taxonomists (Mobberley
1956). In contrast, no DMSP is detected in the tetraploid species analyzed to date.
Although the evolutionary steps giving rise to this physiological novel capacity are
yet to be elucidated, the comparative data generated to date indicate that it may have
been enabled by the transition to the hexaploid condition. If so, this would represent
an example of a presumably important ecological adaptation arising from
polyploidy.
Physiological (and more generally phenotypic) evolution following hybridization and polyploidy is a direct outcome of the genomic consequences of genome
merger and duplication. Although we are still far from obtaining an exhaustive
knowledge of the complex gene and regulatory networks involved in most phenotypic traits of adaptive importance, significant progress has been made in recent
years regarding genetic and genomic processes accompanying polyploid evolution.
EC
272
233
CO
RR
270
271
Book ISBN: 978-3-642-31441-4
Page: 233/242
Polyploid Evolution in Spartina
12.4 Genome Evolution Following Hybridization
and Allopolyploid Speciation in Spartina
UN
Editor Proof
12
Book ID: 272454_1_En
Date: 16-8-2012
It is now well established that hybrid and allopolyploid genomes are not simply
additive with respect to their parental genomes and that myriad novel interactions
at both the structural and functional levels may lead to rapid evolution and
Layout: T1 Standard SC
Chapter No.: 12
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 234/242
318
319
320
321
322
323
324
325
326
327
328
329
330
331
332
333
334
335
336
337
338
339
340
341
342
343
344
345
346
347
348
349
350
351
352
353
354
F
317
PR
OO
316
D
314
315
TE
313
evolutionary novelties. It also may be that these phenomena play a critical role in
the evolutionary success of newly formed species in both short-term and long-term
evolutionary time (e.g., Wendel 2000; Chen 2007; Doyle et al. 2008; Van de Peer
et al. 2009).
The genetic context of the hybrid or allopolyploid species formation is an
important parameter to consider for subsequent evolution. Recurrent hybridization
events in natural populations provide new lineages with an expanded genetic base,
as observed in the introgressant populations resulting from multiple hybridization
and recurrent backcrosses involving different S. alterniflora and S. foliosa genotypes in California (Ayres et al. 2007). In contrast, low genetic diversity is
encountered in the parental populations of S. maritima and S. alterniflora that
hybridized in southern England and southern France (Baumel et al. 2003; Yannic
et al. 2004; Ainouche et al. Ainouche et al. 2004a). As a result, the F1 hybrids S. x
townsendii and S. x neyrautii share very similar parental genotypes and exhibit
similar, additive genetic composition. A strong genetic bottleneck seems to have
affected the new allopolyploid S. anglica as a result of a unique origin from S. x
townsendii: a very low interindividual genetic diversity is encountered within and
among populations in both its native and introduced ranges (Guénégou 1988;
Raybould et al. 1991b; Baumel et al. 2001; Ainouche et al. 2004a), although some
genetic variants may be encountered (Ayres et al. 2001).
Contrasting with most other young or experimentally resynthesized allopolyploid systems that exhibit rapid genome structural evolution (Ozkan et al. 2001;
Skalika et al. 2005; Gaeta et al. 2007; Lim et al. 2008; Tate et al. 2009; Szadowski
et al. 2010; Buggs et al. 2012; Chester et al. 2012), the new allododecaploid
S. anglica exhibits relative genome stability (Baumel et al. 2002b; Ainouche et al.
2004a). Most of the early evolutionary changes following allopolyploid speciation
in Spartina seem to affect the regulation of gene expression, including epigenetic
and transcriptome changes (Fig. 12.3). DNA methylation alterations revealed by
Methylation Sensitive AFLP (MSAP) appear triggered by hybridization in both
S. x townsendii and S. x neyrautii (Salmon et al. 2005). Genome duplication does
not entail significant additional changes, as S. anglica has inherited most of the
changes observed in S. x towsendii but exhibits few specific methylation
alterations. Parisod et al. (2009) have shown that an important fraction of these
methylation changes affect regions flanking transposable elements, which agrees
with the general view of methylation having evolved to control transposable
elements in eukaryotic genomes (Slotkin and Martienssen 2007) and with the fact
that no burst of transposition was detected following allopolyploid speciation
(Baumel et al. 2002b; Parisod et al. 2010). The hexaploid parental species
S. maritima and S. alterniflora have a genome size of 2C = 3.8 pg and 4.3 pg,
respectively (Fortune et al. 2008), which suggest that the basic haploid genome
ranges between 600 and 700 MB in these species. A preliminary investigation
from 454 pyrosequencing of genomic DNA in S. maritima revealed that about
27 % of the analyzed sequences were recognized as repetitive, with Gypsy-like
elements being mostly represented.
EC
312
CO
RR
311
M. Ainouche et al.
UN
Editor Proof
234
Layout: T1 Standard SC
Chapter No.: 12
Book ISBN: 978-3-642-31441-4
Page: 235/242
Polyploid Evolution in Spartina
235
TE
D
PR
OO
F
Editor Proof
12
Book ID: 272454_1_En
Date: 16-8-2012
357
358
359
360
361
362
363
364
365
366
367
368
369
Transcriptome evolution was first investigated in Spartina using oligo-microarrays. As no Expressed Sequence Tag (EST) database was then available for
Spartina, heterologous hybridization was performed using the related model
species Oryza sativa (Chelaifa et al. 2010a, b). The hexaploid species S. maritima
and S. alterniflora that exhibit high exon sequence identity (94.0–99.7 %) at
homologous loci displayed 1,247 differentially expressed genes in leaves from
plants grown in the same controlled conditions (Chelaifa et al. 2010a). Most of
these genes were found to be up-regulated in S. alterniflora. Similar levels of nonadditive parental patterns of gene expression were observed in both of the hybrids
S. x townsendii and S. x neyrautii (6.1 and 6.4 % of the analyzed genes, respectively, Fig. 12.3), including parental (mostly maternal) gene expression dominance
and transgressively expressed genes. However, the dominance of maternal
expression appeared more pronounced in S. x townsendii than in S. x neyrautii.
About 8.7 % of the analyzed genes were found differentially expressed between
these two F1 hybrids (Fig. 12.3), and interestingly, most transgressively expressed
UN
355
356
CO
RR
EC
Fig. 12.3 Genome evolution following recent hybridization and allopolyploidy in Spartina.
Percentages in red represent transcriptome changes evaluated using microarrays as described in
Chelaifa et al. (2010a, b). Effects of hybridization were estimated by comparing expression
profiles in the two independently formed natural F1 hybrids to a theoretical mid-parent value
representing additive parental expression. Effects of genome duplication were evaluated by
comparing S. anglica to S. x townsendii. Percentages in black represent DNA methylation
alterations evaluated using methylation sensitive AFLP, following the procedure employed by
Salmon et al. (2005) and Parisod et al. (2009)
Layout: T1 Standard SC
Chapter No.: 12
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 236/242
376
377
378
379
380
381
382
383
384
385
386
387
388
389
390
391
392
393
394
395
396
397
398
399
400
401
402
403
404
405
406
407
408
409
410
411
412
413
414
F
375
PR
OO
374
D
373
TE
372
genes were different, with genes up-regulated in S. x townsendii being related to
transport, detoxification and stress, and genes up-regulated in S. x neyrautii being
related to cellular growth and development. The two independent hybridization
events involving the same parental species appear to have generated differential
consequences in terms of gene expression. The functions of these differentially
expressed genes are consistent with the phenotypic differences previously mentioned between the two hybrids. Genome duplication in S. anglica entailed additional transcriptome changes (Fig. 12.3), consisting of the attenuation of the
maternal dominance observed in the F1 hybrid and an increased number of
transgressively overexpressed genes (Chelaifa et al. 2010b). Thus, both hybridization and genome duplication appear to have important, though different, effects
on the Spartina transcriptome, occurring shortly after genome merger and polyploidization. For the first time, these decoupled effects were analyzed during the
allopolyploid speciation process, by comparing the actual (naturally formed) F1
hybrid to its immediately derived allopolyploid that formed and survived in natural
conditions. Interestingly, our findings seem to parallel the conclusions emerging
from similar comparisons involving natural, more or less recent allopolyploids
and/or synthetic F1 hybrids (Hegarty et al. 2006; Flagel et al. 2008; Flagel and
Wendel 2010; Buggs et al. 2011).
The microarray analyses allowed the first large-scale investigation of the
Spartina transcriptome. However, it should be kept in mind that the expression
changes reported in these studies are most likely underestimated: only global
expression of the genes that hybridized on the rice microarrays were examined,
and homoeologous gene expression could not be distinguished. Similar levels of
expression might be attained via biased parental expression, which represents an
important component of the functional plasticity of polyploid genomes (Adams
et al. 2003; Chaudhary et al. 2009; Flagel et al. 2009; Buggs et al. 2011).
Distinction of homoeologous gene expression has been studied in several allopolyploid models where diploid representatives of the parental species are identified
(e.g. cotton: Udall et al. 2006; Flagel et al. 2008; Arabidopis: Chang et al. 2010;
Tragopogon: Buggs et al. 2010; Buggs et al. 2011). In Spartina, this task is particularly challenging as the parents of the young allopolyploid S. anglica are hexaploids
(expected to have retained up to three more-or-less divergent duplicated homoeologs
per locus) and because no diploid species is known in the genus (Fig. 12.4a).
Next-Generation Sequencing (NGS) technologies offer unique avenues to distinguish homoeologous copies in highly redundant genomes from natural, nonmodel species that have experienced successive polyploidization events. This
procedure is being developed for Spartina as follows (Fig. 12.4b): (1) construction
of a Spartina reference transcriptome for the hexaploid parental species using 454
Roche GS-FLX pyrosequencing; (2) Single Nucleotide Polymorphism (SNP)
detection among reads per annotated contig; (3) haplotype assembly to discriminate homoeoalleles; and (4) Illumina RNA-Sequencing (RNA-Seq) to explore
variation of homoeolog expression in the parental species.
Using the procedure outlined above, 38,000 contigs representing *17,000
unigenes were annotated for S. maritima and S. alterniflora from leaf and root
EC
371
CO
RR
370
M. Ainouche et al.
UN
Editor Proof
236
Layout: T1 Standard SC
Chapter No.: 12
Book ISBN: 978-3-642-31441-4
Page: 237/242
Polyploid Evolution in Spartina
237
D
PR
OO
F
Editor Proof
12
Book ID: 272454_1_En
Date: 16-8-2012
417
418
419
420
421
422
423
424
425
426
427
428
429
430
431
432
cDNA libraries, which represent the first reference transcriptome for the hexaploid
Spartina species (Ferreira et al. submitted ). An example of SNP detection among
reads is presented in Fig. 12.4b for an aligned portion of two homologous contigs
annotated in S. maritima and S. alterniflora as HECT-domain-containing protein
(Oryza sativa annotation LOC_Os12g24080.1|13112.m02448|cDNA). Reads were
assembled using a 95 % identity threshold, to avoid potential comparisons
involving paralogs. Six polymorphic sites are detected in this region, including
four polymorphic sites shared between S. maritima and S. alterniflora and two
species-specific polymorphic sites. The shared polymorphisms allow distinction of
two divergent haplotypes (1 and 2, Fig. 12.4b) present in both hexaploids, and one
(in S. maritima) or two (in S. alterniflora) additional minor variants. Screening of a
larger number of polymorphic sites and loci will provide information about the
number and divergence of (homoeo) alleles encountered in the hexaploids and
shed light on the evolutionary history of this lineage. A variable number of
retained copies per homologous locus may be expected. For instance, Fortune et al.
(2007) analyzed the low-copy nuclear gene Waxy that is present in two paralogs
(A and B) in Spartina. Only one B copy was encountered in S. maritima, whereas
three distinct B copies were found in S. alterniflora. These two species have
CO
RR
416
UN
415
EC
TE
Fig. 12.4 a Expected nuclear homoeologous gene copies per locus in the hexaploid parents, F1
hybrid and allododecaploid Spartina species b Procedure for sequence heterogeneity detection at
homologous nuclear coding loci; example from two homologous contigs in the hexaploid S.
maritima (contig length = 4,294, number of read = 127) and S. alterniflora (contig
length = 3,961, number of reads = 85) assembled from 454 Roche pyrosequencing of cDNA
libraries. Arrows of similar color (green or pink) represent divergent alleles (1 and 2) present in
the two hexaploids and detected from shared polymorphic sites between the two species, in the
central zone of the compared region. Other haplotypes represent slight variants of these two
copies
Layout: T1 Standard SC
Chapter No.: 12
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 238/242
M. Ainouche et al.
464
465
466
467
468
Acknowledgments This work benefited from the financial support of the Centre National de la
Recherche Scientifique (CNRS), University of Rennes 1, the French National Research Agency
(ANR), Region Bretagne, Genoscope, and from the Biogenouest (Transcriptomics and Environmental Genomics) platform facilities. P. Wincker, J. Poulain, Corinne Da Silva, O. Lima, S.
Coudouel, D. Naquin, A. Deillhy are thanked for their contribution to the 454 pyrosequencing
469
References
439
440
441
442
443
444
445
446
447
448
449
450
451
452
453
454
455
456
457
458
459
460
461
462
470
471
472
473
PR
OO
438
D
437
TE
436
EC
435
CO
RR
434
F
463
apparently lost the A copy that is still retained in the hexaploid S. foliosa (Fortune
et al. 2007). The large-scale detection of homoeologs from massive parallel
sequencing will provide a genome-wide view of the retention-loss process at
various evolutionary time scales (in the hexaploid parents and the nascent allododecaploid). Transcript loss might result from either homoeolog silencing as
observed in the various cases of subfunctionalization reported in allopolyploids
(reviewed in Osborn et al. 2003; Chen 2007; Doyle et al. 2008), physical loss of
the duplicated copies that may occur more or less rapidly following polyploid
speciation (e.g. Gaeta et al. 2007; Tate et al. 2009; Koh et al. 2010; Buggs et al.
2012) or from homoeologous recombination (Cifuentes et al. 2010; Salmon et al.
2010; Gaeta et al. 2010). High-throughput sequencing of genomic DNA and targeted sequencing (e.g. Grover et al. 2012) offer new possibilities to differentiate
the effects of these alternatives. Finally, distinction among Spartina homoeologs
will allow more accurate analysis of homoeologous expression in the F1 hybrids
and the allododecaploid using RNA-Seq data for various organs from plants collected in different ecological conditions.
In conclusion, the well-established framework now available for Spartina offers
important opportunities to elucidate the phenotypic, ecological, and genomic consequences of recurrent hybridization and polyploidy. Based on the dramatic increase
in knowledge that has accumulated for various polyploid systems in recent years
(Ainouche and Jenczewski 2010), it has become clear that hybridization and polyploidy generate a range of possible responses that vary among genera. Recent allopolyploidy in S. anglica was not accompanied by rapid restructuring of the parental
genomes as has occurred in other polyploid systems. The major evolutionary events
in S. anglica appear to affect the regulation of gene expression (including epigenetic
regulation); these appear profoundly altered by the merger of different genomes. The
rapid advances of NGS technologies will allow more exhaustive exploration of
highly redundant genomes (which until now suffered from severe technical limitations) such as those of Spartina. These will provide a better understanding of the
genetic and epigenetic mechanisms underlying expression plasticity, and their effect
on adaptive and ecologically relevant functions.
433
UN
Editor Proof
238
Adams KL, Cronn R, Percifield R, Wendel JF (2003) Genes duplicated by polyploidy show
unequal contributions to the transcriptome and organ-specific reciprocal silencing. Proc Natl
Acad Sc USA 100(8):4649–4654
Ainouche ML, Jenczewski E (2010) Focus on polyploidy. New Phytol 186:1–4
Layout: T1 Standard SC
Chapter No.: 12
239
EC
TE
D
PR
OO
F
Ainouche ML, Baumel A, Salmon A (2004a) Spartina anglica Schreb. A natural model system
for analysing early evolutionary changes that affect allopolyploid genomes. Biol J Linn Soc
82:475–484
Ainouche ML, Baumel A, Salmon A, Yannic G (2004b) Hybridization, polyploidy and speciation
in Spartina Schreb (Poaceae). New Phytol 161:165–172
Ainouche ML, Fortune PM, Salmon A, Parisod C, Grandbastien M-A, Fukunaga K, Ricou M,
Misset M-T (2009) Hybridization, polyploidy and invasion: lessons from Spartina (Poaceae).
Biol Invasion. doi:10.1007s10530-0089383-2
Antilla CK, King RA, Ferris C, Ayres DR, Strong DR (2000) Reciprocal hybrid formation of
Spartina in San Francisco Bay. Mol Ecol 9:765–770
Ayres DR, Strong DR (2001) Origin and genetic diversity of Spartina anglica (Poaceae) using
nuclear DNA markers. Am J Bot 88:1863–1867
Ayres DR, Garcia-Rossi D, Davis HG, Strong DR (1999) Extent and degree of hybridization
between exotic (Spartina alterniflora) and native (S. foliosa) cordgrass (Poaceae) in
California, USA determined by randomly amplified polymorphic DNA (RAPDs). Mol Ecol
8:1179–1186
Ayres DR, Smith DL, Zaremba K, Klohr S, Strong DR (2004) Spread of exotic cordgrass and
hybrids (Spartina sp) in the tidal marshes of San-Francisco Bay CA, USA. Biol Invasions
6:221–231
Ayres DA, Zaremba K, Sloop CM, Strong DR (2007) Sexual reproduction of cordgrass hybrids
(Spartina foliosa 9 alterniflora) invading tidal marshes in San Francisco Bay. Divers Distrib
14:187–195
Ayres DR, Grotkopp E, Zaremba C, Sloop CM, Bloom MJ, Bailey JP, Anttila CK, Strong DR
(2008) Hybridization between invasive Spartina densiflora (Poaceae) and native S. foliosa in
San Francisco Bay. Am J Bot 95:713–719
Baumel A, Ainouche ML, Levasseur JE (2001) Molecular investigations in populations of
Spartina anglica C.E. Hubbard (Poaceae) invading coastal Brittany (France). Mol Ecol
10:1689–1701
Baumel A, Ainouche ML, Bayer RJ, Ainouche AK, Misset M-T (2002a) Molecular phylogeny of
hybridizing species from the genus Spartina Schreb. (Poaceae). Mol Phylogenet Evol
22:303–314
Baumel A, Ainouche ML, Kalendar R, Schulman AH (2002b) Retrotransposons and genomic
stability in populations of the young allopolyploid species Spartina anglica C.E. Hubbard
(Poaceae). Mol Biol Evol 19:1218–1227
Baumel A, Ainouche ML, Misset MT, Gourret JP, Bayer RJ (2003) Genetic evidence for
hybridization between the native Spartina maritima and the introduced Spartina alterniflora
(Poaceae) in South-West France: Spartina x neyrautii re-examined. Plant Syst Evol 237:87–97
Bortolus A (2006) The austral cordgrass Spartina densiflora Brong.: its taxonomy, biogeography
and natural history. J Biogeogr 33:158–168
Buggs RJA, Chamala S, Wu W, Gao L, May GD, Schnable PS, et al. (2010) Characterization of
duplicate gene evolution in the recent natural allopolyploid Tragopogon miscellus by next
generation sequencing and Sequenom iPLEX MassARRAY genotyping. Mol Ecol 19:132–146
Buggs RJA, Zhang L, Miles N, Gao L, Wu W, Schnable P, Barbazuk WB, Soltis PS, Soltis DE
(2011) Transcriptomic shock generates evolutionary novelty in a newly formed, natural
allopolyploid plant. Curr Biol 21:551–556
Buggs RJA, Chamala S, Wu W, Tate JA, Schnable PS, Soltis DS, Soltis PS, Barbazuk WB (2012)
Rapid, repeated, and clustered loss of duplicated genes in allopolyploid Tragopogon
populations of independent origin. Curr Biol 22:248–252
Castillo JM, Ayres DR, Leira-Doce1 P, Bailey J, Blum M, Strong DR, Luque T and Figueroa E
(2010) The production of hybrids with high ecological amplitude between exotic Spartina
densiflora and native S. maritima in the Iberian Peninsula. Diversity and Distributions, 1–12
doi: 10.1111/j.1472-4642.2010.00673.x
Chalupska D, Lee HY, Faris JD, Evrard A, Chalhoub B, Kaselkorn R, Gorniki P (2008) Acc
Homeoloci and the evolution of wheat genomes. Proc Natl Acad Sci U S A 105:9691–9696
CO
RR
474
475
476
477
478
479
480
481
482
483
484
485
486
487
488
489
490
491
492
493
494
495
496
497
498
499
500
501
502
503
504
505
506
507
508
509
510
511
512
513
514
515
516
517
518
519
520
521
522
523
524
525
526
527
Book ISBN: 978-3-642-31441-4
Page: 239/242
Polyploid Evolution in Spartina
UN
Editor Proof
12
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 12
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 240/242
EC
TE
D
PR
OO
F
Chang PL, Dilkes B, McMahon M, Comai L, Nuzhdin SV (2010) Homoeolog-specific retention
and use in allotetraploid Arabidopsis suecica depends on parent of origin and network
partners. Genome Biol 11:R125. doi:10.1186/gb-2010-11-12-r125
Chaudhary B, Flagel L, Stupar RM, Udall JA, Verma N, Springer NM, Wendel JF (2009)
Reciprocal silencing, transcriptional bias and functional divergence of homeologs in
polyploid cotton (Gossypium). Genetics 182: 503–517
Chelaifa H, Mahe F, Ainouche M (2010a) Transcriptome divergence between the hexaploid saltmarsh sister species Spartina maritima and Spartina alterniflora (Poaceae). Mol Ecol
19:2050–2063
Chelaifa H, Monnier A, Ainouche M (2010b) Transcriptomic changes following recent natural
hybridization and allopolyploidy in the salt marsh species Spartina x townsendii and Spartina
anglica (Poaceae). New Phytol 186:161–174
Chen ZJ (2007) Genetic and epigenetic mechanisms for gene expression and phenotypic variation
in plant polyploids. Annu Rev Plant Biol 58:377–406
Chester M, Gallagher JP, Symonds VV, Veruska Cruz da Silva A, da Silva A, Mavrodiev EV,
Leitch AR, Soltis PS, Soltis DE (2012) Extensive chromosomal variation generated in a
recently formed natural allopolyploid species, Tragopogon miscellus (Asteraceae). Proc Natl
Acad Sci U S A 109:1176–1181
Christin P-A, Besnard G, Samaritani E, Duvall MR, Hodkinson TR, Savolainend V, Salamini N
(2008) Oligocene CO2 Decline Promoted C4 Photosynthesis in Grasses. Curr Biol 18:37–43
Cifuentes M, Grandont L, Moore G, Chèvre AM, Jenczewski E (2010) Genetic regulation of
meiosis in polyploid species: new insights into an old question. New Phytol 186:37–45
Civille JC, Sayce K, Smith SD, Strong DR (2005) Reconstructing a century of Spartina
alterniflora invasion with historical records and contemporary remote sensing. Ecoscience
12:330–338
Columbus JT, Cerros-Tlatilpa R, Kinney MS, Siqueiros-Delgado M-E, Bell HL, Griffith MP,
Refulio-Rodrigez NF (2007) Phylogenetics of Chloridoideae (Gramineae): a preliminary study
based on nuclear internal transcribed spacer and chloroplast trnL-F sequences. Aliso 23:565–579
Corbière L (1926) La Spartine de Townsend en Normandie. Bulletin de la société Linéenne de
Normandie, 7e série 9:92–117
Crooks JA (2002) Characterizing ecosystem-level consequences of biological invasions: the role
of ecosystem engineers. Oikos 97:153–166
Daehler CC, Strong DR (1997) Hybridization between introduced smooth cordgrass (Spartina
alterniflora; Poaceae) and native California cordgrass (S. foliosa) in San Francisco Bay,
California. U S A Am J Bot 81:307–313
Doyle JJ, Negan EN (2009) Dating the origins of polyploidy events. New Phytol 186:73–85
Doyle JJ, Flagel LE, Paterson AH, Rapp RA, Soltis DE, Soltis PS, Wendel JF (2008)
Evolutionary genetics of genome merger and doubling in plants. Annu Rev Genet 42:443–461
Ferreira de Carvalho J, Poulain J, Da Silva C, Wincker P, Michon-Coudouel S, Dheilly A, Naquin
D, Boutte J, Salmon A, Ainouche M. Transcriptome de novo assembly and comparative
analysis of the hexaploid salt marsh species Spartina maritima and Spartina alterniflora
(Poaceae) using high-throughput 454 Roche pyrosequencing. Heredity, submitted
Flagel LE, Wendel JF (2010) Evolutionary rate variation, genomic dominance and duplicate gene
expression evolution during allotetraploid cotton speciation. New Phytol 186:184–193
Flagel L, Udall J, Nettleton D, Wendel JF (2008) Duplicate gene expression in allopolyploid
Gossypium reveals two temporally distinct phases of expression evolution. BMC Biol 6:16
Flagel LE, Chen L, Chaudhary B, Wendel JF (2009) Coordinated and fine-scale control of
homoeologous gene expression in allotetraploid cotton. J Hered 100:487–490
Fortuné PM, Schierenbeck K, Ainouche A, Jacquemin J, Wendel JF, Ainouche ML (2007)
Evolutionary dynamics of waxy and the origin of hexaploid Spartina species. Mol Phylogenet
Evol 43:1040–1055
Fortuné PM, Schierenbeck K, Ayres D, Bortolus A, Clatrice O, Ainouche ML (2008) The
enigmatic invasive Spartina densiflora: a history of hybridizations in a polyploidy context.
Mol Ecol 17:4304–4316
CO
RR
528
529
530
531
532
533
534
535
536
537
538
539
540
541
542
543
544
545
546
547
548
549
550
551
552
553
554
555
556
557
558
559
560
561
562
563
564
565
566
567
568
569
570
571
572
573
574
575
576
577
578
579
580
581
M. Ainouche et al.
UN
Editor Proof
240
Layout: T1 Standard SC
Chapter No.: 12
241
EC
TE
D
PR
OO
F
Foucaud (1897) Un Spartina inédit. Ann Soc Sci Nat Char Inf 32:220–222
Gaeta RT, Pires J (2010) Homoeologous recombination in allopolyploids: the polyploidy rachet.
New Phytol 186:18–27
Gaeta RT, Pires JC, Iniguez-Luy F, Leon E, Osborn TC (2007) Genomic changes in
resynthesized Brassica napus and their effect on gene expression and phenotype. Plant Cell
19:3403–3417
Gaut BS (2002) Evolutionary dynamics of grass genomes. New Phytol 154:15–28
Gedye K, Gonzalez-Hernandez J, Ban Y, Thimmapuram J, Sun F, Wright C, Ali S, Boe A,
Owens V (2010) Investigation of the transcriptome of prairie cordgrass, a new cellulosic
biomass crop. The Plant Genome 3, 2:69–80
Grass Phylogeny Working Group (GPWG) (2001) Phylogeny and subfamilial classification of the
grasses (Poaceae). Ann Missouri Bot Gard 88:373–457
Gray AJ, Benham PEM, Raybould AF (1990) Spartina anglica-the evolutionary and ecological
background. In: Gray AJ, Benham PEM (eds) Spartina anglica-a research review. Institute of
terrestrial ecology, Natural environment research council, pp 5–10
Grover CE, Salmon A, and Wendel JF (2012) Targeted sequence capture as a powerful tool for
evolutionary analysis. Am J Bot (in press)
Groves H, Groves J (1880) Spartina townsendii nobis. Rep Bot Soc Exch Club Br Id 1:37
Guénégou MC, Levasseur JE (1993) La nouvelle espèce amphidiplo Spartina anglica C.E.
Hubbard: son origine, argumentation et implications. Biogeographica 69:125–133
Guénégou MC, Citharel J, Levasseur JE (1988) The hybrid status of Spartina anglica (Poaceae).
Enzymatic analysis of the species and the presumed parents. Can J Bot 66:1830–1833
Hegarty MJ, Barker GL, Wilson ID, Abbott RJ, Edwards KJ, Hiscock SJ (2006) Transcriptome
shock after Interspecific hybridization in Senecio is ameliorated by genome duplication. Curr
Biol 16:1652–1659
Hilu KW, Alice LA (2001) A phylogeny of Chloridoideae (Poaceae) based on matK sequences.
Syst Bot 26:386–405
Hubbard JCE (1968) Grasses, 2nd edn. Penguin Books, London
Huskin CL (1930) The origin of S. x townsendii. Genetica 12:531–538
IUCN (2000) World’s worst invasive alien species. In: IUCN The World Conservation Union.
http://iucn.org
Jovet P (1941) Notes systématiques et écologiques sur les Spartines du Sud-Ouest. Bull Soc Bot
Fr 88:115–123
Kim S, Rayburn AL, Lee DK (2010) Genome size and chromosome analyses in prairie cordgrass.
Crop Sci 50:2277–2282
Kocsis MG, Hanson AD (2000) Biochemical evidence for two novel enzymes in the biosynthesis
of 3-dimethylsulphoniopropionate in Spartina alterniflora. Plant Physiol 123:1153–1161
Kocsis MG, Nolte KD, Rhodes D, Shen TL, Gage DA, Hanson AD (1998) Dimethylsulfoniopropionate biosynthesis in Spartina alterniflora. Plant Physiol 117:273–281
Koh J, Soltis P, Soltis DE (2010) Homoeolog loss and expression changes in natural populations
of the recently formed allotetraploid Tragopogon mirus (Asteraceae). BMC Genomics 11:97
Lambrinos G (2008) Managing invasive ecosystem engineers: the case of Spartina in Pacific
estuaries. Theor Ecol Ser 4:299–322
Larher F, Hamelin J, Steward GR (1977) L’acide diméthylsulphonium-3-propano de Spartina
anglica. Phytochemistry 16:2019–2020
Le Comber SC, Ainouche ML, Kovarik A, Leitch AR (2010) Making a functional diploid: from
polysomic to disomic inheritance. New Phytol 186:113–122
Lee RW (2003) Physiological adaptations of the invasive cordgrass Spartina anglica to reducing
sediments: rhizome metabolic gas fluxes and enhanced O2 and H2S transport. Mar Biol
143:9–15
Lim KY, Soltis DE, Soltis PS, Tate J, Matyasek R, Srubarova H, Kovarik A, Pires JC, Xiong Z,
Leitch AR (2008) Rapid chromosome evolution in recently formed polyploids in Tragopogon
(Asteraceae). PLoS ONE 3(10):e3353. doi:10.1371/journal.pone.0003353
CO
RR
582
583
584
585
586
587
588
589
590
591
592
593
594
595
596
597
598
599
600
601
602
603
604
605
606
607
608
609
610
611
612
613
614
615
616
617
618
619
620
621
622
623
624
625
626
627
628
629
630
631
632
633
634
Book ISBN: 978-3-642-31441-4
Page: 241/242
Polyploid Evolution in Spartina
UN
Editor Proof
12
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 12
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 242/242
EC
TE
D
PR
OO
F
Marchant CJ (1963) Corrected chromosome numbers for Spartina x townsendii and its parent
species. Nature 199:929
Marchant CJ (1967) Evolution in Spartina (Gramineae): I. the history and morphology of the
genus in Britain. Bot J Linn Soc 60(381):1–24
Marchant CJ (1968a) Evolution in Spartina (Graminae). III species chromosome numbers and
their taxonomic signifiance. Bot J Linn Soc 60(383):411–417
Marchant CJ (1968b) Evolution in Spartina (Graminae) II. chromosome basic relationships and
the problem of S. x towsendii agg. Bot J Linn Soc 60(383):381–409
Marchant CJ (1977) Hybrid characteristics in Spartina x neyrautii Fouc., a taxon rediscovered in
northern Spain. Bot J Lin Soc. 74:289–296
Maricle BR, Lee RW (2002) Aerenchyma development and oxygen transport in the estuarine
cordgrasses Spartina alterniflora and S. anglica. Aquat Bot 74:109–120
Maricle BR, Crosier JJ, Bussiere BC, Lee RW (2006) Respiratory enzyme activities correlate
with anoxia tolerance in saltmarsh grasses. J Exp Mar Biol Ecol 337:30–37
Maricle BR, Koteyeva NK, Voznesenskaya EV, Thomasson JR, Edwards GE (2009) Diversity in
leaf anatomy, and stomatal distribution and conductance, between salt marsh and freshwater
species in the C4 genus Spartina (Poaceae). New Phytol 184:216–233
Mobberley DG (1956) Taxonomy and distribution of the genus Spartina. Iowa State Coll J Sci
30:471–574
Nasrallah JB, Liu P, Sherman-Broyles S, Schmidt R, Nasrallah ME (2007) Epigenetic
mechanisms for breakdown of self-incompatibility in interspecific hybrids. Genetics
175:1965–1973
Osborn TC, Pires JC, Birchler JA, Auger DL, Chen ZJ, Lee HS, Comai L, Madlung A, Doerge
RW, Colot V, Martienssen RA (2003) Understanding mechanisms of novel gene expression in
polyploids. Trends Genet 19:141–147
Otte ML, Wilson G, Morris JT, Moran BM (2004) Dimethylsulphoniopropionate (DMSP) and
related compounds in higher plants. J Exp Bot 55:919–925
Otto SP (2007) The evolutionary consequences of polyploidy. Cell 131:452–462
Ozkan H, Levy AA, Feldman M (2001) Allopolyploidy-induced rapid genome evolution in the
wheat (Aegilops-Triticum) group. Plant Cell 13:1735–1747
Parisod C, Salmon A, Zerjal T, Tenaillon M, Grandbastien M-A, Ainouche M (2009) Rapid
structural and epigenetic reorganization near transposable elements in hybrid and allopolyploid genomes in Spartina. New Phytol 184:1003–1015
Parisod C, Alix K, Just J, Petit M, Sarilar V, Mhiri C, Ainouche M, Chalhoub B, Grandbastien
MA (2010) Impact of transposable elements on the organization and function of allopolyploid
genomes. New Phytol 186:37–45
Prasad V, Stroemberg CAE, Alimohammadian H, Sahni A (2005) Dinosaur coprolites and the
early evolution of grasses and grazers. Science 310:1177–1180
Prasad V, Stroemberg CAE, Leaché AD, Samant B, Patnaik R, Tang L, Mohabey DM, Ge S,
Sahni A (2011) Late Cretaceous origin of the rice tribe provides evidence for early
diversification in Poaceae. Nat Communication 2:480. doi:10.1038/ncomms1482
Raybould AF, Gray AJ, Lawrence MJ, Marshall DF (1991a) The evolution of Spartina anglica
C.E. Hubbard (Gramineae): genetic variation and status of the parental species in Britain. Biol
J Linn Soc 44:369–380
Raybould AF, Gray AJ, Lawrence MJ, Marshall DF (1991b) The evolution of S. anglica C.E.
Hubbard (Gramineae): origin and genetic variability. Biol J Linnean Soc 43:111–126
Renny-Byfield S, Ainouche M, Leitch IJ, Lim KY, Le Comber SC, Leitch AR (2010) Flow
cytometry and GISH reveal mixed ploidy populations and Spartina nonaploids with genomes
of S. alterniflora and S. maritima origin. Ann Bot 105:527–533
Saint-Yves A (1932) Monographia Spartinarum. Cand 5:19–100
Salmon A, Ainouche ML, Wendel JF (2005) Genetic and epigenetic consequences of recent
hybridization and polyploidy in Spartina (Poaceae). Mol Ecol 14:1163–1175
Salmon A, Flagel L, Ying B, Udall JA, Wendel JF (2010) Homoeologous nonreciprocal
recombination in polyploid cotton. New Phytol 186:123–134
CO
RR
635
636
637
638
639
640
641
642
643
644
645
646
647
648
649
650
651
652
653
654
655
656
657
658
659
660
661
662
663
664
665
666
667
668
669
670
671
672
673
674
675
676
677
678
679
680
681
682
683
684
685
686
687
688
M. Ainouche et al.
UN
Editor Proof
242
Layout: T1 Standard SC
Chapter No.: 12
243
EC
TE
D
PR
OO
F
Skalická K, Lim KY, Matyasek R, Matzke M, Leitch AR, Kovarik A (2005) Preferential
elimination of repeated DNA sequences from the paternal, Nicotiana tomentosiformis genome
donor of a synthetic, allotetraploid tobacco. New Phytol 166:291–303
Sloop C, Ayres DR, Strong DR (2009) The rapid evolution of self-fertility in Spartina
hybrids (Spartina alterniflora 9 foliosa) invading San Francisco Bay, CA. Biol Invasions
11:1131–1144
Slotkin K, Martienssen R (2007) Transposable elements and the epigenetic regulation of the
genome. Nat Rev Genet 8:272–285
Soltis DE et al (2013) Polyploidy and genome evolution. In: Soltis PS, Soltis DE (eds) The early
stages of polyploidy: rapid and repeated evolution in Tragopogon. Springer, Heidelberg
Stebbins GL (1950) Variation and evolution in plants. Columbia University Press, New York
Straub SCK, Pfeil BE, Doyle JJ (2003) Testing the polyploid past of soybean using a low-copy
nuclear gene-Is Glycine (Fabaceae: Papilionoideae) an auto- or allopolyploid? Mol
Phylogenet Evol 39:580–584
Szadkowski E, Eber F, Huteau V, Lode M, Huneau C, Belcram H, Coriton O, ManzanaresDauleux MJ, Delourme R, King GJ et al (2010) The first meiosis of resynthesized Brassica
napus, a genome blender. New Phytol 186:102–112
Tate JA, Joshi P, Soltis KA, Soltis PS, Soltis DE (2009) On the road to diploidization?
Homoeolog loss in independently formed populations of the allopolyploid Tragopogon
miscellus (Asteraceae). BMC Plant Biol 9:80. doi:10.1186/1471-2229-9-80
Thompson JD, McNeilly T, Gray AJ (1991) Population variation in Spartina anglica C.
E. Hubbard. I. Evidence from a common garden experiment. New Phytol 117:115–128
Triplet P, Gallicé A (2008) Les plantes envahissantes du littoral atlantique: le cas de la Spartine
anglaise (Spartina anglica). Aestuaria 13 Aestuarium– Le Forum des Marais Atlantiques eds
Udall JA, Swanson JM, Nettleton D, Percifield RJ, Wendel JF (2006) A novel approach for
characterizing expression levels of genes duplicated by polyploidy. Genetics 173:1823–1827
Van de Peer Y, Maere S, Meyer A (2009) OPINION the evolutionary significance of ancient
genome duplications. Nat Rev Genet 10:725–732
Warner DA, Edwards GE (1993) Effects of polyploidy on photosynthesis. Photosynthetis
research 35:135–147
Wendel JF (2000) Genome evolution in polyploids. Plant Mol Biol 42:225–249
Wolfe KH, Gouy M, Yang Y-W, Sharpt PM, Li W-H (1989) Date of the monocot-dicot
divergence estimated from chloroplast DNA sequence data. Proc Natl Acad Sci U S A
86:6201–6205
Yannic G, Baumel A, Ainouche ML (2004) Uniformity of the nuclear and chloroplast genomes of
Spartina maritima (Poaceae) a salt marshes species in decline along the Western European
Coast. Heredity 93:182–188
CO
RR
689
690
691
692
693
694
695
696
697
698
699
700
701
702
703
704
705
706
707
708
709
710
711
712
713
714
715
716
717
718
719
720
721
722
723
724
725
Book ISBN: 978-3-642-31441-4
Page: 243/242
Polyploid Evolution in Spartina
UN
Editor Proof
12
Book ID: 272454_1_En
Date: 16-8-2012
Metadata of the chapter that will be visualized in
SpringerLink
Book Title
Polyploidy and Genome Evolution
Series Title
Chapter Title
Allopolyploid Speciation in Action: the Origins and Evolution of Senecio cambrensis
Copyright Year
2012
Copyright HolderName
Springer-Verlag Berlin Heidelberg
Corresponding Author
Family Name
Hegarty
Particle
Given Name
Matthew J.
Suffix
Author
Division
Institute of Biological, Environmental and Rural Sciences
Organization
Aberystwyth University
Address
Penglais Campus, SY23 3DA, Aberystwyth, Ceredigion, UK
Email
ayh@aber.ac.uk
Family Name
Abbott
Particle
Given Name
Richard J.
Suffix
Author
Division
School of Biology
Organization
University of St. Andrews
Address
Harold Mitchell Building, KY16 9TH, St. Andrews, Fife, UK
Email
rja@st-andrews.ac.uk
Family Name
Hiscock
Particle
Given Name
Simon J.
Suffix
Abstract
Division
School of Biological Sciences
Organization
University of Bristol
Address
Woodland Road, BS8 1UG, Bristol, UK
Email
simon.hiscock@bristol.ac.uk
Senecio cambrensis is one of a few allopolyploid plant species known to have originated in the recent past
and, therefore, provides excellent material for analysing allopolyploid speciation. This allohexaploid species
originated in the UK within the last 100 years following hybridization between diploid S. squalidus and
tetraploid S. vulgaris. In this chapter, we first describe the events leading up to hybridization between these
two species, focusing mainly on the origin and spread of S. squalidus in the UK. We then consider alternative
pathways by which S. cambrensis might have originated and conclude that current evidence suggests an origin
via formation of the triploid hybrid (S. x baxteri) followed by chromosome doubling. We next review our
investigations into levels of genetic diversity and also changes to gene expression and the possible causes of
this (epigenetic effects) during the origin of S. cambrensis. High levels of genetic diversity, assessed by
surveys of allozyme and AFLP variation, have been recorded in S. cambrensis, and it is likely that
intergenomic recombination was an important generator of this diversity. Our studies of ‘resynthesized’ S.
cambrensis have shown that the initial genome merger (hybridization) producing S. x baxteri generates
genome-wide, non-additive alterations to parental patterns of gene expression and DNA methylation, with
genome duplication resulting in a secondary burst of both transcriptional and epigenetic modification. In
synthetic allohexaploid lines of S. cambrensis phenotypic changes become apparent from the second to fifth
generations, possibly as a consequence of recombination or epigenetic effects; these include changes in ray
flower form and emergence of self-incompatible individuals. We conclude by considering the future of S.
cambrensis from the standpoint of it being a model species for further study of allopolyploid speciation, and
second its long-term success in the wild. Ongoing work to produce a draft reference genome for S.
squalidus will underpin future research in S. cambrensis, enabling a more thorough survey of changes to DNA
methylation, small RNA activity and promoter binding in the hybrids, as well as comparison with the related
allotetraploid S. eboracensis to determine the effects of genome dosage. The future of the species in the wild
is currently uncertain. The population in Edinburgh that represented a separate origin of the species in the
wild during the 1970s is now extinct, and there has been a marked decline in the number of populations and
individuals of the species in its heartland, North Wales, since the 1980s. An analysis of how its ecology
compares with those of its parents is lacking. However, it appears to share the same habitats in the wild with
its parents, which might have contributed to its decline. Although S. cambrensis may become extinct in the
wild in the near future, the potential will remain for it to originate again in the UK providing that conditions
prevail for its parents to hybridize.
Book ISBN: 978-3-642-31441-4
Page: 245/270
Chapter 13
5
Matthew J. Hegarty, Richard J. Abbott and Simon J. Hiscock
9
10
11
12
13
14
15
16
17
18
19
20
21
22
D
8
Abstract Senecio cambrensis is one of a few allopolyploid plant species known to
have originated in the recent past and, therefore, provides excellent material for
analysing allopolyploid speciation. This allohexaploid species originated in the
UK within the last 100 years following hybridization between diploid S. squalidus
and tetraploid S. vulgaris. In this chapter, we first describe the events leading up to
hybridization between these two species, focusing mainly on the origin and spread
of S. squalidus in the UK. We then consider alternative pathways by which S.
cambrensis might have originated and conclude that current evidence suggests an
origin via formation of the triploid hybrid (S. x baxteri) followed by chromosome
doubling. We next review our investigations into levels of genetic diversity and
also changes to gene expression and the possible causes of this (epigenetic effects)
during the origin of S. cambrensis. High levels of genetic diversity, assessed by
surveys of allozyme and AFLP variation, have been recorded in S. cambrensis, and
it is likely that intergenomic recombination was an important generator of this
diversity. Our studies of ‘resynthesized’ S. cambrensis have shown that the initial
genome merger (hybridization) producing S. x baxteri generates genome-wide,
non-additive alterations to parental patterns of gene expression and DNA
TE
7
EC
6
CO
RR
3
PR
OO
4
Allopolyploid Speciation in Action:
the Origins and Evolution of Senecio
cambrensis
2
F
1
Book ID: 272454_1_En
Date: 16-8-2012
M. J. Hegarty (&)
Institute of Biological, Environmental and Rural Sciences, Aberystwyth University,
Penglais Campus, Aberystwyth, Ceredigion SY23 3DA, UK
e-mail: ayh@aber.ac.uk
R. J. Abbott
School of Biology, University of St. Andrews, Harold Mitchell Building,
St. Andrews, Fife KY16 9TH, UK
e-mail: rja@st-andrews.ac.uk
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: 13
S. J. Hiscock
School of Biological Sciences, University of Bristol,
Woodland Road, Bristol BS8 1UG, UK
e-mail: simon.hiscock@bristol.ac.uk
P. S. Soltis and D. E. Soltis (eds.), Polyploidy and Genome Evolution,
DOI: 10.1007/978-3-642-31442-1_13, Springer-Verlag Berlin Heidelberg 2012
245
Layout: T1 Standard SC
Chapter No.: 13
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 246/270
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
F
26
PR
OO
25
methylation, with genome duplication resulting in a secondary burst of both
transcriptional and epigenetic modification. In synthetic allohexaploid lines of S.
cambrensis phenotypic changes become apparent from the second to fifth generations, possibly as a consequence of recombination or epigenetic effects; these
include changes in ray flower form and emergence of self-incompatible individuals. We conclude by considering the future of S. cambrensis from the standpoint
of it being a model species for further study of allopolyploid speciation, and
second its long-term success in the wild. Ongoing work to produce a draft reference genome for S. squalidus will underpin future research in S. cambrensis,
enabling a more thorough survey of changes to DNA methylation, small RNA
activity and promoter binding in the hybrids, as well as comparison with the
related allotetraploid S. eboracensis to determine the effects of genome dosage.
The future of the species in the wild is currently uncertain. The population in
Edinburgh that represented a separate origin of the species in the wild during the
1970s is now extinct, and there has been a marked decline in the number of
populations and individuals of the species in its heartland, North Wales, since the
1980s. An analysis of how its ecology compares with those of its parents is
lacking. However, it appears to share the same habitats in the wild with its parents,
which might have contributed to its decline. Although S. cambrensis may become
extinct in the wild in the near future, the potential will remain for it to originate
again in the UK providing that conditions prevail for its parents to hybridize.
D
24
TE
23
M. J. Hegarty et al.
44
13.1 Introduction
46
13.1.1 General Introduction
48
49
50
51
52
53
54
55
56
57
58
59
60
61
Polyploidization (genome duplication) is an important evolutionary process in plants
(Grant 1981) that appears to have accompanied major transitions in land plant
evolution, including the evolution of the seed habit and the evolution of angiosperms
(Jiao et al. 2011). All angiosperms are thought to have a polyploid ancestry (Jiao et al.
2011) but most of these are paleopolyploids that now function essentially as diploids.
Nevertheless, there are also numerous examples of recently formed polyploids
(Adams and Wendel 2005; Wood et al. 2009). Most recent polyploids have formed in
association with interspecific hybridization (Grant 1981; Leitch and Bennett 1997;
Soltis and Soltis 1999; Otto and Whitton 2000; Leitch and Leitch 2008; Hegarty and
Hiscock 2008)—allopolyploidy, which is now recognised as perhaps the most
important mechanism of abrupt speciation in plants (Grant 1981; Leitch and Leitch
2008; Hegarty and Hiscock 2008). Allopolyploidy confers rapid fertility to hybrids
because duplicated parental chromosomes pair ‘normally’ during meiosis; it also
confers reproductive isolation of the hybrids from their parental species because of
aberrant or failed chromosome pairing (Soltis and Soltis 1999; Rieseberg et al. 2003;
AQ1
CO
RR
47
EC
45
UN
Editor Proof
246
AQ2
Layout: T1 Standard SC
Chapter No.: 13
Book ISBN: 978-3-642-31441-4
Page: 247/270
Allopolyploid Speciation in Action
247
73
13.1.2 The Genus Senecio and the Origins of UK Allopolyploids
71
74
75
76
77
78
79
80
81
82
83
84
85
86
87
88
89
90
91
92
93
94
95
96
97
98
PR
OO
70
The genus Senecio (Asteraceae), which includes ragworts and groundsels, is one of
the largest and most morphologically diverse genera of flowering plants. With
between 1000 and 3000 species, the genus has a worldwide distribution with species
described from almost every land mass on Earth (Vincent 1996). Recent revisions,
following molecular phylogenetic analysis, of the genus have assigned many former
Senecio species to other genera leaving a conservative 1200 species of Senecio sensu
stricto (Pelser et al. 2007). Within the genus there are numerous examples of
hybridization and polyploidy, and many species have been proposed to have an
allopolyploid origin (Abbott and Lowe 1996; Coleman et al. 2001; Kadereit et al.
2006; Pelser et al. 2012). In the UK, three new polyploid taxa have arisen within the
last 100 years as a consequence of hybridization between native tetraploid Senecio
vulgaris (common groundsel) and the introduced invasive diploid species S.
squalidus. The origins of these new polyploid taxa (see below) provide one of the best
examples of ‘evolution in action’ (Abbott and Lowe 1996, Hegarty and Hiscock
2008). The introduction and rapid spread of alien S. squalidus across the UK was the
main catalyst for this burst of hybrid speciation, which was further facilitated by the
obligate outcrossing mating system of S. squalidus increasing the frequency of
interspecies pollinations (Brennan and Hiscock 2010).
D
68
69
TE
66
67
EC
64
65
CO
RR
63
F
72
Rieseberg and Willis 2007). Numerous examples of allopolyploid speciation have
been described in the literature, including at least six new plant species that have
arisen in the last century: Tragopogon mirus and T. miscellus (Novak et al. 1991,
Chap. 14, this volume); Spartina anglica (Gray et al. 1991, Chap. 12, this volume),
Senecio cambrensis and S. eboracensis (Ashton and Abbott 1992, Abbott and Lowe
1996, 2004) and Cardamine schulzii (Urbanska et al. 1997). In this chapter, we
review the origins of the Senecio allopolyploid species S. cambrensis (Welsh
groundsel). We show how resynthesised forms of this allopolyploid can reveal
important insights into the genetic and genomic consequences of allopolyploidization and how parental traits, both morphological and physiological, recombine in
neopolyploids.
62
13.1.3 Oxford Ragwort in the British Isles: Introduction
and Spread
UN
Editor Proof
13
Book ID: 272454_1_En
Date: 16-8-2012
Senecio squalidus (2n = 2x = 20), commonly known as Oxford ragwort, is itself
a recently evolved (homoploid) hybrid species (James and Abbott 2005; Abbott
et al. 2010; Brennan et al. 2012) that originated on Mount Etna, Sicily, as a result
of hybridization between S. aethnensis (2n = 2x = 20, a Mount Etna endemic of
higher altitudes) and S. chrysanthemifolius (2n = 2x = 20, a native Sicilian
AQ3
Layout: T1 Standard SC
Chapter No.: 13
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 248/270
106
107
108
109
110
111
112
113
114
115
116
117
118
119
120
121
122
123
124
125
126
127
128
129
130
131
132
133
134
135
136
137
138
139
F
PR
OO
105
D
104
TE
103
EC
101
102
species of lower altitudes). At mid-altitudes on the volcano the distribution of
these species overlaps, leading to the formation of a stable hybrid zone (Brennan
et al. 2009). Material from this hybrid zone was introduced to the Oxford Botanic
Garden in the early 1700s, but records of the collection locality or localities,
methods of cultivation, and numbers of plants cultivated in the garden have been
lost. Plants subsequently escaped from the Botanic Garden and colonised the
masonry of the old college walls from the end of the eighteenth century (Harris
2002). During the industrial revolution of the nineteenth and early twentieth
centuries S. squalidus moved rapidly out of Oxford by colonising the clinker beds
of the expanding UK railway, of which Oxford was a key hub. The chronology of
this famous plant invasion is meticulously documented in a set of papers by Kent
(reviewed in Abbott et al. 2009), which record that it began to spread northwards
in the late nineteenth century, reaching different parts of northern England during
the middle of the twentieth century, before becoming established in the Central
Belt of Scotland by the mid-1950s. Today, S. squalidus continues to spread north
in Scotland and across Northern Ireland.
The rapidity of this invasion is intriguing because S. squalidus is self-incompatible (Abbott and Forbes 1993; Hiscock 2000a, b, and obligate outcrossers are generally thought not to make good colonizers or invasives (Baker 1967). Most invasive
species tend to have uniparental reproduction, either sexual (selfing) or asexual
(apomixis or vegetative reproduction), although there are exceptions, most notably
other species of Asteraceae, such as yellow star thistle, Centaurea solstitialis (Sun
and Ritland 1998). According to Baker’s ‘rule’ (Baker 1967), successful colonising
and invasive plants are usually self-fertile (self-compatible [SC]) (Stebbins 1957;
Baker 1967). Studies of the mating system of S. squalidus, however, have shown that
individuals exhibit strong self-incompatibility (SI) across its entire British range,
and, as in other species of Asteraceae, this SI is regulated sporophytically by a single
polymorphic S locus (Hiscock 2000a, b; Brennan et al. 2002, 2005, 2006). The
finding of strong SI in S. squalidus is intriguing, particularly in the light of the
extreme population bottleneck that its ancestors must have experienced during its
introduction and early colonisation. Following a population bottleneck, allelic
diversity at the S locus will be lowered and opportunities for mating (between
individuals carrying different S alleles) correspondingly reduced (Hiscock 2000b;
Brennan et al. 2002). An extensive survey of SI in S. squalidus across the UK showed
that a combination of substantial between-population sharing of the seven S alleles
contained in the entire UK population and low levels of selfing (‘pseudo-self-compatibility’) were the most likely cause of S. squalidus’ reproductive success as a
colonizer (Brennan et al. 2005, 2006).
CO
RR
99
100
M. J. Hegarty et al.
UN
Editor Proof
248
13.1.4 The Origins of Senecio cambrensis
During its spread across the UK, S. squalidus (2n = 2x = 20) hybridized with the
self-compatible native groundsel, S. vulgaris (2n = 4x = 40), resulting in the recent
Layout: T1 Standard SC
Chapter No.: 13
Book ISBN: 978-3-642-31441-4
Page: 249/270
Allopolyploid Speciation in Action
249
PR
OO
F
Editor Proof
13
Book ID: 272454_1_En
Date: 16-8-2012
146
147
148
149
150
151
152
153
154
155
156
157
158
159
160
161
162
163
164
165
TE
EC
144
145
CO
RR
142
143
origin of three hybrid taxa. These are the allohexaploid S. cambrensis
(2n = 6x = 60), the recombinant tetraploid S. eboracensis (2n = 4x = 40), and the
stabilized introgressant radiate form of S. vulgaris, S. vulgaris var. hibernicus
(2n = 4x = 40). Interestingly, all three new hybrid taxa are self-compatible, suggesting that this SC mating system, inherited from S. vulgaris, is ‘dominant’ over the
SI mating system present in S. squalidus. Detailed descriptions of these new taxa and
what is known about their origins are presented elsewhere (see Abbott et al. 1992;
Lowe and Abbott 2003; Abbott and Lowe, 2004; Kim et al. 2008).
Here, we briefly summarise the information available on the origin of S. cambrensis (Fig. 13.1) focussing particularly on the possible pathways of its origin.
Knowing the pathway of origin of a polyploid taxon is helpful in regard to
understanding the species’ potential to generate genetic diversity during its initial
stages of development, and also for accurate production of synthetic forms of the
polyploid used to study possible genetic and epigenetic changes that occurred in
the taxon immediately following its origin in the wild (Hegarty et al. 2006, 2008,
2011; Lukens et al. 2006; Buggs et al. 2009).
Senecio cambrensis was described by Rosser (1955) from material provided by
H.E. Green, who first observed the plant growing at Ffrith and Ceffn-y-bedd, North
Wales, UK, in 1948. The plant was described as an annual or short-lived perennial
herb that was hexaploid with flower heads (capitula) containing ray florets having
short ligules (*4.8 mm in length). Rosser (in Crisp 1972) later determined a herbarium specimen, collected in Denbigh, North Wales in 1925 and originally named
as S. squalidus x S. vulgaris, to be S. cambrensis. However, in the absence of a
chromosome count there remains some doubt as to whether this specimen is
S. cambrensis or, alternatively, a fertile hybrid of S. squalidus and S. vulgaris
(see below). Fertile, hexaploid plants with similar morphology to the wild form of
UN
140
141
D
Fig. 13.1 The neoallohexaploid Senecio cambrensis (centre) flanked by its parents, tetraploid S.
vulgaris (left) and diploid S. squalidus (right). Here, S. cambrensis has flower heads (capitula) of
intermediate type to its parents; however, its ray florets can vary in length, and occasionally nonradiate forms (lacking ray florets) are also found in the wild
Layout: T1 Standard SC
Chapter No.: 13
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 250/270
174
175
176
177
178
179
180
181
182
183
184
185
186
187
188
189
190
191
192
193
194
195
196
197
198
199
200
201
202
203
204
205
206
207
208
209
210
F
PR
OO
172
173
D
170
171
TE
169
EC
168
S. cambrensis can be produced by treating synthetic triploid hybrids between S.
squalidus and S. vulgaris with colchicine (Harland 1955, Weir and Ingram 1980,
Hegarty et al. 2005). On this basis, Rosser (1955) concluded that S. cambrensis was a
new species that originated by hybridization between native S. vulgaris and introduced S. squalidus followed by chromosome doubling. The species is likely to have
originated shortly before it was first recorded in North Wales and after S. squalidus
had spread to the region in the early part of the twentieth century (Kent 1963).
In 1982, S. cambrensis was found growing in Edinburgh, UK (Abbott et al.
1983), and subsequent molecular analysis involving surveys of allozyme and
chloroplast DNA variation showed that it had originated independently in Edinburgh rather than being dispersed there from North Wales (Ashton and Abbott
1992; Harris and Ingram 1992). Herbarium records indicate that the Edinburgh
lineage may have existed since at least 1974; however, it is now thought to be
extinct as the species has not been recorded in the Edinburgh area or nearby since
1993 (Abbott and Forbes 2002).
Because S. cambrensis is readily synthesised by treating the triploid hybrid
between S. squalidus and S. vulgaris with colchicine, it has been assumed that
chromosome doubling of the triploid hybrid was the likely pathway of origin of the
allopolyploid species in the wild (Rosser 1955). In theory, however, the species could
have originated along several possible pathways (Table 13.1) with the first step
involving formation of a triploid, tetraploid, pentaploid or hexaploid hybrid. Of these
alternatives, the formation of a triploid hybrid is more likely in that it results from
fusion of normal haploid gametes produced by each parent. In contrast, formation of
higher ploidy hybrids relies on the production of unreduced gametes, which will be
generated at a much lower frequency in each parent species.
There are many records of the triploid hybrid (Senecio x baxteri Druce)
occurring in the wild (Crisp 1972; Benoit et al. 1975; Marshall and Abbott 1980).
It is easily recognised because of its intermediate morphology and its almost
complete seed sterility. Progeny tests of S. vulgaris plants have shown that this
hybrid is generated regularly but at very low frequencies in the wild where S.
vulgaris and S. squalidus co-occur (Marshall and Abbott 1980). In contrast, the
tetraploid hybrid has never been reported unequivocally in the wild, although
Crisp (1972) described a plant likely to have been such a hybrid based on morphology and an analysis of its offspring. Although no chromosome count was
made of the plant, all of its offspring were approximately tetraploid and segregated
for a range of morphological, reproductive and disease resistance traits. Crisp
(1972) suggested that such offspring could become stabilized in the wild to form
distinct taxa, and it is feasible that the tetraploid S. eboracensis originated in this
way (Lowe and Abbott 2000). Whether such a hybrid might have contributed to
the origin of S. cambrensis as detailed in Table 13.1 remains unknown, but seems
less likely than an origin involving the triploid hybrid, given the apparent rarity of
the tetraploid hybrid in the wild. Similarly, origins involving the formation of
either a pentaploid or hexaploid hybrid are less parsimonious than one involving
the triploid hybrid, although cannot be ruled out entirely. Further support for the
hypothesis that formation of a triploid hybrid was the first step in the origin of
CO
RR
166
167
M. J. Hegarty et al.
UN
Editor Proof
250
Layout: T1 Standard SC
Chapter No.: 13
13
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 251/270
Allopolyploid Speciation in Action
251
213
214
215
216
217
218
219
220
221
222
223
224
225
226
227
S. cambrensis comes from reports by Vosa (in Crisp 1972) and Ingram (1978) that
rare allohexaploid offspring were produced spontaneously by natural selfing of the
synthetic triploid hybrids they made. However, Weir and Ingram (1980) also
reported the production of an allohexaploid plant directly from a cross between S.
vulgaris and S. squalidus. This could have been formed by fusion of unreduced
gametes or alternatively by chromosome doubling of a triploid hybrid early in its
development. We shall never know exactly how the different lineages of S.
cambrensis originated in the UK, but given that the triploid hybrid is regularly
encountered in the wild and is capable of producing allohexaploid offspring
spontaneously, an origin involving doubling of the chromosome number of a
triploid hybrid seems the most likely route of origin.
Because S. cambrensis is self-fertile, one newly formed individual of the species would have been able to reproduce sexually and successfully following the
species’ origin, i.e. without need of a mate. Moreover, if the species had originated
through chromosome doubling of a triploid hybrid, it would be expected initially
to be homozygous at all loci within its parental genomes, but to exhibit frequent
fixed heterozygosity at duplicated loci among parental genomes. Somewhat
CO
RR
212
UN
211
EC
TE
D
PR
OO
F
Editor Proof
Table 13.1 Some possible pathways of origin for Senecio cambrensis in the wild
(1) Via Triploid hybrid (2n=30)
• Step 1 - Formation of triploid hybrid (2n=30) through fusion of haploid gametes
produced by each parent species.
• Step 2 - Chromosome doubling of triploid hybrid by: (i) fusion of ‘unreduced’ triploid gametes
(n=30); or (ii) doubling of chromosome number of a somatic cell ancestral to a floret or flower
head producing ‘reduced’ triploid gametes (n=30).
(2) Via Tetraploid hybrid (2n=40)
• Step 1 - Formation of tetraploid hybrid (2n=40) by: (i) fusion of a haploid gamete (n=20)
of S. vulgaris and a diploid (unreduced) gamete (n=20) of S. squalidus; or (ii) fusion
of an ‘unreduced’ triploid gamete of triploid hybrid (n=30) and haploid gamete of
S. squalidus (n=10).
• Step 2 - Production of hexaploid hybrid by: (i) fusion of an ‘unreduced’ tetraploid gamete
(n=40) generated by tetraploid hybrid and a ‘reduced’ diploid gamete (n=20) of same hybrid
or of S. vulgaris or an unreduced gamete of S. squalidus; or (ii) fusion of ‘balanced’ triploid
gametes (n=30) produced by same hybrid.
(3) Via Pentaploid hybrid (2n=50)
• Step 1 - Formation of pentaploid hybrid by: (i) fusion of a haploid gamete of S. squalidus
(n=10) and an unreduced gamete of S. vulgaris (n=40); or (ii) fusion of unreduced gamete
of triploid hybrid (n=30) with haploid gamete of S. vulgaris (n=20) or unreduced gamete
of S. squalidus (n=20).
• Step 2 - Production of hexaploid hybrid by: (i) fusion of gametes with same or different
‘balanced’ chromosome numbers (i.e. n=10, n=20, n=30, n=40, n=50) generated by
pentaploid hybrid such that the zygote produced is hexaploid (2n=60); (ii) fusion of tetraploid
‘balanced’ gamete (2n=40) produced by pentaploid hybrid with reduced gamete (n=20) of
S. vulgaris or unreduced gamete of S. squalidus. (iii) fusion of diploid ‘balanced’ gamete
(n=20) produced by pentaploid hybrid with unreduced gamete (n=40) of S. vulgaris.
4) Direct formation of Hexaploid hybrid (2n=60)
• Step 1 - Formation of hexaploid hybrid by fusion of unreduced gametes from both
S. squalidus (n=20) and S. vulgaris (n=40).
Layout: T1 Standard SC
Chapter No.: 13
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 252/270
234
235
236
237
238
239
240
241
242
243
244
245
246
247
248
249
250
F
233
PR
OO
232
D
230
231
surprisingly, however, the species has been shown through surveys of allozyme
variation (Ashton and Abbott 1992) and particularly AFLP variation (Abbott et al.
2007) to contain high levels of genetic diversity, indicating that it rapidly generated this diversity following its origin. Abbott et al. (2007) considered the ways in
which such genetic diversity was produced and concluded that intergenomic
recombination would most likely have been an important mechanism, although
other mechanisms such as aneuploidy, gene conversion, activation of transposons
and retroelements, other forms of mutation and gene flow from parental species
could not be ruled out. The occurrence of radiate and non-radiate forms of S.
cambrensis as well as variation in the ligule length of ray florets have been
attributed to intergenomic recombination (Ingram and Noltie 1984), while the
observation of multivalent formation occurring at low frequency in meiotic cells of
the species (Ingram and Noltie, 1989) provides a mechanism for such recombination to occur. Although not reported by Ingram and Noltie (1989), Crisp (1972)
observed up to eight chromosomes with subterminal centromeres in the somatic
complement of S. cambrensis plants (based on root tip squashes). He pointed out
that as neither parent species possessed such chromosomes they were probably the
products of chromosome rearrangements following meiotic abnormalities. In
addition to generating genetic diversity, intergenomic recombination resulting in
chromosome rearrangements could lead to the formation of reproductive barriers
between divergent lineages of S. cambrensis. Clearly, further work on the frequency of intergenomic recombination and its possible effects in S. cambrensis
would be worthwhile.
TE
229
EC
228
M. J. Hegarty et al.
13.2 Consequences of Hybridization and Polyploidy
in Natural and Resynthesised Senecio cambrensis
253
13.2.1 Transcriptome Shock
254
255
256
257
258
259
260
261
262
263
264
265
266
CO
RR
252
251
In common with many other allopolyploid species, the merger of two divergent
genomes during the formation of Senecio cambrensis has had a dramatic impact at
the level of gene expression. As part of our investigation of the allopolyploid
origins of S. cambrensis, we conducted gene expression analysis using a custom
cDNA microarray platform (Hegarty et al. 2005) to survey the transcript levels of
floral genes in both the intermediate triploid hybrid S. x baxteri and wild S.
cambrensis, relative to their progenitor species. The experimental design of this
comparison is shown in Fig. 13.2. This experiment revealed an initial large change
in floral gene expression in S. x baxteri, with approximately 475 cDNA clones
showing up- or down-regulation relative to its parental taxa or, also importantly,
relative to natural S. cambrensis, from which it differs primarily by a change in
ploidal level (Hegarty et al. 2005). Thus, the greatest changes in gene expression
relative to the parents appeared to be associated with the hybridization step to form
UN
Editor Proof
252
Layout: T1 Standard SC
Chapter No.: 13
Book ISBN: 978-3-642-31441-4
Page: 253/270
Allopolyploid Speciation in Action
253
PR
OO
F
Editor Proof
13
Book ID: 272454_1_En
Date: 16-8-2012
273
274
275
276
277
278
279
280
281
282
283
284
285
286
287
288
289
290
291
TE
EC
271
272
CO
RR
269
270
S. x baxteri. This initial burst of altered gene expression we termed ‘‘transcriptome
shock’’, after the phenomenon of ‘‘genome shock’’ described by McClintock in her
seminal work on transposable elements in plant hybrids (McClintock 1984). The
‘transcriptome shock’ effect in S. x baxteri was confirmed in our later analysis of
resynthesised S. cambrensis, which further showed that the polyploidization event
(here induced by colchicine) had an immediate calming (ameliorating) effect on
altered patterns of gene expression detected in S. x baxteri (Hegarty et al. 2006).
Importantly, this altered pattern of gene expression, apparent in first-generation
allopolyploids, was preserved in four successive generations of the synthetic allopolyploids and in wild S. cambrensis (Hegarty et al. 2006). Previous research in
resynthesised wheat (Feldman and Levy 2005) identified separate effects of
hybridization and polyploidization on the genome and transcriptome, but our
findings in S. cambrensis represented one of the first indications that these changes
in gene expression were genome-wide. Interestingly, the putative functional
classes of genes affected by hybridization and allopolyploidization were remarkably similar, with no functional class of genes being overly affected by hybridization or allopolyploidization (Fig. 13.3). However, perhaps not surprisingly,
when compared with functional classes of genes not affected by either process,
there was a greater representation of genes potentially involved in flower/inflorescence and pollen developments, which may reflect the transitions in floral
phenotypes observed after hybridization and allopolyploidization.
We later reassessed the data (Hegarty et al. 2008) in light of a new approach
used by Wang et al. (2006a) in their studies of allotetraploid Arabidopsis suecica.
In this study, they focused on the identification of genes whose expression in
hybrids differed from the additive expression midpoint of the two different parental
UN
267
268
D
Fig. 13.2 Experimental design employed in microarray comparisons of gene expression between
the allopolyploid S. cambrensis and its progenitor taxa, S. vulgaris and S. squalidus, and their
sterile triploid F1 hybrid, S. x baxteri. Experimental details can be found in Hegarty et al. (2005),
but, briefly, mature flower bud tissue was harvested from a mixed population of approximately 30
plants and pooled prior to RNA extraction to create an ‘average’ for each taxon. Labelled cDNA
for each taxon was hybridized to a custom floral cDNA microarray. Two taxa were differentially
labelled and compared per array hybridization (with 10 replicate hybridizations performed per
comparison) using dye swaps to account for any bias in labelling efficiency. Each taxon was
compared with the other three, for a total of 30 array hybridizations per taxon. Raw expression
data for each taxon were extracted from these 30 replicates and imported separately into the
GENESPRING microarray analysis software (Silicon Genetics) to enable comparison between all
four taxa. Figure reproduced from Hegarty et al. (2008)
AQ5
Layout: T1 Standard SC
Chapter No.: 13
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 254/270
M. J. Hegarty et al.
D
PR
OO
F
Editor Proof
254
294
295
296
297
298
299
300
301
302
303
304
305
306
307
gene copies. A similar approach had also been used to analyse gene expression
change in maize hybrids (Stupar et al. 2007). Such an approach provides a consistent and unified methodology for identifying genes affected by hybridization
and/or polyploidization in different model study systems. We therefore reanalysed
the microarray data from our original study to identify specific genes and classes of
genes affected by hybridization and polyploidization. Using methods similar to
Stupar et al. (2007), we tested whether changes in gene expression observed in
synthetic S. x baxteri and wild allohexaploid S. cambrensis were additive or nonadditive (Hegarty et al. 2008). By averaging the parental expression values for
each feature on the array showing differential expression in the hybrids, a parental
midpoint expression value (MPV) was obtained. The derived midpoint values were
then used to calculate a ratio of hybrid and parental expression values compared to
the MPV for each array feature. A ratio of -0.33 indicates additive gene
expression, whilst ratios below -1 or above 1 represent expression in the hybrid
outside the range of either parent. Statistical analysis of differentially expressed
genes from our previous microarray experiment (Hegarty et al. 2005) showed that,
CO
RR
293
UN
292
EC
TE
Fig. 13.3 Functional classes of genes affected by allopolyploidization and hybridization. Basic
gene ontologies for cDNA clones displaying a conserved expression changes in both wild and
synthetic Senecio cambrensis relative to S. x baxteri (genes affected by allopolyploidy, 540
clones), b expression changes relative to the parental taxa S. squalidus and S. vulgaris in both
hybrid taxa (genes affected by hybridization, 99 clones) and c genes showing no expression
difference between the parental and hybrid taxa (unaffected by hybridization or polyploidy, 289
clones). With the exception of a higher proportion of floral/pollen-related genes in (a, b)
compared with (c), there are no substantial differences between the classes of affected genes
(adapted with permission from Hegarty et al. 2006)
Layout: T1 Standard SC
Chapter No.: 13
315
316
317
318
319
320
321
322
323
324
325
326
327
328
329
330
331
332
333
334
335
336
337
338
339
340
341
342
343
344
345
346
347
348
349
350
351
352
F
314
PR
OO
313
D
311
312
for both hybrids, the median ratio was significantly different from -0.33, allowing
us to reject the null hypothesis of largely additive gene expression changes.
Instead, for both hybrids, the majority of the data were skewed towards one of the
parents; in the case of S. x baxteri, expression was skewed towards that of the
lower expressing parent, whereas in S. cambrensis it was skewed towards that of
the higher expressing parent. Further analysis of the data showed that for both
hybrids S. vulgaris was the lower expressing parent in 70 % of cases. Expression
outside the parental range was observed in a substantial proportion of cases in both
hybrids: 7.42 % in S. x baxteri and 3.03 % S. cambrensis (Hegarty et al. 2008).
Having identified a pool of cDNA clones displaying non-additive changes to
gene expression in both hybrid taxa, we then tested these clones for evidence of
expression beyond the parental ranges, i.e. transgressive gene expression. In S. x
baxteri, 80.4 % of non-additively expressed clones differed from the MPV by
[1.5-fold, with 42.2 % of clones in S. cambrensis showing the same effect. Within
both of these groups, the majority of cases involved upregulation compared with
the MPV (66.9 and 70.4 % in S. x baxteri and S. cambrensis, respectively). Aside
from the genes for which no functional class could be ascribed (49.2 %), the major
functional groups affected in S. x baxteri were genes involved in development
(6.6 %), nucleotide binding (6.1 %), mitochondrial activity (4.76 %) and cell wall
function (3.97 %). Within the development category, a high proportion of clones
(32 %) were found to encode tubulins, profilins or senescence-associated proteins.
Of the clones involved in nucleotide binding, 34 % were transcription factors. In S.
cambrensis, the majority of clones (58 %) could not be assigned to a functional
category. Of the remainder, the largest categories were defense (11.1 %) and cell
wall-related genes (6.17 %).
Our reanalysis of the microarray data therefore revealed a relatively high
proportion of non-additive gene expression change in the hybrids relative to their
parental expression levels (Hegarty et al. 2008). In addition, the degree of nonadditive gene expression was lower in allohexaploid S. cambrensis compared with
its triploid intermediate S. x baxteri. This finding was consistent with our previous
observation that the ‘‘transcriptome shock’’ resulting from allopolyploidization is
largely due to hybridization, with polyploidization resulting in a distinct secondary
shift (amelioration) in gene expression (Hegarty et al. 2006). The fairly diverse
nature of the genes affected was consistent with other findings in Arabidopsis
(Wang et al. 2006b), cotton (Adams et al. 2004) and maize (Stupar et al. 2007) that
non-additive changes to gene expression are genome-wide.
Interestingly, similar functional classes of genes were affected by hybridization
in Senecio, Arabidopsis and maize (Fig. 13.4), suggesting that certain gene networks may be particularly susceptible to perturbation by hybridization; the functional categories of nucleotide binding, defense and mitochondria being good
examples. In terms of the classes of genes affected in Senecio, it is also noteworthy
that one of the major affected groups in S. x baxteri was nucleotide binding. In
addition to a number of (primarily down-regulated) transcription factors that have
shown similar alterations in expression pattern in the polyploid Arabidopsis suecica (Wang et al. 2006b), this category also contained clones encoding cytidine
TE
310
255
EC
309
CO
RR
308
Book ISBN: 978-3-642-31441-4
Page: 255/270
Allopolyploid Speciation in Action
UN
Editor Proof
13
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 13
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 256/270
M. J. Hegarty et al.
PR
OO
F
Editor Proof
256
355
356
357
358
359
360
361
362
363
364
365
366
367
368
369
deaminase (CDA) and 8-oxoG-DNA glycosylase (OGG1). OGG1 has been
implicated in DNA base excision repair (García-Ortiz et al. 2001), while CDA has
been suspected to be involved in RNA editing, although it now appears that
pentatricopeptide repeat proteins that contain CDA-like domains are the more
likely candidates (Salone et al. 2007). These genes were of interest, given that we
also observed a relatively high number of clones encoding proteins involved in
either DNA modification or cell division. In addition to cytidine deaminase and
OGG1 (both of which were upregulated compared with the parental midpoint in S.
x baxteri), there was also up-regulation of adenosylhomocysteinase and adenosyl
kinase, the genes involved in S-adenosylmethionine (SAM) dependent methylation
(Moffatt et al. 2002; Mull et al. 2006). The expression of SAM synthetase was also
increased relative to both parents. SAM-dependent methylation is used in gene
silencing and also in pectin methylation (Pereira et al. 2006), and, indeed, we
observed an increase in the expression of pectin methylesterase as well as another
SAM-dependent enzyme, caffeic acid 3-O-methyltransferase (Hegarty et al. 2008).
It is clear from these analyses that hybridization and polyploidy have separate,
distinct effects on gene expression in Senecio. These changes in gene expression
CO
RR
354
UN
353
EC
TE
D
Fig. 13.4 Comparison of non-additive expression changes resulting from hybridization in maize
and hybridization/polyploidy in Senecio and Arabidopsis. The formation of the hybrids is shown
in each case, together with the level of non-additive gene expression in the hybrids expressed as a
percentage of the features on the microarray platform used. Finally, the top five functional gene
classes affected (ignoring unknowns) for each hybrid are displayed for comparison. Red indicates
a functional gene class affected in all four hybrid systems, blue indicates a functional class
affected in at least one of the Senecio hybrids and one of the other two hybrid taxa, and green
indicates a functional class affected in both Senecio hybrids but not in either Arabidopsis or
maize. For Arabidopsis suecica, gene function data were taken from Wang et al. (2006b), and for
maize from an extrapolation of the supplementary data given in Stupar et al. (2007). Sb, S. x
baxteri; Sc, S. cambrensis. Reproduced from Hegarty et al. (2008)
Layout: T1 Standard SC
Chapter No.: 13
Book ISBN: 978-3-642-31441-4
Page: 257/270
Allopolyploid Speciation in Action
257
376
13.2.2 Epigenetic Modification
378
379
380
381
382
383
384
385
386
387
388
389
390
391
392
393
394
395
396
397
398
399
400
401
402
403
404
405
406
407
408
409
410
PR
OO
377
While we were unable to demonstrate complete silencing of any clones in our
microarray analyses, it is important to note that the cDNA-based microarrays used
could not distinguish between different parental homeologues. Therefore, in the
cases where a hybrid showed down-regulation of a gene relative to its parental
taxa, it may have been due to either homeologue loss or silencing as a consequence
of DNA methylation. Both phenomena were observed using cleaved amplified
polymorphic sequence (CAPS) analysis in natural and resynthesised allotetraploid
Tragopogon miscellus (Buggs et al. 2009) and suggested that homeologue loss was
not an immediate consequence of hybridization or polyploidy but occurs slightly
later due to recombinational events. Silencing was also not observed to occur
immediately in Tragopogon synthetics, but observations in natural populations
suggest that silencing of one homoeologue is more prevalent than sequence loss.
Studies in other allopolyploid systems such as wheat, Arabidopsis and Spartina
have demonstrated that methylation status in F1 hybrids and first-generation
allopolyploids displays similar changes to those observed for gene expression.
To determine whether methylation in Senecio x baxteri and S. cambrensis was
affected in a similar manner, we undertook a methylation-sensitive AFLP (MSAP)
analysis of the parental taxa, three triploid lines and their S0–S1 allohexaploid
derivatives (Hegarty et al. 2011). The MSAP technique (Xiong et al. 1999)
involves digestion of genomic DNA with a standard, rare-cutter enzyme (i.e.,
EcoRI) and one of a pair of isoschizomeric enzymes that share a restriction site but
are either sensitive or insensitive to cytosine methylation. In this case, we
employed HpaII, which is sensitive to methylation of either cytosine in its recognition site (CCGG) and MspI, which is sensitive only to methylation of the
external cytosine. Comparing the two AFLP profiles produced from each isoschizomer enables identification of the methylation status of a given locus, as
described in Table 13.2. Five selective primer combinations were used to screen
33 plants as detailed in Table 13.3.
We successfully amplified MSAP products from all 33 individuals and obtained
a total of 408 reliable MSAP loci, which were then surveyed to determine levels of
non-additivity in the hybrids. Of the 408 loci, 264 (64.7 %) showed polymorphisms between the parental taxa S. squalidus and S. vulgaris. In the remaining
144 loci that were monomorphic in the parents, 75.7 % were monomorphic across
all hybrid samples tested. Surveying the polymorphic loci, it was found that the
D
374
TE
373
EC
372
CO
RR
371
F
375
affect a wide spectrum of transcripts from a number of functional groups and are
non-additive in nature. A good proportion of these changes are also transgressive—outside the range of either parent species—representing a possible source of
heterotic success in interspecific hybrids. To investigate possible mechanisms for
these changes in gene expression, we focused our attention on epigenetic modification in Senecio hybrids.
370
UN
Editor Proof
13
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 13
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 258/270
258
M. J. Hegarty et al.
F
Editor Proof
Table 13.2 HpaII% MspI banding patterns and locus methylation state (reproduced from Hegarty et al, 2011)
HpaII
MspI
Methylation status
+
CCGG (unmethylated)
+
CmCGG (methylation of internal cytosine)
m
CCGG or mCmCGG (methylation of external cytosine)
hm
+
CCGG (hemimethylation of external cytosine)
PR
OO
Note: In the case of methylation of the external cytosine, it is not possible from MSAP alone to
determine whether the internal cytosine is also methylated
414
415
416
417
418
419
420
421
422
423
424
425
426
427
428
429
430
CO
RR
413
triploid hybrids each displayed similar overall methylation patterns (Table 13.4),
with a strong bias in favour of the S. vulgaris (maternal) methylation state for each
locus (an average of 57.4 % of loci between the three triploid lines). In addition,
the triploids also displayed approximately equivalent levels of non-additive
methylation (13.4 % on average). To determine whether methylation patterns were
maintained following genome duplication, the methylation status of the S0 allohexaploids was compared to the triploid lines from which they were derived. In the
vast majority of cases, the synthetic allohexaploids retained the methylation state
observed in the triploid. On average, 78.2 % of these cases involved additive
methylation, whereas 9.7 % of loci involved retention of a non-additive methylation profile. An average of 10.1 % of loci displayed a shift relative to the triploid
in the specific parental methylation state favoured (4.5 % shift to S. squalidus,
3.8 % shift to S. vulgaris, 1.8 % shift to both). Finally, an average of 2 % of loci
showed novel non-additive methylation not observed in the triploids.
The analysis was then extended to the S1 allohexaploids, comparing them to
both their triploid ancestors and the preceding S0 generation. As with the S0
allohexaploids, overall methylation state was highly similar among the three lines.
Again, the most common result was retention of methylation status compared to
both the triploid and the S0 allohexaploid, with an average of 70.3 % of loci
showing retention of additive methylation patterns and 7.3 % of non-additive
UN
411
412
EC
TE
D
Table 13.3 Oligonucleotides for methylation-sensitive AFLP and number of loci (reproduced
from Hegarty et al, 2011)
Oligo
Sequence (5’-3’
Primer Combination
# Loci
EcoRI adaptor 1
CTCGTAGACTGCGTACC
Eco+AGC/Hpa+CTG
118
EcoRI adaptor 2
AATTGGTACGCAGTC
Eco+AAC/Hpa+CTT
88
HpaII adaptor 1
GATCATGAGTCCTGCT
Eco+AGC/Hpa+CTT
80
HpaII adaptor 2
CGAGCAGGACTCATGA
Eco+AAC/Hpa+AAG
Failed
Eco+A
GACTGCGTACCAATTCA
Eco+ACG/Hpa+AAG
69
Hpa
ATCATGAGTCCTGCTCGG
Eco+AAC/Hpa+CTT
53
Eco+AAC (NED)
GACTGCGTACCAATTCAAC
Eco+ACG (FAM)
GACTGCGTACCAATTCACG
Hpa+CTG
ATCATGAGTCCTGCTCGGCTG
Hpa+CTT
ATCATGAGTCCTGCTCGGCTT
Hpa+AAG
ATCATGAGTCCTGCTCGGAAG
Layout: T1 Standard SC
Chapter No.: 13
13
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 259/270
Allopolyploid Speciation in Action
259
TE
D
PR
OO
F
Editor Proof
Table 13.4 Summarised methylation status of hybrid lines (reproduced from Hegarty et al.
2011)
Methylation
Percentage of Loci (triploids)
State
Line 1
Line 2
Line 3
Additive (SS)
24.58%
24.58%
22.26%
Additive (SV)
56.15%
57.14%
58.80%
Additive (monomorphic)
4.65%
6.98%
4.65%
Nonadditive
14.62%
11.30%
14.29%
Percentage of Loci (S0 allohexaploids)
Same as triploid (additive)
76.17%
78.86%
79.53%
Same as triploid (nonadditive)
9.73%
9.73%
9.73%
Differs from triploid (additive SS)
5.03%
4.36%
4.03%
Differs from triploid (additive SV)
5.37%
3.69%
2.35%
Differs from triploid (monomorphic)
2.01%
0.67%
2.68%
Novel nonadditive methylation
1.68%
2.68%
1.68%
Percentage of Loci (S1 allohexaploids)
Same as triploid+S0 (additive)
72.06%
72.38%
66.35%
Same as triploid+S0 (nonadditive)
7.62%
7.94%
6.35%
7.62%
5.71%
3.49%
Same as S0 not triploid (additive)
Same as S0 not triploid (nonadditive)
0.32%
1.27%
0.63%
Same as triploid not S0 (additive)
3.81%
3.17%
1.90%
0.95%
0.00%
2.54%
Same as triploid not S0 (nonadditive)
Differs from triploid+S0 (additive SS)
2.54%
4.76%
7.62%
Differs from triploid+S0 (additive SV)
3.49%
2.22%
7.62%
Novel nonadditive methylation
1.59%
2.54%
3.49%
433
434
435
436
437
438
439
440
441
442
443
444
445
446
447
448
methylation. A further 5.6 % of loci showed retention of additive methylation in
the two allohexaploid generations, whereas the triploid had displayed non-additivity. Methylation status was not always consistent between the two allohexaploid
generations; however, in 4.1 % of cases, loci displayed a shift in methylation state
to that seen in the triploid but not in the S0 allohexaploid. Furthermore, the S1
generation showed a return to additivity in 9.4 % of cases, where both the triploid
and the S0 allohexaploids had been non-additive. As with the S0 generation,
though, there was a small (2.5 % average) degree of novel non-additivity.
The results were in accordance with our previous studies of gene expression
(Hegarty et al. 2006, 2008): we found that, while cytosine methylation in both
hybrid taxa was largely additive between the two parental patterns, a significant
degree of non-additivity also exists. Overall methylation status was well conserved
between different hybrid lines; while individual loci displayed differences, the
global percentages of different methylation states were highly similar between
lines (Hegarty et al. 2011). In all three triploid lines, approximately 13.4 % of loci
showed non-additive methylation, although the precise type of methylation was
not identical between lines in all cases. Levels of non-additive methylation
observed in other allopolyploid systems are variable: 8.3 % in Arabidopsis
CO
RR
432
UN
431
EC
Note: SS = S. squalidus, SV = S. vulgaris. ‘‘Monomorphic’’ refers to loci with a common
methlyation state in the parents, rather than across hybrid lines
Layout: T1 Standard SC
Chapter No.: 13
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 260/270
456
457
458
459
460
461
462
463
464
465
466
467
468
469
470
471
472
473
474
475
476
477
478
479
480
481
482
483
484
485
486
487
488
489
490
491
492
493
F
455
PR
OO
454
D
453
TE
451
452
(Madlung et al. 2002), 9 % in Brassica (Gaeta et al. 2007), 13–20 % in wheat
(Dong et al. 2005; Pumphrey et al. 2009) and as high as 30 % in Spartina (Salmon
et al. 2005). It has been speculated that the higher genome copy number in wheat
and Spartina might explain their greater levels of methylation, although Doyle
et al. (2008) point out that both species are monocots, which tend to possess a
higher GC content (and thus greater potential for methylation) than eudicots. The
fact that conserved methylation changes between the Senecio hybrids are more on
a par with the levels seen in Arabidopsis and Brassica suggests that this latter
hypothesis may be correct, as S. cambrensis exhibits the same ploidy as wheat. We
should note, however, that the wheat genome is significantly larger than that of
Senecio (2C genome sizes of 33.96 Gbp wheat; 5.05 Gbp S. cambrensis) and is
known to contain a significant amount of repetitive DNA including large numbers
of retroelements. It is therefore probable that alterations to methylation are more
necessary to prevent widespread activation of these genetic regions in wheat and
similar polyploid monocots. Indeed, studies of methylation in Spartina (Parisod
et al. 2010) showed that such changes frequently occur in the vicinity of transposable elements and, perhaps as a result, no transposition burst was detected in
the Spartina hybrids analysed. Methylation change therefore appears to play a
frequent role in genome mergers, but there are exceptions: despite significant
differential gene expression in the allotetraploid Gossypium hirsutum, almost no
differences in methylation could be observed between the hybrid and its parental
taxa (Liu et al. 2001) nor do the parental genomes undergo any significant rearrangement. In this situation, it appears that subfunctionalization of the two genomes is the primary cause of phenotypic variation (Adams et al. 2004; Liu and
Adams 2007), although a recent study by Chaudhary et al. (2009) demonstrated
that neofunctionalization and divergence in parental cis-regulatory sequences also
play a significant role. Exactly what factors determine the response of the parental
genomes to hybridization are largely unknown, although the degree of parental
divergence is speculated to play a large role (Chapman and Burke 2007; Buggs
et al. 2008; Paun et al. 2009).
In further accordance with our expression analyses, we observed that nonadditive methylation in S. x baxteri triploids was maintained, on average, in only
73.6 and 55.6 % of cases in the S0 and S1 allohexaploids, respectively (Hegarty
et al. 2011). In approximately 73 % of cases observed in our microarray studies,
the resynthesised allohexaploid lines (S0–S4) displayed either an immediate or
gradual shift towards an expression pattern similar to that of wild S. cambrensis
(Hegarty et al. 2006). It seems likely, therefore, that our previous observation that
non-additivity results from hybridization but can be partially reduced by genome
duplication, also holds true when applied to DNA methylation. This again matches
observations from MSAP analysis of Spartina allopolyploids (Salmon et al. 2005),
which showed that the allopolyploid S. anglica retained 71.4 % of the non-additive
methylation patterns observed in the F1 hybrid. These findings were again confirmed when assessing methylation changes associated with transposable elements
(Parisod et al. 2010), with the additional observation that many of the changes seen
in the F1 hybrid involved loss of parental markers (usually in the maternal
EC
450
CO
RR
449
M. J. Hegarty et al.
UN
Editor Proof
260
Layout: T1 Standard SC
Chapter No.: 13
501
502
503
504
505
506
507
508
509
510
511
512
513
514
515
516
517
518
519
520
521
522
523
524
525
526
527
528
529
530
531
532
533
534
535
536
537
538
F
PR
OO
499
500
D
497
498
genome), indicating that such changes involved structural rearrangements to the
parental genomes. A similar process may be at play in S. x baxteri, because MSAP
markers also detect structural changes: indeed, wild populations of S. cambrensis
show evidence of intergenomic recombination (Abbott et al. 2007, and above),
again favouring the S. vulgaris genome as with our triploid lines here. By contrast,
most of the differences observed in allopolyploid Spartina involved alterations to
methylation status, rather than structural changes. Similar findings have also been
identified in Brassica, where most of the methylation changes identified in the S0
allotetraploid were maintained in S5 lines, but with a number of revertants and
novel changes present as well (Gaeta et al. 2007).
A key finding from our analysis was that the global patterns of DNA methylation change observed in our experiments strongly mirror the global changes in
gene expression observed in our earlier microarray analyses, indicating a possible
underlying causation. Whilst further investigation of specific loci showing methylation differences is required to make a definitive case, the similarities between
changes in gene expression and DNA methylation are nevertheless striking. For
example, we noted that a number of loci displayed novel non-additivity in both the
S0 and S1 allohexaploids (2.01 % on average in the S0 lines, 2.54 % in the S1),
again a point of consistency between the methylation study and our earlier
microarray expression analysis (Hegarty et al. 2008). The overall level of nonadditive methylation may therefore not actually decrease as a consequence of
genome duplication, but instead the level of conserved methylation may be lessened. A proportion of loci also displayed unstable methylation patterns across
generations in the hexaploids, with an average of 16.08 % of loci showing differences between the S0 and S1 lines (including the aforementioned novel nonadditive methylation). Of these, the majority consisted of cases where the S1
allohexaploids revert to additivity or favoured a different parental methylation
state to the S0 line. This reversion to an additive profile was also observed between
the triploids and the S0 plants and agreed with observations from the microarray
data that wild S. cambrensis often showed an opposing expression pattern to S. x
baxteri.
However, approximately one-quarter of loci also displayed a shift relative to the
S0 allohexaploids to favour the methylation state seen in the triploid. These
findings suggest that the methylation state of some loci may vary as a consequence
of segregation. This may explain the novel changes observed by Gaeta et al. (2007)
in their S5 allopolyploids of Brassica napus. Similarly, an analysis of natural
populations of the allopolyploid Tragopogon miscellus (Buggs et al. 2009), where
hybridization occurred at least 40 generations ago, identified a random loss of one
parental homoeologue at a rate of 3.2 % across 10 loci in 57 natural hybrids from
five populations. In addition, a further 6.8 % of loci showed evidence of gene
silencing in one parental copy. The loci lost/silenced were not consistent across
populations or individuals, although within populations, there was some conservation in the loci affected. Whilst Buggs et al. (2009) did not note any homoeologue loss/silencing in resynthesised S0 hybrids, the variability in silencing after
such an extended period of time since hybrid formation suggests that
TE
496
261
EC
495
CO
RR
494
Book ISBN: 978-3-642-31441-4
Page: 261/270
Allopolyploid Speciation in Action
UN
Editor Proof
13
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 13
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 262/270
M. J. Hegarty et al.
559
13.2.3 Phenotypic Change
546
547
548
549
550
551
552
553
554
555
556
557
560
561
562
563
564
565
566
567
568
569
570
571
572
573
574
575
576
577
578
579
PR
OO
545
D
544
TE
542
543
EC
541
Senecio x baxteri F1 hybrids generated by crossing S. vulgaris, as female, with S.
squalidus were all self-sterile in contrast to previous studies which reported some
self-fertility (Crisp 1972; Ingram 1978). Treating shoots of S. x baxteri plants with
colchicine produced ‘chimeric’ plants with allohexaploid branches that produced
flower heads that were fully self-fertile. Seed from these flower heads was then
used to found the synthetic S. cambrensis lines (S0–S5) used in the transcriptomic
and epigenetic analyses described above. The first wholly allohexaploid plants
generated from this seed (S0 lines), and their progeny (S1 lines), showed similar
ray flower structure and self-compatibility (Hiscock and Hegarty, unpublished).
However, in subsequent lines, from the S2 onwards, variation in ray flower form
was detected between individuals within and between the nine independent lineages of synthetic S. cambrensis. Some individuals were observed with no ray
flowers, some had short partially tubular ray flowers, while in others ray flowers
were observed of different lengths and number (Fig. 13.5.) Observations on the
progeny of these different individuals showed that these various forms of ray
flower are heritable (Hiscock, unpublished). Comparable variation in S. cambrensis ray flower form was previously attributed to intergenomic recombination
(Ingram and Noltie 1984), but here we suggest another possibility for such abrupt
changes to ray flower phenotype—epigenetic effects associated with the
epigenetic instability observed in early-generation synthetic S. cambrensis
CO
RR
540
F
558
independently formed hybrids can still display significant levels of epigenetic
variation. This study, as well as that of Gaeta et al. (2007), was based on a survey
of a limited number of loci using cDNA-AFLP or CAPS assay. A later study by
Buggs et al. (2011) used the Sequenom MassARRAY allelotyping technology to
survey a much larger set of loci in Tragopogon miscellus and confirmed that
natural hybrids displayed altered patterns of tissue-specific gene expression, whilst
resynthesised hybrids demonstrated relaxed control of tissue-specificity. This latter
finding suggests a possible mechanism for the ‘transcriptome shock’ effect we
observed in Senecio x baxteri and S. cambrensis, resulting from a loss of tissuespecific expression patterns seen in the parent taxa. Further work will be needed to
confirm if this is the case in Senecio. The Sequenom assay used by Buggs et al.
(2011) shows the benefits of new molecular tools for studies of allopolyploid
systems: with the advent of new technologies for global analysis of DNA methylation such as MSAP and next-generation sequencing (Salmon and Ainouche
2010), it would also be interesting to analyse our later-generation allohexaploid
derivatives at a global scale to investigate the longer term changes in methylation
as hybrid genomes undergo recombination and adaptation. Such studies may
therefore provide further insights into which epigenetic changes are mandated by
hybridity, and which may vary between populations and serve as a source of
novelty upon which selection may act.
539
UN
Editor Proof
262
Layout: T1 Standard SC
Chapter No.: 13
Book ISBN: 978-3-642-31441-4
Page: 263/270
Allopolyploid Speciation in Action
263
PR
OO
F
Editor Proof
13
Book ID: 272454_1_En
Date: 16-8-2012
584
585
586
587
588
589
590
591
592
593
594
595
596
597
598
599
600
601
602
603
EC
583
CO
RR
582
(Hegarty et al. 2011). Genes shown to be involved in ray flower development in
Senecio (RAY1 and RAY2) are orthologues of CYCLOIDEA (CYC) (Kim et al.
2008), which has been shown to occur as a stable mutant (hypermethylated) epiallele that in Linaria (toadflax, Plantaginaceae) manifests itself in a change in
flower symmetry from bilateral to radial symmetry (Cubas et al. 1999). We
therefore hypothesise that the observed changes in ray flower form may in part be
associated with epigenetic modifications to RAY1 and/or RAY2. Testing this
hypothesis will be a focus for future work.
Another unexpected observation in the synthetic S. cambrensis lines was the
appearance of self-sterile individuals in otherwise SC lines, again from the S2
generation onwards. Subsequent analyses using controlled self- and crosspollinations confirmed that these self-sterile individuals possessed functional
sporophytic self-incompatibility (Brennan and Hiscock 2010). This is the first time
that sporophytic SI has been shown to be inherited and expressed in an allopolyploid and raises intriguing questions about the mechanism regulating this
important trait. All S0 and S1 lines of synthetic S. cambrensis were highly selffertile (SC) indicating that the SI system, present in parental S. squalidus, was
repressed in these allopolyploids, only to be reactivated/derepressed later. The
emergence of SI individuals may be a consequence of recombination or might also
be associated with epigenetic changes observed in the early-generation synthetic
allopolyploids. Most observations of wild S. cambrensis have reported it to be SC
(Abbott and Lowe 2004), but the finding of SI in synthetic S. cambrensis prompted
a re-examination of the mating system of wild S. cambrensis. An analysis of
selfing rates in 41 wild S. cambrensis individuals from Edinburgh and North Wales
UN
580
581
TE
D
Fig. 13.5 Variation in ray flower morphology observed in individuals of synthetic lines of
allohexaploid S. cambrensis from the S2 to S5 generation
Layout: T1 Standard SC
Chapter No.: 13
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 264/270
M. J. Hegarty et al.
608
identified five SI individuals (Brennan and Hiscock 2010) implying, albeit from a
relatively small sample size, that SI may be present in wild S. cambrensis at a
frequency of *12 %. This important finding means that S. cambrensis should now
be considered as possessing a mixed mating system that has the potential to evolve
towards either outcrossing or selfing.
609
13.3 Future Prospects
604
605
606
615
616
617
618
619
620
621
622
623
624
625
626
627
628
629
630
631
632
633
634
635
636
637
638
639
D
614
We are currently engaged in the generation of a draft reference sequence of the Senecio
squalidus ‘‘gene-space’’ (low-copy, non-methylated regions of the genome). Nextgeneration sequencing (NGS) platforms enable a variety of potential experiments to
examine the consequences of polyploidy and hybridization in Senecio further. For
example, we intend to identify promoter regions using chromatin-immunoprecipitation sequencing (ChIPseq) to enrich for DNA fragments bound by enzymes involved in
transcription. We can then determine whether hybrids and polyploids display alterations in promoter sequence/binding that may explain the altered patterns of expression
observed in our microarray experiments. Further, once a reference sequence is available, we can consider bisulphite sequencing to identify genomic regions which show
differential methylation in hybrids relative to their progenitors. One key target for
bisulphite sequencing will be the RAY1 and RAY2 genes (Kim et al. 2009) which we
suspect may show differential methylation associated with the observed variation in
ray flower morphology that appears in synthetic S. cambrensis lines. Identification of
small interfering RNAs (siRNAs) and their targets will enable analysis of changes to
epigenetic regulation of gene expression in hybrids. At the structural level, resequencing of hybrid ‘‘gene-space’’ and comparative sequence analysis may allow us to
detect genomic rearrangements and sequence loss, plus the activity of transposable
elements. The increasing capability of genotyping-by-sequencing (GBS) approaches
such as restriction-associated DNA (RAD) sequencing (Baird et al. 2008) may also
prove useful in detecting structural rearrangements in hybrid genomes. Finally,
comparative sequencing of Senecio cambrensis and the two other hybrid derivatives of
S. vulgaris and S. squalidus, i.e. S. vulgaris var. hibernicus and S. eboracensis, may
enable identification of dosage effects, since the hybrids share parental genomes but
differ in the specific combinations thereof.
TE
612
613
13.3.1 Next-Generation Approaches to Studying Evolution
in S. cambrensis
EC
611
CO
RR
610
PR
OO
F
607
UN
Editor Proof
264
13.3.2 S. cambrensis in the Wild
Senecio cambrensis now exists in the wild only in North Wales, UK, following
extinction of the Edinburgh population in 1993 (Abbott and Forbes 2002).
AQ6
Layout: T1 Standard SC
Chapter No.: 13
Editor Proof
13
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 265/270
Allopolyploid Speciation in Action
(a)
265
40
30
25
F
20
15
10
5
0
1982
1984
1987
Year
3500
3000
2500
2000
2002
2003
2004
2003
2004
D
1500
1000
500
0
1982
1983
TE
Number of individuals
(b)
1983
PR
OO
Number of sites
35
1984
1987
2002
Year
642
643
644
645
646
647
648
649
650
651
652
653
654
655
656
Although the species was recorded at numerous sites and in relatively high
numbers in North Wales in the early 1980s (Ingram and Noltie 1995; Fig. 13.6), it
has undergone a significant decline in this region since then. In the most recent
census of the species undertaken in 2004, it was found at only nine sites in North
Wales with a total of 349 individuals recorded across all sites (Abbott et al. 2007;
Fig. 13.6). It has been speculated that this decline in numbers has been caused by a
reduction of available sites for colonisation by the species (e.g., waste ground) and
also to increased use of herbicide to control weed populations (Abbott et al. 2007).
Its disappearance from the thin layer of soil that collects along roads between the
road edge and verge, where it was found often in the past, is most likely due to
increased use of herbicides on plants growing along road sides. However, it has
also been noted that plants of the species are frequently heavily infected with the
rust Puccinia lagenophorae. Although this rust also infects both parent species, it
is possible that its effects on S. cambrensis are more dramatic in terms of production of offspring for colonising new sites, given that numbers of S. squalidus
and S. vulgaris that reproduce each year in North Wales and elsewhere in the UK
are vast relative to those of S. cambrensis. Although no detailed analysis has been
CO
RR
641
UN
640
EC
Fig. 13.6 Decline in the number of a sites and b flowering individuals across sites, recorded for
S. cambrensis in North Wales in different years from 1982 to 2004. Records are from Abbott et al.
(2007) and were made in June each year except in 1987 when they were collected in May
Layout: T1 Standard SC
Chapter No.: 13
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 266/270
M. J. Hegarty et al.
679
680
681
682
683
684
685
Acknowledgments Our research on Senecio cambrensis and its parents has been supported by
grants from the NERC, BBSRC and Leverhulme Trust to whom we are grateful. We would also
like to acknowledge our long-term collaboration with Keith Edwards and the considerable efforts
of former research students, technicians and postdocs who contributed to the research over the
past 25 years. These include Alexandra Allen, Paul Ashton, Garry Barker, Tom Batstone, Adrian
Brennan, Mark Chapman, David Forbes, Amanda Gillies, Helen Ireland, Juliet James, Joanna
Jones, Andrew Lowe, David Tabah, and Ian Wilson.
686
References
687
688
689
690
691
692
693
694
695
696
697
698
699
Abbott RJ, Forbes DG (1993) Outcrossing rate and self-incompatibility in the colonizing species
Senecio squalidus L. Heredity 71:155–159
Abbott RJ, Forbes DG (2002) Extinction of the edinburgh lineage of the allopolyploid
neospecies, Senecio cambrensis Rosser (Asteraceae). Heredity 88:267–269
Abbott RJ, Lowe AJ (1996) A review of hybridization and evolution in British Senecio In:Hind
DJN, Beentje HJ (eds.) Compositae: systematics. Proceedings of the international compositae
conference Kew 1994, Royal Botanic Gardens, Kew, pp 679–689
Abbott RJ, Lowe AJ (2004) Origins, establishment and evolution of new polyploid species:
Senecio cambrensis and S. eboracensis in the British isles. Biol J Linn Soc 82:467–474
Abbott RJ, Ashton PA, Forbes DG (1992) Introgressive origin of the radiate groundsel Senecio
vulgaris L. var. hibernicus Syme: Aat-3 evidence. Heredity 68:425–435
Abbott RJ, Ingram R, Noltie HJ (1983) Discovery of Senecio cambrensis Rosser in edinburgh.
Watsonia 14:407–408
663
664
665
666
667
668
669
670
671
672
673
674
675
676
677
PR
OO
662
D
661
TE
660
EC
659
CO
RR
658
F
678
undertaken on whether S. cambrensis is ecologically divergent from its two parents, it tends to grow in sympatry with one or both parent species in the wild. Thus,
there is likely competition between the three species for occupation of available
open sites, and this might place S. cambrensis at a disadvantage if the seed it
produces is less numerous relative to that of S. vulgaris and/or S. squalidus
occurring in the same area. Whatever the cause of its marked decline in numbers in
North Wales over the last 25 years or so, it is clear that S. cambrensis has reached
the stage where its presence in the wild is endangered and that a conservation plan
is required to prevent it from possibly becoming extinct in the near future.
Given the decline in numbers of S. cambrensis in recent years it is important to
gain a better understanding of the nature of its mating system and mating
dynamics, especially in the light of our findings of SI in wild populations (Brennan
and Hiscock 2010). If there has been a recent shift in mating system from predominantly SC towards SI, it is possible that mating potential has become compromised due to obligate outcrossing. Given the possibility of a single hybrid
origin for S. cambrensis in Wales, it is likely that wild populations possess very
few S alleles. Whilst a small number of shared S alleles are not necessarily a
problem when populations are large, it becomes a problem when populations are in
decline and when stochastic effects can lead to S allele loss (Brennan et al. 2003;
Pickup and Young 2008). This can then exacerbate decline, leading to an
uncontrollable spiral of extinction. A reappraisal of the mating system of S.
cambrensis is thus urgently needed.
657
UN
Editor Proof
266
Layout: T1 Standard SC
Chapter No.: 13
267
EC
TE
D
PR
OO
F
Abbott RJ, Ireland HE, Rogers HJ (2007) Population decline despite high genetic diversity in the
new allopolyploid species Senecio cambrensis (Asteraceae). Mol Ecol 16:1023–1033
Abbott RJ, Brennan AC, James JK, Forbes DG, Hegarty MJ, Hiscock SJ (2009) Recent hybrid
origin and invasion of the British isles by a self-incompatible species, Oxford ragwort
(Senecio squalidus L., Asteraceae). Biol Invasions 11:1145–1158
Abbott RJ, Hegarty MJ, Hiscock SJ, Brennan AC (2010) Homoploid hybrid speciation in action.
Taxon 59:1375–1386
Adams KL, Percifield R, Wendel JF (2004) Organ-specific silencing of genes in a newly
synthesized cotton allotetraploid. Genetics 168:2217–2226
Adams KL, Wendel JF (2005) Polyploidy and genome evolution in plants. Curr Opin Plant Biol
8:135–141
Ashton PA, Abbott RJ (1992) Multiple origins and genetic diversity in the newly arisen
allopolyploid species, Senecio cambrensis Rosser (Compositae). Heredity 68:25–32
Baird NA, Etter PD, Atwood TS, Currey MC, Shiver AL, Lewis ZA, Selker WA, Cresko WA,
Johnson EA (2008) Rapid SNP discovery and genetic mapping using sequenced RAD
markers. PLoS ONE 3:e3376
Baker HG (1967) Support for Baker’s’ Law—as a rule. Evolution 21:853–856
Benoit PM, Crisp PC, Jones BMG (1975) Senecio L. In: Stace CA (ed) Hybridization and the
flora of the British isles. Academic, London, pp 404–410
Brennan AC, Hiscock SJ (2010) Expression and inheritance of sporophytic self-incompatibility in
synthetic allohexaploid Senecio cambrensis (Asteraceae). New Phytol 186:251–261
Brennan ACE, Harris SA, Tabah DA, Hiscock SJ (2002) The population genetics of sporophytic
self-incompatibility in Senecio squalidus L. (Asteraceae) I: S allele diversity in a natural
population. Heredity 89:430–438
Brennan ACE, Harris SA, Hiscock SJ (2003) The population genetics of sporophytic selfincompatibility in Senecio squalidus L. (Asteraceae): avoidance of mating constraints
imposed by low S-allele number. Phil Trans R. Soc Lond B 358:1047–1050
Brennan AC, Bridle JR, Wang A-L, Hiscock SJ, Abbott RJ (2009) Adaptation and selection in the
Senecio (Asteraceae) hybrid zone on Mount Etna, Sicily. New Phytol 183:702–717
Brennan ACE, Harris SA, Hiscock SJ (2006) The population genetics of sporophytic selfincompatibility in Senecio squalidus L. (Asteraceae): S allele diversity across the British
range. Evolution 60:213–224
Brennan ACE, Harris SA, Hiscock SJ (2005) Modes and rates of selfing and associated
inbreeding depression in the self-incompatible plant Senecio squalidus (Asteraceae): a
successful colonizing species in the British Isles. New Phytol 168:475–486
Brennan AC, Barker D, Hiscock SJ, Abbott RJ (2012) Molecular genetic and quantitative trait
divergence associated with recent homoploid hybrid speciation: a study of Senecio squalidus
(Asteraceae). Heredity 109:87–95
Buggs RJA, Soltis PS, Mavrodiev EV, Symonds VV, Soltis DE (2008) Does phylogenetic
distance between parental genomes govern the success of polyploids? Castanea 73:74–93
Buggs RJA, Doust AN, Tate JA, Koh J, Soltis K, Feltus FA, Paterson A, Soltis PS, Soltis DE
(2009) Gene loss and silencing in Tragopogon miscellus (Asteraceae): comparison of natural
and synthetic allotetraploids. Heredity 103:73–81
Buggs RJA, Zhang L, Miles N, Tate JA, Gao L, Wei W, Schnable PS, Barbazuk WB, Soltis PS,
Soltis DE (2011) Transcriptomic shock generates evolutionary novelty in a newly formed
natural allopolyploid plant. Curr Biol 21:551–556
Chaudhary B, Flagel L, Stupar RM, Udall JA, Verma N, Springer NM, Wendel JH (2009)
Reciprocal silencing, transcriptional bias and functional divergence of homeologs in
polyploid cotton (gossypium). Genetics 182:503–517
Chapman MA, Burke JM (2007) Genetic divergence and hybrid speciation. Evolution 61:1773–1780
Coleman M, Forbes DG, Abbott RJ (2001) A new subspecies of Senecio mohavensis
(Compositae) reveals an old–new World species disjunction. Edinburgh J Botany 58:384–403
Crisp PC (1972) Cytotaxonomic studies in the section Annui of Senecio. Ph.D Thesis, University
of London
CO
RR
700
701
702
703
704
705
706
707
708
709
710
711
712
713
714
715
716
717
718
719
720
721
722
723
724
725
726
727
728
729
730
731
732
733
734
735
736
737
738
739
740
741
742
743
744
745
746
747
748
749
750
751
752
753
Book ISBN: 978-3-642-31441-4
Page: 267/270
Allopolyploid Speciation in Action
UN
Editor Proof
13
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 13
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 268/270
EC
TE
D
PR
OO
F
Cubas P, Vincent C, Coen E (1999) An epigenetic mutation responsible for natural variation in
floral symmetry. Nature 401:157–161
Dong YZ, Liu ZL, Shan XH, Qiu T, He MY, Liu B (2005) Allopolyploidy in wheat induces rapid
and heritable alterations in DNA methylation patterns of cellular genes and mobile elements.
Genetika 41:1089–1095
Doyle JJ, Flagel LE, Paterson AH, Rapp RA, Soltis DE, Soltis PS, Wendel JF (2008)
Evolutionary genetics of genome merger and doubling in plants. Annu Rev Genet 42:443–461
Feldman M, Levy AA (2005) Allopolyploidy—a shaping force in the evolution of wheat
genomes. Cytogenet Genome Res 109:250–258
Garcı0 a-Ortiz MV, Ariza RR, Rolda0 n-Arjona T (2001) An OGG1 orthologue encoding a
functional 8-oxoguanine DNA glycosylase/lyase in Arabidopsis thaliana. Plant Mol Biol
47:795–804
Gaeta RT, Pires JC, Iniguez-Luy F, Leon E, Osborn TC (2007) Genomic changes in
resynthesized Brassica napus and their effect on gene expression and phenotype. Plant Cell
19:3403–3417
Gray AJ, Marshall DF, Raybould AF (1991) A century of evolution in Spartina anglica. Adv
Ecol Res 21:1–62
Grant V (1981) Plant speciation. Columbia University Press, New York
Harland SC (1955) The experimental approach to the species problem. In: Lousley JE (ed)
Species studies in the British flora. Botanical Society of the British Isles, London, pp 16–20
Harris SA (2002) Introduction of Oxford ragwort, Senecio squalidus L. (Asteraceae), to the
United Kingdom. Watsonia 24:31–43
Harris SA, Ingram R (1992) Molecular systematics of the genus Senecio L. I. hybridization in a
British polyploid complex. Heredity 69:1–10
Hegarty MJ, Hiscock SJ (2008) Genomic clues to the evolutionary success of polyploid plants.
Curr Biol 18:R435–R444
Hegarty MJ, Jones JM, Wilson ID, Barker GL, Coghill JA, Sanchez-Baracaldo P, Liu G, Buggs
RJA, Abbott RJ, Edwards KJ, Hiscock SJ (2005) Development of anonymous cDNA
microarrays to study changes to the Senecio floral transcriptome during hybrid speciation.
Mol Ecol 14:2493–2510
Hegarty MJ, Barker GL, Wilson ID, Abbott RJ, Edwards KJ, Hiscock SJ (2006) Transcriptome
shock after interspecific hybridization in Senecio is ameliorated by genome duplication. Curr
Biol 16:1652–1659
Hegarty MJ, Barker GL, Brennan AC, Edwards KJ, Abbott RJ, Hiscock SJ (2008) Changes to
gene expression associated with hybrid speciation in plants: further insights from transcriptomic studies in Senecio. Philos Trans R Soc Lond B 363:3055–3069
Hegarty MJ, Batstone T, Barker GL, Edwards KJ, Abbott RJ, Hiscock SJ (2011) Nonadditive
changes to cytosine methylation as a consequence of hybridization and genome duplication in
Senecio (Asteraceae). Mol Ecol 20:105–113
Hiscock SJ (2000a) Genetic control of self-incompatibility in Senecio squalidus L. (Asteraceae) –
a successful colonising species. Heredity 84:10–19
Hiscock SJ (2000b) Self-incompatibility in Senecio squalidus L. (Asteraceae). Ann Bot
85(Supplement A):181–190
Ingram R (1978) The genomic relationship of Senecio squalidus L. and Senecio vulgaris L. and
the significance of genomic balance in their hybrid S. x baxteri Druce. Heredity 40:459–462
Ingram R, Noltie HJ (1984) Ray floret morphology and the origin of variability in Senecio
cambrensis Rosser, a recently established allopolyploid species. New Phytol 96:601–607
Ingram R, Noltie HJ (1989) Early adjustment of patterns of metaphase association in the
evolution of polyploid species. Genetica 78:21–24
Ingram R, Noltie HJ (1995) Biological flora of the British isles: Senecio cambrensis Rosser.
J Ecol 83:537–546
James JK, Abbott RJ (2005) Recent, allopatric, homoploid hybrid speciation: the origin of
Senecio squalidus (Asteraceae) in the British Isles from a hybrid zone on Mount Etna, Sicily.
Evolution 59:2533–2547
CO
RR
754
755
756
757
758
759
760
761
762
763
764
765
766
767
768
769
770
771
772
773
774
775
776
777
778
779
780
781
782
783
784
785
786
787
788
789
790
791
792
793
794
795
796
797
798
799
800
801
802
803
804
805
806
807
M. J. Hegarty et al.
UN
Editor Proof
268
Layout: T1 Standard SC
Chapter No.: 13
269
EC
TE
D
PR
OO
F
Jiao L, Wickett NJ, Ayyampalayam S, Chanderbali et al (2011) Ancestral polyploidy in seed
plants and angiosperms. Nature 473:97–100
Kadereit JW, Uribe-Convers S, Westberg E, Comes HP (2006) Reciprocal hybridization at
different times between Senecio flavus and Senecio glaucus gave rise to two polyploidy
species in north Africa and south-west Asia. New Phytol 169:431–441
Kent DH (1963) Senecio squalidus L. in the British isles. 7 Wales. Nat Wales 8:175–178
Kim M, Cui M-L, Cubas P, Gillies A, Lee K, Chapman MA, Abbott RJ, Coen E (2008)
Regulatory genes control a key morphological and ecological trait transferred between
species. Science 322:1116–1119
Leitch IJ, Bennett MD (1997) Polyploid in angiosperms. Trends Plant Sci 2:470–476
Leitch AR, Leitch IJ (2008) Genomic plasticity and diversity of polyploid plants. Science
320:481–483
Liu Z, Adams KL (2007) Expression partitioning between genes duplicated by polyploidy under
abiotic stress and during organ development. Curr Biol 17:1669–1674
Liu B, Brubaker CL, Mergeai G, Cronn RC, Wendel JF (2001) Polyploid formation in cotton is
not accompanied by rapid genomic changes. Genome 44:321–330
Lowe AJ, Abbott RJ (2000) Routes of origin of two recently evolved hybrid taxa: Senecio
vulgaris var. hibernicus and York radiate groundsel (Asteraceae). Am J Bot 87:1159–1167
Lowe AJ, Abbott RJ (2003) A new British species, Senecio eboracensis (Asteraceae), another
hybrid derivative of S. vulgaris L. and S. squalidus L. Watsonia 24:375–388
Lukens LN, Pires JC, Leon E, Vogelzang R, Oslach L, Osborn T (2006) Patterns of sequence loss
and cytosine methylation within a population of newly resynthesized Brassica napus
allopolyploids. Plant Physiol 140:336–348
McClintock B (1984) The significance of responses of the genome to challenge. Science
226:792–801
Madlung A, Masuelli RW, Watson B, Reynolds SH, Davison J, Comai L (2002) Remodeling of
DNA methylation and phenotypic and transcriptional changes in synthetic Arabidopsis
allotetraploids. Plant Physiol 129:733–746
Marshall DF, Abbott RJ (1980) On the frequency of introgression of the radiate (Tr) allele from
Senecio squalidus L. into Senecio vulgaris L. Heredity 45:133–135
Moffatt BA, Allen M, Snider S, Pereira LA, Todorova M, Summers PS, Weretilnyk EA, MartinMcCaffrey L, Wagner C (2002) Adenosine kinase deficiency is associated with developmental
abnormalities and reduced transmethylation. Plant Physiol 128:812–821
Mull L, Ebbs ML, Bender J (2006) A histone methylation-dependent DNA methylation pathway
is uniquely impaired by deficiency in Arabidopsis S-adenosylhomocysteine hydrolase.
Genetics 174:1161–1171
Novak SJ, Soltis DE, Soltis PS (1991) Ownbey’s Tragopogons: 40 years later. Am J Bot 78:1586–1600
Otto SP, Whitton J (2000) Polyploidy: incidence and evolution. Annu Rev Genet 34:401–437
Parisod C, Salmon A, Zerjal T, Tenaillon M, Grandbastien MA, Ainouche M (2010) Rapid
structural and epigenetic reorganization near transposable elements in hybrid and allopolyploid genomes in Spartina. New Phytol 184:1003–1015
Paun O, Forest F, Fay MF, Chase MW (2009) Hybrid speciation in angiosperms: parental
divergence drives ploidy. New Phytol 182:507–518
Pelser PB, Nordenstam B, Kadereit JW, Watson LE (2007) An ITS phylogeny of tribe
Senecioneae (Asteraceae) and a new delimitation of Senecio L. Taxon 56:1062–1077
Pelser PB, Abbott RJ, Comes HP, Milton JJ, Möller M, Looseley ME, Cron GV, Barcelona JF,
Kennedy AH, Watson LE, Barone R, Hernández F, Kadereit JW (2012) The genetic ghost of
an invasion past: colonization and extinction revealed by historical hybridization in Senecio.
Mol Ecol 21:369–387
Pereira LA, Schoor S, Goubet F, Dupree P, Moffatt BA (2006) Deficiency of adenosine kinase
activity affects the degree of pectin methyl-esterification in cell walls of Arabidopsis thaliana.
Planta 224:1401–1414
Pickup M, Young AG (2008) Population size, self-incompatibility and genetic rescue in diploid
and tetraploid races of Rutidosis leptorrhynchoides (Asteraceae). Heredity 100:268–274
CO
RR
808
809
810
811
812
813
814
815
816
817
818
819
820
821
822
823
824
825
826
827
828
829
830
831
832
833
834
835
836
837
838
839
840
841
842
843
844
845
846
847
848
849
850
851
852
853
854
855
856
857
858
859
860
861
Book ISBN: 978-3-642-31441-4
Page: 269/270
Allopolyploid Speciation in Action
UN
Editor Proof
13
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 13
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 270/270
EC
TE
D
PR
OO
F
Pumphrey M, Bai J, Laudencia-Chingcuanco D, Anderson O, Gill BS (2009) Nonadditive
expression of homoeologous genes is established upon polyploidization in hexaploid wheat.
Genetics 181:1147–1157
Rieseberg LH, Raymond O, Rosenthal DM, Lai Z, Livingstone K, Nakazato T, Durphy JL,
Schwarzbach AE, Donovan LA, and Lexer C (2003) Major ecological transitions in wild
sunflowers facilitated by hybridization. Science 301:1211–1216
Rieseberg LH, Willis JH (2007) Plant speciation. Science 317:910–914
Rieseberg LH, Archer MA, Wayne RK (1999) Transgressive segregation, adaptation and
speciation. Heredity 83:363–372
Rosser EM (1955) A new British species of Senecio. Watsonia 3:228–232
Salone V, Rudinger M, Polsakiewicz M, Hoffmann B, Groth-Malonek M, Szurek B, Small I,
Knoop V, Lurin C (2007) A hypothesis on the identification of the editing enzyme in plant
organelles. FEBS Lett 581:4132–4138
Salmon A, Ainouche ML (2010) Polyploidy and DNA methylation: new tools available. Mol
Ecol 19:213–215
Salmon A, Ainouche ML, Wendel JF (2005) Genetic and epigenetic consequences of recent
hybridization and polyploidy in Spartina (Poaceae). Mol Ecol 14:1163–1175
Soltis PS, Soltis DE (1999) Polyploidy: recurrent formation and genome evolution. Trends Ecol
Evol 14:348–352
Stebbins GL (1957) Self fertilization and population variability in higher plants. Am Nat
91:337–354
Stupar RM, Hermanson PJ, Springer NM (2007) Nonadditive expression and parent-of-origin
effects identified by microarray and allele-specific expression profiling of maize endosperm.
Plant Physiol 145:411–425
Sun M, Ritland K (1998) Mating system of yellow starthistle (Centaurea solstitialis), a successful
colonizer in North America. Heredity 80:225–232
Urbanska KM, Hurka H, Landolt E, Neuffer B, Mummenhoff K (1997) Hybridization and
evolution in Cardamine (Brassicaceae) at Urnerboden, central Switzerland: biosystematics
and molecular evidence. Plant Syst Evol 204:233–256
Vincent PLD (1996) Progress on clarifying the generic concept of Senecio based on an extensive
world-wide sample of taxa. In Hind DJN, Beentje HJ (eds) Compositae: systematics.
proceedings of the international compositae conference Kew 1994, vol 1, Royal Botanic
Gardens, Kew, pp 597–611
Wang J, Tian L, Lee H-Y, Chen ZJ (2006a) Nonadditive regulation of FRI and FLC loci mediates
flowering-time variation in Arabidopsis allopolyploids. Genetics 173:965–974
Wang J, Tian L, Lee H-S, Wei NE, Jiang H, Watson B, Madlung A, Osborn TC, Doerge RW,
Comai L, Chen ZJ (2006b) Genomewide nonadditive gene regulation in Arabidopsis
allotetraploids. Genetics 172:507–517
Weir J, Ingram R (1980) Ray morphology and cytological investigations of Senecio cambrensis
Rosser. Heredity 86:237–241
Wood TE, Takebayashi N, Barker MS, Mayrose I, Greenspoon PB, Rieseberg LH (2009) The
frequency of polyploid speciation in flowering plants. Proc Nat Acad Sci USA 106:13875–13879
Xiong LZ, Xu CG, Saghai Maroof MA, Zhang Q (1999) Patterns of cytosine methylation in an
elite rice hybrid and its parental lines, detected by a methylation-sensitive amplification
polymorphism technique. Mol Gen Genet 261:439–446
CO
RR
862
863
864
865
866
867
868
869
870
871
872
873
874
875
876
877
878
879
880
881
882
883
884
885
886
887
888
889
890
891
892
893
894
895
896
897
898
899
900
901
902
903
904
905
906
M. J. Hegarty et al.
UN
Editor Proof
270
13
F
Chapter No.:
PR
OO
Editor Proof
Author Queries
Details Required
Author’s Response
AQ1
Please provide the index term for this chapter.
AQ2
‘Rieseberg et al (2003)’ has been changed to ‘Rieseberg
et al. (1999)’ so that this citation matches the list.
AQ3
References ’ Hegarty and Hiscock (2002); James and
Abbott (2005); Ingram and Noltie (1989); Hegarty et al.
2009; Hiscock and Hegarty, unpublished are cited in the
text but not provided in the list please provide it or
delete these citations
AQ5
McClintock 1986 has been changed to McClintock 1984
so that this citation matches the list.
AQ6
‘Baird et al. (2008)’ has been changed to ‘Baird et al.
(2009)’ so that this citation matches the list.
AQ7
Kindly check and confirm the updated volume id, issue
id and page range for the reference ’Brennan et al
(2012)’
AQ8
Kindly check and confirm the updated year, volume,
issue id and page range for the reference ‘Pelser etal.
(2011)’.
UN
CO
RR
EC
TE
D
Query Refs.
Metadata of the chapter that will be visualized in
SpringerLink
Book Title
Polyploidy and Genome Evolution
Series Title
Chapter Title
The Early Stages of Polyploidy: Rapid and Repeated Evolution in Tragopogon
Copyright Year
2012
Copyright HolderName
Springer-Verlag Berlin Heidelberg
Corresponding Author
Family Name
Soltis
Particle
Given Name
Douglas E.
Suffix
Author
Division
Department of Biology
Organization
University of Florida
Address
32611, Gainesville, FL, USA
Email
dsoltis@botany.ufl.edu
Family Name
Buggs
Particle
Given Name
Richard J. A.
Suffix
Division
Department of Biology
Organization
University of Florida
Address
32611, Gainesville, FL, USA
Division
Organization
School of Biological and Chemical Sciences, Queen Mary University of
London
Address
E1 4NS, London, UK
Email
Author
Family Name
Barbazuk
Particle
Given Name
W. Brad
Suffix
Division
Department of Biology
Organization
University of Florida
Address
32611, Gainesville, FL, USA
Email
Author
Family Name
Chamala
Particle
Given Name
Srikar
Suffix
Division
Department of Biology
Organization
University of Florida
Address
32611, Gainesville, FL, USA
Email
Author
Family Name
Chester
Particle
Given Name
Michael
Suffix
Division
Department of Biology
Organization
University of Florida
Address
32611, Gainesville, FL, USA
Email
Author
Family Name
Gallagher
Particle
Given Name
Joseph P.
Suffix
Division
Department of Biology
Organization
University of Florida
Address
32611, Gainesville, FL, USA
Division
Department of Ecology, Evolution, and Organismal Biology
Organization
Iowa State University
Address
50011, Ames, IA, USA
Email
Author
Family Name
Schnable
Particle
Given Name
Patrick S.
Suffix
Division
Organization
Center for Plant Genomics, Iowa State University
Address
50011, Ames, IA, USA
Email
Author
Family Name
Soltis
Particle
Given Name
Pamela S.
Suffix
Division
Organization
Florida Museum of Natural History, University of Florida
Address
32611, Gainesville, FL, USA
Email
Abstract
Elucidating the causes and consequences of polyploidy (whole-genome duplication; WGD) is arguably central
to understanding the evolution of most eukaryotic lineages. However, much of what we know about these
processes is derived from the study of crops and synthetic polyploids. Tragopogon provides the unique
opportunity to investigate the genetic and genomic changes that occur across an evolutionary series from
F1 hybrids, synthetic allopolyploids, independently formed natural populations of T. mirus and T.
miscellus that are 60–80 years post-formation, to older Eurasian polyploids that are dated by molecular clocks
at several million years old, and finally to a putative ancient polyploidization thought to have occurred prior
to or early in the history of the Asteraceae (40–43 mya). Tragopogon joins other well-studied natural polyploid
systems (e.g., Glycine, Nicotiana, Gossypium, Spartina, Senecio), but presents a range of research possibilities
that is not available in any other system. We have shown in T. mirus and T. miscellus that upon
allopolyploidization, massive gene loss occurs in patterns that are repeated across populations of independent
origin and with a bias against genes derived from T. dubius, the diploid parent shared by both new
allotetraploids. We have also shown significant changes in gene expression (transcriptomic shock) in the early
generations of allopolyploidy in these species. Massive and repeated patterns of chromosomal variation
(intergenomic translocations and aneuploidy) have been revealed by fluorescence in situ hybridization.
Aneuploidy results in substitutions between homeologous chromosomes, through reciprocal monosomytrisomy (1:3 copies) or nullisomy-tetrasomy (0:4 copies). We propose that substantial chromosomal instability
results in karyotype restructuring, a likely common process following WGD and a driver of allopolyploid
speciation, which has largely unexplored implications for gene losses, gains, and expression patterns. But
gene loss and expression changes as well as karyotypic changes are ongoing in T. mirus and T. miscellus, in
that no population is fixed for any of these events; thus, we have literally caught evolution in the act.
6
7
8
9
10
11
12
13
14
15
16
17
18
F
Douglas E. Soltis, Richard J. A. Buggs, W. Brad Barbazuk, Srikar
Chamala, Michael Chester, Joseph P. Gallagher, Patrick S. Schnable
and Pamela S. Soltis
PR
OO
5
Abstract Elucidating the causes and consequences of polyploidy (whole-genome
duplication; WGD) is arguably central to understanding the evolution of most
eukaryotic lineages. However, much of what we know about these processes is derived
from the study of crops and synthetic polyploids. Tragopogon provides the unique
opportunity to investigate the genetic and genomic changes that occur across an
evolutionary series from F1 hybrids, synthetic allopolyploids, independently formed
natural populations of T. mirus and T. miscellus that are 60–80 years post-formation, to
older Eurasian polyploids that are dated by molecular clocks at several million years
old, and finally to a putative ancient polyploidization thought to have occurred prior to
or early in the history of the Asteraceae (40–43 mya). Tragopogon joins other wellstudied natural polyploid systems (e.g., Glycine, Nicotiana, Gossypium, Spartina,
Senecio), but presents a range of research possibilities that is not available in any other
D
4
The Early Stages of Polyploidy: Rapid
and Repeated Evolution in Tragopogon
TE
3
Chapter 14
EC
2
Book ISBN: 978-3-642-31441-4
Page: 271/291
D. E. Soltis (&) R. J. A. Buggs W. B. Barbazuk S. Chamala
M. Chester J. P. Gallagher
Department of Biology, University of Florida, Gainesville,
FL 32611, USA
e-mail: dsoltis@botany.ufl.edu
CO
RR
1
Book ID: 272454_1_En
Date: 16-8-2012
Present Address:
J. P. Gallagher
Department of Ecology, Evolution, and Organismal Biology,
Iowa State University, Ames, IA 50011, USA
R. J. A. Buggs
School of Biological and Chemical Sciences, Queen Mary University of London,
London, E1 4NS, UK
P. S. Schnable
Center for Plant Genomics, Iowa State University, Ames, IA 50011, USA
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: 14
P. S. Soltis
Florida Museum of Natural History, University of Florida, Gainesville,
FL 32611, USA
P. S. Soltis and D. E. Soltis (eds.), Polyploidy and Genome Evolution,
DOI: 10.1007/978-3-642-31442-1_14, Springer-Verlag Berlin Heidelberg 2012
271
Layout: T1 Standard SC
Chapter No.: 14
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 272/291
23
24
25
26
27
28
29
30
31
32
33
F
21
22
system. We have shown in T. mirus and T. miscellus that upon allopolyploidization,
massive gene loss occurs in patterns that are repeated across populations of independent origin and with a bias against genes derived from T. dubius, the diploid parent
shared by both new allotetraploids. We have also shown significant changes in gene
expression (transcriptomic shock) in the early generations of allopolyploidy in these
species. Massive and repeated patterns of chromosomal variation (intergenomic
translocations and aneuploidy) have been revealed by fluorescence in situ hybridization. Aneuploidy results in substitutions between homeologous chromosomes, through
reciprocal monosomy-trisomy (1:3 copies) or nullisomy-tetrasomy (0:4 copies).
We propose that substantial chromosomal instability results in karyotype restructuring,
a likely common process following WGD and a driver of allopolyploid speciation,
which has largely unexplored implications for gene losses, gains, and expression
patterns. But gene loss and expression changes as well as karyotypic changes are
ongoing in T. mirus and T. miscellus, in that no population is fixed for any of these
events; thus, we have literally caught evolution in the act.
PR
OO
19
20
D. E. Soltis et al.
35
14.1 Introduction
36
14.1.1 General Introduction
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
Polyploidy, or whole-genome duplication (WGD), is currently recognized as a major
evolutionary force in eukaryotes (e.g., Mable 2003; Gregory and Mable 2005; Mable
et al. 2011). Polyploidy generally results in instant speciation, increasing biodiversity
and providing new genetic material for evolution (e.g., Levin 1983, 2002). Some
evidence suggests that two polyploid events occurred in ancestors of vertebrates (Ohno
1970; Panapoulou and Poustka 2005; see Chap. 16 of this volume), with subsequent
polyploidy in amphibians and fish (Mable et al. 2011; see Chaps. 18 and 17, respectively, this volume). The genomes of yeast and other Saccharomyces also appear to be
anciently duplicated (Wolfe and Shields 1997; Kellis et al. 2004; Dujon et al. 2004; see
Chap. 15 of this volume). Researchers have long recognized that polyploidy is an
inseparable part of angiosperm biology; the polyploidy process has, in fact, been
studied in plants for a little over a century. Early reviews of polyploidy in plants include
now classic papers by Müntzing (1936), Darlington (1937), Clausen et al. (1945), Löve
and Löve (1949), and Stebbins (1950, 1971). However, these seminal works did not
anticipate the huge role for polyploidy in evolution that genomic studies now suggest.
During the past decade there has been a tremendous resurgence of interest in
polyploidy, stimulated in large part by the development of increasingly powerful
genetic and genomic tools. The result has been numerous new insights into the
genomic and genetic consequences of polyploidy (other chapters of this volume).
Recent discoveries have dramatically reshaped traditional views and concomitantly revealed that polyploidy is a highly dynamic and ubiquitous process. For
example, studies of many duplicated genes across genomes suggest that all
EC
38
CO
RR
37
TE
D
34
UN
Editor Proof
272
Layout: T1 Standard SC
Chapter No.: 14
Book ISBN: 978-3-642-31441-4
Page: 273/291
The Early Stages of Polyploidy
273
82
14.1.2 Introduction to Tragopogon
66
67
68
69
70
71
72
73
74
75
76
77
78
79
80
83
84
85
86
87
88
89
90
91
92
93
94
95
96
97
98
99
PR
OO
65
D
64
TE
63
EC
61
62
Tragopogon now provides a well-known, textbook example of recent allopolyploid speciation; two new allotetraploid species originated within the last 80 years
(Soltis et al. 2004, 2009b) (Fig. 14.1). Tragopogon mirus and T. miscellus
(Fig. 14.1) formed repeatedly following the introduction of three diploids from
Europe into the Palouse region of North America in the early 1900s (Ownbey
1950; Soltis et al. 2004); the tetraploids have not formed in Europe. The parentage
of T. mirus (T. dubius and T. porrifolius) and T. miscellus (T. dubius and
T. pratensis) is well documented (Figs. 14.1 and 14.2) and confirmed with multiple
markers and approaches (Soltis et al. 1995, 2004, 2009b).
Ownbey (1950) described the few populations of the newly formed allotetraploids (each consisting of fewer than 100 individuals) as ‘‘small and precarious’’, but noted that they had ‘‘attained a degree of success’’ and they appeared to
be ‘‘competing successfully’’ with their diploid parents. He also stated that it
would be ‘‘important to follow the ecological development of the newly formed
polyploids’’ through time. Significantly, both tetraploids have been highly successful since their formation. Novak et al. (1991) conducted a survey to determine
the distributions of the two polyploids 40 years after Ownbey’s discovery. One or
CO
RR
60
F
81
angiosperms have undergone at least one round of genome doubling. Significantly,
genomic and phylogenetic analyses also associate polyploidy with major diversifications (e.g., within Poales, Solanaceae, Fabaceae; Soltis et al. 2009a), and the
origin of angiosperms and seed plants (Jiao et al. 2011).
As a result of a diverse array of studies, we have learned a great deal about the
interactions that occur among the diploid genomes forced together via allopolyploidy. Because of its apparent prevalence, elucidating the causes and consequences of polyploidy is arguably central to understanding the origin and
diversification of most major lineages of eukaryotes. Significantly, however, most
of what we know about the genetic and genomic consequences of polyploidy is
derived from the study of synthetic polyploids, crops (e.g., cotton, wheat, Brassica,
Nicotiana), and genetic models (Arabidopsis) (see refs above, Chen et al. 2004;
Soltis and Soltis 2009). To understand better how polyploidization impacts genome evolution and gene function in natural populations, we must extend from a
few crops, genetic models, and synthetics to naturally occurring polyploids.
Three systems are known that permit insights into the early stages of polyploidy
in nature: Spartina anglica (Ainouche et al. 2004, 2009; Salmon et al. 2005; see
Chap. 12 of this volume), Senecio cambrensis (Ashton and Abbott 1992; Abbott
and Lowe 2004; Hegarty et al. 2005, 2006; see Chap. 13, this volume), and
Tragopogon (T. mirus, T. miscellus, Soltis et al. , 2009b). Research has progressed
on all, and they are complementary. In addition, a2004s stressed here, Tragopogon
also affords unique opportunities to investigate polyploidy over a continuum of
ages, as well as the consequences of recent and frequently repeated polyploidy.
59
UN
Editor Proof
14
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 14
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 274/291
D. E. Soltis et al.
106
14.1.3 Tragopogon as a Unique Evolutionary Model
111
112
113
114
115
116
117
118
119
120
121
122
123
124
125
126
127
128
129
130
131
132
133
134
135
136
137
138
139
140
PR
OO
109
110
For many reasons Tragopogon is a novel system that affords the opportunity to
examine the early stages of polyploidization in nature. The natural populations are
approximately 80 years (40 generations in these biennials) old—this time frame
and the fact that they have experienced natural selection provide a window into
polyploidization that cannot be matched via the study of synthetic polyploids (e.g.,
crops and genetic models). Furthermore, molecular studies suggest that the diploid
parents diverged *2.5–5 MYA and that there may be as many as 21 lineages of
separate origin of T. miscellus and 11 of T. mirus just in the Palouse (Soltis et al.
1995, 2004, 2009b; Symonds et al. 2010); the polyploids have also formed in
Arizona, Oregon, Wyoming, and Montana (Soltis et al. 2012; Ownbey unpublished
data). Recent studies employing microsatellite markers (Symonds et al. 2010)
reveal multiple origins on a small geographic scale (see Sect. 14.1.4). Given that
multiple polyploidizations are common in plants (Soltis and Soltis 1993, 1999,
2000, 2009), Tragopogon represents in microcosm what occurs in other polyploids
over much larger geographic areas and longer time frames. These repeated origins
in a small geographic area and a narrow time frame also provide the unique
opportunity to ask if evolution repeats itself across these many lineages.
Adding to the utility of the Tragopogon system as an evolutionary model is the recent
production of multiple synthetic lines of both T. mirus and T. miscellus (Tate et al.
2009a), providing the added opportunity of examining both species from polyploidization onward. Morphologically, the synthetics resemble the natural polyploids with
short- and long-liguled forms of T. miscellus resulting when T. pratensis and T. porrifolius are reciprocally crossed (Tate et al. 2009a). In nature, all formations of T. mirus
have T. porrifolius as the maternal parent and T. dubius as the paternal parent, but we
have synthesized T. mirus reciprocally. We also produced allotetraploids between
T. porrifolius and T. pratensis, which are not known from nature (Fig. 14.1). All of these
synthetic lines are now in the fourth generation and offer the unique opportunity for
comparative study of repeated formations of both natural and synthetic polyploids.
Further adding to the allure of Tragopogon are Old World polyploids that are
much older than the recently formed New World polyploids. Tragopogon comprises *150 species, 12 of which are Eurasian polyploids (Mavrodiev et al.
2008a). Of several Eurasian polyploids for which we clarified parentage
(Mavrodiev et al. 2008a, b, c), T. castellanus (2n = 24) from Spain (Blanca and
Díaz de la Guardia 1996) has emerged as a promising new model. Estimating the
D
107
108
TE
104
EC
102
103
CO
RR
101
F
105
both polyploids were found in most towns of the Palouse with populations ranging
from small (fewer than 100 individuals) to many thousands of individuals.
Tragopogon miscellus is now one of the most common weeds in and around
Spokane, WA, as well as in Moscow, ID, and Spangle, WA. Populations of
T. mirus and T. miscellus often form dense stands and are, in fact, displacing their
parents, particularly T. pratensis and T. porrifolius.
100
UN
Editor Proof
274
Layout: T1 Standard SC
Chapter No.: 14
Book ISBN: 978-3-642-31441-4
Page: 275/291
The Early Stages of Polyploidy
275
EC
TE
D
PR
OO
F
Editor Proof
14
Book ID: 272454_1_En
Date: 16-8-2012
142
143
144
145
146
147
148
date of origin of a polyploid is difficult, but given well-known caveats, we used
DNA sequence data to estimate the age of T. castellanus as 0.8–2.8 million years
(Mavrodiev et al. 2008a, b, c). This agrees with other evidence in that T. castellanus occurs in well-known Pleistocene glacial refugia that harbor other paleoendemics (e.g., Petit et al. 2003). Thus, in Tragopogon we have the opportunity to
extend our analyses to older Eurasian polyploids, providing a continuum of ages
from F1 hybrids and raw synthetics, to 80-year-old natural polyploids, to natural
polyploids that are perhaps several million years old.
UN
141
CO
RR
Fig. 14.1 Summary of parentage of tetraploid Tragopogon species comparing natural with
synthetic allopolyploids. The diploid parents (with 2n = 12) are at the corners of the triangle;
polyploids (2n = 24) are in between the corners. Synthetic polyploids are on the outside of the
triangle (connected by white lines); those polyploids forming naturally are to the inside of the
triangle. Polyploids are placed closer to the maternal parent. In nature, T. miscellus has formed
reciprocally, and T. mirus has formed only with T. porrifolius as the maternal parent. However,
we have made reciprocal synthetic lines of both and have also made reciprocal polyploids of
T. pratensis 9 T. porrifolius; this polyploid has not formed in nature. Note that populations of
T. miscellus of reciprocal origin differ in morphology. Those with T. pratensis as the maternal
parent have short ligules, and those with T. dubius as the maternal parent have long ligules
Layout: T1 Standard SC
Chapter No.: 14
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 276/291
149
D. E. Soltis et al.
14.1.4 Origins of Species
174
14.2 Does Evolution Repeat Itself?
156
157
158
159
160
161
162
163
164
165
166
167
168
169
170
171
172
175
176
177
178
179
180
181
182
183
184
185
186
187
PR
OO
155
D
154
TE
153
EC
152
CO
RR
151
F
173
The Tragopogon system is comparable in some ways to an island biogeography
scenario. The diploids and the derivative polyploids only occur in small towns that
are scattered across the Palouse area of eastern Washington and adjacent Idaho and
not in the intervening areas, which are large tracts of agricultural land. This begged
the question—how did the polyploids spread so quickly to many towns? Certainly,
seed dispersal is one possibility given the wind-dispersed nature of the achenes.
Molecular data indicate instead that repeated formation has played the major role
in range expansion. This was suspected by Ownbey and McCollum (1953), who
surmised that T. miscellus had formed reciprocally with the long-liguled form
(found only in Pullman, WA) having T. dubius as the maternal parent and all other
populations having T. pratensis as the maternal parent; this reciprocal parentage
results in distinctive morphologies (Fig. 14.1) and was later confirmed by
molecular methods (Soltis and Soltis 1989).
A suite of molecular markers including allozymes, AFLPs, sequence data, and
microsatellites has now revealed that most populations of the allotetraploids are of
distinct origin (Soltis et al. 2004; Symonds et al. 2010). Furthermore, microsatellites
have documented multiple origins on a fine geographic scale, revealing in several
cases that distinct populations in the same town separated by only 1–2 km are of
separate origin (Symonds et al. 2010). Interestingly, microsatellite data reveal that of
the many genotypes of T. dubius currently in nature, only three general types appear
to have contributed to the repeated formations of both polyploids, and there are no
exact matches to present-day T. dubius genotypes. Hence, the genotypes detected in
the two polyploid species appear to represent a snapshot of the historical population
structure in the diploid progenitors, rather than modern diploid genotypes.
150
Evolutionary biologists have long wondered if evolution would repeat itself, given
the chance. Gould (1994) suggested that, on a broad evolutionary scale, if we
could replay the evolutionary tape of life on Earth, it would play differently—
‘‘history involves too much chaos,’’ and too many chance events are involved for
the evolutionary process to be repetitive. He stated that ‘‘chains of historical events
are so intricate, so imbued with random and chaotic elements, so unrepeatable in
encompassing such a multitude of unique objects, that standard models of simple
prediction and replication do not apply.’’ In contrast, other researchers have argued
that ‘‘within certain limits the outcome of evolutionary processes might be rather
predictable’’ (Morris 1998; see also Stern et al. 2009). However, is this true on a
finer scale? Are certain aspects of the polyploidy process actually ‘‘hard-wired’’?
Preservation of duplicated gene copies following genome duplication appears far
from random, with specific functional categories preferentially retained (Blanc and
UN
Editor Proof
276
Layout: T1 Standard SC
Chapter No.: 14
Book ISBN: 978-3-642-31441-4
Page: 277/291
The Early Stages of Polyploidy
277
TE
D
PR
OO
F
Editor Proof
14
Book ID: 272454_1_En
Date: 16-8-2012
190
191
192
193
194
195
196
197
198
199
200
201
202
Wolfe 2004; Freeling 2009) and reduplicated in subsequent polyploidizations
(Paterson et al. 2006). Independent WGDs in the ancestors of Arabidopsis, Oryza
(rice), Saccharomyces (yeast), and Tetraodon (pufferfish) have been followed by
convergent fates of many gene families (Paterson et al. 2006). Collectively, these
observations indicate that at deep timescales there may exist certain ‘‘principles’’
that govern the fates of gene and genome duplications. But is this true in the early
stages following polyploid formation?
In the sections that follow we review data from different sets of molecular
markers as well as chromosomal data across polyploid populations of separate
origin and ask if aspects of polyploid evolution are indeed hard-wired or if
stochastic processes prevail.
CO
RR
189
UN
188
EC
Fig. 14.2 Maps showing the location of Tragopogon allotetraploid populations that are found
within the Palouse region of the Pacific Northwest of the U.S.A. Populations of T. miscellus
(squares) and T. mirus (circles) located in towns are indicated in the relevant counties of
Washington and Idaho. A contiguous urbanized area containing multiple T. miscellus populations
is indicated (shaded)
14.2.1 rDNA Loci/Concerted and Repeated Evolution
Concerted evolution, which results in the homogenization of gene sequences to
one type, is a common feature of ribosomal RNA genes (e.g., Zimmer et al. 1980).
In both T. mirus and T. miscellus, concerted evolution is ongoing, but incomplete
Layout: T1 Standard SC
Chapter No.: 14
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 278/291
D. E. Soltis et al.
225
14.2.2 Homeolog Loss and Gene Silencing
226
14.2.2.1 One Gene at a Time
211
212
213
214
215
216
217
218
219
220
221
222
223
227
228
229
230
231
232
233
234
235
236
237
238
239
240
241
242
PR
OO
209
210
D
207
208
TE
206
EC
205
Although Tragopogon affords unique opportunities for evolutionary study, it is not
a genetic model organism; hence, until recently, genetic resources have not been
available (but see Sect. 14.2.2.2), which has slowed research progress. As a result,
genetic and genomic changes in the newly formed tetraploids were initially
examined one gene at a time for upto 29 loci (Tate et al. 2006, 2009b; Buggs et al.
2009, 2010b; Koh et al. 2010). Initially, we used AFLP-cDNA display to screen
plants of T. miscellus, T. mirus, and parental diploids to look for promising candidate genes—that is, fragments that did not show additivity in the allopolyploids
as would be expected (Tate et al. 2006; Koh et al. 2010). Additional genes were
surveyed because they were orthologous to genes that were singletons in other
Asteraceae species (Buggs et al. 2009; Koh et al. 2010); the fate of such genes
seemed of particular interest in new polyploids.
The results of these one-gene-at-a-time surveys are presented in detail elsewhere (Tate et al. 2006, 2009b; Buggs et al. 2009, 2010b; Koh et al. 2010).
Significantly, most of the changes observed in populations of both young polyploids are homeolog loss events, which far outnumber gene-silencing events in all
CO
RR
204
F
224
(Kovarik et al. 2005). In contrast to 80-year-old natural polyploids, F1 hybrids
have equal contributions of the diploid parents, as do raw (S0) synthetic polyploids, as well as the earliest natural populations of T. mirus and T. miscellus (based
on DNA from herbarium specimens). But in all modern-day natural populations
except one, each representing a distinct origin, the rDNA type of T. dubius is
consistently in very low abundance, with either the T. pratensis rDNA type (in
T. miscellus) or T. porrifolius rDNA type (in T. mirus) in much greater abundance.
In only one population of T. mirus are the parental contributions balanced (Malinska et al. 2011). Thus, concerted evolution has consistently occurred in these
new polyploid lines of separate origin, and it has repeatedly operated ‘‘against’’
T. dubius, homogenizing those copies in the direction of the other parent. Surprisingly, despite being the least abundant in terms of rDNA gene copy number,
T. dubius is by far the most abundant transcript in natural polyploid populations
(Matyasek et al. 2007).
This bias against the rDNA cistron in natural polyploid populations is already
apparent in the S1 generation of synthetic polyploids (Malinska et al. 2010, 2011). For
example, in four lines of synthetic T. miscellus, only three individuals (4 %) had
balanced parental gene ratios while 65 individuals (92 %) inherited more T. pratensisorigin units than would be expected under additivity. In seven lines of synthetic
T. mirus, 32 individuals (29 %) exhibited balanced rDNA genotypes, 69 individuals
(63 %) showed more 35S rDNA of T. porrifolius origin than expected, and only 9
plants (8 %) had more T. dubius-origin rDNA (Malinska et al. 2010, 2011).
203
UN
Editor Proof
278
Layout: T1 Standard SC
Chapter No.: 14
Book ISBN: 978-3-642-31441-4
Page: 279/291
The Early Stages of Polyploidy
279
D
PR
OO
F
Editor Proof
14
Book ID: 272454_1_En
Date: 16-8-2012
TE
Fig. 14.3 Considerable variation from plant to plant within and among populations in the
amount of homeolog loss detected in natural populations of T. miscellus. Population and plant
designations are given on the x-axis; proportion of loci on the y-axis. Colors designate genotype
as indicated in the figure. D = T. dubius allele; P = T. pratensis allele
247
248
249
250
251
252
253
254
255
256
257
258
259
260
261
262
EC
245
246
populations examined of both polyploids. Furthermore, most of the homeolog
losses in both polyploids were from T. dubius, the diploid parent that is shared by
both T. mirus and T. miscellus.
It is also noteworthy that the same suite of genes consistently shows additivity
of the parental gene copies (no loss or silencing) in polyploid populations of
separate origin, whereas some of the genes analyzed consistently show some
evidence of loss across at least some of the populations surveyed. Thus, these early
analyses of Tragopogon polyploids are also in agreement with the hypothesis that
there may be some underlying ‘‘principles’’ to polyploidization at the genetic or
biochemical level.
But stochasticity is operating as well in these young polyploids. Although
homeolog loss is present in the polyploid populations, the process is ongoing, and
appears to be stochastic within individual populations. In no population examined
has silencing or loss been complete (i.e., observed in all individuals of a population). The amount of gene loss within populations varies (Fig. 14.3). Furthermore, these losses and gene silencing events were not detected in F1 hybrids or
early-generation synthetic lines (S1). Hence, loss of homeologs and gene silencing
are not immediate consequences of hybridization or polyploidization in Tragopogon, but appear to occur several generations after polyploid formation and for
certain genes.
CO
RR
244
UN
243
Layout: T1 Standard SC
Chapter No.: 14
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 280/291
280
D. E. Soltis et al.
263
F
PR
OO
Editor Proof
Fig. 14.4 Chromosome
diagram for the alloetraploid
T. miscellus (representative of
other allotetraploids)
illustrating terminology used
throughout this chapter—
chromosome, homeolog, and
allele
14.2.2.2 Tragopogon Goes Genomic
285
14.2.2.3 Genomic Insights
269
270
271
272
273
274
275
276
277
278
279
280
281
282
283
286
287
288
289
290
291
TE
268
EC
267
CO
RR
266
Using Sequenom genotyping, we examined patterns of homeolog presence/
absence in 70 sets of homeologs in 59 plants from five independently formed
populations of allotetraploid T. miscellus (Buggs et al. 2012). Extensive gene loss
occurred in the B40 generations since polyploidization in T. miscellus. An average
of *20 % of the 70 loci investigated in each plant of T. miscellus was missing one
or both alleles of one homeolog (excluding assays where neither homeolog was
UN
265
D
284
Although important insights were obtained in Tragopogon, as well as other nonmodel systems, using the methods described above, these approaches have
shortcomings. These analyses were based on examination of specific homeolog
pairs using cleaved amplified polymorphic sequence analysis (CAPS). This
approach uses restriction enzymes and hence has limitations, as discussed by
Gaeta and Pires (2010). In addition, such surveys are slow and labor-intensive
(reviewed in Soltis et al. 2009b; Buggs et al. 2009). Because of these limitations,
we have sought ways to quickly and inexpensively build a framework for
addressing genome-scale questions in Tragopogon.
Newly developed genomic resources quickly facilitated the use of Tragopogon as
a model for the study of recent and repeated polyploidization. Using a combination of
454 and Illumina sequencing of genomic and cDNA, we identified SNPs (singlenucleotide polymorphisms) between the parental species T. dubius and T. pratensis
for analysis of the allotetraploids (Buggs et al. 2010a, b). Thousands of SNPs
distinguish the three parental diploids. Following SNP discovery, we designed
primers for Sequenom analysis (see Buggs et al. 2010a). Briefly, Sequenom iPLEx
genotyping uses mass spectrometry to carry out high-throughput and highly accurate
genotyping with multiple SNPs multiplexed in one reaction (Jurinke et al. 2005). The
assays permitted detection of gene loss and changes in allele number (‘allele’ is used
here to designate alleles at both homologous and homeologous loci; Fig. 14.4) within
polyploids versus changes in gene expression (see Sect. 14.2.3 below).
264
Layout: T1 Standard SC
Chapter No.: 14
298
299
300
301
302
303
304
305
306
307
308
309
310
311
312
313
314
315
316
317
318
319
320
321
322
323
324
325
326
327
328
329
330
331
332
F
297
PR
OO
296
D
295
TE
294
detected). Approximately one-third of these cases were single-allele absences.
There was an overall bias toward loss of paternally derived homeologs. Assuming
that immediately after polyploidization each individual had two alleles from
T. dubius and two from T. pratensis at each pair of homeologous loci, on average
at least 7.7 % of the original 280 allele copies have been lost in an individual
plant’s lineage since polyploidization occurred. In only one case did we find that
absence of a homeolog was fixed in a population. The fact that few gene losses
were fixed in populations, and many individuals were missing single alleles,
indicates that allele loss is ongoing in these populations (Buggs et al. 2012).
These genomic data confirmed and extended our previous results and provided
more insight into the processes involved. Certain loci were repeatedly missing in
independently formed populations; the loci studied could be grouped into 12
clusters that followed recurrent patterns of presence/absence in populations with
unique origins. Therefore, evolution is repeated in separate lineages of independent origin.
Homeolog loss is also repeated at deeper phylogenetic scales. We compared
patterns of gene loss found in T. miscellus with patterns found in 12 other species
of Asteraceae that are considered ancient polyploids (Barker et al. 2008). Eighteen
genes in our study had GO categories that tended to be lost in Asteraceae (Barker
et al. 2008), and these had median homeolog absence of 7.0 % in T. miscellus. A
comparison of these 18 genes versus all other genes (i.e., those that tend to be
retained in Asteraceae, plus those with no preferential loss or retention: 45 genes,
with a median absence of 4.0 %) showed significantly higher homeolog absence in
the 18 genes. Thus, gene loss in T. miscellus, a very young Asteraceae polyploid,
appears to repeat patterns occurring in older Asteraceae polyploids.
One hypothesis for the repeated patterns of duplicate gene retention is the
gene balance hypothesis (Freeling 2009; Freeling and Thomas 2006; Birchler
et al. 2005; Papp et al. 2003). This hypothesis maintains that genes coding for
products that are highly connected (i.e., within protein complexes or biochemical
pathways) are dosage-sensitive in that they must be present in the nucleus in the
same number of copies as the genes for products with which they interact. Thus,
‘‘connected’’ genes are hypothesized to be retained together as duplicate copies
so as to preserve stoichiometry rather than reverting to singleton status one by
one over time. In contrast, genes whose products are less connected are dosageinsensitive and are expected to revert gradually to singleton status. Our analyses
so far suggest that dosage sensitivity may in fact be playing a role in Tragopogon
(Buggs et al. 2012).
EC
293
281
CO
RR
292
Book ISBN: 978-3-642-31441-4
Page: 281/291
The Early Stages of Polyploidy
UN
Editor Proof
14
Book ID: 272454_1_En
Date: 16-8-2012
14.2.3 Tissue-Specific Silencing
Divergence of duplicate gene expression patterns among tissues has been suggested as a precursor of future evolution (Ohno 1970). Expression of a gene
duplicate in a tissue where the progenitor copy was not expressed may indicate
Layout: T1 Standard SC
Chapter No.: 14
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 282/291
340
341
342
343
344
345
346
347
348
349
350
351
352
353
354
355
356
357
358
359
360
361
362
363
364
365
366
367
368
369
370
371
372
373
374
375
376
F
339
PR
OO
338
D
336
337
TE
335
neofunctionalization (Ohno 1970; Duarte et al. 2006; see Chap. 1 of this volume),
while division of ancestral patterns of tissue-specific expression among duplicates
suggests subfunctionalization (Lynch and Conery 2000; Rodin and Riggs 2003;
Duarte et al. 2006) and silencing of a gene duplicate in all tissues points to
nonfunctionalization (Duarte et al. 2006). Both neofunctionalization and subfunctionalization will lead to long-term retention of duplicated genes, whereas
silencing/nonfunctionalization will generally lead to loss of a duplicate. Tissuespecific expression of duplicated genes has been studied in older gene duplicates in
model organisms and crops (Adams et al. 2003; Duarte et al. 2006; Ganko et al.
2007; Semon and Wolfe 2008; Chaudhary et al. 2009), but in these species the
ancestral patterns of gene expression and the ages of duplicates are not known
precisely. Cases of tissue-specific expression patterns of very young gene duplicates are restricted to a few synthetic polyploids (Adams et al. 2003, 2004; Wang
et al. 2004; Chaudhary et al. 2009).
We examined the expression of 13 homeolog pairs in seven tissues of 10 plants
of T. mirus from two natural populations of independent origin (Buggs et al.
2010b). Of the 910 assays in T. mirus, 63 % showed expression of both homeologs, 7 % showed no expression of either homeolog, 20 % showed non-expression
of one homeolog across all tissues of a plant, and 8 % showed non-expression of a
homeolog in a particular tissue within a plant. We found two cases of reciprocal
tissue-specific expression between homeologs, potentially indicative of subfunctionalization. This study therefore showed that tissue-specific silencing, and even
apparent subfunctionalization, can arise rapidly in the early generations of natural
allopolyploidy. Similar results were found for 18 homeolog pairs using the same
approaches in T. miscellus populations (Buggs et al. 2011b).
In T. miscellus we also examined tissue-specific gene expression using the
previously described Sequenom assays (Buggs et al. 2011b). Tissue-specific
expression of 144 homeolog pairs in two natural populations was compared with
patterns of allelic expression in both in vitro ‘‘hybrids’’ and hand-crossed F1
hybrids between the parental diploids T. dubius and T. pratensis, and with patterns
of homeolog expression in synthetic (S1) allotetraploids. Tissue-specific homeolog
expression was frequent in natural allopolyploids, but F1 hybrids and S1 allopolyploids showed less tissue-specific homeolog expression than the natural allopolyploids and the in vitro ‘‘hybrids’’ of diploid parents. These results suggest that
‘‘transcriptomic shock’’ upon hybridization (McClintock 1984) includes the activation of allele/homeolog expression in all tissues, causing a loss of tissue-specific
expression patterns seen in the diploid parents. Such activation has seldom been
considered in terms of the tissue-specific activation of protein-coding genes.
Activation of homeologs has also been found in cotton F1 hybrids and allopolyploids (Chaudhary et al. 2009), who termed it ‘‘transcriptional neofunctionalization’’. We showed this to be widespread in Tragopogon. This may fit a scenario
in which activity of small interfering RNA molecules, which influence gene
expression, is temporarily lost in F1 hybrids and early allopolyploids, but restored
subsequently.
EC
334
CO
RR
333
D. E. Soltis et al.
UN
Editor Proof
282
Layout: T1 Standard SC
Chapter No.: 14
383
384
385
386
387
388
389
390
391
392
393
394
395
396
397
398
399
400
401
402
403
404
405
406
407
408
409
410
411
412
413
414
415
416
417
418
F
382
PR
OO
381
D
380
Despite enormous progress in our understanding of many aspects of polyploidy,
little attention has been paid to chromosomal constitution, structure, and organization. The New World Tragopogon allotetraploids illustrate the valuable role of
cytology in examining hybridization and polyploidy in plants. Our recent studies
have revealed high levels of chromosomal variation in *80-year-old natural
populations, as well as the newly synthesized allotetraploids T. mirus and T. miscellus. The earliest studies of T. mirus and T. miscellus identified mitotic complements of 2n = 24 and typically 12 bivalents at meiosis (Ownbey 1950;
Ownbey and McCollum 1954). This supported the expectation that allotetraploids
were chromosomally additive of the diploid progenitors, which are both 2n = 12.
Lim et al. (2008) used genomic and fluorescence in situ hybridization (GISH/
FISH) in a preliminary survey of several T. mirus and T. miscellus plants and
found a few plants of each species that were not chromosomally additive of the
diploid parents. Several natural T. mirus and T. miscellus plants were found to be
aneuploid, with intergenomic translocations. Although most of the aneuploid
individuals examined were 2n = 24, not all chromosomes were present in two
copies as expected. GISH conducted on synthetic T. mirus pollen mother cells at
the diplotene stage of meiosis showed allosyndetic pairing within multivalents.
Meiotic instability, in the form of anaphase bridges and lagging chromosomes, was
also observed by Tate et al. (2009a) in the first synthetic polyploid generation (S1)
in T. mirus and T. miscellus lines. Thus, gametes with aneuploid and/or rearranged
chromosome complements can be potentially generated as early as the first meiosis
following genome doubling, which has been shown to be the case for synthetic
allotetraploid B. napus (Szadkowski et al. 2010).
The preliminary work of Lim et al. (2008) prompted a detailed examination
using GISH and FISH of the chromosomal variation generated in six T. miscellus
populations of independent origin (Chester et al. 2012). In all six populations, both
aneuploidy and translocations were common (Fig. 14.5). Only 3 of the 58 plants
exhibited the expected additivity of the diploid parental karyotypes (with neither
aneuploidy nor translocations). Although approximately 70 % of polyploid plants
were aneuploid for one or more chromosomes, variation in copy number appears
to be constrained. Most plants were 2n = 24, and the total copy number for each
homeologous group of chromosomes was typically four as a result of aneuploidy
being reciprocal between homeologous chromosomes (Fig. 14.5). Thus, most
deviations from disomy were in the form of monosomy-trisomy or nullisomytetrasomy, between homeologous chromosomes. This pattern of extensive aneuploidy while maintaining the overall copy number closely resembles cytological
changes in synthetic neoallotetraploid Brassica napus (Xiong et al. 2011). Gene
dosage has been implicated as a major factor constraining chromosomal changes
such that imbalances, which arise, require compensation by chromosomes (compensatory aneuploidy) or homeologous segments (compensatory translocations).
TE
379
EC
378
283
14.2.4 Cytogenetic Insights
CO
RR
377
Book ISBN: 978-3-642-31441-4
Page: 283/291
The Early Stages of Polyploidy
UN
Editor Proof
14
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 14
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 284/291
D. E. Soltis et al.
PR
OO
F
Editor Proof
284
424
425
426
427
428
429
430
431
432
433
434
435
436
437
438
439
440
441
TE
422
423
EC
421
In T. miscellus, genome rearrangements via intergenomic translocations were
common; 76 % of the plants showed evidence of at least one translocation. The two
largest chromosomes (groups A and B) showed the highest incidence of translocations.
Across all T. miscellus plants at least six different translocation breakpoint positions
were observed along the A group chromosomes. In one T. miscellus population in
Spokane, WA, an A-chromosome intergenomic translocation appeared close to fixation, being homozygous in 8 of the 10 individuals analyzed. Individuals from six
populations also appear to have undergone reciprocal translocations on the B-group
chromosomes, but they mostly occur at a similar position, near the end of the long arm.
The observed chromosomal changes in the recently formed Tragopogon allotetraploids provide two possible explanations for the loss of DNA that has been
observed using molecular SNP-based assays. (1) If a translocation was in a nonreciprocal state and homozygous, this could lead to the loss of DNA from one of
the parental diploids in the translocated region. (2) Nullisomy would lead to the
complete loss of DNA from one of the parental chromosomes. This may have
contributed to the clustering of patterns of gene loss and retention found using
Sequenom assays (see above). However, the diverse patterns of homeolog losses
that have been detected are not readily explainable by only these large-scale
chromosomal changes detectable with GISH.
CO
RR
420
UN
419
D
Fig. 14.5 Mitotic GISH karyotypes of three aneuploid T. miscellus individuals. Chromosomes
derived from each parental genome are shown, i.e., the T. dubius genome (D-subgenome) and the
T. pratensis genome (P-subgenome). Chromosomes deviating from disomy are as follows: top,
monosomy: trisomy for E chromosomes; middle, nullisomy: tetrasomy for E chromosomes;
bottom, four cases of monosomy: trisomy for chromosomes B, C, D, and E. All three individuals
are 2n = 24 and have four copies of each homeologous group (A–F). Scale bar: 5 lm
14.3 Comparing Tragopogon to Other Well-Studied Systems
Genome evolution in other well-studied polyploid systems (e.g., Gossypium,
Triticum, Nicotiana, Arabidopsis, Brassica, Senecio, Spartina) exhibits important
similarities and differences. Tragopogon is noteworthy in that initial studies show
Layout: T1 Standard SC
Chapter No.: 14
449
450
451
452
453
454
455
456
457
458
459
460
461
462
463
464
465
466
467
468
469
470
471
472
473
474
475
476
477
478
479
480
481
482
483
484
485
486
F
448
PR
OO
447
D
445
446
that while true expression changes play a major role, homeolog losses are also very
prominent in these young polyploids (Tate et al. 2006, 2009b; Koh et al. 2010;
Buggs et al. 2009, 2010a, b, 2011b, 2012). Of the initial 23 genes analyzed in
T. miscellus, 15 showed homeolog loss in one or more plants from nature, and 8
showed true expression changes (results for T. mirus are comparable); these patterns were confirmed via genomic analyses of many more genes. In contrast, in
synthetic wheat (Triticum) and synthetic Arabidopsis thaliana and A. suecica
polyploids, as well as in tetraploid cotton (Gossypium), expression changes
dominate (Adams et al. 2003, 2004; Hovav et al. 2008a, b; Udall et al. 2006; Flagel
et al. 2008; Madlung et al. 2004; Kashkush et al. 2002; Wang et al. 2006).
Tragopogon may be most similar to the allotetraploid B. napus, in which most of
the apparent gene silencing events observed in later generations were due to gene
losses, most likely resulting from genomic rearrangements (Song et al. 1995; Gaeta
et al. 2007). Across 50 lines of B. napus, genetic changes are equally distributed
between the parental diploid genomes (Gaeta et al. 2007). The system is dynamic—
some lines become more ‘‘oleracea like’’ and others more ‘‘rapa like’’ in terms of
losses and corresponding expression differences. In Arabidopsis suecica allopolyploids, silencing of homeologs from one parent (A. thaliana) was observed more
frequently than the silencing of homeologs from the other parent, Arabidopsis
arenosa (Wang et al. 2006). In synthetic polyploids in Triticale, the contribution of
the rye genome (Secale cereale) is preferentially silenced (Ma et al. 2004; Ma et al.
2006). In cotton, tissue-specific subfunctionalization occurs for some loci (Adams
et al. 2003, 2004; Adams and Wendel 2004), but overall gene expression is biased
toward one parent (Udall et al. 2006; Flagel et al. 2008; Rapp et al. 2009; Flagel and
Wendel 2010). In both T. miscellus and T. mirus, homeologs of one diploid genome
(T. dubius) are more often lost or not expressed; that is, T. dubius is often the ‘‘loser
genome’’ based on the set of genes surveyed to date.
Tragopogon also exhibits cytogenetic similarities to Brassica. Both T. mirus
and T. miscellus exhibit numerous translocations as well as extensive aneuploidy
while maintaining the overall copy number expected in an allotetraploid (reciprocal monosomy: trisomy and nullisomy: tetrasomy); this closely resembles
cytological changes in synthetic neoallotetraploid B. napus (Xiong et al. 2011).
It is also now possible to compare results for Tragopogon with several of the
other recently formed natural polyploids. Great research progress has now been
made on four of the five polyploids known to have formed in the past two centuries. In addition to T. mirus and T. miscellus, numerous insights into recent
polyploidy have been obtained for Senecio cambrensis (also in the Asteraceae) and
Spartina anglica (Poaceae). Both these model systems are reviewed elsewhere in
this volume (see Chaps. 13 and 12, respectively). Interestingly, all of these recent
polyploids have formed following introductions of one or both progenitors into a
completely new geographic area. Some of these new polyploids have been successful in nature, particularly Spartina anglica, which is now distributed worldwide with major ecological impact (Ainouche et al. 2004, 2009; see Chap. 12 of
this volume). The Tragopogon polyploids are now major weeds in the Palouse
region of northwestern USA. In contrast, following initial range expansion,
TE
444
285
EC
443
CO
RR
442
Book ISBN: 978-3-642-31441-4
Page: 285/291
The Early Stages of Polyploidy
UN
Editor Proof
14
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 14
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 286/291
494
495
496
497
498
499
500
501
502
503
504
505
506
507
508
509
510
511
512
513
514
515
516
517
518
519
520
521
522
523
524
525
526
527
528
529
530
531
F
493
PR
OO
492
D
491
TE
489
490
Senecio cambrensis now seems to be disappearing from parts of its original range
(Abbott et al. 2007; see Chap. 13 of this volume).
Senecio cambrensis formed from parents of different ploidal levels (one parent
is itself a polyploid), whereas in Spartina anglica both parents are polyploids but
of the same ploidal level. In Senecio cambrensis there is an additional level of
complexity in that the diploid parent is itself of homoploid hybrid origin. In
contrast, both parents of the recent Tragopogon polyploids are diploid. The
multiple layers of recent hybridization and polyploidization in Senecio and
Spartina could influence the genetic and expression changes detected in the recent
polyploids in those genera.
Molecular studies have revealed striking genetic and genomic changes in all of
these recently formed polyploids. Transcriptomic shock has now been shown
following hybridization in both Senecio (Hegarty et al. 2005, 2006) and Tragopogon (Buggs et al. 2011b), and substantial expression changes have also been
reported in F1 hybrids and polyploids in Spartina (Cheilafa et al. 2010a, b) Thus,
in recent polyploids in all three genera, hybridization seems to result in a major
burst of altered gene expression.
Changes in gene expression have been shown to be important in all of the
systems, but have been best characterized in Spartina (Ainouche et al. 2009, Chap.
12 of this volume) and Senecio (Hegarty et al. 2008, Chap. 13 of this volume).
Methylation alterations have been detected in both Spartina (Salmon et al. 2005)
and Senecio (Hegarty et al. 2011), with epigenetic changes now considered to play
a major role in these new polyploids (Parisod et al. 2009; Ainouche et al. 2009;
Hegarty et al. 2011). However, epigenetic changes have not yet been analyzed in
Tragopogon. Dramatic changes in gene expression have been documented in
Spartina anglica, Senecio cambrensis, and the recently formed Tragopogon
polyploids. However, homeolog loss seems to have played a more prominent role
in young Tragopogon polyploids than in these other recent polyploids.
In both the recently formed Tragopogon polyploids and in S. cambrensis, genomic studies reveal that one parental genome predominates over the other in terms of
global patterns of expression and gene retention. In the allohexaploid S. cambrensis,
gene expression is more similar to that of the tetraploid parent (S. vulgaris) than to the
diploid parent, S. squalidus (Hegarty et al. 2006). In the young Tragopogon polyploids, homeologs of the diploid parent T. dubius are more often lost or not expressed.
That is, following polyploidy there are clear winner and loser parental genomes, and
these patterns are established quickly and repeatedly.
Chromosomal changes have been investigated in detail in both Tragopogon and
Spartina using FISH/GISH; recent polyploids in both genera exhibit substantial
change, but different types of variation are evident. In Spartina, nonaploid plants
(2n = ca. 90) were detected (rather than the expected 2n = 120–124), most likely
resulting from backcrosses between S. anglica and its maternal parent S. alterniflora
(Renny-Byfield et al. 2010). In Tragopogon, frequent translocations were detected as
well as frequent reciprocal monosomy: trisomy and nullisomy: tetrasomy—these
have not been previously reported from nature (Chester et al. 2012). Chromosomal
pairing abnormalities (multivalents) have been detected in both Tragopogon
EC
488
CO
RR
487
D. E. Soltis et al.
UN
Editor Proof
286
Layout: T1 Standard SC
Chapter No.: 14
Book ISBN: 978-3-642-31441-4
Page: 287/291
The Early Stages of Polyploidy
287
539
14.4 Conclusions
541
542
543
544
545
546
547
548
549
550
551
552
553
554
555
556
557
558
559
560
561
562
563
564
565
566
567
568
569
570
571
572
PR
OO
540
A diverse array of experimental approaches has helped us to understand the
evolution of young, natural Tragopogon polyploids at the genomic and transcriptomic levels. These have demonstrated: (1) the allopolyploids have multiple
origins; (2) substantial gene loss occurs within the first 40 generations in nature,
and is still ongoing; (3) patterns of gene loss are repeated among independent
origins of the allopolyploids; (4) transcriptomic shock occurs upon allopolyploidization, involving a reduction in tissue-specific expression; (5) major chromosomal changes have occurred and are ongoing in the natural allopolyploids.
The dynamic evolutionary processes that appear to be underway in Tragopogon
polyploids may be representative of those that have occurred in other groups, and
after past polyploidization events. Care is needed when extrapolating from a single
system, because patterns of change may be influenced by the genetic background
of the polyploid, such as the level of genetic differentiation between the two
parental species of an allopolyploid (reviewed in
Buggs et al. 2011a). Studies of polyploids in many plant families, of different
ages and of different parental combinations, are needed to provide a comprehensive understanding of the evolution of allopolyploids.
Tragopogon has great potential for further development as a model system for
polyploidy. Older Eurasian polyploids will allow placement of another time point
in the evolutionary series. Research is also underway on natural hybrids between
T. pratensis and T. porrifolius (T. 9 mirabilis) that occur in the United Kingdom.
Further development of genetic resources—sequencing of the T. dubius genome
and production of a genetic map—will allow new questions to be addressed.
In recent years, progress in understanding the genetic consequences of polyploidy has outstripped progress in understanding the ecological background in
which evolution occurs (but see Ramsey 2011). As a genus that occurs almost
exclusively in the wild, Tragopogon affords a system in which relevant ecological
studies can be carried out, comparing the fitness of polyploids and diploids, and the
different chromosomal variants of the polyploids. Such studies will contribute to a
comprehensive view of polyploid evolution.
D
537
TE
535
536
EC
534
CO
RR
533
F
538
allopolyploids as well as in synthetic polyploids (Ownbey and McCollum 1953; Tate
et al. 2009a). FISH/GISH has been problematic in Senecio, but traditional cytogenetic analysis suggests possible structural chromosomal changes, as well as some
meiotic pairing irregularities (Crisp 1972). In addition to generating genetic diversity, intergenomic recombination resulting in chromosome rearrangements could
lead to the formation of reproductive barriers between lineages of separate or
independent origin, which is under investigation in Tragopogon.
532
UN
Editor Proof
14
Book ID: 272454_1_En
Date: 16-8-2012
Acknowledgments Funding for this research was provided by the University of Florida and
NSF grants MCB-0346437, DEB-0614421, DEB-0919254, DEB-0922003, and DEB-0919348.
R.J.A.B. has been supported since March 2010 by NERC Fellowship NE/G01504X/1.
Layout: T1 Standard SC
Chapter No.: 14
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 288/291
D. E. Soltis et al.
References
574
575
576
577
578
579
580
581
582
583
584
585
586
587
588
589
590
591
592
593
594
595
596
597
598
599
600
601
602
603
604
605
606
607
608
609
610
611
612
613
614
615
616
617
618
619
620
621
622
623
624
Abbott RJ, Ireland HE, Rogers HJ (2007) Population decline despite high genetic diversity in the
new allopolyploid species Senecio cambrensis (Asteraceae). Mol Ecol 16:1023–1033
Abbott RJ, Lowe AJ (2004) Origins, establishment, and evolution of new polyploids species:
Senecio cambrensis and S. eboracensis in the British Isles. Biol J Linn Soc 82:467–474
Adams KL, Cronn R, Percifield R, Wendel JF (2003) Genes duplicated by polyploidy show
unequal contributions to the transcriptome and organ-specific reciprocal silencing. Proc Nat
Acad Sci USA 100:4649–4654
Adams KL, Percifield R, Wendel JF (2004) Organ-specific silencing of duplicated genes in a
newly synthesized cotton allotetraploid. Genetics 168:2217–2226
Adams KL, Wendel JF (2004) Exploring the genomic mysteries of polyploidy in cotton. Biol J
Linn Soc 82:573–581
Ainouche ML, Baumel A, Salmon A (2004) Spartina anglia C. E. Hubbard: a natural model
system for analyzing early evolutionary changes that affect allopolyploid genomes. Biol J
Linn Soc 82:475–484
Ainouche ML, Fortune PM, Salmon A, Parisod C, Grandbastien M-A, Ricou K, Fukunaga M,
Misset M-T (2009) Hybridization, polyploidy and invasion: Lessons from Spartina (Poaceae).
Biol Invasion. doi:10.1007s10530-0089383-2
Ashton PA, Abbott RJ (1992) Multiple origins and genetic diversity in the newly arisen
allopolyploid species Senecio cambrensis Rosser (Compositae). Heredity 68:25–32
Barker MS, Kane NC, Matvienko M, Kozik A, Michelmore RW, Knapp SJ, Rieseberg LH (2008)
Multiple Paleopolyploidizations during the evolution of the compositae reveal parallel
patterns of duplicate gene retention after millions of years. Mol Biol Evol 25:2445–2455
Birchler JA, Riddle NC, Auger DL, Veitia R (2005) Dosage balance in gene regulation:
biological implications. Trends Genet 21:219–226
Blanc G, Wolfe KH (2004) Functional divergence of duplicated genes formed by polyploidy
during Arabidopsis divergence. Plant Cell 16:1679–1691
Blanca G, Díaz de la Guardia C (1996) Sinopsis del género Tragopogon L. (Asteraceae) en la
Peninsula Ibérica. Anales del Jardín Botánico de Madrid 54:358–363
Buggs RJA, Doust AN, Tate JA, Koh J, Soltis K, Feltus FA, Paterson AH, Soltis PS, Soltis DE
(2009) Gene loss and silencing in Tragopogon miscellus (Asteraceae): comparison of natural
and synthetic allotetraploids. Heredity 103:73–81
Buggs RJA, Chamala S, Wu W, Gao L, May GD, Schnable PS, Soltis DE, Soltis PS, Barbazuk
WB (2010a) Characterization of duplicate gene evolution in the recent natural allopolyploid
Tragopogon miscellus by next-generation sequencing and Sequenom iPLEX genotyping. Mol
Ecol 19(1):1–15
Buggs RJA, Elliott NM, Zhang L, Koh J, Viccini LF, Soltis DE, Soltis PS (2010b) Tissue-specific
silencing of homoeologs in natural populations of the recent allopolyploid Tragopogon mirus.
New Phytol 186:175–183
Buggs RJA, Soltis PS, Soltis DE (2011a) Biosystematic relationships and the formation of
polyploids. Taxon 60:324–332
Buggs RJA, Zhang L, Miles N, Tate JA, Gao L, Schnable PS, Barbazuk WB, Soltis PS, Soltis DE
(2011b) Genomic and transcriptomic shock generate evolutionary novelty in a newly formed,
natural allopolyploid plant. Curr Biol 21:1–6
Buggs RJA, Gao L, Wu W, Chamala S, Tate JA, Schnable PS, Soltis DE, Soltis PS, Barbazuk WB
(2012) Rapid and repeated gene loss in a young polyploidy species. Curr Biol 22:248–252
Chaudhary B, Flagel L, Stupar RM, Udall JA, Verma N, Springer NM, Wendel JF (2009)
Reciprocal silencing, transcriptional bias and functional divergence of homoeologs in
polyploid cotton (Gossypium). Genetics 182:503–517
Chelaifa H, Mahe F, Ainouche M (2010a) Transcriptome divergence between the hexaploid saltmarsh sister species Spartina maritima and Spartina alterniflora (Poaceae). Mol Ecol
19:2050–2063
CO
RR
EC
TE
D
PR
OO
F
573
UN
Editor Proof
288
Layout: T1 Standard SC
Chapter No.: 14
289
EC
TE
D
PR
OO
F
Chelaifa H, Monnier A, Ainouche M (2010b) Transcriptomic changes following recent natural
hybridization and allopolyploidy in the salt marsh species Spartina x townsendii and Spartina
anglica (Poaceae). New Phytol 186:161–174
Chen ZJ, Wang J, Tian L, Lee HS, Wang JJ, Chen M, Lee JJ, Josefsson C, Madlung A, Watson B,
Pires JC, Lippman Z, Vaughn M, Colot V, Birchler JA, Doerge RW, Martienssen RA, Comai
L, Osborn TC (2004) The development of an Arabidopsis model system for genome-wide
analysis of polyploidy effects. Biol J Linn Soc 82:689–700
Chester M, Gallagher JP, Symonds VV, da Veruska Cruz Silva A, Mavrodiev EV, Leitch AR,
Soltis PS, Soltis DE (2012) Extensive and repeated patterns of chromosomal variation in
natural populations of a recently formed polyploid plant species. Proc Nat Acad Sci USA
109:1176–1181
Clausen J, Keck DD, Hiesey WM. 1945. Experimental studies on the nature of species II. Plant
evolution through amphiploidy and autopolyploidy, with examples from the Madiinae.
Publication 564, Carnegie Institute of Washington, Washington, DC
Crisp PC (1972) Cytotaxonomic studies in the section Annui of Senecio. Ph. D Thesis, University
of London
Darlington CD (1937) Recent advances in cytology, 2nd edn. The Blakiston Company,
Philadelphia
Duarte JM, Cui L, Wall PK, Zhang Q, Zhang X, Leebens-Mack J, Ma H, Altman N, dePamphilis
CW (2006) Expression pattern shifts following duplication indicative of subfunctionalization
and neofunctionalization in regulatory genes of Arabidopsis. Mol Biol Evol 23:469–478
Dujon B, Sherman D, Fischer G, Durrens P, Casaregola S, Lafontaine I, De Montigny J, Marck C,
Neuvéglise C, Talla E et al (2004) Genome evolution in yeasts. Nature 430:35–44
Flagel L, Udall J, Nettleton D, Wendel J (2008) Duplicate gene expression in allopolyploid
Gossypium reveals two temporally distinct phases of expression evolution. BMC Biol 6:11
Flagel LE, Wendel JF (2010) Evolutionary rate variation, genomic dominance and duplicate gene
expression during allotetraploid cotton speciation. New Phytol 186:184–193
Freeling M (2009) Bias in plant gene content following different sorts of duplication: tandem,
whole-genome, segmental, or by transposition. Annu Rev Plant Biol 60:433–453
Freeling M, Thomas BC (2006) Gene-balanced duplications, like tetraploidy, provide predictable
drive to increase morphological complexity. Genome Res 16:805–814
Gaeta RT, Pires JC (2010) Homoeologous recombination in allopolyploids: the polyploid ratchet.
New Phytol 186:18–28
Gaeta RT, Pires JC, Iniguez-Luy F, Leon E, Osborn TC (2007) Genomic changes in
resynthesized Brassica napus and their effect on gene expression and phenotype. Plant Cell
19:3403–3417
Ganko EW, Meyers BC, Vision TJ (2007) Divergence in expression between duplicated genes in
Arabidopsis. Mol Biol Evol 24:2298–2309
Gould SJ (1994) The evolution of life on Earth. Sci Am 271:85–86
Gregory TR, Mable BK (2005) Polyploidy in animals. In: Gregory TR (ed) The evolution of the
Genome. Elsevier/Academic, San Diego, pp 428–501
Hegarty MJ, Jones JM, Wilson ID, Barker GL, Coghill JA, Sanchez-Baracaldo P, Liu G, Buggs
RJA, Abbott RJ, Edwards KJ, Hiscock SJ (2005) Development of anonymous cDNA
microarrays to study changes to the Senecio floral transcriptome during hybrid speciation.
Mol Ecol 14:2493–2510
Hegarty MJ, Barker GL, Wilson ID, Abbott RJ, Edwards KJ, Hiscock SJ (2006) Transcriptome
shock after interspecific hybridization in Senecio is ameliorated by genome duplication. Curr
Biol 16:1652–1659
Hegarty MJ, Barker GL, Brennan AC, Edwards KJ, Abbott RJ, Hiscock SJ (2008) Changes to
gene expression associated with hybrid speciation in plants: further insights from transcriptomic studies in Senecio. Philos Trans R Soc London B series 363:3055–3069
Hegarty MJ, Batstone T, Barker GL, Edwards KJ, Abbott RJ, Hiscock SJ (2011) Nonadditive
changes to cytosine methylation as a consequence of hybridization and genome duplication in
Senecio (Asteraceae). Mol Ecol 20:105–113
CO
RR
625
626
627
628
629
630
631
632
633
634
635
636
637
638
639
640
641
642
643
644
645
646
647
648
649
650
651
652
653
654
655
656
657
658
659
660
661
662
663
664
665
666
667
668
669
670
671
672
673
674
675
676
677
678
Book ISBN: 978-3-642-31441-4
Page: 289/291
The Early Stages of Polyploidy
UN
Editor Proof
14
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 14
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 290/291
EC
TE
D
PR
OO
F
Hovav R, Udall J, Chaudhary B, Flagel L, Rapp R, Wendel J (2008a) Partitioned expression of
duplicated genes during development and evolution of a single cell in a polyploid plant. Proc
Nat Acad Sci USA 105:6191
Hovav R, Udall JA, Chaudhary B, Hovav E, Flagel L, Hu G, Wendel JF (2008b) The evolution of
spinnable cotton fiber entailed prolonged development and a novel metabolism. PLoS Genet 4:e2
Jiao Y, Wickett N, Ayyampalayam S, Chanderbali A, Landherr L, Ralph PE, Soltis PS, Soltis DE,
Clifton SE, Ma H, Leebens-Mack J, dePamphilis CW (2011) Phylogenomic analysis reveals
ancient genome duplications in seed plant and angiosperm history. Nature 473:97–100
Jurinke C, Denissenko MF, Oeth P, Ehrich M, van den Boom D, Cantor CR (2005) A single
nucleotide polymorphism based approach for the identification and characterization of gene
expression modulation using MassARRAY. Mutat Res 573:83–95
Kashkush K, Feldman M, Levy AA (2002) Gene loss, silencing, and activation in a newly
synthesized wheat allotetraploid. Genetics 160:1651–1659
Kellis M, Birren BW, Lander ES (2004) Proof and evolutionary analysis of ancient genome
duplication in the yeast Saccharomyces cerevisiae. Nature 428:617–624
Koh J, Tate JA, Soltis PS, Soltis DE (2010) Genomic and expression differences in natural
populations of the recently formed allotetraploid Tragopogon mirus (Asteraceae). BMC
Genomics 11:97
Kovarik A, Pires JC, Leitch AR, Lim KY, Sherwood A, Matyasek R, Rocca J, Soltis DE, Soltis
PS (2005) Rapid concerted evolution in two allopolyploids of recent and recurrent origin.
Genetics 169:931–944
Levin DA (1983) Polyploidy and novelty in flowering plants. Am Nat 122:1–25
Levin DA (2002) The role of chromosomal change in plant evolution. Oxford University Press,
New York
Lim KY, Soltis DE, Soltis PS, Tate JA, Matyasek R, Srubarova H, Kovarik A, Pires JC, Xiong
ZY, Leitch AR (2008) Rapid chromosome evolution in recently formed polyploids in
Tragopogon (Asteraceae). PLoS One 3:e3353
Löve A, Löve D (1949) The geobotanical significance of polyploidy. I. Polyploidy and latitude.
Portugaliae Acta Biologica Serie A, Suppl vol. pp 273–352
Lynch M, Connery JS (2000) The evolutionary fate and consequences of duplicate genes. Science
290:1151–1155
Ma XF, Fang P, Gustafson JP (2004) Polyploidization-induced genome variation in Triticale.
Genome 47:839–848
Ma XF, Fang P, Gustafson JP (2006) Timing and rate of genome variation in Triticale following
allopolyploidization. Genome 49:950–958
Mable B (2003) Breaking down taxonomic barriers in polyploidy research. Trends Plant Sci
8:582–590
Mable BK, Alexandrou MA, Taylor MI (2011) Genome duplications in amphibians and fish: an
extended synthesis. J Zool 284:151–182
Madlung A, Comai L (2004) The effect of stress on genome regulation and structure. Ann Bot
94:481 495
Malinska H, Tate JA, Matyasek R, Leitch AR, Soltis DE, Soltis PS, Kovarik A (2010) Similar
patterns of rDNA evolution in synthetic and recently formed natural populations of
Tragopogon (Asteraceae) allotetraploids. BMC Evol Biol 10:291
Malinska H, Tate JA, Mavrodiev E, Matyasek R, Lim KY, Leitch AR, Soltis DE, Soltis PS,
Kovarik A (2011) Ribosomal RNA genes evolution in Tragopogon: A story of New and Old
World allotetraploids and synthetic lines. Taxon 60:348–354
Matyasek R, Tate J, Lim YK, Srubaraova H, Koh J, Leitch A, Soltis DE, Soltis PS (2007)
Concerted evolution of rDNA in recently formed Tragopogon allotetraploids is typically
associated with an inverse correlation between gene copy number and expression. Genetics
176:2509–2519
Mavrodiev E, Soltis PS, Soltis DE (2008a) Parentage of six Old World polyploids in Tragopogon
L. (Asteraceae: Scorzonerinae) based on ITS. ETS and plastid sequence data. Taxon
57:1217–1232
CO
RR
679
680
681
682
683
684
685
686
687
688
689
690
691
692
693
694
695
696
697
698
699
700
701
702
703
704
705
706
707
708
709
710
711
712
713
714
715
716
717
718
719
720
721
722
723
724
725
726
727
728
729
730
731
732
D. E. Soltis et al.
UN
Editor Proof
290
Layout: T1 Standard SC
Chapter No.: 14
291
EC
TE
D
PR
OO
F
Mavrodiev E, Nawchoo I, Soltis DE, Soltis PS (2008b) Molecular data reveal that the tetraploid
Tragopogon kashmirianus Singh, a narrow endemic of Kashmir, is distinct from the North
American T. mirus M. Ownbey. Bot J Linn Soc 158:391–398
Mavrodiev EV, Albach DC, Speranza P (2008c) A new polyploid species of the genus
Tragopogon (Asteraceae, Cichorieae) from Russia. Novon 18:229–232
McClintock B (1984) The significance of responses of the genome to challenge. Science
226:792–801
Morris SC (1998) The crucible of creation: the Burgess shale and the rise of animals. Oxford
University Press, Oxford
Müntzing A (1936) The evolutionary significance of autopolyploidy. Hereditas 21:263–378
Novak SJ, Soltis DE, Soltis PS (1991) Ownbey Tragopogons—40 Years later. Am J Bot
78:1586–1600
Ohno S (1970) Evolution by gene duplication. Springer, Berlin
Ownbey M (1950) Natural hybridization and amphiploidy in the genus Tragopogon. Am J Bot
37:487–499
Ownbey M, McCollum G (1954) The chromosomes of Tragopogon. Rhodora 56:7–21
Ownbey M, McCollum GD (1953) Cytoplasmic inheritance and reciprocal amphiploidy in
Tragopogon. Am J Bot 40:788–796
Panopoulou G, Poustka AJ (2005) Timing and mechanism of ancient vertebrate genome
duplications—the adventure of a hypothesis. Trends Genet 10:559–567
Papp B, Pal C, Hurst LD (2003) Dosage sensitivity and the evolution of gene families in yeast.
Nature 424:194–197
Parisod C, Salmon A, Zerjal T, Tenaillon M, Grandbastien M-A, Ainouche M (2009) Rapid
structural and epigenetic reorganization near transposable elements in hybrid and allopolyploid genomes in Spartina. New Phytol 184:1003–1015
Paterson AH, Chapman BA, Kissinger J, Bowers JE, Feltus FA, Estill JC, Marler BS (2006)
Many gene and domain families have convergent fates following independent whole-genome
duplication events in Arabidopsis, Oryza, Saccharomyces and Tetraodon. Trends Genet
22:597–602
Petit RJ, Aguinagalde, JL de Beaulieu, C Bittkau, S Brewer, R Cheddadi, R Ennos, S Fineschi, D
Grivet, M Lascoux, A Mohanty, G Müller-Starck, B Musch, A Palmé, S Rendell, GG.
Vendramin (2003) Glacial refugia: hotspots but not melting pots of genetic diversity. Science
300:1563–1565
Ramsey J (2011) Polyploidy and ecological adaptation in wild yarrow. Proc Nat Acad Sci USA
108:6697–6698
Rapp RA, Udall JA, Wendel JF (2009) Genomic expression dominance in allopolyploids. BMC
Biol. 7:18
Renny-Byfield S, Ainouche M, Leitch IJ, Lim KY, Le Comber SC, Leitch AR (2010) Flow
cytometry and GISH reveal mixed ploidy populations and Spartina nonaploids with genomes
of S. alterniflora and S. maritima origin. Ann Bot 105:527–533
Rodin SN, Riggs AD (2003) Epigenetic silencing may aid evolution by gene duplication. J Mol
Evol 56:718–729
Salmon A, Ainouche ML, Wendel JF (2005) Genetic and epigenetic consequences of recent
hybridization and polyploidy in Spartina (Poaceae). Mol Ecol 14:1163–1175
Semon M, Wolfe KH (2008) Preferential subfunctionalization of slow-evolving genes after
allopolyploidization in Xenopus laevis. Proc Nat Acad Sci USA 105:8333–8338
Soltis DE, Soltis PS (1995) The dynamic nature of polyploid genomes. Proc Nat Acad Sci USA
92:8089–8091
Soltis DE, Soltis PS, Pires JC, Kovarik A, Tate JA, Mavrodiev E (2004) Recent and recurrent
polyploidy in Tragopogon (Asteraceae): cytogenetic, genomic and genetic comparisons. Biol
J Linn Soc 82:485–501
Soltis DE, Albert VA, Leebens-Mack J, Bell CD, Paterson A, Zheng C, Sankoff D, Wall PK,
Soltis PS (2009a) Polyploidy and angiosperm diversification. Am J Bot 96:336–348
CO
RR
733
734
735
736
737
738
739
740
741
742
743
744
745
746
747
748
749
750
751
752
753
754
755
756
757
758
759
760
761
762
763
764
765
766
767
768
769
770
771
772
773
774
775
776
777
778
779
780
781
782
783
784
785
Book ISBN: 978-3-642-31441-4
Page: 291/291
The Early Stages of Polyploidy
UN
Editor Proof
14
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 14
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 292/291
EC
TE
D
PR
OO
F
Soltis DE, Buggs RJA, Barbazuk WB, Schnable PS, Soltis PS (2009b) On the origins of species:
does evolution repeat itself in polyploid populations of independent origin? Cold spring
harbor symposia on quantitative biology, Vol. LXXIV
Soltis DE, Mavrodiev EV, Meyers, SC, Severns PM, Zhang L, Gitzendanner MA, Ayers T,
Chester M, Soltis PS (2012) Additional origins of Ownbey’s Tragopogon mirus. Bot Linn Soc
169:297–311
Soltis DE, Soltis PS (1989) Allopolyploid speciation in Tragopogon: Insights from chloroplast
DNA. Am J Bot. 76:1119–1124
Soltis DE, Soltis PS (1993) Molecular data and the dynamic nature of polyploidy. Crit Rev Plant
Sci 12:243–273
Soltis DE, Soltis PS (1999) Polyploidy: recurrent formation and genome evolution. Trends Ecol
Evol 14:348–352
Soltis PS, Soltis DE (2000) The role of genetic and genomic attributes in the success of
polyploids. Proc Nat Acad Sci (USA) 97:7051–7057
Soltis PS, Soltis DE (2009) The role of hybridization in plant speciation. Annu Rev Plant Biol
60:561–588
Soltis PS, Plunkett GM, Novak SJ, Soltis DE (1995) Genetic variation in Tragopogon species:
additional origins of the allotetraploids T. mirus and T. miscellus (Compositae). Am J Bot
82:1329–1341
Song KM, Lu P, Tang KL, Osborn TC (1995) Rapid genome change in synthetic polyploids of
Brassica and its implications for polyploid evolution. Proc Nat Acad Sci USA 92:7719–7723
Stebbins GL (1950) Variation and evolution in plants. Columbia, New York
Stebbins GL (1971) Chromosomal evolution in higher plants. Addison-Wesley, London
Stern DL, Orgogozo V (2009) Is genetic evolution predictable? Science 323:746–751
Symonds VV, Soltis PS, Soltis DE (2010) Dynamics of polyploid formation in Tragopogon
(Asteraceae): recurrent formation, gene flow, and population structure. Evolution 64:1984–2003
Szadkowski E, Eber F, Huteau V, Lod M, Huneau C, Belcram H, Coriton O, ManzanaresDauleux M, Delourme R, King G (2010) The first meiosis of resynthesized Brassica napus, a
genome blender. New Phytol 186:102–112
Tate JA, Ni Z, Scheen AC, Koh J, Gilbert CA, Lefkowitz D, Chen ZJ, Soltis PS, Soltis DE (2006)
Evolution and expression of homoeologous loci in Tragopogon miscellus (Asteraceae), a
recent and reciprocally formed allopolyploid. Genetics 173:1599–1611
Tate JA, Symonds VV, Doust AN, Buggs RJA, Mavrodiev EV, Soltis PS, Soltis DE (2009a)
Synthetic polyploids of Tragopogon miscellus and T. mirus (Asteraceae): 50 ? years after
Ownbey’s discovery. Am J Bot 96:979–988
Tate JA, Joshi P, Soltis K, Soltis PS, Soltis DE (2009b) On the road to diploidization?
Homoeolog loss in independently formed populations of the allopolyploid Tragopogon
miscellus (Asteraceae). BMC Plant Biol 9:80
Udall JA, Swanson JM, Nettleton D, Percifield RJ, Wendel JF (2006) A novel approach for
characterizing expression levels of genes duplicated by polyploidy. Genetics 173(3):1823–1827
Xiong Z, Gaeta RT, Pires JC (2011) Homoeologous shuffling and chromosome compensation
maintain genome balance in resynthesized allopolyploid Brassica napus. Proc Nat Acad Sci
USA 108:7908–7913
Wang JL, Tian L, Madlung A, Lee HS, Chen M, Lee JJ, Watson B, Kagochi T, Comai L, Chen ZJ
(2004) Stochastic and epigenetic changes of gene expression in Arabidopsis polyploids.
Genetics 167:1961–1973
Wang JL, Tian L, Lee HS, Wei NE, Jiang HM, Watson B, Madlung A, Osborn TC, Doerge RW,
Comai L, Chen ZJ (2006) Genomewide nonadditive gene regulation in Arabidopsis
allotetraploids. Genetics 172:507–517
Wolfe KH, Shields DC (1997) Molecular evidence for an ancient duplication of the entire yeast
genome. Nature 387:708–713
Zimmer EA, Martin SL, Beverley SM, Kan YW, Wilson AC (1980) Rapid duplication and loss of
genes coding for the q chains of hemoglobin. Proc Nat Acad Sci USA 77:2158–2162
CO
RR
786
787
788
789
790
791
792
793
794
795
796
797
798
799
800
801
802
803
804
805
806
807
808
809
810
811
812
813
814
815
816
817
818
819
820
821
822
823
824
825
826
827
828
829
830
831
832
833
834
835
836
837
838
D. E. Soltis et al.
UN
Editor Proof
292
Metadata of the chapter that will be visualized in
SpringerLink
Book Title
Polyploidy and Genome Evolution
Series Title
Chapter Title
Yeast as a Window into Changes in Genome Complexity Due to Polyploidization
Copyright Year
2012
Copyright HolderName
Springer-Verlag Berlin Heidelberg
Corresponding Author
Family Name
Conant
Particle
Given Name
Gavin C.
Suffix
Author
Division
MU Informatics Institute
Organization
University of Missouri
Address
65211, Columbia, MO, USA
Division
Division of Animal Sciences
Organization
University of Missouri
Address
Columbia, MO, USA
Email
conantg@missouri.edu
Family Name
Hudson
Particle
Given Name
Corey M.
Suffix
Division
MU Informatics Institute
Organization
University of Missouri
Address
65211, Columbia, MO, USA
Email
Abstract
Due to the long history of genetic analyses in yeasts and their experimental tractability, the yeast genome
duplication provides important perspectives on the genome and population-level processes that follow wholegenome duplication (WGD). We discuss the history of the discovery of the Saccharomyces cerevisiae WGD,
with special emphasis on the role of comparative genomics in its analysis. We then explore models of the
WGD shaped population and species divergence, both at a gene level (e.g., Dobzhansky-Muller
incompatibility) and from the perspective of recent work on secondary allopolyploidy in Saccharomyces
pastorianus. Finally, we explore the selective forces that act on the WGD-produced paralogs and shape their
patterns of loss and retention. In addition to discussing the dosage balance hypothesis as it applies to the yeast
WGD, we explore the role of the WGD in shaping several complex metabolic and regulatory phenotypes.
Book ISBN: 978-3-642-31441-4
Page: 293/307
Chapter 15
5
Corey M. Hudson and Gavin C. Conant
9
10
11
12
13
14
15
16
17
D
8
Abstract Due to the long history of genetic analyses in yeasts and their experimental tractability, the yeast genome duplication provides important perspectives
on the genome and population-level processes that follow whole-genome duplication (WGD). We discuss the history of the discovery of the Saccharomyces
cerevisiae WGD, with special emphasis on the role of comparative genomics in its
analysis. We then explore models of population and species divergence, both at a
gene level (e.g., Dobzhansky-Muller incompatibility) and from the perspective of
recent work on secondary allopolyploidy in Saccharomyces pastorianus. Finally,
we explore the selective forces that act on the WGD-produced paralogs and shape
their patterns of loss and retention. In addition to discussing the dosage balance
hypothesis as it applies to the yeast WGD, we explore the role of the WGD in
shaping several complex metabolic and regulatory phenotypes.
TE
6
7
EC
3
20
21
22
23
24
CO
RR
18
19
PR
OO
4
Yeast as a Window into Changes
in Genome Complexity Due
to Polyploidization
2
F
1
Book ID: 272454_1_En
Date: 16-8-2012
15.1 Introduction
Researchers have found remnants of ancient whole-genome duplications (WGDs)
preserved in the genomes of many and diverse eukaryotes. This book, in fact, is a
testament to that diversity, illustrating the sheer number of independent events in
plants as well as the evolutionarily basal events in vertebrates and other, more
recent WGDs in teleost fishes and frogs. Although we have still not fully validated
C. M. Hudson G. C. Conant
MU Informatics Institute, University of Missouri, Columbia, MO 65211, USA
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: 15
G. C. Conant (&)
Division of Animal Sciences, University of Missouri, Columbia, MO, USA
e-mail: conantg@missouri.edu
P. S. Soltis and D. E. Soltis (eds.), Polyploidy and Genome Evolution,
DOI: 10.1007/978-3-642-31442-1_15, Springer-Verlag Berlin Heidelberg 2012
293
Layout: T1 Standard SC
Chapter No.: 15
Book ID: 272454_1_En
Date: 16-8-2012
C. M. Hudson and G. C. Conant
48
15.2 Evidence for WGD in Yeast
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
49
50
51
52
53
54
55
56
57
58
59
60
61
62
63
64
65
PR
OO
30
D
29
TE
28
EC
27
Although early analyses of genes potentially created by the vertebrate 2R events
used phylogenetic approaches (Hughes 1999; Furlong and Holland 2002), most
current studies of WGD rely on one or both of two methods: (1) finding numerous
blocks of paralogous genes in multiple chromosomes with similar gene orders and
(2) clustering homologs into groups by measuring the rate of synonymous substitutions (Ks or dS). This second method assumes that the gene pairs created by
WGD cluster about some mean Ks value (Lynch and Conery 2000). The simultaneous application of both methods has been used to group multiple WGD events
within species (e.g., Arabidopsis thaliana and Tetraodon nigroviridis; Jaillon et al.
2004; Van de Peer et al. 2009). However, the yeast genomes present an interesting
challenge in this respect because the synonymous substitutions between yeast
paralogs produced by WGD (hereafter ohnologs; Wolfe 2000) are often saturated
(Byrne and Wolfe 2007). In other words, identical synonymous positions between
two ohnologs occur almost as often due to repeated convergent substitutions as due
to common ancestry, a fact pointed out by Smith (1987), who attempted to date
histone gene duplicates in yeast. While the genomic structure of the core histone
genes suggested that they were all duplicated simultaneously, these genes show
CO
RR
26
F
47
Susumo Ohno’s claim for the primacy of polyploidization in the generation of new
adaptations (Ohno 1970), it is clear that WGD events have had a massive influence
on the content and structure of the genomes of their possessors. The next step in
exploring Ohno’s hypothesis is to link genome evolution to known changes in
function. This goal, however, remains challenging, primarily because our
knowledge of how genotype links to phenotype remains woefully incomplete
(Pigliucci 2010). However, one group of organisms in which we can at least begin
to make such associations is in the polyploid yeasts. Our knowledge of the
functional genomics of yeast is drawn primarily from Saccharomyces cerevisiae,
which has a well-annotated genome, decades of biochemical, genetic, and cell
biology research, a relatively small genome, and a life cycle that lends itself to
scalable laboratory analyses. Taken together, these facts have allowed yeast
researchers not only to understand the structure of the genome following WGD but
also to experimentally evaluate hypotheses regarding the evolution of particular
complex phenotypes. As we will stress throughout this chapter, one of the themes
that emerges from all of these analyses is the degree to which the outcome of a
WGD depends as much on the interactions between genes as on the role of any
particular locus. Of equal importance evolutionarily, we now also have data from
other polyploid yeast species, which are valuable both as a point of comparison to
S. cerevisiae and for their own sakes. This wealth of data affords us insight into the
mechanisms that drive the preferential loss and preservation of gene duplicates
after polyploidy, lead to the functional divergence of genes, and are behind the
evolutionary origins of complex phenotypes.
25
UN
Editor Proof
294
Book ISBN: 978-3-642-31441-4
Page: 294/307
Layout: T1 Standard SC
Chapter No.: 15
73
74
75
76
77
78
79
80
81
82
83
84
85
86
87
88
89
90
91
92
93
94
95
96
97
98
99
100
101
102
103
104
105
F
71
72
PR
OO
70
15.2.1 Synteny-Based Evidence for WGD in Saccharomyces
cerevisiae
D
69
TE
68
295
considerable variation in the numbers of synonymous substitutions separating
them. This observation led Smith (1987) to hypothesize that S. cerevisiae underwent a WGD ancient enough that the duplicates surviving from it had saturated.
However, it was not until the genome sequence of S. cerevisiae became available
that this speculation could be confirmed (see below). Another similar but subtler
problem in using Ks as a means of dating duplicate genes is the issue of gene
conversion. Gene conversion was presumed to be quite common in yeast (Petes
and Hill 1988), even prompting some authors (Gao and Innan 2004) to suggest that
estimates of duplication rates based on duplicate divergences were inapplicable
due to the homogenization of duplicate loci by conversion. Fortunately, although
gene conversion is very common among yeast ribosomal proteins, it does not
appear to be a general characteristic of the genome (Evangelisti and Conant 2010).
Nonetheless, these various issues collectively meant that comparisons of paralogous sequence divergence were deemed unhelpful as a means to detect WGD in
yeast.
Given that paralogous sequence comparisons were generally unhelpful in finding
WGD relics, another tactic was to consider gene order. In fact, even before the S.
cerevisiae genome sequence was completed in 1996, it was clear to many
researchers that it contained numerous, long, homologous clusters of ordered
genes (Goffeau et al. 1996). Melnick and Sherman (1993) found ordered homologous gene clusters in chromosomes V and X covering 7.5 kb. Lalo et al. (1993)
similarly found ordered homologous gene clusters in chromosomes XIV and III
covering 15 kb. When the genome was sequenced, researchers found 18 ordered
homologous genes in chromosomes IV and II that covered 120 and 170 kb,
respectively (Goffeau et al. 1996). Just how to interpret these redundant regions
remained a challenge at that time (Goffeau et al. 1996; Oliver 1996), and, in spite
of Smith’s prior hypothesis (Smith 1987), few, if any, of the contemporaneous
explanations included an ancient WGD.
However, opinions changed the next year when Wolfe and Shields (1997)
presented a thorough, genome sequence-based, analysis that gave strong evidence
for WGD in S. cerevisiae. To find syntenic regions, they conducted a BLASTP
search of amino acid sequences throughout the yeast genome and made a dot plot
of the results. They then created gene blocks from these data, where each block
was required to have at least three homologous pairs with intergenic distances B
50 kb and conservation of gene order and orientation. This analysis yielded 55
duplicated regions containing a total of 376 pairs of ohnologs. The large number of
duplicated regions led Wolfe and Shields to posit two explanations: (1) successive
independent gene duplications, and (2) a single duplication of the entire genome,
EC
67
CO
RR
66
Book ISBN: 978-3-642-31441-4
Page: 295/307
Yeast as a Window into Changes in Genome Complexity
UN
Editor Proof
15
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 15
Book ID: 272454_1_En
Date: 16-8-2012
113
114
115
116
117
118
119
120
121
122
123
124
125
126
127
128
129
130
131
132
133
134
135
136
137
138
139
140
141
142
143
144
145
F
112
PR
OO
110
111
15.2.2 Comparative Genomics and Proof of WGD
in S. cerevisiae
D
109
A number of researchers disputed the claims of Wolfe and Shields (1997), arguing
that, because the syntenic regions identified made up only a small part of the
genome, independent duplications better explained the genomic structure of S.
cerevisiae (Coissac et al. 1997; Mewes et al. 1997; Hughes et al. 2000; Llorente
et al. 2000a; Llorente et al. 2000b; Friedman and Hughes 2001; Piskur 2001;
Koszul et al. 2004). However, this independent duplication hypothesis became
untenable following the genome sequencing of other yeasts that proved to lack
these syntenic paralog blocks. These sequences were described by three independent groups. The comparison of S. cerevisiae with Kluyveromyces waltii
(Kellis et al. 2004) and the comparison of S. cerevisiae with Ashbya gossypii
(Dietrich et al. 2004) involved different genomes, but effectively made the same
argument: that the 2:1 mapping of blocks of paralogs from S. cerevisiae to
homologous single-copy genes in K. waltii/A.gossypii could best be explained by
WGD. This explanation was particularly striking because the doubly conserved
synteny blocks cover 90 % of the genome in K. waltii (Kellis et al. 2004) and 96 %
of that in A. gossypii (Dietrich et al. 2004). Furthermore, both studies found a large
number of 2:1 pairing of centromeres in the species-respective chromosomes.
There were 16:8 such pairings between S. cerevisiae and K. waltii and 14:7
between S. cerevisiae and A. gossypii with a subsequent break at the expected
centromere position in S. cerevisiae chromosomes X and XII that are syntenic with
regions in A. gossypii chromosomes I and III. Finally, and perhaps most strikingly,
both groups also showed that the single-copy orthologs of genes from A. gossypii
or K. waltii in the genome of S. cerevisiae are interleaved between two paralogous
chromosomes in S. cerevisiae that nonetheless retain the relative gene order of the
single chromosome in the non-WGD yeast (see Fig. 15.1). Such a pattern is only
explicable under the hypothesis of a WGD event followed by massive gene losses.
TE
108
followed by massive gene loss. There were two lines of evidence discounting the
first possibility. First, 90 % (50/55) of the gene regions shared the same orientation
with respect to the centromeres of the duplicated regions when we would expect
independent duplications to be instead randomly distributed about the centromeres. Second, there were no examples of triplicated regions in the S. cerevisiae
genome. If the duplications involved several distinct events separated in time, such
a pattern would be highly unlikely, because it would require that later duplication
events never overlapped with prior ones. Given these arguments, Wolfe and
Shields (1997) argued for a single ancient WGD, which they dated to be hundreds
of millions of years old (note that attempts to conclusively date this event have
been difficult, due to a lack of fossils and the previously mentioned saturation of
substitutions; see Taylor and Berbee 2006; Rolland and Dujon 2011).
EC
107
CO
RR
106
C. M. Hudson and G. C. Conant
UN
Editor Proof
296
Book ISBN: 978-3-642-31441-4
Page: 296/307
Layout: T1 Standard SC
Chapter No.: 15
Book ISBN: 978-3-642-31441-4
Page: 297/307
Yeast as a Window into Changes in Genome Complexity
297
CO
RR
EC
TE
D
PR
OO
F
Editor Proof
15
Book ID: 272454_1_En
Date: 16-8-2012
UN
Fig. 15.1 Yeast gene order browser (YGOB) screenshots with a window size of six. Each box
represents a gene; each color, a chromosome. The gene in focus, the A. gossypii gene ABR086W,
is highlighted by an orange border. Each vertical column (‘‘pillar’’) represents a single gene prior
to the WGD (hence, all genes in a column are homologs, and the paired upper and lower genes,
when present, are paralogs). The ancestral order of these genes (pink boxes) just prior to the
WGD has also been exhaustively inferred (Gordon et al. 2011). Connectors join nearby genes: a
solid bar for adjacent genes, two bars for loci less than five genes apart, and one bar for loci \20
genes apart. The connectors are extended in gray over intervening space. The end of a
chromosome or contig is denoted by a brace. Arrows denote transcriptional orientation. The
browser also includes a control panel that allows users to select the window size and the gene to
focus on. This panel also has buttons for running BLAST searches against YGOB’s database,
outputting YGOB data in tabular format, obtaining pairwise Ka and Ks values among genes, and
computing multiple sequence alignments and phylogenetic trees of individual pillars. Species
names for each track are labeled at right (Byrne and Wolfe 2005)
Layout: T1 Standard SC
Chapter No.: 15
Book ID: 272454_1_En
Date: 16-8-2012
C. M. Hudson and G. C. Conant
157
15.2.3 Yeast Gene Order Browser
148
149
150
151
152
153
154
155
PR
OO
147
F
156
The argument of Dujon et al. (2004) is subtly different. They sequenced and analyzed four other genomes. One genome that of Candida glabrata, shares the genome
duplication with S. cerevisiae. This was determined by comparing syntenic blocks in S.
cerevisiae and C. glabrata with the other three sequenced genomes, Kluveromyces
lactis, Debaryomyces hansenii, and Yarrowia lipolytica. Dujon et al. (2004) found 20
distinct blocks of paralogs shared by both S. cerevisiae and C. glabrata. These blocks
allowed them to map the WGD onto a phylogeny, rather than do a simple pairwise
comparison. Mapping this WGD phylogenetically creates distinct hypotheses as to
where in the tree we expect to find polyploid yeasts (c.f., Fig. 15.1), predictions that
have been confirmed with each of the subsequently sequenced genomes of known
phylogenetic position (Wapinski et al. 2007; Scannell et al. 2011).
146
170
15.2.4 Additional Non-Saccharomyces-Specific WGDs
162
163
164
165
166
167
168
TE
161
EC
160
CO
RR
159
D
169
One of the major benefits of studying the yeast WGD is that the relatively slow rates of
gene order change in yeast genomes and the compactness of their genomes means that
an exhaustive enumeration of all WGD-produced ohnologs is possible. Just such a
project was carried out, with the results presented as the web-based Yeast Gene Order
Browser (YGOB) (Byrne and Wolfe 2005), which illustrates a number of non- and
post-WGD yeasts in a graphical framework (Fig. 15.2). This work has been followed
by a reconstruction of the set of genes and their relative orders that existed just prior to
the WGD (Gordon et al. 2009) and by a likelihood-based model of post-WGD
duplicate loss that attempts to quantify the orthology inferences made by YGOB
(Conant and Wolfe 2008a). On the basis of these three projects, the post-WGD evolutionary history of virtually every locus in the S. cerevisiae genome can be traced
(Fig. 15.1 is thus illustrative of the predominant pattern seen across the genome).
158
174
In addition to the ancient WGD that characterizes the Saccharomyces clade
(Fig. 15.2), several cases of allopolyploidy have been discovered in yeasts. Some
of these occur in species within the Saccharomyces sensu stricto clade (Scannell
et al. 2011), while others are independent.
175
15.2.4.1 Secondary Allopolyploidy in Saccharomyces pastorianus
171
172
173
176
177
178
179
UN
Editor Proof
298
Book ISBN: 978-3-642-31441-4
Page: 298/307
Several cases of allopolyploidy are known from within S. sensu stricto (Dequin
and Casaregola 2011). One of the most well studied is that of the lager yeast,
S. pastorianus (syn. S. carlsbergensis). It has long been known that the polyploid
S. pastorianus and other members of the complex of related lager yeasts are
Layout: T1 Standard SC
Chapter No.: 15
Book ISBN: 978-3-642-31441-4
Page: 299/307
Yeast as a Window into Changes in Genome Complexity
299
PR
OO
F
Editor Proof
15
Book ID: 272454_1_En
Date: 16-8-2012
184
185
186
187
188
189
190
191
192
193
194
195
196
197
198
199
200
201
EC
183
CO
RR
182
allotetraploids of diploid S. cerevisiae and some other unknown diploid species
(Martini and Kurtzman 1985; Kielland-Brandt et al. 1995). However, aside from
the general difficulties facing anyone interested in identifying the origins of hybrid
genomes, the debate surrounding the origin of the second parental diploid species
was further complicated by a difficulty in delimiting species within these groups
(Rainieri et al. 2006). The tetraploid S. pastorianus belongs to a group of yeast
species which, until recently, was represented as a phylogenetically unresolved
species complex including S. pastorianus, S. monacensis (S. pastorianus strain
CBS 1503), S. bayanus, and S. bayanus var. uvarum (Casaregola et al. 2001;
Rainieri et al. 2006). This taxonomic confusion has recently been partially
resolved through the sequencing of the genomes of both S. pastorianus and one of
its presumed parental diploid species, S. eubayanus (Nakao et al. 2009). The
genome history that has emerged is a complicated story of allopolyploidy followed
by genomic transformation forming the related species S. bayanus (Libkind et al.
2011). As summarized by Libkind et al. (2011), S. cerevisiae hybridized
with S. eubayanus (a species recently recovered in Patagonia) with subsequent
genome doubling producing the allotetraploid progenitor of modern S. pastorianus. Following domestication, smaller regions of the S. pastorianus genome were
then apparently transferred into the genome of the diploid parent S. eubayanus
(which is nonetheless a descendant of the ancient polyploidy). This hybrid form of
S. eubayanus, with contributions from S. pastorianus, then proceeded to interbreed
with diploid S. uvarum to produce the modern, diploid, S. bayanus (Fig. 15.3).
UN
180
181
TE
D
Fig. 15.2 Consensus view of the evolutionary relationships between the yeast taxa discussed.
Black branches indicate relationships described by both Kurtzman and Robnett (2003) and
Fitzpatrick et al. (2006). Red branches indicate conflicts between the two phylogenies, in which
case Fitzpatrick et al. (2006) is presented. Curved grey branches illustrate the allopolyploidy
events between two species (S. cerevisiae and S. bayanus, Z. rouxii, and Z. pseudorouxii). Taxa in
blue are reported in text (e.g., S. pastorianus and Z. rouxii ATCC 42981). Stars mark wholegenome duplications. Note that genus names are an imperfect guide to the relationships
Layout: T1 Standard SC
Chapter No.: 15
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 300/307
Editor Proof
300
C. M. Hudson and G. C. Conant
natural environments
brewing environment
early
S. pastorianus
1 50:50 hybrid of
S. eubayanus and
S.cerevisiae
modern
S. pastorianus
3 S. pastorianus
cells lyse and
release large
DNA fragments
4 transformation
of S. eubayanus
by S. pastorianus
2 SUL1 inactivation,
loss-of-heterozygosity
on the right arm of chromosome VII adds an extra copy
of ScerIMA1, aneuploidy and
chromosome rearrangements
S. bayanus
F
S. eubayanus
brewing contaminants
domestication
PR
OO
ale-type
S. cerevisiae
NBRC 1948
CBS 380
S. uvarum
5 independent
hybridizations
with S. uvarum
functional SUL1
inactive SUL1
202
TE
D
Fig. 15.3 Genome evolution in S. pastorianus. A model of the formation of S. pastorianus and the
hybrid strains of S. bayanus. First, wild S. eubayanus and ale-type S. cerevisiae hybridized to form
an allotetraploid that became the ancestor of the modern (doubly paleopolyploid) S. pastorianus.
Second, domestication imposed strong selective pressure for strains with the most desirable brewing
properties. Third, in the brewing vats with high densities of S. pastorianus, cell lysis releases large
DNA fragments that occasionally transform, fourth, contaminating wild strains of S. eubayanus
(which possesses only the ancient WGD shared with S. cerevisiae) because of the lack of pure
culture techniques. Fifth, multiple hybridization events between S. eubayanus and wild strains of
S. uvarum gave rise to CBS 380T and NBRC 1948. This model does not exclude prior or parallel
involvement of S. uvarum in brewing or contamination. Reprinted from Libkind et al. (2011)
15.2.4.2 Allopolyploidy in Zygosaccharomyces rouxii
215
15.3 WGD and Speciation
206
207
208
209
210
211
212
213
216
217
218
219
220
CO
RR
205
UN
204
EC
214
The spoilage agent and industrial yeast Zygosaccharomyces rouxii strain ATCC
42981 was identified as another allopolyploid by James et al. (2005) and Gordon
and Wolfe (2008). This hybridization/polyploidy event is significant for two reasons. First, unlike all of the previous examples, it occurs outside of S. sensu stricto.
Second, Gordon and Wolfe (2008) determined that most of the paralogs produced
by WGD are still present, presumably due to the recentness of the event. Thus,
while other yeast genome duplications are ancient and show considerable gene loss
and rearrangement (Wolfe and Shields 1997), the Z. rouxii genome retains most of
the ‘‘new’’ genes produced by its WGD. Since the survival time of ohnologs has
been modeled to follow a power law, most of the duplicates are expected to be lost
very rapidly (Maere et al. 2005), suggesting that Z. rouxii represents an example of
the early features of genome evolution following WGD.
203
An important potential outcome of polyploidy is in altering patterns of speciation.
This change can happen in at least two ways. First, the WGD can relax selective
constraints resulting in an adaptive radiation by means of ecological speciation.
Another, more neutral mechanism, is a special case of the Dobzhansky-Muller
(DM) process of speciation, in which species lose reciprocal paralogs following
Layout: T1 Standard SC
Chapter No.: 15
227
228
229
230
231
232
233
234
235
236
237
238
239
240
241
242
243
244
245
246
247
248
249
250
251
252
253
254
255
256
257
258
259
260
261
262
263
264
F
226
PR
OO
225
D
224
TE
223
301
some period of isolation (Lynch and Force 2000; Werth and Windham 1991; see
also Chap. 1, this volume). WGD potentially increases the probability of this
simply by increasing the number of paired genes in a genome. The fertility of
hybrids is 0.75n, where n is the number of reciprocal losses of essential genes
among populations (Werth and Windham 1991). Clearly, for any significant
number of reciprocal losses (such as occur after WGD), the number of viable,
fertile offspring of a crossing of two such populations is negligible. Both phylogenetic and experimental studies of the DM process after WGD have been carried
out in yeast. Scannell et al. (2006) showed that the number of reciprocal gene
losses in several species of yeast sharing the S. cerevisiae WGD was sufficient to
induce such inviability. This observation suggests that a DM mechanism was
partly responsible for the multiple speciation events among the Saccharomyces
species (e.g., S. cerevisiae, S. bayanus, and C. glabrata) following WGD.
An advantage of studying the DM process in yeast is the ability to experimentally create and cross artificial polyploids. This possibility has been highlighted in experimental studies of reproductive isolation. Polyploid yeasts have
been allowed to evolve in different selective environments (Dettman et al. 2007)
and in neutral environments subject to random mutagenesis (Maclean and Greig
2011). These two experiments have shown that moderate reproductive isolation,
coupled with reciprocal gene loss, results in a clear loss of fitness when independently derived polyploids are crossed. Similarly, Lee et al. (2008) showed that
hybrids of S. cerevisiae and S. bayanus were less fit than their parental phenotypes,
due primarily to incompatibility between their nuclear and mitochondrial genomes. Chou et al. (2010) extended this analysis, providing another pair of mitochondrial and nuclear genes and posited nuclear-mitochondrial incompatibility as
a common mechanism in species formation. In another twist, Anderson et al.
(2010) demonstrated the existence of alleles with depressed hybrid fitness in lowglucose environments, which argues for a model in which neutral changes in
paired genes are followed by strong selection, a sequence of events that promotes
rapid reproductive isolation. Kao et al. (2010), however, argue against the existence of a small number of so-called speciation genes, instead claiming that
genome scans provide no evidence of any single paired dominant or recessive
genic incompatibilities. They instead argue that following WGD, many changes in
loci of little effect resulted in lowered fitness due, in part, to the rewiring of
transcriptional and metabolic networks.
Another debate that has emerged in this field is whether these changes are due
primarily to the decrease in the fertility of hybrids (Xu and He 2011) or a decrease
in their viability (Greig 2008). This question ultimately amounts to a debate about
what stage in the yeast life cycle the genetic incompatibilities occur—sporulation
or clonal growth, and whether the decrease in fitness is the result of competition
for resources or offspring. The discontinuity between these ideas likely represents
an opportunity to explain speciation as a process across different genomic and
temporal scales, and we would speculate that the process of DM incompatibility
induces selection for the evolution of some form of prezygotic barrier.
EC
222
CO
RR
221
Book ISBN: 978-3-642-31441-4
Page: 301/307
Yeast as a Window into Changes in Genome Complexity
UN
Editor Proof
15
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 15
Book ID: 272454_1_En
Date: 16-8-2012
271
272
273
274
275
276
277
278
279
280
281
282
283
284
285
286
287
288
289
290
291
292
293
294
295
296
297
298
299
300
301
302
303
304
305
306
307
F
270
PR
OO
269
D
268
Duplicate retention and evolutionary models. In addition to such population
processes as speciation, WGD also altered many other aspects of the S. cerevisiae
lifestyle. For instance, several pairs of ohnologs have been shown to have
undergone various types of functional divergence, allowing the study of some of
the proposed mechanisms of duplicate divergence after duplication (Conant and
Wolfe 2008b). In an elegant series of experiments, van Hoof (2005) showed that
two ohnologs, ORC1 and SIR3, have distinct and non-overlapping functions (in
DNA replication and gene silencing, respectively). Strikingly, however, the mutual
ortholog of these genes from the non-WGD yeast S. kluyveri is able to complement
both functions, constituting a clear example of subfunctionalization. An apparently
similar case, involving the S. cerevisiae ohnolog pair GAL1 and GAL3, which
presently functions, respectively, as an enzyme and as a transcriptional regulator,
was complicated by the discovery of an adaptive conflict between the shared
regulator and enzymatic function of their ortholog in the non-WGD K. lactis.
Thus, although the K. lactis GAL1 gene does indeed serve the functions of both
GAL1 and GAL3 in S. cerevisiae, it does so in a suboptimal way, being unable to
tune its expression to both roles simultaneously (Hittinger and Carroll 2007). This
conflict illustrates an important point about subfunctionalization, namely that the
original neutral model of subfunctionalization proposed by Force and coauthors
(Force et al. 1999) is not the only possible mechanism for such functional partitioning (Des Marais and Rausher 2008). Other examples of divergence among
ohnologs where the mechanism of that divergence is less clear include ribosomal
proteins (Ni and Snyder 2001; Komili et al. 2007; Kim et al. 2009), glucose
sensors (Özcan et al. 1998), and glycolysis enzymes (Boles et al. 1997).
The dosage balance hypothesis (DBH). In addition to facilitating the above
work, the wealth of functional data from S. cerevisiae also provides an excellent
opportunity to test hypotheses explaining the differences in gene retention patterns
after WGD and small-scale duplications (hereafter SSD). Chief among these is
probably the DBH (Papp et al. 2003; Freeling and Thomas 2006; Birchler and
Veitia 2007; Freeling 2009), which states that, in eukaryotes, there is selection
operating to disfavor duplications of central network genes due to the imbalance in
network stoichiometry that results. This situation is reversed for WGD because in
that case, the loss of a second copy of a gene introduces imbalances relative to the
remaining duplicated genes. In keeping with the DBH, several classes of genes are
over-retained after several evolutionarily ancient WGD events, including that in
yeast. They include ribosomal proteins, protein kinases and transcription factors
(Seoighe and Wolfe 1999; Blanc and Wolfe 2004; Maere et al. 2005; Aury et al.
2006; Conant and Wolfe 2008a). Similarly, genes that tend to have been fixed by
WGD are less likely to have undergone SSD in other yeast species (Wapinski et al.
2007). However, duplicates produced by WGD have more protein interactions
(Guan et al. 2007; Hakes et al. 2007), more phosphorylation sites (Amoutzias et al.
2010), and tend to be highly expressed (Seoighe and Wolfe 1999) than those from
TE
267
EC
266
15.4 Changes in Genome Content and Complexity Post-WGD
CO
RR
265
C. M. Hudson and G. C. Conant
UN
Editor Proof
302
Book ISBN: 978-3-642-31441-4
Page: 302/307
Layout: T1 Standard SC
Chapter No.: 15
314
315
316
317
318
319
320
321
322
323
324
325
326
327
328
329
330
331
332
333
334
335
336
337
338
339
340
341
342
343
344
345
346
347
348
349
350
351
352
F
313
PR
OO
312
D
311
TE
310
303
SSD. Although genes retained in duplicate after WGD are rarely essential on an
individual basis (Guan et al. 2007), this dispensability appears to be due to
functional compensation by the other ohnolog (DeLuna et al. 2008). Thus, it
appears that while ohnologs are less likely to be essential than their SSD counterparts today, their ancestral genes were actually at least as essential as current
single-copy genes (DeLuna et al. 2008).
System-level changes produced by WGD. Of course, one of the unique features of
polyploidy relative to SSD is the possibility of coordinated changes among multiple
sets of ohnologs. At the simplest level, we have previously illustrated examples of
what appears to be network subfunctionalization where a number of ohnologs collectively divided two expression domains among themselves (Conant and Wolfe
2006). A more complex and interesting example is the role of the WGD (Piškur et al.
2006) in shaping S. cerevisiae’s propensity for aerobic glucose fermentation
(the Crabtree effect; Geladé et al. 2003; Johnston and Kim 2005), a novel and
somewhat paradoxical phenotype. There is a general association between the presence of the WGD and the Crabtree effect across yeast species (Merico et al. 2007).
As a result, we and others have argued that dosage effects among the glycolysis
enzymes post-WGD helped to increase flux through glycolysis (Blank et al. 2005;
Kuepfer et al. 2005; Conant and Wolfe 2007; Merico et al. 2007; van Hoek and
Hogeweg 2009). Such increased flux likely could only be accommodated through
fermentation pathways, given the complex spatial organization of the competing
respiratory pathway (Conant and Wolfe 2007). Supporting this hypothesis is an
elegant computational analysis by van Hoek and Hogeweg (2009) showing that
future WGD events in modern S. cerevisiae could also be expected to provide a
selective advantage in glucose-rich environments through the preferential retention
of duplicated glycolysis enzymes. Note that the apparently ‘‘wasteful’’ fermentation
can actually be selectively advantageous in the context of rich but ephemeral
resource patches (Pfeiffer et al. 2001; Pfeiffer and Schuster 2005), a phenomenon that
has been experimentally confirmed in yeast (MacLean and Gudelj 2006). Such a
change in the yeast lifestyle likely led to other, later changes in the genome.
One suggestive example concerns the decoupling of cytosolic and mitochondrial
ribosomal protein expression post-WGD (Ihmels et al. 2005). Prior to WGD, bakers’
yeast was likely similar to other yeasts in having a strong association in the
expression of the two types of ribosomal proteins. After WGD, however, cisregulatory element evolution diverged in the two groups of genes (Ihmels et al.
2005), allowing S. cerevisiae to express only cytosolic proteins at high levels during
fermentation, an important refinement in a fermentative lifestyle.
Connecting the DBH to large-scale evolutionary changes following WGD,
Conant (2010) and Fusco et al. (2010) found transcriptional regulatory motifs to be
over-retained in ohnologs. Modeling network evolution after WGD, these authors
find the network enriched for transcription factors and particular network motifs.
Duplicated transcription factors still show some relics of the WGD, being more
likely to share targets than are random transcription factors, but on the whole show
considerable divergence post-WGD (Conant 2010). Given this rapid regulatory
evolution, it may not be easy to ascertain the role of WGD in the evolution of
EC
309
CO
RR
308
Book ISBN: 978-3-642-31441-4
Page: 303/307
Yeast as a Window into Changes in Genome Complexity
UN
Editor Proof
15
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 15
Book ID: 272454_1_En
Date: 16-8-2012
C. M. Hudson and G. C. Conant
357
the modern S. cerevisiae regulatory network. Nonetheless, the retention of many
transcription factors that have acquired distinct sets of target genes may imply that
the WGD served to ‘‘relax’’ the regulatory complexity of this organism, which
may have implications for its future ability to adapt (as seen for the GAL1/GAL3
example).
358
15.5 Conclusions
353
354
355
PR
OO
F
356
370
371
372
373
Acknowledgments We would like to thank Michaël Bekaert, Patrick Edger, and Chris Pires for
helpful discussions. This work was supported by the National Library of Medicine Biomedical and
Health Informatics Training Fellowship [LM007089-19] (CMH) and the Reproductive Biology
Group of the Food for the twenty-first century program at the University of Missouri (GCC).
374
References
375
376
377
378
379
380
381
382
383
384
385
386
387
388
389
390
391
Amoutzias GD, He Y, Gordon J, Mossialos D, Oliver SG, Van de Peer Y (2010) Posttranslational
regulation impacts the fate of duplicated genes. Proc Natl acad sci U S A 107:2967–2971
Anderson JB, Funt J, Thompson DA et al (2010) Determinants of divergent adaptation and
Dobzhansky-Muller interaction in experimental yeast populations. Curr Biol 20:1383–1388
Aury JM, Jaillon O, Duret L et al (2006) Global trends of whole-genome duplications revealed by
the ciliate Paramecium tetraurelia. Nature 444:171–178
Birchler JA, Veitia RA (2007) The gene balance hypothesis: from classical genetics to modern
genomics. Plant Cell 19:395–402
Blanc G, Wolfe KH (2004) Functional divergence of duplicated genes formed by polyploidy
during Arabidopsis evolution. Plant Cell 16:1679–1691
Blank LM, Lehmbeck F, Sauer U (2005) Metabolic-flux and network analysis of fourteen
hemiascomycetous yeasts. FEMS Yeast Res 5:545–558
Boles E, Schulte F, Miosga T, Freidel K, Schlüter E, Zimmermann FK, Hollenberg CP, Heinisch JJ
(1997) Characterization of a glucose-repressed pyruvate kinase (Pyk2p) in Saccharomyces
cerevisiae that is catalytically insensitive to fructose-1-6-biphosphate. J Bacteriol 179:2987–2993
Byrne KP, Wolfe KH (2005) The yeast gene order browser: combining curated homology and
syntenic context reveals gene fate in polyploid species. Genome Res 15:1456–1461
364
365
366
367
368
TE
362
363
EC
361
CO
RR
360
D
369
The S. cerevisiae WGD has been implicated in a number of evolutionarily complex
events. At a minimum, a set of duplicated genes of identical age is a powerful
system for exploring duplicate gene evolution (van Hoof 2005; Conant and Wolfe
2006; Fares et al. 2006; Kim and Yi 2006). However, we also suggest that, as with
the GAL1/GAL3 example, we will not fully understand the biology of S. cerevisiae
until we account for how the WGD has altered both the individual roles of particular
genes and their relationships to each other. We have outlined some of the areas of
yeast biology that we think were altered by this genome-doubling event: there
remain others yet to be discovered. Similarly, the presence of other WGD events, of
varying ages, allows us to study how these events unfold over various timescales,
including, potentially, on the timescale of laboratory experiments in evolution.
359
UN
Editor Proof
304
Book ISBN: 978-3-642-31441-4
Page: 304/307
Layout: T1 Standard SC
Chapter No.: 15
305
EC
TE
D
PR
OO
F
Byrne KP, Wolfe KH (2007) Consistent patterns of rate asymmetry and gene loss indicate
widespread neofunctionalization of yeast genes after whole-genome duplication. Genetics
175:1341–1350
Casaregola S, Nguyen HV, Lapathitis G, Kotyk A, Gaillardin C (2001) Analysis of the
constitution of the beer yeast genome by PCR, sequencing and subtelomeric sequence
hybridization. Int J Syst Evol Microbiol 51:1607–1618
Chou J-Y, Hung Y-S, Lin K-H, Lee H-Y, Leu J-Y (2010) Multiple molecular mechanisms cause
reproductive isolation between three yeast species. PLoS Biol 8:e1000432
Coissac E, Maillier E, Netter P (1997) A comparative study of duplications in bacteria and
eukaryotes: the importance of telomeres. Mol Biol Evol 14:1062–1074
Conant GC (2010) Rapid reorganization of the transcriptional regulatory network after genome
duplication in yeast. Proc R Soc B 277:869–876
Conant GC, Wolfe KH (2006) Functional partitioning of yeast co-expression networks after
genome duplication. PLoS Biol 4:e109
Conant GC, Wolfe KH (2007) Increased glycolytic flux as an outcome of whole-genome
duplication in yeast. Mol Syst Biol 3:129
Conant GC, Wolfe KH (2008a) Probabilistic cross-species inference of orthologous genomic
regions created by whole-genome duplication in yeast. Genetics 179:1681–1692
Conant GC, Wolfe KH (2008b) Turning a hobby into a job: how duplicated genes find new
functions. Nat Rev Genet 9:938–950
DeLuna A, Vetsigian K, Shoresh N, Hegreness M, Colón-González M, Chao S, Kishony R (2008)
Exposing the fitness contribution of duplicated genes. Nat Genet 40:676–681
Dequin S, Casaregola S (2011) The genomes offermentative Saccharomyces. CR Biol 334:687–693
Des Marais DL, Rausher MD (2008) Escape from adaptive conflict after duplication in an
anthocyanin pathway gene. Nature 454:762–765
Dettman JR, Sirjusingh C, Kohn LM, Anderson JB (2007) Incipient speciation by divergent
adaptation and antagonistic epistasis in yeast. Nature 447:585–588
Dietrich FS, Voegeli S, Brachat S et al (2004) The Ashbya gossypii genome as a tool for mapping
the ancient Saccharomyces cerevisiae genome. Science 304:304–307
Dujon B, Sherman D, Fischer G et al (2004) Genome evolution in yeasts. Nature 430:35–44
Evangelisti AM, Conant GC (2010) Nonrandom survival of gene conversions among yeast
ribosomal proteins duplicated through genome doubling. Genome Biol Evol 2:826–834
Fares MA, Byrne KP, Wolfe KH (2006) Rate asymmetry after genome duplication causes
substantial long-branch attraction artifacts in the phylogeny of Saccharomyces species. Mol
Biol Evol 23:245–253
Fitzpatrick D, Logue M, Stajich J, Butler G (2006) A fungal phylogeny based on 42 complete
genomes derived from supertree and combined gene analysis. BMC Evol Biol 6:99
Force A, Lynch M, Pickett FB, Amores A, Yan Y, Postlethwait J (1999) Preservation of duplicate
genes by complementary, degenerative mutations. Genetics 151:1531–1545
Freeling M (2009) Bias in plant gene content following different sorts of duplication: tandem,
whole-genome, segmental, or by transposition. Annu Rev Plant Biol 60:433–453
Freeling M, Thomas BC (2006) Gene-balanced duplications, like tetraploidy, provide predictable
drive to increase morphological complexity. Genome Res 16:805–814
Friedman R, Hughes AL (2001) Gene duplication and the structure of eukaryotic genomes.
Genome Res 11:373–381
Furlong RF, Holland PWH (2002) Were vertebrates octoploid? Philos Trans R Soc Lond B
357:531–544
Fusco D, Grassi L, Bassetti B, Caselle M, Lagomarsino MC (2010) Ordered structure of the
transcription network inherited from the yeast whole-genome duplication. BMC Syst Biol 4:77
Gao LZ, Innan H (2004) Very low gene duplication rate in the yeast genome. Science 306:
1367–1370
Geladé R, Van De Velde S, Van Dijck P, Thevelein JM (2003) Multi-level response of the yeast
genome to glucose. Genome Biol 4:233
Goffeau A, Barrell B, Russey H et al (1996) Life with 6000 genes. Science 274:562–567
CO
RR
392
393
394
395
396
397
398
399
400
401
402
403
404
405
406
407
408
409
410
411
412
413
414
415
416
417
418
419
420
421
422
423
424
425
426
427
428
429
430
431
432
433
434
435
436
437
438
439
440
441
442
443
444
445
Book ISBN: 978-3-642-31441-4
Page: 305/307
Yeast as a Window into Changes in Genome Complexity
UN
Editor Proof
15
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 15
Book ID: 272454_1_En
Date: 16-8-2012
EC
TE
D
PR
OO
F
Gordon JL, Byrne KP, Wolfe KH (2009) Additions, losses and rearrangements on the
evolutionary route from a reconstructed ancestor to the modern Saccharomyces cerevisiae
genome. PLoS Genet 5:e1000485
Gordon JL, Byrne KP, Wolfe KH (2011) Mechanisms of chromosome number evolution in yeast.
PLoS Genet 7:e1002190
Gordon JL, Wolfe KH (2008) Recent allopolyploid origin of Zygosaccharomyces rouxii strain
ATCC 42981. Yeast 25:449–456
Greig D (2008) Reproductive isolation in Saccharomyces. Heredity 102:39–44
Guan Y, Dunham MJ, Troyanskaya OG (2007) Functional analysis of gene duplications in
Saccharomyces cerevisiae. Genetics 175:933–943
Hakes L, Pinney JW, Lovell SC, Oliver SG, Robertson DL (2007) All duplicates are not equal:
the difference between small-scale and genome duplication. Genome Biol 8:R209
Hittinger CT, Carroll SB (2007) Gene duplication and the adaptive evolution of a classic genetic
switch. Nature 449:677–681
Hughes AL (1999) Phylogenies of developmentally important proteins do not support the
hypothesis of two rounds of genome duplication early in vertebrate history. J Mol Evol
48:565–576
Hughes TR, Roberts CJ, Dai H et al (2000) Widespread aneuploidy revealed by DNA microarray
expression profiling. Nat Genet 25:333–337
Ihmels J, Bergmann S, Gerami-Nejad M, Yanai I, McClellan M, Berman J, Barkai N (2005)
Rewiring of the yeast transcriptional network through the evolution of motif usage. Science
309:938–940
Jaillon O, Aury J-M, Brunet F et al (2004) Genome duplication in the teleost fish Tetraodon
nigroviridis reveals the early vertebrate proto-karyotype. Nature 431:946–957
James SA, Bond CJ, Stratford M, Roberts IN (2005) Molecular evidence for the existence of
natural hybrids in the genus Zygosaccharomyces. FEMS Yeast Res 5:747–755
Johnston M, Kim J-H (2005) Glucose as a hormone: Receptor-mediated glucose sensing in the
yeast Saccharomyces cerevisiae. Biochem Soc Trans 33:247–252
Kao KC, Schwartz K, Sherlock G (2010) A genome-wide analysis reveals no nuclear
Dobzhansky-Muller pairs of determinants of speciation between S. cerevisiae and S.
paradoxus, but suggests more complex incompatibilities. PLoS Genet 6:e1001038
Kellis M, Birren BW, Lander ES (2004) Proof and evolutionary analysis of ancient genome
duplication in the yeast Saccharomyces cerevisiae. Nature 428:617–624
Kielland-Brandt MC, Nilsson-Tillgren T, Gjermansen C, Holmberg S, Pedersen MB (1995)
Genetics of brewing yeasts. In: Rose AH, Wheals AE, Harrison JS (eds) The Yeasts, vol 6,
2nd edn. Academic, London, pp 223–254
Kim S-H, Yi SV (2006) Correlated asymmetry of sequence and functional divergence between
duplicate proteins of Saccharomyces cerevisiae. Mol Biol Evol 23:1068–1075
Kim T-Y, Ha CW, Huh W-K (2009) Differential subcellular localization of ribosomal protein L7
paralogs in Saccharomyces cerevisiae. Mol Cells 27:539–546
Komili S, Farny NG, Roth FP, Silver PA (2007) Functional specificity among ribosomal proteins
regulates gene expression. Cell 131:557–571
Koszul R, Caburet S, Dujon B, Fischer G (2004) Eucaryotic genome evolution through the
spontaneous duplication of large chromosomal segments. EMBO J 23:234–243
Kuepfer L, Sauer U, Blank LM (2005) Metabolic functions of duplicate genes in Saccharomyces
cerevisiae. Genome Res 15:1421–1430
Kurtzman C, Robnett C (2003) Phylogenetic relationships among yeasts of the ‘Saccharomyces
complex’ determined from multigene sequence analyses. FEMS Yeast Res 3:417–432
Lalo D, Stettler S, Mariotte S, Slominski PP, Thuriaux P (1993) Two yeast chromosomes are
related by a fossil duplication of their centromeric regions. C R Acad Sci 316:367–373
Lee H-Y, Chou J-Y, Cheong L, Chang N-H, Yang S-Y, Leu J-Y (2008) Incompatibility of
nuclear and mitochondrial genomes causes hybrid sterility between two yeast species. Cell
135:1065–1073
CO
RR
446
447
448
449
450
451
452
453
454
455
456
457
458
459
460
461
462
463
464
465
466
467
468
469
470
471
472
473
474
475
476
477
478
479
480
481
482
483
484
485
486
487
488
489
490
491
492
493
494
495
496
497
498
C. M. Hudson and G. C. Conant
UN
Editor Proof
306
Book ISBN: 978-3-642-31441-4
Page: 306/307
Layout: T1 Standard SC
Chapter No.: 15
307
EC
TE
D
PR
OO
F
Libkind D, Hittinger CT, Valério E, Gonçalves C, Dover J, Johnston M, Gonçalves P, Sampaio JP
(2011) Microbe domestication and the identification of the wild genetic stock of lagerbrewing yeast. Proc Nat Acad Sci 108:14539–14544
Llorente B, Durrens P, Malpertuy A et al (2000a) Genomic exploration of the hemiascomycetous
yeasts: 20. Evolution of gene redundancy compared to Saccharomyces cerevisiae. FEBS Lett
487:122–133
Llorente B, Malpertuy A, Neuvéglise C et al (2000b) Genomic exploration of the hemiascomycetous yeasts: 18. Comparative analysis of chromosome maps and synteny with
Saccharomyces cerevisiae. FEBS Lett 487:101–112
Lynch M, Conery JS (2000) The evolutionary fate and consequences of duplicate genes. Science
290:1151–1155
Lynch M, Force AG (2000) The origin of interspecific genomic incompatibility via gene
duplication. Am Nat 156:590–605
Maclean CJ, Greig D (2011) Reciprocal gene loss following experimental whole-genome
duplication causes reproductive isolation in yeast. Evolution 65:932–945
MacLean RC, Gudelj I (2006) Resource competition and social conflict in experimental
populations of yeast. Nature 441:498–501
Maere S, De Bodt S, Raes J, Casneuf T, Van Montagu M, Kuiper M, Van de Peer. Y (2005) Modeling
gene and genome duplications in eukaryotes. Proc Nat Acad Sci U S A 102:5454–5459
Martini AV, Kurtzman CP (1985) Deoxyribonucleic acid relatedness among species of the genus
Saccharomyces sensu stricto. Int J Syst Bacteriol 35:508–511
Melnick L, Sherman F (1993) The gene clusters ARC and COR on chromosomes 5 and 10,
respectively, of Saccharomyces cerevisiae share a common ancestry. J Mol Biol 233:372–388
Merico A, Sulo P, Piškur J, Compagno C (2007) Fermentative lifestyle in yeasts belonging to the
Saccharomyces complex. FEBS J 274:976–989
Mewes H, Albermann K, Bähr M et al (1997) Overview of the yeast genome. Nature 387:7–65
Nakao Y, Kanamori T, Itoh T, Kodama Y, Rainieri S, Nakamura N, Shimonaga T, Hattori M,
Ashikari T (2009) Genome sequence of the lager brewing yeast, an interspecies hybrid. DNA
Res 16:115–129
Ni L, Snyder M (2001) A genomic study of the bipolar bud site selection pattern in
Saccharomyces cerevisiae. Mol Biol Cell 12:2147–2170
Ohno S (1970) Evolution by gene duplication. Springer, New York
Oliver SG (1996) From DNA sequence to biological function. Nature 379:597–600
Özcan S, Dover J, Johnston M (1998) Glucose sensing and signaling by two glucose receptors in
the yeast Saccharomyces cerevisiae. EMBO J 17:2566–2573
Papp B, Pal C, Hurst LD (2003) Evolution of cis-regulatory elements in duplicated genes of
yeast. Trends Genet 19:417–422
Petes T, Hill C (1988) Recombination between repeated genes in microorganisms. Annu Rev
Genet 22:147–168
Pfeiffer T, Schuster S (2005) Game-theoretical approaches to studying the evolution of
biochemical systems. Trends Biochem Sci 30:20–25
Pfeiffer T, Schuster S, Bonhoeffer S (2001) Cooperation and competition in the evolution of
ATP-producing pathways. Science 292:504–507
Pigliucci M (2010) Genotype-phenotype mapping and the end of the ‘genes as blueprint’
metaphor. Philos Trans R Soc Lond B 365:557–566
Piskur J (2001) Origin of the duplicated regions in the yeast genomes. Trends Genet 17:302–303
Piškur J, Rozpedowska E, Polakova S, Merico A, Compagno C (2006) How did Saccharomyces
evolve to become a good brewer? Trends Genet 22:183–186
Rainieri S, Kodama Y, Kaneko Y, Mikata K, Nakao Y, Ashikari T (2006) Pure and mixed genetic
lines of Saccharomyces bayanus and Saccharomyces pastorianus and their contribution to the
lager brewing strain genome. Appl Environ Microbiol 72:3968–3974
Rolland T, Dujon B (2011) Yeasty clocks: dating genomic changes in yeasts. CR Biol 334:620–628
Scannell DR, Byrne KP, Gordon JL, Wong S, Wolfe KH (2006) Multiple rounds of speciation
associated with reciprocal gene loss in polyploid yeasts. Nature 440:341–345
CO
RR
499
500
501
502
503
504
505
506
507
508
509
510
511
512
513
514
515
516
517
518
519
520
521
522
523
524
525
526
527
528
529
530
531
532
533
534
535
536
537
538
539
540
541
542
543
544
545
546
547
548
549
550
551
552
Book ISBN: 978-3-642-31441-4
Page: 307/307
Yeast as a Window into Changes in Genome Complexity
UN
Editor Proof
15
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 15
Book ID: 272454_1_En
Date: 16-8-2012
EC
TE
D
PR
OO
F
Scannell DR, Zill OA, Rokas A, Payen C, Dunham MJ, Eisen MB, Rine J, Johnston M, Hittinger CT
(2011) The awesome power of yeast evolutionary genetics: new genome sequences and strain
resources for the Saccharomyces sensu stricto genus. G3: Genes, Genomes, Genetics 1:11–25
Seoighe C, Wolfe KH (1999) Yeast genome evolution in the post-genome era. Curr Opin
Microbiol 2:548–554
Smith M (1987) Molecular evolution of the Saccharomyces cerevisiae histone gene loci. J Mol
Evol 24:252–259
Taylor JW, Berbee ML (2006) Dating divergences in the fungal tree of life: review and new
analyses. Mycologia 98:838–849
Van de Peer Y, Fawcett J, Proost S, Sterk L, Vandepoele K (2009) The flowering world: a tale of
duplications. Trends Plant Sci 14:680–688
van Hoek MJ, Hogeweg P (2009) Metabolic adaptation after whole genome duplication. Mol Biol
Evol 26:2441–2453
van Hoof A (2005) Conserved functions of yeast genes support the duplication, degeneration and
complementation model for gene duplication. Genetics 171:1455–1461
Wapinski I, Pfeffer A, Friedman N, Regev A (2007) Natural history and evolutionary principles
of gene duplication in fungi. Nature 449:54–61
Werth CR, Windham MD (1991) A model for divergent, allopatric speciation of polyploid
pteridophytes resulting from silencing of duplicate-gene expression. Am Nat 137:515–526
Wolfe KH (2000) Robustness-it’s not where you think it is. Nat Genet 25:3–4
Wolfe KH, Shields D (1997) Molecular evidence for an ancient duplication of the entire yeast
genome. Nature 387:708–713
Xu M, He X (2011) Genetic incompatibility dampens hybrid fertility more than hybrid viability:
yeast as a case study. PLoS ONE 6:e18341
CO
RR
553
554
555
556
557
558
559
560
561
562
563
564
565
566
567
568
569
570
571
572
573
574
575
576
C. M. Hudson and G. C. Conant
UN
Editor Proof
308
Book ISBN: 978-3-642-31441-4
Page: 308/307
Metadata of the chapter that will be visualized in
SpringerLink
Book Title
Polyploidy and Genome Evolution
Series Title
Chapter Title
Two Rounds of Whole-Genome Duplication: Evidence and Impact on the Evolution of Vertebrate Innovations
Copyright Year
2012
Copyright HolderName
Springer-Verlag Berlin Heidelberg
Corresponding Author
Family Name
Cañestro
Particle
Given Name
Cristian
Suffix
Abstract
Division
Departament de Genètica, Facultat de Biologia
Organization
Universitat de Barcelona
Address
Av. Diagonal, 643, edifici Prevosti, 2a planta, 08028, Barcelona, Spain
Email
canestro@ub.edu
The origin and evolution of the vertebrates have been linked to the study of genome duplications since Susumo
Ohno ventured the 2R-hypothesis, suggesting that the successful diversification of complex vertebrates was
facilitated by polyploidization in the stem vertebrate ancestor due to two rounds of whole-genome duplication
(2R-WGD). This chapter first reviews evidence supporting Ohno’s 2R-hypothesis and gathers information
about the timing and mechanisms underlying the 2R-WGD. Second, this chapter describes the impact of the
2R-WGD on the evolution of the vertebrate genome structure, gene number, and the evolutionary dynamics
of the functional fate of vertebrate ohnologs (paralogous genes that originated by WGD) in comparison with
non-vertebrate chordate gene homologs. Finally, this review discusses the functional consequences of the
2R-WGD on the origin and evolution of vertebrate innovations compared with urochordates and
cephalochordates, paying special attention to the origin of neural crest cells, placodes, and the big complex
brain, key features that probably facilitated the transition from ancestral filter-feeding non-vertebrate
chordates to voracious vertebrate predators. Currently available data, however, seem to suggest that these
putative key features were present to at least some extent in stem Olfactores; hence, the impact of the 2RWGD may not be related to the immediate origin of vertebrate innovations, but to the subsequent
diversification of a wide variety of complex structures that facilitated the successful radiation of vertebrates.
Book ISBN: 978-3-642-31441-4
Page: 309/338
Chapter 16
6
Cristian Cañestro
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
D
9
10
Abstract The origin and evolution of the vertebrates have been linked to the study of
genome duplications since Susumo Ohno ventured the 2R-hypothesis, suggesting that
the successful diversification of complex vertebrates was facilitated by polyploidization in the stem vertebrate ancestor due to two rounds of whole-genome duplication
(2R-WGD). This chapter first reviews evidence supporting Ohno’s 2R-hypothesis and
gathers information about the timing and mechanisms underlying the 2R-WGD.
Second, this chapter describes the impact of the 2R-WGD on the evolution of the
vertebrate genome structure, gene number, and the evolutionary dynamics of the
functional fate of vertebrate ohnologs (paralogous genes that originated by WGD) in
comparison with non-vertebrate chordate gene homologs. Finally, this review
discusses the functional consequences of the 2R-WGD on the origin and evolution of
vertebrate innovations compared with urochordates and cephalochordates, paying
special attention to the origin of neural crest cells, placodes, and the big complex
brain, key features that probably facilitated the transition from ancestral filter-feeding
non-vertebrate chordates to voracious vertebrate predators. Currently available data,
however, seem to suggest that these putative key features were present to at least some
extent in stem Olfactores; hence, the impact of the 2R-WGD may not be related to the
immediate origin of vertebrate innovations, but to the subsequent diversification of a
wide variety of complex structures that facilitated the successful radiation of
vertebrates.
TE
7
8
EC
4
CO
RR
3
PR
OO
5
Two Rounds of Whole-Genome
Duplication: Evidence and Impact
on the Evolution of Vertebrate
Innovations
2
F
1
Book ID: 272454_1_En
Date: 16-8-2012
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: 16
C. Cañestro (&)
Departament de Genètica, Facultat de Biologia, Universitat de Barcelona,
Av. Diagonal, 643, edifici Prevosti, 2a planta, 08028 Barcelona, Spain
e-mail: canestro@ub.edu
P. S. Soltis and D. E. Soltis (eds.), Polyploidy and Genome Evolution,
DOI: 10.1007/978-3-642-31442-1_16, Springer-Verlag Berlin Heidelberg 2012
309
Layout: T1 Standard SC
Chapter No.: 16
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 310/338
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
61
62
63
64
65
66
67
68
69
70
F
33
PR
OO
32
D
31
The study of the origin and evolution of vertebrates, the subphylum to which we
belong, has stood at the crossroad between genome evolution and molecular
developmental biology since the late 1960s, when Susumo Ohno published his
famous work on Evolution by Gene Duplication and proposed his hypothesis about
the pivotal role of genome duplication in the origin of vertebrates and their
diversification (Ohno et al. 1968; Ohno 1970). Vertebrates comprise all animals
that have a backbone and include mammals, birds, reptiles, amphibians, fishes, and
agnathans—the jawless lampreys and hagfishes. Vertebrates together with
urochordates (tunicates) form the Olfactores, which together with cephalochordates (amphioxus or lancelets) constitute the chordates (Fig. 16.1). All chordates
share a common basic body plan at least during the larval stage of their life cycle,
consisting of a notochord running through a post-anal tail, with a dorsal hollow
nerve cord, longitudinal blocks of muscle along the notochord, and ciliated
pharyngeal gill slits (Brusca and Brusca 2002). Recent phylogenomic analyses
have dethroned cephalochordates from the long-assumed position as sister group
of the vertebrates; this position is now occupied by urochordates, which include
ascidians, larvaceans and thaliaceans (Wada et al. 2006; Oda et al. 2002; Bourlat
et al. 2006; Delsuc et al. 2006; Putnam et al. 2008) (Fig. 16.1).
Ohno’s 2R-hypothesis was based on comparative analyses of genome sizes and
isozyme complexity among chordate taxa. Ohno found that basally divergent
chordate subphyla had smaller genomes and less isozyme complexity than
vertebrate lineages. This observation led him to suggest that the combination of
tandem gene duplication and in particular an octoploidization event involving two
rounds of whole-genome duplication were key to the invertebrate–vertebrate
transition, and for the subsequent successful vertebrate diversification (Ohno et al.
1968).
Ohno was one of the pioneers in conceiving the evolutionary significance of
whole-genome duplication (see also Chap. 1, this volume). Ohno emphasized the
importance of gene duplication as probably the main source of raw genetic material
for the evolution of new gene functions (reviewed in Taylor and Raes 2004). In
Ohno’s classical model, one of the duplicated genes retains the original function
whereas its duplicate either disappears by accumulation of detrimental mutations
(called pseudogenization or nonfunctionalization) or it is preserved after gaining
advantageous mutations that confer positively selected novel functions (neofunctionalization) (Ohno 1970; Nowak et al. 1997; Force et al. 1999). The duplication,
degeneration, complementation hypothesis (or DDC model) suggests a third
possibility for duplicate gene preservation: subfunctionalization, the complementary partitioning of ancestral structural and/or regulatory subfunctions between two
duplicate genes, so that the sum of their functions provides at least that of the
original pre-duplication gene (Force et al. 1999). The DDC model predicts that
subfunctionalized genes will have lower pleiotropy than the original pre-duplicated
gene and lower evolutionary constraints, and thereby will be more permissive to the
TE
30
EC
29
16.1 Introduction
CO
RR
28
C. Cañestro
UN
Editor Proof
310
Layout: T1 Standard SC
Chapter No.: 16
Editor Proof
16
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 311/338
Two Rounds of Vertebrate Whole-Genome Duplication
311
CHORDATES
OLFACTORES
UROCHORDATES
teleost fish
sharks - rays
amphibians
birds
mammals
F
CEPHALOCHORDATES
lamprey
VERTEBRATES
PR
OO
amphioxus
ascidianslarvaceans
tetrapods
cartilaginous
vertebrates
agnathans
(jawless vert.)
R1
R2
R1+R2 gnathostomes
(jawed vert.)
R1+R2
3rd. Scenario:
R1+R2 after the agnathan-gnathostome split
?
2nd. Scenario: (Panvertebrate Quadruplication PV4)
R1+R2 prior to agnathan-gnathostome split
1st. Scenario:
R1 prior to agnathan-gnathostome split
R2 in stem jawed vert. prior to cartilagenous-bony vert. split
D
Vertebrate Innovations:
Big Brain
Neural Crest Cells
Placodes
bony vertebrates
73
74
75
76
77
78
79
80
81
82
83
84
85
86
87
accumulation of mutations that might confer novel functions. The acquisition of
new functions is favored if the duplication affects the entire genome at once, as
opposed to multiple individual gene duplications, because when entire gene
networks are duplicated, gene stoichiometry is maintained, and therefore deleterious gene dosage effects can be counteracted (Birchler and Veitia 2007, 2010; Van de
Peer et al. 2009; Makino and McLysaght 2010; see also Chap. 2, this volume).
Genes that originated by gene duplication are called paralogs. Genes that have been
duplicated via genome duplication, however, are a special type of paralogs referred
as ohnologs, a term suggested by Wolfe (2000) in honor of Ohno’s contribution.
This term is useful because of the special properties that ohnologs possess at their
birth compared to duplications that arise by other local mechanisms such as unequal
crossing-over, tandem gene duplication, or retrotransposition.
The sudden creation and evolution of ohnologs by the 2R-WGD that occurred
in the stem vertebrate lineage has been suggested as one of the potential key events
underlying the increase of morphological complexity, facilitating the acquisition
of genetic and developmental innovations of vertebrates (Shimeld and Holland
2000; Aburomia et al. 2003). Genome duplication doubles the number of genes,
CO
RR
72
UN
71
EC
TE
Fig. 16.1 Evolutionary tree of chordates representing the three possible scenarios for the timing
of the 2R-WGD and the origins for vertebrate innovations. Evidence suggests that the 2R-WGD
might have had a significant impact on the diversification (black star) of vertebrate innovations,
including structures derived from neural crest cells and placodes, as well as the development of a
big complex brain in the stem vertebrate. But whether the 2R-WGD was crucial for the
evolutionary origin of these structures remains unclear, and that the hypothesis that the origin of
these innovations dated back to at least stem olfactores (gray star) cannot be dismissed. (Lamprey
picture courtesy of Juan Pascual-Anaya)
Layout: T1 Standard SC
Chapter No.: 16
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 312/338
96
97
98
99
100
101
102
103
104
105
106
107
108
109
110
111
112
113
114
115
116
117
118
119
120
121
122
123
124
125
126
127
128
129
130
131
132
F
94
95
PR
OO
93
D
92
TE
90
91
many of which have the chance of evolving ‘novel’ functions that might provide
new selectable advantages promoting species diversification (Lynch et al. 2001;
Van de Peer et al. 2009). Some vertebrate species that have undergone recent
polyploidization, such as the frog Xenopus laevis that experienced tetraploidization *40 million years ago (Hellsten et al. 2007), show a higher adaptability to a
variety of different environments, such as drought, salt, cold, and disease resistance, than closely related diploid species, such as Silurana tropicalis (for further
details on frog polyploidization see also Chap. 18, this volume). Interestingly, only
a limited number of retained ohnologs present evidence of neofunctionalization or
subfunctionalization in X. laevis, suggesting that additional selective mechanisms
might act on preserving gene duplicates that could promote species diversification
(Chain and Evans 2006; Semon and Wolfe 2008).
In addition to the evolutionary significance of novel fates of duplicate genes on
species biology, another mechanism that might contribute to species diversification
after genome duplication is reciprocal ohnolog loss between different populations, a
concept known as ‘divergent resolution’ that can lead to reproductive isolation
(Lynch and Force 2000). Reciprocal ohnolog loss is likely to occur in the period of
relaxed selection that duplicate genes experience while they are functionally
redundant (Werth and Windham 1991; Lynch and Conery 2000; Lynch and Force
2000; Scannell et al. 2006; Taylor et al. 2001; Semon and Wolfe 2007). The
divergent resolution of gene redundancies, such that one population loses one
ohnolog copy while the second population loses the other ohnolog copy, leads to
chromosomal restructuring such that gametes produced by hybrid individuals can be
completely lacking in functional genes for a duplicate pair. In addition to the
isolation due to reciprocal gene losses, this model can be further expanded to
isolation due to independent processes of gene duplicate subfunctionalization
between two populations, in which hybrids will lack one or more subfunctions
(Force et al. 1999; Lynch and Force 2000). Hence, large-scale reciprocal ohnolog
loss and independent subfunctionalization of ohnologs can be the cause of reproductive isolation of two populations after polyploidization, favoring genetic
divergence of these newly incipient future species. This hypothesis is supported by
the analyses of both fish and angiosperm lineages that have undergone polyploidization and include more species diversity (e.g. salmonids, catostomids, eudicots,
grasses) than their sister groups that did not go through polyploidization and include
a lower number of species (Nelson 1994; Ferris et al. 1979; Soltis et al. 2009).
Recent integrated approaches of comparative genomics and gene expression analyses in teleosts, however, provide limited evidence supporting the significance of
differential ohnolog loss in reproductive isolation and diversification (Kassahn et al.
2009) (see also Chap. 17, this volume).
Many studies have tackled the central question of whether or not the 2R-WGD
had a significant impact on the origin of vertebrate innovations and their subsequent
diversification. This chapter first reviews evidence supporting the 2R-hypothesis
and information regarding the timing and potential mechanisms underlying the
2R-WGD in vertebrates. Second, this chapter examines the impact that the
2R-WGD may have had on the evolution of vertebrate genome structure, number of
EC
89
CO
RR
88
C. Cañestro
UN
Editor Proof
312
Layout: T1 Standard SC
Chapter No.: 16
Book ISBN: 978-3-642-31441-4
Page: 313/338
Two Rounds of Vertebrate Whole-Genome Duplication
313
142
16.2 Supporting Evidence for the 2R-Hypothesis
139
140
143
144
145
146
147
148
149
150
151
152
153
154
155
156
157
158
159
160
161
162
163
164
165
166
167
168
169
170
171
172
173
PR
OO
138
If two rounds of genome duplication (2R-WGD) have occurred, we would expect
the presence of many paralogs (ohnologs) in conserved, syntenic genomic regions,
which are known as paralogons (Coulier et al. 2000) (or ohnologons (Gout et al.
2009)). Paralogons, therefore, consist of series of linked (but frequently functionally and phylogenetically unrelated) genes on one chromosome region, many
of which have linked paralogs on at least one other chromosome region. The
discovery of paralogy groups made of four paralogons in the genome of human and
mouse was interpreted as remnants of the two events of tetraploidization that
occurred early during vertebrate evolution and therefore provided the earliest
strong evidence supporting Ohno’s 2R-hypothesis (Lundin 1979, 1993; Pebusque
et al. 1998). Possibly one of the best and first examples of a paralogy group
supporting the 2R-hypothesis is the case of the four Hox-bearing regions on human
chromosomes Hsa2, Hsa7, Hsa12, and Hsa17 (Fig. 16.2) (Kappen et al. 1989;
Bailey et al. 1997; Larhammar et al. 2002; Lundin et al. 2003). In contrast, only a
single Hox cluster is present in the cephalochordate amphioxus (Garcia-Fernàndez
and Holland 1994). This 4:1 ratio is consistent with Ohno’s hypothesis of two
tetraploidization events after the cephalochordate–vertebrate split (Holland et al.
1994; Sidow 1996; Garcia-Fernandez 2005). In addition to the Hox paralogy
group, several other similar examples have been identified (e.g. MHC (Katsanis
et al. 1996), Tbx (Ruvinsky and Silver 1997), G-protein-coupled receptors
(Fredriksson et al. 2003), ParaHox clusters (Ferrier et al. 2005), linked receptor
tyrosine kinases (Siegel et al. 2007), endothelin ligands and receptors (Braasch
et al. 2009), Fox cluster (Wotton and Shimeld 2006), and the EGF ligand paralogons (Laisney et al. 2010)).
Several databases of conserved syntenic chromosomal regions between different
species are available, and these provide additional evidence of two rounds of
genome duplication. Popovici et al. (2001), for instance, identified 14 paralogons
containing more than 1600 genes assembled in a human genome paralogy map
(http://u119.marseille.inserm.fr/Db/paralogy.html). The ParaDB (http://abi.mars
eille.inserm.fr/paradb) predicted that the human genome far exceeds 1000 paralogons that contain more than three pairs of duplicated genes (Leveugle et al. 2003).
D
137
TE
136
EC
135
CO
RR
134
F
141
genes, and the fate of retained ohnologs in comparison with non-vertebrate chordate
paralogs. Finally, this chapter discusses how the 2R-WGD might have affected the
origin and evolution of vertebrate innovations, with special emphasis on the vertebrate big complex brain, and structures derived from neural crest cells and placodes. Recent data, however, suggest that neural crest cells and placodes could
already have been present in stem chordates. Hence, the impact of the 2R-WGD
cannot be related to the origin of neural crest cells and placodes, but it could be
related to their subsequent diversification and development of a wide variety of
complex structures.
133
UN
Editor Proof
16
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 16
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 314/338
Editor Proof
314
C. Cañestro
(a)
Genomic Synteny Conservation
A
F
B
C
(b)
Hsa2
R2
HOX-D cluster
THSD7B
UPP2
ITGB6FIGN
COBLL1
GRB14
HOXD1
OSBPL6
HOXD9 PLEKHA3
HNRNPA3
CDCA7HOXD10 DFNB59
FKBP7
PDE1A
SP9SP3 CHN1
FRZB
NEUROD1
ATF2 EVX2
SCRN3
STK17B
IGFBP2 VIL1
ERBB4
ABCA12 IGFBP5 TNS1
BZW1
HECW2 FAM126BICA1L
HOXB1
HOXB3
HOXB5
HOX-B cluster HOXB6
ADAP2
CCT6B
MYO1D
ERBB2RAPGEFL1
CDK12GRB7 IGFBP4
NEUROD2
FKBP10
NT5C3L
HOXB7
ARL4D
NFE2L1 HOXB8
TMEM106A
CRHR1ITGB3SP2 HOXB9
FAM117A
OSBPL7
ETV4SOSTHDAC5
SNX11
SCRN2
MPP2
CALCOCO2
MMD
PRKAR1A
AC113554.1
AMZ2
TTYH2
NACA2
GNA13WIPI1 SDK2
FAM20A
ITGB4
DNAH17
USP36
CYTH1
FOXK2
FSCN2 RAC3
ACTG1 RFNG
MAFG
Hsa7
TE
D
Hsa17
D
PR
OO
2R-WGD: R1
TTYH3 FOXK1
SP4 DFNA5
AMZ1
CREB5PLEKHA8
FSCN1 TMEM106BTSPAN13 DNAH11 NFE2L3
CDK13 STK17APPIAIGFBP1ABCA13
EGFR
ARL4ABZW2
LFNG
TAX1BP1
RAC1GLCCI1
FAM20C MAFK SDK1 WIPI2USP42
SCIN
ITGB8 MPP6SNX10
EVX1 CHN2 AQP1
POU6F2
HECW1
ADCY1 UPP1
CCT6A
PRKAR1B
GNA12 ACTB
ETV1
SCRN1
CRHR2
PDE1C
DPY19L1
INHBA
FAM126A
HOXA1
NACADTNS3FIGNL1
DGKB
NT5C3
SFRP4
SOSTDC1 RAPGEF5
HNRNPA2B1 HOXA3 FKBP14NEUROD6
COBL
AMPH GLI3 MYO1G
MMD2 CYTH3ICA1
ADAP1
IGFBP3 GRB10
HOXA4
SP8 OSBPL3
THSD7A HDAC9 CDCA7L
HOXA5
HOXA6
HOXA7
HOXA9
HOXA10
HOXA11
HOXC11
HOXC9
HOXC8
GLI1
HOXC6
AQP5
ERBB3 INHBE TSPAN31
AQP2
SP1 HOXC5 PDE1B DGKA
INHBC
NFE2
MIP
AVIL
HDAC7ADCY6
BIN2
CALCOCO1
NACA
DPY19L2
NEUROD4
TENC1
POU6F1
ITGB7 HNRNPA1
RAPGEF3
SP7 ATF7
IGFBP6
Hsa12
EC
HOX-A cluster
CO
RR
HOX-C cluster
Fig. 16.2 Conserved synteny in the vertebrate genome generated by the 2R-WGD. a Simplified
representation of a genomic region that has been amplified by the 2R-WGD (R1 and R2) showing
conserved genes (colored boxes) in four syntenic regions (a–d), which have suffered genomic
rearrangements and gene loss (white boxes) and different degrees of conservation (green and red
lines label ohnologs preserved in two or more than two regions, respectively). b Representation of
the four human paralogons containing Hox A-D clusters in chromosomes Hsa2, Hsa7, Hsa12, and
Hsa17, displaying high amounts of conserved synteny between ohnologs in two (green lines) or
at least three (red lines) different paralogons. This representation has been generated using a 100gene sliding window in the Synteny Database (Catchen et al. 2009)
175
176
177
178
The Genomicus database v60.01 (http://www.dyogen.ens.fr/genomicus) predicted
that 18,228 ancestral vertebrate genes were grouped in 2,642 conserved ancestral
synteny blocks with a median N50 size of 5 genes (Muffato et al. 2010). The Synteny
Database (http://teleost.cs.uoregon.edu/synteny_db) predicted 231 paralogy clusters with more than 5 genes, and 102 paralogy clusters with more than 10 genes, in a
UN
174
Layout: T1 Standard SC
Chapter No.: 16
185
186
187
188
189
190
191
192
193
194
195
196
197
198
199
200
201
202
203
204
205
206
207
208
209
210
211
212
213
214
215
216
217
218
219
220
221
222
223
F
184
PR
OO
183
D
182
TE
181
315
more rigorous count using a 100-gene sliding window and taking amphioxus genes
as the outgroup for paralogy assignment in human (Catchen et al. 2009, 2011).
Until recently, however, available evidence did not permit us to discard the
possibility that these groups of paralogous genes originated by multiple independent block duplications, rather than two duplications of the entire genome
(Skrabanek and Wolfe 1998; Wolfe 2001; Larhammar et al. 2002). An initial
hypothesis, based on extensive phylogenetic analysis and dating of the duplications that produced hundreds of vertebrate gene families, proposed a ‘big-bang
mode’ of sudden large-scale gene origin resulting from two waves of gene
duplications, rather than the alternative hypothesis of a constant generation by
small-scale duplications (Gu et al. 2002). Wave-I was suggested to consist of
tandem or segmental duplications that occurred after the mammalian radiation, and
wave-II was interpreted as a rapid increase of paralogs in the early stage of
vertebrate evolution after their split from non-vertebrate chordates, consistent with
one round of whole-genome duplication (Gu et al. 2002). The first analyses of the
human genome draft led to the conclusion that the most parsimonious explanation
of the current structure of the human paralogons was a ‘big-bang’ expansion event
by a paleopolyploidy that included the whole genome or substantial sections of it.
However, no specific evidence was found for two rounds of polyploidy as opposed
to one (Venter et al. 2001; McLysaght et al. 2002; Panopoulou et al. 2003).
Recently, however, several analyses have provided definitive support for the
2R-hypothesis (reviewed in Kasahara 2007). Dehal and Boore (2005) developed
an elegant, compelling approach to test the 2R-hypothesis by plotting the genomic
map position of only those genes that were duplicated prior to the fish–tetrapod
split, which rendered a clear global physical pattern of four-way paralogon organization covering most of the human genome. Dehal and Boore’s work therefore
provided unmistakable evidence of two distinct rounds of genome duplication
during early vertebrate evolution.
Furthermore, the recent sequencing of the whole genome of the chordate
amphioxus Branchiostoma floridae (sister to all other chordates) provided even
more indisputable evidence supporting the 2R-hypothesis (Putnam et al. 2008).
Despite the fact that small-region comparison between human, chicken, teleost
fish, and amphioxus genomes revealed low gene-order conservation at the local
level (microsynteny), striking extensive gene linkage conservation was observed
when entire chromosomes were considered (macrosynteny). Syntenic analysis
reconstructed 17 chordate linkage groups (CLG) that might represent the protochromosomes of the last common chordate ancestor (Putnam et al. 2008).
Exhaustive evaluation of the 17 CLGs revealed that most of the human genome
(112 segments spanning 2.68 Gb, which is the equivalent of 95 % of the
euchromatic genome) was affected by large-scale duplication events that occurred
on the stem vertebrate lineage before the teleost/tetrapod split. Analysis of the
distribution of the human segments among the 17 CLGs showed that nearly all
ancient chordate chromosomes were quadruplicated (Putnam et al. 2008)
(Fig. 16.3). This result robustly demonstrated the occurrence of two rounds of
genome duplication, corroborating previous lines of evidence based on analysis of
EC
180
CO
RR
179
Book ISBN: 978-3-642-31441-4
Page: 315/338
Two Rounds of Vertebrate Whole-Genome Duplication
UN
Editor Proof
16
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 16
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 316/338
C. Cañestro
231
16.2.1 Timing of the Vertebrate 2R-WGD
232
233
234
235
236
237
238
239
240
241
242
243
244
245
246
247
248
249
250
251
252
253
254
255
256
257
258
259
260
261
262
263
264
PR
OO
229
Analyses of the completely sequenced genome of the cephalochordate amphioxus
(Putnam et al. 2008; Holland et al. 2008) and the genomes from various urochordates
(Dehal et al. 2002; Small et al. 2007; Denoeud et al. 2010) validated Ohno’s
hypothesized lower-bound timing for the 2R-WGD as after the split between vertebrates and non-vertebrate chordates. Regarding the upper-bound timing, extensive
analysis of gene duplicates (Robinson-Rechavi et al. 2004) and the identification of
the four clusters in the genome of the elephant shark suggested that the 2R-WGD took
place before the cartilaginous/bony vertebrate split (Venkatesh et al. 2007; Ravi et al.
2009).
Within this time window, the most prevalent hypothesis suggests a scenario in
which the first round (R1) occurred before the split between gnathostome and
jawless vertebrates, and the second (R2) occurred in the stem jawed vertebrates
after their divergence from jawless vertebrates (Fig. 16.1). However, a second
scenario proposes that both rounds (R1 ? R2) of genome duplication took place
before the split between gnathostome and jawless vertebrates (pan-vertebrate
quadruplication (PV4) hypothesis (Kuraku et al. 2009)) (Fig. 16.1). Comparative
analysis of 55 gene families revealed a common expansion in both jawless and
jawed vertebrates, which has been interpreted as evidence supporting this second
scenario (Kuraku et al. 2009). Available information from sea lampreys and
hagfish does not permit us to discern between these two hypothetical scenarios,
because these organisms also appear to have suffered lineage-specific duplications
and reciprocal gene losses compared to vertebrates, which together obscure the
assessments of orthology/paralogy (reviewed in Kuraku 2008, 2010). For instance,
multiple Hox gene surveys in different species of sea lampreys and hagfish
suggested that extensive independent duplications of Hox genes might have
occurred during the evolution of jawless vertebrates (Pendleton et al. 1993;
Sharman and Holland 1998; Takio et al. 2004; Force et al. 2002; Irvine et al. 2002;
Fried et al. 2003; Stadler et al. 2004; Kuraku et al. 2009). Some of the jawless Hox
clusters might have disintegrated, casting doubt as to the usefulness of Hox genes
as reliable markers to trace duplications during genome evolution in stem vertebrates (Kuraku 2011). Finally, recent phylogenetic analysis of the degenerated
ParaHox cluster in hagfish has opened the possibility of a third scenario, in which
both rounds (R1 ? R2) occurred in stem jawed vertebrates after their divergence
D
228
TE
227
EC
226
CO
RR
225
F
230
specific regions of interest, such as the Hox-bearing regions (Garcia-Fernàndez and
Holland 1994; Larhammar et al. 2002) and the major histocompatibility regions
(Vienne et al. 2003; Danchin and Pontarotti 2004). Spring (1997) proposed the
term ‘‘tetralogs’’ to refer groups of quadruplicated vertebrate genes at four
different chromosomal locations formed by the 2R-WGD corresponding to a single
invertebrate gene, with all four more similar to each other than to members of the
other tetralogy group.
224
UN
Editor Proof
316
Layout: T1 Standard SC
Chapter No.: 16
Book ISBN: 978-3-642-31441-4
Page: 317/338
Two Rounds of Vertebrate Whole-Genome Duplication
317
PR
OO
F
Human Chromosomes
X
Y
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
Bfl_V2_12
0Mb
1Mb
D
Amphioxus contig Bfl_V2_12
CO
RR
EC
Human Chromosomes
TE
X
Y
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
Bfl_V2_21
0Mb
2Mb
4Mb
6Mb
8Mb
10Mb
Amphioxus contig Bfl_V2_21
Fig. 16.3 Quadruplicated conserved syntenic pattern between the amphioxus and the human
genome as a result of the 2R-WGD. Dot-plots display the distribution throughout human
chromosomes (y-axes) of human orthologs (blue dots) of amphioxus genes (black dots) located in
two arbitrarily selected genomic regions of approximately 1 Mb (a) and 10 Mb (b) (x-axes). The
dot-plots reveal four major human chromosomes (yellow shadow) of conserved synteny as the
product of the two rounds of whole-genome duplication. In panel (a), the four paralogons
coincide with the Hox-cluster bearing chromosomes 2, 7, 12, and 17, whereas in panel B the four
paralogons coincide with the endothelin receptors and ParaHox-cluster bearing chromosomes
2/5, 4, 13, and X. The dot-plots were generated as described in Canestro et al. (2009) using the
Synteny Database (Catchen et al. 2009)
UN
Editor Proof
16
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 16
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 318/338
C. Cañestro
275
16.2.2 Mechanisms Underlying the Vertebrate 2R-WGD
272
273
276
277
278
279
280
281
282
283
284
285
286
287
288
289
290
291
292
293
294
295
296
297
298
299
300
301
302
303
304
305
PR
OO
270
271
A question that still remains is how did the stem vertebrate genome become
octoploid by two rounds of tetraploidization. There are two main mechanisms of
tetraploidization observed in many species of plants and animals (Van de Peer et al.
2009). The first mechanism is allotetraploidy, which occurs when two related but
not identical genomes are combined by hybridization of closely related species and
associated (often subsequent) genome duplication. In the case of allotetraploidy, the
pairs of distinct ‘homologous’ chromosomes that are sufficiently different due to
their separated origin are called homeologs. The second mechanism is autotetraploidy, which occurs when the genomes are not sufficiently diverged into
homeologous sets; autotetraploidy therefore ranges from the combination of
genomes of two conspecific individuals (perhaps from different populations) to the
combination of identical genomes from a single individual. The genetic attributes
of allo- and autotetraploids differ and may have substantial effects at individual,
population, and species levels (see also Chap. 2, this volume). Both allotetraploidy
and autotetraploidy could be generated by several processes such as: (i) an
abnormal non-disjunction of sister chromatids at meiosis; (ii) the uncoupling of
mitotic DNA replication and cell division during early development of the germ
line (this process, for instance, occurs normally during the endoreplication of
megakaryocytic bone-marrow precursors of blood platelets, or during the development of the oikoblastic epithelia that secrete the house in basal urochordate
larvaceans); (iii) potential cell fusion during early embryo development or in germline precursors in syncytial gametogenesis (cell fusion is observed naturally, for
instance, in skeletal muscle cells and placenta) (reviewed in Storchova and Pellman
2004; Shemer and Podbilewicz 2000).
In the case of allotetraploids, each pair of homologous chromosomes should
segregate normally during meiosis, and genetic interchange between homeologous
chromosomes is rare. If two consecutive events of allotetraploidization occurred in
stem vertebrates, we would predict that in the ideal situation in the absence of gene
losses, a phylogenetic tree of homeologs will render a symmetrical (A,B) (C,D)
topology (Furlong and Holland 2002).
D
269
TE
268
EC
267
CO
RR
266
F
274
from jawless vertebrates (Furlong et al. 2007) (Fig. 16.1). The validation of this
third scenario could have a significant impact on our understanding of vertebrate
evolution, because it would imply that the 2R-WGD would have not been
important for the origin of vertebrate innovations (i.e. big brain, neural crest cells,
and placodes, which clearly exist in jawless vertebrates (Kuratani and Ota 2008;
Kuratani 2009)). According to this third scenario, however, the 2R-WGD would
have been important for the radiation of gnathostomes into cartilaginous fish, bony
fishes, and tetrapods. A solid answer about the timing of the 2R-WGD may have to
wait until larger-scale comparisons of the whole-genome organization of hagfish
and lampreys are available.
265
UN
Editor Proof
318
Layout: T1 Standard SC
Chapter No.: 16
312
313
314
315
316
317
318
319
320
321
322
323
324
325
326
327
328
329
330
331
332
333
334
335
336
337
338
339
340
341
342
343
344
345
F
311
PR
OO
310
16.3 Consequences of the 2R-WGD on the Evolution
of Vertebrate Genome Structure
D
309
It has been suggested that polyplodization events, at least in plants, can trigger
genomic stress associated with major genomic rearrangements, in many cases
mediated by a burst of mobilization of transposable elements (Matzke and Matzke
1998; Comai 2000). Transposable elements can be substrates for unequal and
illegitimate recombination and can be responsible for a variety of genome
reorganizations associated with the transposition, including chromosomal insertions, deletions, inversions, translocations, and duplications. Lineage-specific
genome rearrangements mediated by transposable elements might facilitate rapid
evolution, reproductive isolation of different populations, and consequently species
diversification (Parisod et al. 2010).
Contrary to possible genome reorganization after polyploidization, as noted
above for plants, in stem vertebrates, recent work based on proximate gene pair
methods and measurement of syntenic clustering conservation found that the 2RWGD in vertebrates were not followed by an increase of genome rearrangement
(Hufton et al. 2008). Unexpectedly, this work measured massive genome rearrangements prior to the 2R-WGD, which has been interpreted as a pre-existing
‘disposition’ toward genomic structural change (Hufton et al. 2008). Interestingly, in contrast to the archetypal condition that has been described in the
organization of particular genomic regions (e.g. Hox-cluster region (GarciaFernàndez and Holland 1994)), the amphioxus genome structure is not exceptionally well conserved, evolving its own particular type of repetitive elements
(e.g. ‘mirage’ minisatellites (Cañestro et al. 2002b; Ebner et al. 2010)), undergoing extensive local tandem gene duplications (see section below), and experiencing a moderate rate of synteny loss similar to that of sea urchin or sea
TE
308
319
In autopolyploids, however, meiotic pairing might occur between any of the four
identical chromosomes at meiosis I, facilitating genetic interchanges freely among
the four alleles, and leading to ‘tetrasomic inheritance’. Eventually the alleles, and
chromosomes, might diverge, starting a process of diploidization that reestablishes
diploidy. Randomly one of the chromosomes will diverge first and no longer form
homologous structures, while the other three will keep pairing until another further
divergence. Hence, if two consecutive events of autotetraploidy occurred in quick
succession (pseudo-octoploidy) in stem vertebrates, we would predict that in the
absence of gene losses, gene family phylogenetic trees will likely render
asymmetrical (((A,B),C),D) topologies (Furlong and Holland 2002). Because many
vertebrate gene families do render asymmetrical tree topologies (Friedman and
Hughes 2001; Hughes 1999; Hughes and Friedman 2003), two quick consecutive
events of autotetraploidy have been considered a likely mechanism for the 2R-WGD
in stem vertebrates (Furlong and Holland 2002; Lynch and Wagner 2009).
EC
307
CO
RR
306
Book ISBN: 978-3-642-31441-4
Page: 319/338
Two Rounds of Vertebrate Whole-Genome Duplication
UN
Editor Proof
16
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 16
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 320/338
354
355
356
357
358
359
360
361
362
363
364
365
366
367
368
369
370
371
372
373
374
375
376
377
378
379
380
381
382
383
384
385
386
387
F
PR
OO
353
D
351
352
TE
349
350
EC
348
anemone (Hufton et al. 2008). Therefore, the amphioxus genome structure
cannot be considered a fossil genome representing the pre-duplication condition,
at least in terms of genome structure (Garcia-Fernàndez et al. 2001; Hufton et al.
2008), although it is far less divergent from the vertebrate genome structure than
is any known urochordate genome (Dehal et al. 2002; Denoeud et al. 2010;
Louis et al. 2012).
There have been several attempts to infer the karyotype and genome structure
from common chordate ancestors and to reconstruct the evolutionary history
leading to present chromosome structures. The first comparisons of conserved
syntenic associations in different vertebrate karyotypes, using an in silico chromosome painting approach, allowed reconstructions of the ancestral vertebrate
genome containing 10–13 ancestral proto-chromosomes (Kohn et al. 2006;
Nakatani et al. 2007). Recently, the sequencing of the amphioxus genome has
allowed researchers to reconstruct the ancestral chordate genome as consisting of
17 conserved syntenic blocks, which might represent the ancient chordate protochromosomes (Putnam et al. 2008).
After the 2R-WGD, under the naive assumption of absence of loss or fusions of
chromosomes, we would expect 68 (17 9 4) proto-vertebrate segments, but
parsimonious reconstruction of chromosome history revealed that numerous
chromosomal fusions and translocations have occurred. These reconstructions
predict at least 20 fusions that led to 37–49 chromosomes in the bony vertebrate
ancestor, which became 12 chromosomes in the stem teleost ancestor due to many
additional fusions, and 33–45 chromosomes in the stem tetrapod ancestor due to at
least 4 fusions shared between human and chicken genomes (Putnam et al. 2008;
Naruse et al. 2004; Nakatani et al. 2007). An excellent example of chromosomal
rearrangement after the 2R-WGD has been recently provided by a phylogenetic
analysis of members of the four Hox paralogons that resulted in a (B(A(C,D)))
topology. These results suggest that two chromosomal rearrangements between
protochromosomes 11 and 4, and 7 and 5 occurred after the clusters duplicated but
before the diversification of extant vertebrates 450 million years ago (Lynch and
Wagner 2009). These chromosomal rearrangements resolve conflicting data
regarding the order of linked genes and support the hypothesis that the 2R-WGD
occurred by two consecutive events of autotetraploidy, and thereby the ancestral
vertebrate might have been ‘‘pseudo-octoploid’’. Interestingly, the asymmetrical
(B(A(C,D))) topology of the vertebrate Hox cluster (Lynch and Wagner 2009)
contrasts with the symmetrical (A,B) (C,D) topology inferred from the cartilaginous elephant shark using the amphioxus Hox cluster as the outgroup (Ravi et al.
2009). Further extensive analyses including HoxA-D clusters from a broader
representation of cartilaginous and bony vertebrates will be required to resolve
these conflicting topologies, which could suggest that the Hox-cluster rearrangement took place after the cartilaginous/bony vertebrate split and not immediately
subsequent to the 2R-WGD.
CO
RR
346
347
C. Cañestro
UN
Editor Proof
320
Layout: T1 Standard SC
Chapter No.: 16
Book ISBN: 978-3-642-31441-4
Page: 321/338
Two Rounds of Vertebrate Whole-Genome Duplication
390
16.4.1 Function of Gene Duplicates After 2R-WGD
321
F
389
16.4 Consequences of the 2R-WGD on the Evolution
of Vertebrate Gene Fate
388
415
16.4.2 Gene Network Rewiring by Tranposons After 2R-WGD
396
397
398
399
400
401
402
403
404
405
406
407
408
409
410
411
412
413
416
417
418
419
420
421
422
423
424
425
D
395
TE
394
EC
393
CO
RR
392
PR
OO
414
After polyploidization, a period of transilience may follow in which genes might
enjoy extra ‘degrees of freedom’ to mutate without selective penalty (reviewed in
Soltis and Soltis 1999; Otto 2007). Understanding the processes by which genome
duplication might influence the fate of duplicated genes is crucial to evaluate how
the 2R-WGD might have impacted the evolution of vertebrate innovations.
Neofunctionalization and subfunctionalization are the two main processes driving
the functional fate of newly generated ohnologs after the 2R-WGD and have been
extensively discussed in the literature (Hughes 1994; Force et al. 1999; Lynch and
Conery 2000; Durand 2003; Postlethwait et al. 2004; Hoekstra and Coyne 2007;
Conant and Wolfe 2008; Semon and Wolfe 2007, 2008; Jimenez-Delgado et al.
2009). A prominent example of neofunctionalization related to the 2R-WGD
occurred during the expansion of the vertebrate retinoic acid receptor (RAR)
family, which acquired new functions in both their expression domains and in their
structural protein activities (Escriva et al. 2006). There are also examples,
however, in which neofunctionalization and subfunctionalization are related to
both 2R-WGD and local tandem duplications (e.g. the expansion of the vertebrate
globin superfamily, which promotes the vertebrate innovation related to oxygen
transport and storage (Hoffmann et al. 2011)), or merely related to local tandem
duplications, and not the 2R-WGD, such as the expansion of the vertebrate
Alcohol Dehydrogenase (Adh) family, which promotes the acquisition of new
enzymes for the synthesis of retinoic acid (Cañestro et al. 2000, 2002a, 2003b).
Therefore, not all vertebrate innovations can be exclusively attributed to the
2R-WGD, and the global weight of the impact of the 2R-WGD on the evolution of
vertebrate gene functions remains unknown.
391
In many cases, neofunctionalization and subfunctionalization can be due to
alterations in cis-regulatory elements that might lead to adaptative changes in
duplicated genes (Force et al. 1999). Many of the cis-regulatory elements appear to
be embedded in distinct repeat families, especially in transposable elements (TE)
(Thornburg et al. 2006; Polak and Domany 2006; Bourque et al. 2008). Analysis of
the distribution of 10,000 TEs in the human genome, for instance, revealed that
most TEs are concentrated under strong purifying selection near regulatory and
developmental genes (Lowe et al. 2007). Most of the described examples of TE
mobilization and rewiring of gene regulatory networks have been associated with
relatively recent events of TE mobilizations. For instance, TE-mediated rewiring
UN
Editor Proof
16
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 16
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 322/338
433
434
435
436
437
438
439
440
441
442
443
444
445
446
447
448
449
450
451
452
453
454
455
456
457
458
459
460
461
462
463
F
432
PR
OO
431
D
430
TE
428
429
for neofunctionalization after gene duplication has been recently described for the
sex-determining gene dmrt1bY in medaka fish, in which a novel regulatory
element driving a negative feedback on dmrt1bY has been acquired due to the
insertion of an Izanagi transposon (Herpin et al. 2010), or for the origin of a novel
gene regulatory network dedicated to pregnancy in placental mammals, which was
due to a transposition of the MER20 TE (Lynch et al. 2011).
A massive expansion of TEs appears, therefore, as a powerful mechanism that
could boost a vast redeployment of cis-regulatory elements into new gene
regulatory networks (Feschotte 2008), promoting large-scale events of neofunctionalization and subfunctionalization (van de Lagemaat et al. 2003; Bennetzen
2005; Bejerano et al. 2006). Polyploidization can trigger the mobilization of
transposable elements (Matzke and Matzke 1998; Parisod et al. 2010), because
recently duplicated genomes contain many redundant genes and substantial
repetitive DNA, which serve as buffer against TE insertional mutagenesis (Matzke
et al. 2000). According to this expectation, bursts of TE mobilization have been
described after polyploidization in different organisms (Matzke and Matzke 1998;
Comai 2000; SanMiguel et al. 1996, 1998).
A question that remains unclear is whether there was or was not a massive
TE mobilization after the 2R-WGD that could have favored a significant
redeployment of cis-regulatory elements into new gene regulatory networks in the
stem vertebrate lineage. Recent comparison of the diversity and content of TEs
between vertebrates and amphioxus has provided some insights that might help to
answer this question (Canestro and Albalat 2012). The dynamics of the TE content
within a genome follows a competition model in which the expansion of a
particular TE might cause the reduction of other types of TEs, consequently
reducing the TE diversity, until a new equilibrium that preserves the functionality
of the genome is reached (Abrusán and Krambeck 2006). According to this model,
if a massive expansion of TEs occurred after the 2R-WGD, we expect that the
diversity of TEs shared among vertebrates should be smaller than in cephalochordates. Consistent with this prediction, a recent comparative study reveals that
the shared TE diversity of vertebrates (14 superfamilies in lampreys, 28 in
ray-finned fishes, 20 in amphibians, 14 in reptiles, 10 in birds, and 15 in mammals)
is lower than the TE diversity in amphioxus (33 superfamilies), which makes
plausible the hypothesis that a TE burst could have occurred after the 2R-WGD in
the stem vertebrate lineage (Canestro and Albalat 2012). Further comparative
genomic analysis between different vertebrates and cephalochordates will be
required to test this hypothetical burst of TEs, and especially to evaluate its
putative impact on the evolution of gene functions after the 2R-WGD.
EC
427
CO
RR
426
C. Cañestro
UN
Editor Proof
322
Layout: T1 Standard SC
Chapter No.: 16
470
471
472
473
474
475
476
477
478
479
480
481
482
483
484
485
486
487
488
489
490
491
492
493
494
495
496
497
498
499
500
501
502
503
504
505
506
F
469
PR
OO
468
While several studies focus on the functional fate of retained gene duplicates, less
attention has been paid to how losses of paralogs or ohnologs might impact the
evolution of the functions of other genes (reviewed in Cañestro et al. 2007). Loss
of one copy of two fully redundant gene duplicates should not usually have
significant impact, but loss of one of the paralogs after functional divergence likely
has evolutionary consequences. Recent analyses of gene losses by comparative
genomics have led to the unexpected finding that significant components of the
developmental toolkit might be lost without major changes to the body plan
(Cañestro and Postlethwait 2007; Holland 2007), which suggests the presence of
compensatory mechanisms or the acquisition of innovations that have preserved
unaltered the ancestral condition (Cañestro et al. 2007). Tracing the evolution of
gene families throughout ancestral proto-chromosomes using blocs of conserved
synteny has become a powerful tool to clarify uncertain phylogenies, to detect
ohnologs gone missing (OGM) (Postlethwait 2007; Catchen et al. 2009, 2011), to
provide robust assessments of orthology and paralogy between different species,
and to discern evolutionary innovations from losses of ancestral features in sister
lineages (Canestro et al. 2009).
There are cases in which different ohnologs in different species acquire the same
expression pattern, which has been called function shuffling (McClintock et al.
2001) and synfunctionalization (Gitelman 2007), and in some cases the convergence
of expression patterns between paralogs can be related to OGM (Postlethwait 2007;
Canestro et al. 2009). The evolution of the vertebrate retinaldehyde dehydrogenease
Aldh1a family provides a paradigmatic example of how uncovering the evolution of
gene family members through the 2R-WGD has been fundamental to illuminating
how gene functions evolve among newly generated paralogs after genome duplications in the face of loss of ohnologs (Canestro et al. 2009). For instance, analysis
of conserved synteny revealed that the presence of Aldh1a1 in tetrapods and its
absence in teleost fish was not due to a tetrapod innovation, but to an OGM in the
teleost stem lineage, which was accompanied by a re-acquisition of ancestral
functions by surviving paralogs (Canestro et al. 2009). Medaka provides a more
radical example in which aldh1a2, the only survivor of the aldh1a family in this
species, recapitulates the expression pattern of all other aldh1a paralogs that have
been lost in medaka. This result is in agreement with a model of functional evolution
in which surviving genes re-acquire ancestral gene family roles in the face of loss of
ohnologs. Other examples that illustrate the importance of identifying OGMs
ohnologs are shown in the endothelin and agouti systems, in which the exclusive
presence of endothelin 4 (edn4) and the agouti-signaling protein 2 genes (asip2a/
b) in teleost fish was not due to a fish innovation related to the teleost-specific wholegenome duplication, but instead to a loss of ohnologs that originated in the 2R-WGD
in the tetrapod lineage (Braasch et al. 2009; Braasch and Postlethwait 2011). To
understand acquisition of functions of vertebrate ohnologs that were generated by
D
467
TE
466
EC
465
323
16.4.3 Ohnologs Gone Missing After 2R-WGD and Impact
on Surviving Ohnologs
CO
RR
464
Book ISBN: 978-3-642-31441-4
Page: 323/338
Two Rounds of Vertebrate Whole-Genome Duplication
UN
Editor Proof
16
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 16
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 324/338
512
513
514
515
516
517
518
519
520
521
522
523
524
525
526
527
528
529
530
531
532
533
534
535
536
537
538
539
540
541
542
543
544
545
546
F
PR
OO
511
16.5 Consequences of the 2R-WGD on Vertebrate Gene
Number and Functional Evolution
How many of the genes that were part of the original fourfold increase in genes
generated by 2R-WGD in the stem vertebrate have actually survived nonfunctionalization? And importantly, how significant have the functional consequences of
those retained genes been for promoting the origin and evolution of vertebrate
innovations? Estimates on gene retention in other organisms that have experienced a
WGD have reported *13 % retention over *100 million years (MY) in yeast
(Wolfe and Shields 1997), *72 % in maize over *11 MY (Ahn and Tanksley 1993;
Gaut and Doebley 1997), and *77 % in Xenopus over *40 MY (Hellsten et al.
2007). In vertebrates, a *33 % retention of divergent functional genes after the
2R-WGD over *500 MY was inferred initially based on theoretical models applied
to 270 gene families of the human genome (Nadeau and Sankoff 1997). More recent
and broader analyses based in the complete catalog of human ohnologs estimated a
rate of retention between 20 and 30 % (Putnam et al. 2008; Huminiecki and Heldin
2010; Makino and McLysaght 2010).
But how can we assess the impact of the 2R-WGD on the origin of vertebrate
complex features? A naive approach to estimating this impact could be to perform a
comparison of the total number of retained paralogs and their distribution among
functional categories in vertebrates and non-vertebrate chordates that did not undergo
any WGD since their split from our last common chordate ancestor. Comparison of
the gene catalog of the three chordate subphyla (i.e. cephalochordates, urochordates,
and vertebrates) has allowed us to identify a lower bound of 8,437 gene families with
members that descend from a single gene in the last common chordate ancestor
(Putnam et al. 2008). Through subsequent genome or local duplication, these families
account for 13,610 amphioxus genes, 13,401 human genes, and 7,216 ascidian genes,
the latter being a significantly lower number due to the extensive gene losses that have
occurred in urochordate lineages (Dehal et al. 2002; Cañestro et al. 2003a; Edvardsen
et al. 2005; Denoeud et al. 2010). Although it is frequently true that the multiple
ohnologs of a vertebrate gene family are represented by a single gene in amphioxus,
the total number of paralogs derived from a single-copy gene in the last common
ancestor is surprisingly similar between amphioxus (13,610) and human (13,401)
(Putnam et al. 2008). Therefore the mere total numbers of retained genes after the
2R-WGD duplicates might not be the key to explain the gain of complexity during the
evolution of the vertebrate lineage in comparison with amphioxus.
In vertebrates, analysis of the functional categories of the gene families that
have expanded after the 2R-WGD revealed that cell signalers and transcriptional
D
510
TE
509
the 2R-WGD, both the impact of the retention of neo- or subfunctionalized
ohnologs, as well as the impact of OGM, on the functions of other survivor gene
family members should be studied.
EC
508
CO
RR
507
C. Cañestro
UN
Editor Proof
324
Layout: T1 Standard SC
Chapter No.: 16
554
555
556
557
558
559
560
561
562
563
564
565
566
567
568
569
570
571
572
573
574
575
576
577
578
579
580
581
582
583
584
585
586
587
588
589
590
591
F
552
553
PR
OO
551
D
550
TE
549
325
regulators of developmental pathways are generally retained as multiple ohnologs
(Roux and Robinson-Rechavi 2008; Putnam et al. 2008; Hufton et al. 2008;
Huminiecki and Heldin 2010). Genes associated with basic cellular functions (i.e.
translation, replication, splicing, and recombination, with the important exception
of cell cycle), however, have been less successfully retained after the 2R-WGD
(Huminiecki and Heldin 2010) (although see Gout et al. (2009) for different results
in other organisms that have also undergone WGD). Analysis of the human
genome reveals that dosage-balance constraints act on the retention of ohnologs,
resulting in an enrichment of dosage-balanced genes, an observation predicted
following WGD (Birchler and Veitia 2007, 2010) and also reported for other
vertebrates, plants, and yeast (e.g., Paterson et al. 2006). Interestingly, many of
these retained ohnologs in humans are refractory to copy number variation, have
rarely experienced subsequent small-scale duplication, and are frequently associated with diseases related to dosage-imbalance such as down syndrome (Makino
and McLysaght 2010). Analysis of retained genes that have originated in vertebrates by local duplications revealed a strong underrepresentation of genes related
to cell communication, cell cycle, and embryo development (Huminiecki and
Heldin 2010).
In amphioxus, although a thorough analysis of amphioxus-specific gene family
expansions has not been performed, Table 16.1 shows an extensive list of
amphioxus-specific duplicated genes reported in the literature (this list is probably
not complete, and may be biased toward the research with which I am most
familiar). This list shows numerous retained duplicates from a broad array of
functional categories, including metabolic enzymes, members of transduction and
signaling cascades, members of the immunity system, as well as pivotal transcription factors of developmental pathways. Awaiting a more exhaustive analysis,
including different amphioxus species to infer the ancestral cephalochordate
condition, the list in Table 16.1 shows no obvious bias toward any particular
functional category, although it is noticeable that duplicated developmental transcription factors do not account for more than two paralogs (with the exception of
the eight hairy amphioxus paralogs (Minguillon et al. 2003)).
Remarkably, the main difference between the newly acquired paralogs in
amphioxus and ohnologs in vertebrates is the mechanism of duplication. While
approximately 25 % of the ancestral chordate gene families have two or more
ancient vertebrate ohnologs generated by the 2R-WGD, there is strong evidence that
most amphioxus paralogs originated by local tandem duplications rather than largescale chromosomal duplications (Table 16.1). Therefore, considering the functional
bias of retention of genes duplicated by WGD or local duplication, it is reasonable to
speculate that the key influence of the 2R-WGD promoting the successful diversification of vertebrate features resides in the fact that whole networks were
duplicated, in contrast to local duplications such as those that occurred in amphioxus, an organism that seems to have maintained morphological and genetic stasis
during the last 200 million years (Garcia-Fernàndez and Holland 1994; Cañestro
et al. 2002a; Somorjai et al. 2008; Canestro and Albalat 2012; Paps et al. 2012).
Duplication of whole gene networks is dosage-balanced and increases the
EC
548
CO
RR
547
Book ISBN: 978-3-642-31441-4
Page: 325/338
Two Rounds of Vertebrate Whole-Genome Duplication
UN
Editor Proof
16
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 16
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 326/338
C. Cañestro
Table 16.1 List of paralogs originated independently in the amphioxus lineage
Functional category
Gene
Number of
References
amphioxus paralogs
28
39
406
EC
Developmental
transcription factors
F
Toll-like receptors
C1q-like
LRR-containing
gene models
Hairy
Brachyury
bHLH
Emx
Hox13–14
Evx
Mnx
Uncx
Pou3
Lhx2/9
Iro
CO
RR
Immunity system
Cañestro et al. (2006)
Albalat (2009)
Albalat et al. (2011)
PR
OO
5
3
22
12
14
22
6
5
10
8
7
8
3
20
18
3
5
12
TE
Transduction and
signaling cascades
Aldh1a
Cyp26
Hsd11b1
Rdh cluster
Bdh1
Rdh11/12
Hsd17b8
ApoD
Cyp2
Cyp11/24/27
Hsd3b
Fabp
Nos
Opsins
GPR54
GnRH
CRHR
Somatostatine
receptors
LGR7/8
PTHR
Secretin receptors
Estrogen receptors
NOK
ACK
TIE
MARTK
EXTK
HNK-Ras
Calmoduline-like
Andreakis et al. (2011)
Holland et al. (2008)
D
Metabolic proteins
UN
Editor Proof
326
6
4
5
2
22
3
7
8
47
2
3
8
2
2
2
2
2
2
2
2
2
2
Bridgham et al. (2008)
D’Aniello et al. (2008)
Bertrand et al. (2009)
Karabinos and
Bhattacharya (2000)
Holland et al. (2008)
Minguillon et al. (2003)
Holland et al. (1995)
Araki et al. (1996)
Minguillon et al. (2002)
Ferrier et al. (2000)
Ferrier et al. (2001)
Holland et al. (2008)
Layout: T1 Standard SC
Chapter No.: 16
598
599
600
601
602
603
604
605
606
607
608
609
610
611
612
613
614
615
616
617
618
619
620
621
622
623
624
625
626
627
628
629
630
631
632
F
597
PR
OO
596
D
595
TE
594
327
evolvability to generate novel functions, which in the case of the vertebrate 2RWGD could have led to an increase in complexity of the signaling and developmental regulatory networks that facilitated the acquisition of innovations.
In addition to the evolutionary role of coding genes in the acquisition of innovations, microRNAs (miRNAs) also play crucial roles during development and have
been postulated as important players for the evolution of organismal complexity (Lee
et al. 2007; Sempere et al. 2006). Analysis of miRNAs in chordate species showed that
the 2R-WGD has increased the diversity of the inventory of miRNAs in vertebrates,
which correlated with the increase of complex patterns of tissue specificity of
miRNAs (Heimberg et al. 2008; Campo-Paysaa et al. 2011). However, the finding of
41 vertebrate-specific miRNA families, absent in non-vertebrate chordates, suggests
that their origin must have occurred in stem vertebrates after their separation from
urochordates and is not explained by the 2R-WGD (Heimberg et al. 2008). The
appearance of these 41 vertebrate-specific miRNA families has been proposed as a
potential key evolutionary force lying behind the dramatic increase of vertebrate
complexity (Heimberg et al. 2008). Future exhaustive analysis of the expression
patterns of the members of these 41 vertebrate-specific families, and an understanding
of their roles, will allow a reevaluation of the importance that this innovation could
have had on the origin of vertebrate features.
16.6 Consequences of the 2R-WGD on the Innovation
of Vertebrate Features
A major question not resolved yet is the precise impact of the 2R-WGD on the
innovation of particular vertebrate features. Three vertebrate features are perhaps
the most prominent innovations: derivatives from neural crest cells, sensory organs
concentrated in the head derived from ectodermal placodes, and a big complex
brain. When taken together, these features probably allowed the transition from
ancestral, peaceful, filter-feeding, non-vertebrate chordates to active, voracious,
vertebrate predators (Northcutt and Gans 1983; Gans and Northcutt 1983);
reviewed in Yu et al. 2008; Holland 2009) (Fig. 16.1).
Vertebrate neural crest cells are a transient population of developmental cells that
delaminate at the border of the neural plate through an epithelial–mesenchymal
transition, migrate, and differentiate at their final destination into a variety of
structures such as sensory neurons, glial cells, peripheral nervous system, pigment
cells, smooth muscle cells, connective tissue, cranio-facial cartilage, skeletal bones,
and teeth (Weston 1970). Vertebrate crest development depends on four crucial sets
of genes that form what is called the neural crest gene regulatory network
(NC-GRN): (1) patterning signal genes establish the expression of (2) neural plate
border specifier genes, which activate (3) crest specifier genes, which turn on (4)
neural crest effector genes that provide differentiated products (Meulemans and
Bronner-Fraser 2004, 2005; Ota and Kuratani 2007; Sauka-Spengler et al. 2007).
Analysis of the amphioxus genome has revealed the presence of cephalochordate
EC
593
CO
RR
592
Book ISBN: 978-3-642-31441-4
Page: 327/338
Two Rounds of Vertebrate Whole-Genome Duplication
UN
Editor Proof
16
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 16
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 328/338
639
640
641
642
643
644
645
646
647
648
649
650
651
652
653
654
655
656
657
658
659
660
661
662
663
664
665
666
667
668
669
670
671
672
673
674
675
676
677
F
638
PR
OO
637
D
636
TE
635
orthologs from all of these four sets of genes, including (1): Fgf, Wnt, Bmp, Notch
Dlx, AP2, SoxB, Zic, and islet; (2): Pax3/7, Msx, Dlx5, and Zic; (3): Snail, SoxE,
AP2, Twist, Id, FoxD, and Myc; and (4): Rho, cRet, Erbb3, Mitf, tyrosinase, and
tyrosinase-related genes, with the remarkable exception of the tyrosine kinase c-Kit
essential for migration and survival of crest cells, and the gene for myelin protein
P0, consistent with the notion that the glial myelin sheath is a vertebrate innovation
(Meulemans and Bronner-Fraser 2007; Holland et al. 2008; Holland and Short 2008;
Nikitina et al. 2009).
The fact that most of the specifier genes are present as single copy in amphioxus,
but multiple paralogs in vertebrates, presumably due to the 2R-WGD, has led to the
hypothesis that neofunctionalization and subfunctionalization of paralogs may have
facilitated the co-option of ancestral genes into the NC-GRN (Sauka-Spengler et al.
2007; Meulemans and Bronner-Fraser 2007; Holland et al. 2008; Holland and Short
2008). Gene ontology (GO) analysis estimates that 91 % of the neural crest genes in
vertebrates have been co-opted from genes already present in basal metazoans,
while the remaining 9 % of the neural crest genes are vertebrate innovations
(Martinez-Morales et al. 2007), including the assembly of new signaling pathways
like the endothelin system (Braasch et al. 2009). The evolution of the vertebrate
NC-GRN, therefore, appears as the result of a combination of ancestral gene cooption, newly evolving genes, and amplification of these components by the 2RWGDs (Braasch et al. 2009).
Interestingly, however, the recent description in urochordates of neural crest-like
cells that express typical vertebrate crest marker orthologs, migrate, and differentiate into pigments challenges the idea that neural crest cells are a vertebrate
innovation (Jeffery et al. 2004, 2008; Jeffery 2007). Thus, it cannot be discounted
that some types of neural crest cells might have been present in the last common
ancestor of olfactores (urochordates ? vertebrates), followed by losses during the
significant morphological and genetic simplification suffered by urochordate
lineages (Seo et al. 2004; Cañestro et al. 2005; Cañestro and Postlethwait 2007;
Holland 2007). Therefore, it seems plausible to consider that the 2R-WGD might
have not been crucial for the origin of the neural crest cells, but the 2R-WGD might
have been important for increasing the evolvability of the NC-GRN and the
diversification of derivative structures.
Similar conclusions have been reached through studies of the gene regulatory
network underlying placode and brain development. Analysis of placode-marking
genes (e.g. Eye, Pitx, Six, and Pax) in ascidian and larvacean urochordates suggested
that the last common olfactore ancestor already presented multiple placode derivatives, such as olfactory and adenohypophyseal. Additional and independent
proliferation and loss of a variety of placodes probably occurred in both urochordate
and vertebrate lineages (Bassham and Postlethwait 2005; Mazet et al. 2005), in
some cases recruiting paralogs that had been independently duplicated in both
urochordates and vertebrates (Bassham et al. 2008).
Despite the fact that non-vertebrate chordates have a simple brain lacking a
midbrain and a midbrain–hindbrain organizer (MHB), most brain-making gene
orthologs are present in non-vertebrate chordates, suggesting that vertebrate brain
EC
634
CO
RR
633
C. Cañestro
UN
Editor Proof
328
Layout: T1 Standard SC
Chapter No.: 16
684
685
686
687
688
689
690
691
692
693
694
695
696
697
698
699
700
701
702
703
704
705
706
707
708
709
710
711
712
713
714
715
716
717
718
F
683
PR
OO
682
D
681
TE
680
329
features were built on a foundation already present in the ancestral chordate
probably facilitated by the new ohnologs created by the vertebrate 2R-WGD
(reviewed in Holland 2009). Recent analysis of developmental genes in the
ascidian brain revealed that the expression of Fgf8 can reorganize the expression
of other brain genes and transform hindbrain structures into an expanded
mesencephelon, recapitulating the organizing activity of the vertebrate MHB and
therefore suggesting that the MHB was already present at least in the last common
ancestor of olfactores (Imai et al. 2009). Analysis of urochordate genomes
revealed that important genes (i.e. Gbx) for the positioning of the MHB have been
lost in stem urochordates (Cañestro et al. 2005), as has the retinoic acid dependent
anterior–posterior axial patterning of the central nervous system (Cañestro et al.
2006), making plausible the hypothesis that the absence of midbrain in urochordates is not due to a vertebrate innovation of a midbrain, but a simplification in
urochordates of an ancestral tripartite brain structure (Cañestro and Postlethwait
2007; Cañestro et al. 2007). Evolutionary analysis of the origin of the complex
Nova-regulated splice variants of the vertebrate brain genes revealed that many of
these variants were already present in the last common olfactore ancestor (Irimia
et al. 2011). It is possible that the 2R-WGD promoted the increase of the
complexity of Nova-dependent splice variants in the vertebrate brain, although a
simplification of this system during urochordate evolution cannot be discarded.
In conclusion, it is likely that the origin of vertebrate features such as neural
crest cells, placodes, and a complex tripartite brain are not related to the 2R-WGD,
but that these features were already present to some extent in stem non-vertebrate
chordates (Fig. 16.1) (reviewed in Donoghue et al. 2008). However, it is likely that
the subsequent evolution of these three features has been strongly influenced by
the new ohnologs that originated after the 2R-WGD, due to processes of
neofunctionalization, subfunction partitioning and subsequent refinement,
recruitment of cis-regulatory elements driven by genome rearrangement and
transposable element activity, inventions of novel miRNA families, and evolution
of novel splice variants, which overall increased the complexity of duplicated
developmental gene regulatory networks after the 2R-WGD. Future integrative
analysis of comparative genomics, functional evo-devo, and examinations of gene
regulatory networks in a wide variety of non-vertebrate chordates as well as
basally divergent jawless vertebrates will help to narrow down the precise timing
of the 2R-WGD and evaluate its actual impact on the origin and evolution of
vertebrate features.
Probably the new ‘2R, or not 2R’ question (Hughes and Friedman 2003) is now
to ascertain whether the origins of vertebrate innovations were, or were not, the
consequence of the 2R-WGD, and to understand the mechanisms by which the
2R-WGD increased the evolvability of developmental gene regulatory networks
that facilitated the diversification of complex vertebrate features.
EC
679
CO
RR
678
Book ISBN: 978-3-642-31441-4
Page: 329/338
Two Rounds of Vertebrate Whole-Genome Duplication
UN
Editor Proof
16
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 16
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 330/338
C. Cañestro
Acknowledgments This work has been supported by grant BFU2010-14875 from the Ministerio
de Ciencia e Innovación (Spain). I would like to thank Julian Catchen for his generous support on
the Synteny Database, and Ingo Braasch, John H. Postlethwait, Ricard Albalat, and Adriana
Rodriguez for their valuable comments on the chapter, and cheerful and endless discussions on
‘‘2R, or not 2R, that is the question … on vertebrate innovations’’.
724
References
725
726
727
728
729
730
731
732
733
734
735
736
737
738
739
740
741
742
743
744
745
746
747
748
749
750
751
752
753
754
755
756
757
758
759
760
761
762
763
764
765
766
767
Abrusán G, Krambeck H-J (2006) Competition may determine the diversity of transposable
elements. Theor Popul Biol 70(3):364–375
Aburomia R, Khaner O, Sidow A (2003) Functional evolution in the ancestral lineage of
vertebrates or when genomic complexity was wagging its morphological tail. J Struct Funct
Genomics 3(1–4):45–52
Ahn S, Tanksley SD (1993) Comparative linkage maps of the rice and maize genomes. Proc Natl
Acad Sci U S A 90(17):7980–7984
Albalat R (2009) The retinoic acid machinery in invertebrates: ancestral elements and vertebrate
innovations. Mol Cell Endocrinol 313(1–2):23–35
Albalat R, Brunet F, Laudet V, Schubert M (2011) Evolution of retinoid and steroid signaling:
vertebrate diversification from an amphioxus perspective. Genome Biol Evol 3:985–1005
Andreakis N, D’Aniello S, Albalat R, Patti FP, Garcia-Fernandez J, Procaccini G, Sordino P,
Palumbo A (2011) Evolution of the nitric oxide synthase family in metazoans. Mol Biol Evol
28(1):163–179
Araki I, Terazawa K, Satoh N (1996) Duplication of an amphioxus myogenic bHLH gene is
independent of vertebrate myogenic bHLH gene duplication. Gene 171(2):231–236
Bailey W, Kim J, Wagner G, Ruddle F (1997) Phylogenetic reconstruction of vertebrate Hox
cluster duplications. Mol Biol Evol 14:843–853
Bassham S, Cañestro C, Postlethwait JH (2008) Evolution of developmental roles of Pax2/5/8
paralogs after independent duplication in urochordate and vertebrate lineages. BMC Biol 6:35
Bassham S, Postlethwait JH (2005) The evolutionary history of placodes: a molecular genetic
investigation of the larvacean urochordate Oikopleura dioica. Development 132(19):4259–
4272
Bejerano G, Lowe CB, Ahituv N, King B, Siepel A, Salama SR, Rubin EM, Kent WJ, Haussler D
(2006) A distal enhancer and an ultra conserved exon are derived from a novel retroposon.
Nature 441(7089):87–90
Bennetzen JL (2005) Transposable elements, gene creation and genome rearrangement in
flowering plants. Curr Opin Genet Dev 15(6):621–627
Bertrand S, Campo-Paysaa F, Camasses A, Garcia-Fernandez J, Escriva H (2009) Actors of the
tyrosine kinase receptor downstream signaling pathways in amphioxus. Evol Dev 11(1):13–26
Birchler JA, Veitia RA (2007) The gene balance hypothesis: from classical genetics to modern
genomics. Plant Cell 19(2):395–402
Birchler JA, Veitia RA (2010) The gene balance hypothesis: implications for gene regulation,
quantitative traits and evolution. New Phytol 186(1):54–62
Bourlat SJ, Juliusdottir T, Lowe CJ, Freeman R, Aronowicz J, Kirschner M, Lander ES,
Thorndyke M, Nakano H, Kohn AB, Heyland A, Moroz LL, Copley RR, Telford MJ (2006)
Deuterostome phylogeny reveals monophyletic chordates and the new phylum Xenoturbellida. Nature 444(7115):85–88
Bourque G, Leong B, Vega VB, Chen X, Lee YL, Srinivasan KG, Chew JL, Ruan Y, Wei CL, Ng
HH, Liu ET (2008) Evolution of the mammalian transcription factor binding repertoire via
transposable elements. Genome Res 18(11):1752–1762
Braasch I, Postlethwait JH (2011) The teleost agouti-related protein 2 gene is an ohnolog gone
missing from the tetrapod genome. Proc Natl Acad Sci U S A 108(13):E47–E48
CO
RR
EC
TE
D
PR
OO
F
719
720
721
722
723
UN
Editor Proof
330
Layout: T1 Standard SC
Chapter No.: 16
331
EC
TE
D
PR
OO
F
Braasch I, Volff JN, Schartl M (2009) The endothelin system: evolution of vertebrate-specific
ligand–receptor interactions by three rounds of genome duplication. Mol Biol Evol
26(4):783–799
Bridgham JT, Brown JE, Rodriguez-Mari A, Catchen JM, Thornton JW (2008) Evolution of a
new function by degenerative mutation in cephalochordate steroid receptors. PLoS Genet
4(9):e1000191
Brusca RC, Brusca GJ (2002) Invertebrates. Sinauer Associates, Sunderland
Campo-Paysaa F, Semon M, Cameron RA, Peterson KJ, Schubert M (2011) microRNA
complements in deuterostomes: origin and evolution of microRNAs. Evol Dev 13(1):15–27
Cañestro C, Albalat R (2012) Transposon diversity is higher in amphioxus than in vertebrates:
functional and evolutionary inferences. Brief Funct Genomics 11(2):131–141
Cañestro C, Albalat R, Hjelmqvist L, Godoy L, Jornvall H, Gonzalez-Duarte R (2002a) Ascidian
and amphioxus Adh genes correlate functional and molecular features of the ADH family
expansion during vertebrate evolution. J Mol Evol 54(1):81–89
Cañestro C, Bassham S, Postlethwait JH (2003a) Seeing chordate evolution through the Ciona
genome sequence. Genome Biol 4(3):208–211
Cañestro C, Bassham S, Postlethwait JH (2005) Development of the central nervous system in the
larvacean Oikopleura dioica and the evolution of the chordate brain. Dev Biol 285(2):298–
315
Cañestro C, Catchen JM, Rodriguez-Mari A, Yokoi H, Postlethwait JH (2009) Consequences of
lineage-specific gene loss on functional evolution of surviving paralogs: ALDH1A and
retinoic acid signaling in vertebrate genomes. PLoS Genet 5(5):e1000496
Cañestro C, Godoy L, Gonzàlez-Duarte R, Albalat R (2003b) Comparative expression analysis of
Adh3 during arthropod, urochordate, cephalochordate and vertebrate development challenges
its predicted housekeeping role. Evol Dev 5(2):157–162
Cañestro C, Gonzàlez-Duarte R, Albalat R (2002b) Minisatellite instability at the Adh locus
reveals somatic polymorphism in amphioxus. Nucleic Acids Res 30(13):2871–2876
Cañestro C, Hjelmqvist L, Albalat R, Garcia-Fernàndez J, Gonzàlez-Duarte R, Jörnvall H (2000)
Amphioxus alcohol dehydrogenase is a class 3 form of single type and of structural
conservation but with unique developmental expression. Eur J Biochem 267:6511–6518
Cañestro C, Postlethwait JH (2007) Development of a chordate anterior–posterior axis without
classical retinoic acid signaling. Dev Biol 305(2):522–538
Cañestro C, Postlethwait JH, Gonzàlez-Duarte R, Albalat R (2006) Is retinoic acid genetic
machinery a chordate innovation? Evol Dev 8(5):394–406
Cañestro C, Yokoi H, Postlethwait JH (2007) Evolutionary developmental biology and genomics.
Nat Rev Genet 8(12):932–942
Catchen JM, Braasch I, Postlethwait JH (2011) Conserved synteny and the zebrafish genome.
Methods Cell Biol 104:259–285
Catchen JM, Conery JS, Postlethwait JH (2009) Automated identification of conserved synteny
after whole-genome duplication. Genome Res 19(8):1497–1505
Chain FJ, Evans BJ (2006) Multiple mechanisms promote the retained expression of gene
duplicates in the tetraploid frog Xenopus laevis. PLoS Genet 2(4):e56
Comai L (2000) Genetic and epigenetic interactions in allopolyploid plants. Plant Mol Biol 43
(2–3):387–399
Conant GC, Wolfe KH (2008) Turning a hobby into a job: how duplicated genes find new
functions. Nat Rev Genet 9(12):938–950
Coulier F, Popovici C, Villet R, Birnbaum D (2000) MetaHox gene clusters. J Exp Zool
288(4):345–351
D’Aniello S, Irimia M, Maeso I, Pascual-Anaya J, Jimenez-Delgado S, Bertrand S, GarciaFernandez J (2008) Gene expansion and retention leads to a diverse tyrosine kinase
superfamily in amphioxus. Mol Biol Evol 25(9):1841–1854
Danchin EG, Pontarotti P (2004) Towards the reconstruction of the bilaterian ancestral pre-MHC
region. Trends Genet 20(12):587–591
CO
RR
768
769
770
771
772
773
774
775
776
777
778
779
780
781
782
783
784
785
786
787
788
789
790
791
792
793
794
795
796
797
798
799
800
801
802
803
804
805
806
807
808
809
810
811
812
813
814
815
816
817
818
819
820
Book ISBN: 978-3-642-31441-4
Page: 331/338
Two Rounds of Vertebrate Whole-Genome Duplication
UN
Editor Proof
16
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 16
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 332/338
EC
TE
D
PR
OO
F
Dehal P, Boore JL (2005) Two rounds of whole genome duplication in the ancestral vertebrate.
PLoS Biol 3(10):e314
Dehal P, Satou Y, Campbell RK, Chapman J, Degnan B, De Tomoso A, Davidson B, Di Gregorio A,
Gelpke M, Goodstein DM, Harafuji N, Hastings KEM, Ho I, Hotta K, Huang W,
Kawashima T, Lemaire P, Martinez D, Meinertzhagen IA, Necula S, Nonaka M, Putnam N,
Rash S, Saiga H, Satake M, Terry A, Yamada L, Wang H-G, Awazu S, Azumi K, Boore J,
Branno M, Chin-bow S, DeSantis R, Doyle S, Francino P, Keys DN, Haga S, Hayashi H,
Hino K, Imai KS, Inaba K, Kano S, Kobayashi K, Kobayashi M, Lee B-I, Makabe KW,
Manohar C, Matassi G, Medina M, Mochizuki Y, Mount S, Morishita T, Miura S, Nakayama A,
Nishizaka S, Nomoto H, Ohta F, Oishi K, Rigoutsos I, Sano M, Sasaki A, Sasakura Y,
Shoguchi E, Shin-i T, Spagnuolo A, Stainier D, Suzuki MM, Tassy O, Takatori N, Tokuoka M,
Yagi K, Yoshizaki F, Wada S, Zhang C, Hyatt PD, Larimer F, Detter C, Doggett N, Glavina T,
Hawkins T, Richardson P, Lucas S, Kohara Y, Levine M, Satoh N, Rokhsar DS (2002) The draft
genome of Ciona intestinalis: insights into chordate and vertebrate origins. Science
298(5601):2157–2167
Delsuc F, Brinkmann H, Chourrout D, Philippe H (2006) Tunicates and not cephalochordates are
the closest living relatives of vertebrates. Nature 439(7079):965–968
Denoeud F, Henriet S, Mungpakdee S, Aury JM, Da Silva C, Brinkmann H, Mikhaleva J, Olsen LC,
Jubin C, Cañestro C, Bouquet JM, Danks G, Poulain J, Campsteijn C, Adamski M, Cross I,
Yadetie F, Muffato M, Louis A, Butcher S, Tsagkogeorga G, Konrad A, Singh S, Jensen MF,
Cong EH, Eikeseth-Otteraa H, Noel B, Anthouard V, Porcel BM, Kachouri-Lafond R,
Nishino A, Ugolini M, Chourrout P, Nishida H, Aasland R, Huzurbazar S, Westhof E,
Delsuc F, Lehrach H, Reinhardt R, Weissenbach J, Roy SW, Artiguenave F, Postlethwait JH,
Manak JR, Thompson EM, Jaillon O, Du Pasquier L, Boudinot P, Liberles DA, Volff JN,
Philippe H, Lenhard B, Roest Crollius H, Wincker P, Chourrout D (2010) Plasticity of animal
genome architecture unmasked by rapid evolution of a pelagic tunicate. Science
330(6009):1381–1385
Donoghue PC, Graham A, Kelsh RN (2008) The origin and evolution of the neural crest.
BioEssays 30(6):530–541
Durand D (2003) Vertebrate evolution: doubling and shuffling with a full deck. Trends Genet
19(1):2–5
Ebner B, Panopoulou G, Vinogradov SN, Kiger L, Marden MC, Burmester T, Hankeln T (2010)
The globin gene family of the cephalochordate amphioxus: implications for chordate globin
evolution. BMC Evol Biol 10:370
Edvardsen RB, Seo HC, Jensen MF, Mialon A, Mikhaleva J, Bjordal M, Cartry J, Reinhardt R,
Weissenbach J, Wincker P, Chourrout D (2005) Remodelling of the homeobox gene
complement in the tunicate Oikopleura dioica. Curr Biol 15(1):R12–R13
Escriva H, Bertrand S, Germain P, Robinson-Rechavi M, Umbhauer M, Cartry J, Duffraisse M,
Holland L, Gronemeyer H, Laudet V (2006) Neofunctionalization in vertebrates: the example
of retinoic acid receptors. PLoS Genet 2(7):e102
Ferrier DE, Dewar K, Cook A, Chang JL, Hill-Force A, Amemiya C (2005) The chordate
ParaHox cluster. Curr Biol 15(20):R820–R822
Ferrier DE, Minguillon C, Holland PW, Garcia-Fernandez J (2000) The amphioxus Hox cluster:
deuterostome posterior flexibility and Hox14. Evol Dev 2(5):284–293
Ferrier DEK, Minguillón C, Cebrián C, Garcia-Fernàndez J (2001) Amphioxus Evx genes:
implications for the evolution of the midbrain–hindbrain boundary and the chordate tailbud.
Dev Biol 237(2):270–281
Ferris SD, Portnoy SL, Whitt GS (1979) The roles of speciation and divergence time in the loss of
duplicate gene expression. Theor Popul Biol 15(1):114–139
Feschotte C (2008) Transposable elements and the evolution of regulatory networks. Nat Rev
Genet 9(5):397–405
Force A, Amores A, Postlethwait JH (2002) Hox cluster organization in the jawless vertebrate
Petromyzon marinus. J Exp Zool 294(1):30–46
CO
RR
821
822
823
824
825
826
827
828
829
830
831
832
833
834
835
836
837
838
839
840
841
842
843
844
845
846
847
848
849
850
851
852
853
854
855
856
857
858
859
860
861
862
863
864
865
866
867
868
869
870
871
872
873
C. Cañestro
UN
Editor Proof
332
Layout: T1 Standard SC
Chapter No.: 16
333
EC
TE
D
PR
OO
F
Force A, Lynch M, Pickett FB, Amores A, Yan Y-L, Postlethwait J (1999) Preservation of
duplicate genes by complementary, degenerative mutations. Genetics 151(4):1531–1545
Fredriksson R, Lagerstrom MC, Lundin LG, Schioth HB (2003) The G-protein-coupled receptors
in the human genome form five main families. Phylogenetic analysis, paralogon groups, and
fingerprints. Mol Pharmacol 63(6):1256–1272
Fried C, Prohaska SJ, Stadler PF (2003) Independent Hox-cluster duplications in lampreys. J Exp
Zoolog Part B Mol Dev Evol 299(1):18–25
Friedman R, Hughes AL (2001) Pattern and timing of gene duplication in animal genomes.
Genome Res 11(11):1842–1847
Furlong R, Holland P (2002) Were vertebrates octoploid? Philos T Roy Soc B 357:531–544
Furlong RF, Younger R, Kasahara M, Reinhardt R, Thorndyke M, Holland PW (2007)
A degenerate ParaHox gene cluster in a degenerate vertebrate. Mol Biol Evol 24(12):
2681–2686
Gans C, Northcutt RG (1983) Neural crest and the origin of vertebrates: a new head. Science
220:268–274
Garcia-Fernandez J (2005) The genesis and evolution of homeobox gene clusters. Nat Rev Genet
6(12):881–892
Garcia-Fernàndez J, Ferrier DE, Minguillón C, Cebrián C (2001) The amphioxus genome in evodevo: archetype or ‘‘cul de sac’’. Int J Dev Biol 45(S1):S137–S138
Garcia-Fernàndez J, Holland PW (1994) Archetypal organization of the amphioxus Hox gene
cluster. Nature 370(6490):563–566
Gaut BS, Doebley JF (1997) DNA sequence evidence for the segmental allotetraploid origin of
maize. Proc Natl Acad Sci U S A 94(13):6809–6814
Gitelman I (2007) Evolution of the vertebrate twist family and synfunctionalization: a mechanism
for differential gene loss through merging of expression domains. Mol Biol Evol 24(9):
1912–1925
Gout JF, Duret L, Kahn D (2009) Differential retention of metabolic genes following wholegenome duplication. Mol Biol Evol 26(5):1067–1072
Gu X, Wang Y, J G (2002) Age distribution of human gene families shows significant roles of
both large- and small-scale duplications in vertebrate evolution. Nat Genet 31:205–209
Heimberg AM, Sempere LF, Moy VN, Donoghue PC, Peterson KJ (2008) microRNAs and the
advent of vertebrate morphological complexity. Proc Natl Acad Sci U S A 105(8):2946–2950
Hellsten U, Khokha MK, Grammer TC, Harland RM, Richardson P, Rokhsar DS (2007)
Accelerated gene evolution and subfunctionalization in the pseudotetraploid frog Xenopus
laevis. BMC Biol 5:31
Herpin A, Braasch I, Kraeussling M, Schmidt C, Thoma EC, Nakamura S, Tanaka M, Schartl M
(2010) Transcriptional rewiring of the sex determining dmrt1 gene duplicate by transposable
elements. PLoS Genet 6(2):e1000844
Hoekstra HE, Coyne JA (2007) The locus of evolution: evo devo and the genetics of adaptation.
Evolution 61(5):995–1016
Hoffmann FG, Opazo JC, Storz JF (2012) Whole-genome duplications spurred the functional
diversification of the globin gene superfamily in vertebrates. Mol Biol Evol 29:303-312
Holland LZ (2007) Developmental biology: a chordate with a difference. Nature 447(7141):153–
155
Holland LZ (2009) Chordate roots of the vertebrate nervous system: expanding the molecular
toolkit. Nat Rev Neurosci 10(10):736–746
Holland LZ, Albalat R, Azumi K, Benito-Gutierrez E, Blow MJ, Bronner-Fraser M, Brunet F,
Butts T, Candiani S, Dishaw LJ, Ferrier DE, Garcia-Fernandez J, Gibson-Brown JJ, Gissi C,
Godzik A, Hallbook F, Hirose D, Hosomichi K, Ikuta T, Inoko H, Kasahara M, Kasamatsu J,
Kawashima T, Kimura A, Kobayashi M, Kozmik Z, Kubokawa K, Laudet V, Litman GW,
McHardy AC, Meulemans D, Nonaka M, Olinski RP, Pancer Z, Pennacchio LA, Pestarino M,
Rast JP, Rigoutsos I, Robinson-Rechavi M, Roch G, Saiga H, Sasakura Y, Satake M, Satou Y,
Schubert M, Sherwood N, Shiina T, Takatori N, Tello J, Vopalensky P, Wada S, Xu A, Ye Y,
Yoshida K, Yoshizaki F, Yu JK, Zhang Q, Zmasek CM, de Jong PJ, Osoegawa K, Putnam
CO
RR
874
875
876
877
878
879
880
881
882
883
884
885
886
887
888
889
890
891
892
893
894
895
896
897
898
899
900
901
902
903
904
905
906
907
908
909
910
911
912
913
914
915
916
917
918
919
920
921
922
923
924
925
926
927
Book ISBN: 978-3-642-31441-4
Page: 333/338
Two Rounds of Vertebrate Whole-Genome Duplication
UN
Editor Proof
16
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 16
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 334/338
EC
TE
D
PR
OO
F
NH, Rokhsar DS, Satoh N, Holland PW (2008) The amphioxus genome illuminates vertebrate
origins and cephalochordate biology. Genome Res 18(7):1100–1111
Holland LZ, Short S (2008) Gene duplication, co-option and recruitment during the origin of the
vertebrate brain from the invertebrate chordate brain. Brain Behav Evol 72(2):91–105
Holland PWH, Garcia-Fernàndez J, Williams NA, Sidow A (1994) Gene duplications and the
origins of vertebrate development. Development (Suppl.):125–133
Holland PWH, Koschorz B, Holland LZ, Herrmann BG (1995) Conservation of Brachyury
(T) genes in amphioxus and vertebrates: developmental and evolutionary implications.
Development 121:4283–4291
Hufton AL, Groth D, Vingron M, Lehrach H, Poustka AJ, Panopoulou G (2008) Early vertebrate
whole genome duplications were predated by a period of intense genome rearrangement.
Genome Res 18(10):1582–1591
Hughes AL (1994) The evolution of functionally novel proteins after gene duplication. Proc Roy
Soc Lond B 256:119–124
Hughes AL (1999) Phylogenies of developmentally important proteins do not support the
hypothesis of two rounds of genome duplication early in vertebrate history. J Mol Evol
48(5):565–576
Hughes AL, Friedman R (2003) 2R or not 2R: testing hypotheses of genome duplication in early
vertebrates. J Struct Funct Genomics 3(1–4):85–93
Huminiecki L, Heldin CH (2010) 2R and remodeling of vertebrate signal transduction engine.
BMC Biol 8:146
Imai KS, Stolfi A, Levine M, Satou Y (2009) Gene regulatory networks underlying the
compartmentalization of the Ciona central nervous system. Development 136(2):285–293
Irimia M, Denuc A, Burguera D, Somorjai I, Martin-Duran JM, Genikhovich G, JimenezDelgado S, Technau U, Roy SW, Marfany G, Garcia-Fernandez J (2011) Stepwise assembly
of the Nova-regulated alternative splicing network in the vertebrate brain. Proc Natl Acad Sci
U S A 108(13):5319–5324
Irvine SQ, Carr JL, Bailey WJ, Kawasaki K, Shimizu N, Amemiya CT, Ruddle FH (2002)
Genomic analysis of Hox clusters in the sea lamprey Petromyzon marinus. J Exp Zool
294:47–62
Jeffery WR (2007) Chordate ancestry of the neural crest: new insights from ascidians. Semin Cell
Dev Biol 18:481-491
Jeffery WR, Chiba T, Krajka FR, Deyts C, Satoh N, Joly JS (2008) Trunk lateral cells are neural
crest-like cells in the ascidian Ciona intestinalis: insights into the ancestry and evolution of
the neural crest. Dev Biol 324(1):152–160
Jeffery WR, Strickler AG, Yamamoto Y (2004) Migratory neural crest-like cells form body
pigmentation in a urochordate embryo. Nature 431(7009):696–699
Jimenez-Delgado S, Pascual-Anaya J, Garcia-Fernandez J (2009) Implications of duplicated cisregulatory elements in the evolution of metazoans: the DDI model or how simplicity begets
novelty. Brief Funct Genomic Proteomic 8(4):266–275
Kappen C, Schughart K, Ruddle FH (1989) Two steps in the evolution of Antennapedia-class
vertebrate homeobox genes. Proc Natl Acad Sci U S A 86(14):5459–5463
Karabinos A, Bhattacharya D (2000) Molecular evolution of calmodulin and calmodulin-like
genes in the cephalochordate Branchiostoma. J Mol Evol 51(2):141–148
Kasahara M (2007) The 2R hypothesis: an update. Curr Opin Immunol 19(5):547–552
Kassahn KS, Dang VT, Wilkins SJ, Perkins AC, Ragan MA (2009) Evolution of gene function
and regulatory control after whole-genome duplication: comparative analyses in vertebrates.
Genome Res 19(8):1404–1418
Katsanis N, Fitzgibbon J, Fisher EMC (1996) Paralogy mapping: identification of a region in the
human MHC triplicated onto human chromosomes 1 and 9 allows the prediction and isolation
of novel PBX and NOTCH loci. Genomics 35:101–108
Kohn M, Hogel J, Vogel W, Minich P, Kehrer-Sawatzki H, Graves JA, Hameister H (2006)
Reconstruction of a 450-My-old ancestral vertebrate protokaryotype. Trends Genet
22(4):203–210
CO
RR
928
929
930
931
932
933
934
935
936
937
938
939
940
941
942
943
944
945
946
947
948
949
950
951
952
953
954
955
956
957
958
959
960
961
962
963
964
965
966
967
968
969
970
971
972
973
974
975
976
977
978
979
980
981
C. Cañestro
UN
Editor Proof
334
Layout: T1 Standard SC
Chapter No.: 16
335
EC
TE
D
PR
OO
F
Kuraku S (2008) Insights into cyclostome phylogenomics: pre-2R or post-2R. Zoolog Sci
25(10):960–968
Kuraku S (2010) Palaeophylogenomics of the vertebrate ancestor—impact of hidden paralogy on
hagfish and lamprey gene phylogeny. Integr Comp Biol 50(1):124–129
Kuraku S (2011) Hox gene clusters of early vertebrates: do they serve as reliable markers for
genome evolution? Genomics Proteomics Bioinformatics 9(3):97–103
Kuraku S, Meyer A, Kuratani S (2009) Timing of genome duplications relative to the origin of
the vertebrates: did cyclostomes diverge before or after? Mol Biol Evol 26(1):47–59
Kuratani S (2009) Insights into neural crest migration and differentiation from experimental
embryology. Development 136(10):1585–1589
Kuratani S, Ota KG (2008) Hagfish (cyclostomata, vertebrata): searching for the ancestral
developmental plan of vertebrates. BioEssays 30(2):167–172
Laisney JA, Braasch I, Walter RB, Meierjohann S, Schartl M (2010) Lineage-specific
co-evolution of the Egf receptor/ligand signaling system. BMC Evol Biol 10:27
Larhammar D, Lundin L, Hallbook F (2002) The human Hox-bearing chromosome regions did
arise by block or chromosome (or even genome) duplications. Genome Res 12(12):
1910–1920
Lee CT, Risom T, Strauss WM (2007) Evolutionary conservation of microRNA regulatory
circuits: an examination of microRNA gene complexity and conserved microRNA-target
interactions through metazoan phylogeny. DNA Cell Biol 26(4):209–218
Leveugle M, Prat K, Perrier N, Birnbaum D, Coulier F (2003) ParaDB: a tool for paralogy
mapping in vertebrate genomes. Nucleic Acids Res 31(1):63–67
Louis A, Roest Crollius H, Robinson-Rechavi M (2012) How much does the amphioxus genome
represent the ancestor of chordates? Brief Funct Genomics 11(2):89–95
Lowe CB, Bejerano G, Haussler D (2007) Thousands of human mobile element fragments
undergo strong purifying selection near developmental genes. Proc Natl Acad Sci U S A
104(19):8005–8010
Lundin LG (1979) Evolutionary conservation of large chromosomal segments reflected in
mammalian gene maps. Clin Genet 16(2):72–81
Lundin LG (1993) Evolution of the vertebrate genome as relected in paralogous chromosomal
regions in man and the house mouse. Genomics 16:1–19
Lundin LG, Larhammar D, Hallbook F (2003) Numerous groups of chromosomal regional
paralogies strongly indicate two genome doublings at the root of the vertebrates. J Struct
Funct Genomics 3(1–4):53–63
Lynch M, Conery J (2000) The evolutionary fate and consequences of gene duplication. Science
290(5494):1151–1155
Lynch M, Force A (2000) The origin of interspecific genomic incompatibility via gene
duplication. Am Nat 156:590–605
Lynch M, O’Hely M, Walsh B, Force A (2001) The probability of preservation of a newly arisen
gene duplicate. Genetics 159(4):1789–1804
Lynch VJ, Leclerc RD, May G, Wagner GP (2011) Transposon-mediated rewiring of gene
regulatory networks contributed to the evolution of pregnancy in mammals. Nat Genet
43:1154-1159
Lynch VJ, Wagner GP (2009) Multiple chromosomal rearrangements structured the ancestral
vertebrate Hox-bearing protochromosomes. PLoS Genet 5(1):e1000349
Makino T, McLysaght A (2010) Ohnologs in the human genome are dosage balanced and
frequently associated with disease. Proc Natl Acad Sci U S A 107(20):9270–9274
Martinez-Morales JR, Henrich T, Ramialison M, Wittbrodt J (2007) New genes in the evolution
of the neural crest differentiation program. Genome Biol 8(3):R36
Matzke MA, Matzke AJ (1998) Polyploidy and transposons. Trends Ecol Evol 13(6):241
Matzke MA, Mette MF, Matzke AJ (2000) Transgene silencing by the host genome defense:
implications for the evolution of epigenetic control mechanisms in plants and vertebrates.
Plant Mol Biol 43(2–3):401–415
CO
RR
982
983
984
985
986
987
988
989
990
991
992
993
994
995
996
997
998
999
1000
1001
1002
1003
1004
1005
1006
1007
1008
1009
1010
1011
1012
1013
1014
1015
1016
1017
1018
1019
1020
1021
1022
1023
1024
1025
1026
1027
1028
1029
1030
1031
1032
1033
1034
Book ISBN: 978-3-642-31441-4
Page: 335/338
Two Rounds of Vertebrate Whole-Genome Duplication
UN
Editor Proof
16
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 16
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 336/338
EC
TE
D
PR
OO
F
Mazet F, Hutt JA, Milloz J, Millard J, Graham A, Shimeld SM (2005) Molecular evidence from
Ciona intestinalis for the evolutionary origin of vertebrate sensory placodes. Dev Biol
282(2):494–508
McClintock JM, Carlson R, Mann DM, Prince VE (2001) Consequences of Hox gene duplication
in the vertebrates: an investigation of the zebrafish Hox paralogue group 1 genes.
Development 128(13):2471–2484
McLysaght A, Hokamp K, Wolfe KH (2002) Extensive genomic duplication during early
chordate evolution. Nat Genet 31:200–204
Meulemans D, Bronner-Fraser M (2004) Gene-regulatory interactions in neural crest evolution
and development. Dev Cell 7(3):291–299
Meulemans D, Bronner-Fraser M (2005) Central role of gene cooption in neural crest evolution.
J Exp Zoolog B Mol Dev Evol 304(4):298–303
Meulemans D, Bronner-Fraser M (2007) Insights from amphioxus into the evolution of vertebrate
cartilage. PLoS One 2(8):e787
Minguillon C, Ferrier DE, Cebrian C, Garcia-Fernandez J (2002) Gene duplications in the
prototypical cephalochordate amphioxus. Gene 287(1–2):121–128
Minguillon C, Jimenez-Delgado S, Panopoulou G, Garcia-Fernandez J (2003) The amphioxus
Hairy family: differential fate after duplication. Development 130(24):5903–5914
Muffato M, Louis A, Poisnel CE, Roest Crollius H (2010) Genomicus: a database and a browser
to study gene synteny in modern and ancestral genomes. Bioinformatics 26(8):1119–1121
Nadeau JH, Sankoff D (1997) Comparable rates of gene loss and functional divergence after
genome duplications early in vertebrate evolution. Genetics 147(3):1259–1266
Nakatani Y, Takeda H, Kohara Y, Morishita S (2007) Reconstruction of the vertebrate ancestral
genome reveals dynamic genome reorganization in early vertebrates. Genome Res
17(9):1254–1265
Naruse K, Tanaka M, Mita K, Shima A, Postlethwait J, Mitani H (2004) A medaka gene map: the
trace of ancestral vertebrate proto-chromosomes revealed by comparative gene mapping.
Genome Res 14(5):820–828
Nelson JS (1994) Fishes of the world, 3rd edn. Wiley-Interscience, New York
Nikitina N, Sauka-Spengler T, Bronner-Fraser M (2009) Chapter 1. Gene regulatory networks in
neural crest development and evolution. Curr Top Dev Biol 86:1–14
Northcutt RG, Gans C (1983) The genesis of neural crest and epidermal placodes: a
reinterpretation of vertebrate origins. Quart Rev Biol 58:1–28
Nowak MA, Boerlijst MC, Cooke J, Smith JM (1997) Evolution of genetic redundancy. Nature
388(6638):167–171
Oda H, Wada H, Tagawa K, Akiyama-Oda Y, Satoh N, Humphreys T, Zhang S, Tsukita S (2002)
A novel amphioxus cadherin that localizes to epithelial adherens junctions has an unusual
domain organization with implications for chordate phylogeny. Evol Dev 4(6):426–434
Ohno S (1970) Evolution by gene duplication. Springer, New York
Ohno S, Wolf U, Atkins NB (1968) Evolution from fish to mammals by gene duplication.
Hereditas 59(1):169–187
Ota KG, Kuratani S (2007) Cyclostome embryology and early evolutionary history of vertebrates.
Integr Comp Biol 47(3):329–337
Otto SP (2007) The evolutionary consequences of polyploidy. Cell 131(3):452–462
Panopoulou G, Hennig S, Groth D, Krause A, Poustka AJ, Herwig R, Vingron M, Lehrach H
(2003) New evidence for genome-wide duplications at the origin of vertebrates using an
amphioxus gene set and completed animal genomes. Genome Res 13(6A):1056–1066
Paps J, Holland PW, Shimeld SM (2012) A genome-wide view of transcription factor gene
diversity in chordate evolution: less gene loss in amphioxus? Brief Funct Genomics
11(2):177–186
Parisod C, Alix K, Just J, Petit M, Sarilar V, Mhiri C, Ainouche M, Chalhoub B, Grandbastien
MA (2010) Impact of transposable elements on the organization and function of allopolyploid
genomes. New Phytol 186(1):37–45
CO
RR
1035
1036
1037
1038
1039
1040
1041
1042
1043
1044
1045
1046
1047
1048
1049
1050
1051
1052
1053
1054
1055
1056
1057
1058
1059
1060
1061
1062
1063
1064
1065
1066
1067
1068
1069
1070
1071
1072
1073
1074
1075
1076
1077
1078
1079
1080
1081
1082
1083
1084
1085
1086
1087
C. Cañestro
UN
Editor Proof
336
Layout: T1 Standard SC
Chapter No.: 16
337
EC
TE
D
PR
OO
F
Paterson AH, Chapman BA, Kissinger JC, Bowers JE, Feltus FA, Estill JC (2006) Many gene and
domain families have convergent fates following independent whole-genome duplication
events in Arabidopsis, Oryza, Saccharomyces and Tetraodon. Trends Genet 22(11):597–602
Pebusque MJ, Coulier F, Birnbaum D, Pontarotti P (1998) Ancient large-scale genome
duplications: phylogenetic and linkage analyses shed light on chordate genome evolution.
Mol Biol Evol 15(9):1145–1159
Pendleton JW, Nagai BK, Murtha MT, Ruddle FH (1993) Expansion of the Hox gene family and
the evolution of chordates. Proc Natl Acad Sci U S A 90:6300–6304
Polak P, Domany E (2006) Alu elements contain many binding sites for transcription factors and
may play a role in regulation of developmental processes. BMC Genomics 7:133
Popovici C, Leveugle M, Birnbaum D, Coulier F (2001) Coparalogy: physical and functional
clusterings in the human genome. Biochem Biophys Res Commun 288(2):362–370
Postlethwait J, Amores A, Cresko W, Singer A, Yan YL (2004) Subfunction partitioning, the
teleost radiation and the annotation of the human genome. Trends Genet 20(10):481–490
Postlethwait JH (2007) The zebrafish genome in context: ohnologs gone missing. J Exp Zoolog B
Mol Dev Evol 308(5):563–577
Putnam NH, Butts T, Ferrier DE, Furlong RF, Hellsten U, Kawashima T, Robinson-Rechavi M,
Shoguchi E, Terry A, Yu JK, Benito-Gutierrez EL, Dubchak I, Garcia-Fernandez J, GibsonBrown JJ, Grigoriev IV, Horton AC, de Jong PJ, Jurka J, Kapitonov VV, Kohara Y, Kuroki Y,
Lindquist E, Lucas S, Osoegawa K, Pennacchio LA, Salamov AA, Satou Y, Sauka-Spengler
T, Schmutz J, Shin IT, Toyoda A, Bronner-Fraser M, Fujiyama A, Holland LZ, Holland PW,
Satoh N, Rokhsar DS (2008) The amphioxus genome and the evolution of the chordate
karyotype. Nature 453(7198):1064–1071
Ravi V, Lam K, Tay BH, Tay A, Brenner S, Venkatesh B (2009) Elephant shark (Callorhinchus
milii) provides insights into the evolution of Hox gene clusters in gnathostomes. Proc
Natl Acad Sci U S A 106(38):16327–16332
Robinson-Rechavi M, Boussau B, Laudet V (2004) Phylogenetic dating and characterization of
gene duplications in vertebrates: the cartilaginous fish reference. Mol Biol Evol 21(3):
580–586
Roux J, Robinson-Rechavi M (2008) Developmental constraints on vertebrate genome evolution.
PLoS Genet 4(12):e1000311
Ruvinsky I, Silver LM (1997) Newly identified paralogous groups on mouse chromosomes 5 and
11 reveal the age of a T-box cluster duplication. Genomics 40(2):262–266
SanMiguel P, Gaut BS, Tikhonov A, Nakajima Y, Bennetzen JL (1998) The paleontology of
intergene retrotransposons of maize. Nat Genet 20(1):43–45
SanMiguel P, Tikhonov A, Jin YK, Motchoulskaia N, Zakharov D, Melake-Berhan A, Springer
PS, Edwards KJ, Lee M, Avramova Z, Bennetzen JL (1996) Nested retrotransposons in the
intergenic regions of the maize genome. Science 274(5288):765–768
Sauka-Spengler T, Meulemans D, Jones M, Bronner-Fraser M (2007) Ancient evolutionary origin
of the neural crest gene regulatory network. Dev Cell 13(3):405–420
Scannell DR, Byrne KP, Gordon JL, Wong S, Wolfe KH (2006) Multiple rounds of speciation
associated with reciprocal gene loss in polyploid yeasts. Nature 440(7082):341–345
Semon M, Wolfe KH (2007) Consequences of genome duplication. Curr Opin Genet Dev
17(6):505–512
Semon M, Wolfe KH (2008) Preferential subfunctionalization of slow-evolving genes after
allopolyploidization in Xenopus laevis. Proc Natl Acad Sci U S A 105(24):8333–8338
Sempere LF, Cole CN, McPeek MA, Peterson KJ (2006) The phylogenetic distribution of
metazoan microRNAs: insights into evolutionary complexity and constraint. J Exp Zool B
Mol Dev Evol 306(6):575–588
Seo HC, Edvardsen RB, Maeland AD, Bjordal M, Jensen MF, Hansen A, Flaat M, Weissenbach
J, Lehrach H, Wincker P, Reinhardt R, Chourrout D (2004) Hox cluster disintegration with
persistent anteroposterior order of expression in Oikopleura dioica. Nature 431(7004):67–71
Sharman AC, Holland PWH (1998) Estimation of hox gene cluster number in lampreys. Int J Dev
Biol 42:617–620
CO
RR
1088
1089
1090
1091
1092
1093
1094
1095
1096
1097
1098
1099
1100
1101
1102
1103
1104
1105
1106
1107
1108
1109
1110
1111
1112
1113
1114
1115
1116
1117
1118
1119
1120
1121
1122
1123
1124
1125
1126
1127
1128
1129
1130
1131
1132
1133
1134
1135
1136
1137
1138
1139
1140
1141
Book ISBN: 978-3-642-31441-4
Page: 337/338
Two Rounds of Vertebrate Whole-Genome Duplication
UN
Editor Proof
16
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 16
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 338/338
EC
TE
D
PR
OO
F
Shemer G, Podbilewicz B (2000) Fusomorphogenesis: cell fusion in organ formation. Dev Dyn
218(1):30–51
Shimeld SM, Holland PW (2000) Vertebrate innovations. Proc Natl Acad Sci U S A 97(9):
4449–4452
Sidow A (1996) Gen(om)e duplications in the evolution of early vertebrates. Curr Opin Genet
Dev 6:715–722
Siegel N, Hoegg S, Salzburger W, Braasch I, Meyer A (2007) Comparative genomics of ParaHox
clusters of teleost fishes: gene cluster breakup and the retention of gene sets following whole
genome duplications. BMC Genomics 8:312
Skrabanek L, Wolfe KH (1998) Eukaryote genome duplication—where’s the evidence? Curr
Opin Genet Dev 8:694–700
Small KS, Brudno M, Hill MM, Sidow A (2007) A haplome alignment and reference sequence of
the highly polymorphic Ciona savignyi genome. Genome Biol 8(3):R41
Soltis DE, Albert VA, Leebens-Mack J, Bell CD, Paterson AH, Zheng C, Sankoff D, Depamphilis
CW, Wall PK, Soltis PS (2009) Polyploidy and angiosperm diversification. Am J Bot
96(1):336–348
Soltis DE, Soltis PS (1999) Polyploidy: recurrent formation and genome evolution. Trends Ecol
Evol 14(9):348–352
Somorjai I, Bertrand S, Camasses A, Haguenauer A, Escriva H (2008) Evidence for stasis and not
genetic piracy in developmental expression patterns of Branchiostoma lanceolatum and
Branchiostoma floridae, two amphioxus species that have evolved independently over the
course of 200 Myr. Dev Genes Evol 218(11–12):703–713
Spring J (1997) Vertebrate evolution by interspecific hybridization—are we polyploid? Fed Eur
Biol Soc Lett 400:2–8
Stadler PF, Fried C, Prohaska SJ, Bailey WJ, Misof BY, Ruddle FH, Wagner GP (2004) Evidence
for independent Hox gene duplications in the hagfish lineage: a PCR-based gene inventory of
Eptatretus stoutii. Mol Phylogenet Evol 32(3):686–694
Storchova Z, Pellman D (2004) From polyploidy to aneuploidy, genome instability and cancer.
Nat Rev Mol Cell Biol 5(1):45–54
Takio Y, Pasqualetti M, Kuraku S, Hirano S, Rijli FM, Kuratani S (2004) Evolutionary biology:
lamprey Hox genes and the evolution of jaws. Nature 429(6989):1–262
Taylor JS, Raes J (2004) Duplication and divergence: the evolution of new genes and old ideas.
Annu Rev Genet 38:615–643
Taylor JS, van de Peer Y, Meyer M (2001) Genome duplication, divergent resolution and
speciation. Trends Genet 17:299–301
Thornburg BG, Gotea V, Makalowski W (2006) Transposable elements as a significant source of
transcription regulating signals. Gene 365:104–110
van de Lagemaat LN, Landry JR, Mager DL, Medstrand P (2003) Transposable elements in
mammals promote regulatory variation and diversification of genes with specialized
functions. Trends Genet 19(10):530–536
Van de Peer Y, Maere S, Meyer A (2009) The evolutionary significance of ancient genome
duplications. Nat Rev Genet 10(10):725–732
Venkatesh B, Kirkness EF, Loh YH, Halpern AL, Lee AP, Johnson J, Dandona N, Viswanathan
LD, Tay A, Venter JC, Strausberg RL, Brenner S (2007) Survey sequencing and comparative
analysis of the elephant shark (Callorhinchus milii) genome. PLoS Biol 5(4):e101
Venter JC, Adams MD, Myers EW, Li PW, Mural RJ, Sutton GG, Smith HO, Yandell M, Evans
CA, Holt RA et al (2001) The sequence of the human genome. Science 291(5507):1304–1351
Vienne A, Shiina T, Abi-Rached L, Danchin E, Vitiello V, Cartault F, Inoko H, Pontarotti P
(2003) Evolution of the proto-MHC ancestral region: more evidence for the plesiomorphic
organisation of human chromosome 9q34 region. Immunogenetics 55(7):429–436
Wada H, Okuyama M, Satoh N, Zhang S (2006) Molecular evolution of fibrillar collagen in
chordates, with implications for the evolution of vertebrate skeletons and chordate phylogeny.
Evol Dev 8(4):370–377
CO
RR
1142
1143
1144
1145
1146
1147
1148
1149
1150
1151
1152
1153
1154
1155
1156
1157
1158
1159
1160
1161
1162
1163
1164
1165
1166
1167
1168
1169
1170
1171
1172
1173
1174
1175
1176
1177
1178
1179
1180
1181
1182
1183
1184
1185
1186
1187
1188
1189
1190
1191
1192
1193
1194
C. Cañestro
UN
Editor Proof
338
Layout: T1 Standard SC
Chapter No.: 16
339
EC
TE
D
PR
OO
F
Werth CR, Windham MD (1991) A model for divergent, allopatric speciation of polypoid
pteridophytes resulting from silencing of duplicate-gene expression. Am Nat 137:515–526
Weston JA (1970) The migration and differentiation of neural crest cells. Adv Morphog 8:41–114
Wolfe K (2000) Robustness—it’s not where you think it is. Nat Genet 25(1):3–4
Wolfe KH (2001) Yesterday’s polyploids and the mystery of diploidization. Nat Rev Genet
2(5):333–341
Wolfe KH, Shields DC (1997) Molecular evidence for an ancient duplication of the entire yeast
genome. Nature 387(6634):708–713
Wotton KR, Shimeld SM (2006) Comparative genomics of vertebrate Fox cluster loci. BMC
Genomics 7:271
Yu JK, Meulemans D, McKeown SJ, Bronner-Fraser M (2008) Insights from the amphioxus
genome on the origin of vertebrate neural crest. Genome Res 18(7):1127–1132
CO
RR
1195
1196
1197
1198
1199
1200
1201
1202
1203
1204
1205
1206
Book ISBN: 978-3-642-31441-4
Page: 339/338
Two Rounds of Vertebrate Whole-Genome Duplication
UN
Editor Proof
16
Book ID: 272454_1_En
Date: 16-8-2012
Metadata of the chapter that will be visualized in
SpringerLink
Book Title
Polyploidy and Genome Evolution
Series Title
Chapter Title
Polyploidy in Fish and the Teleost Genome Duplication
Copyright Year
2012
Copyright HolderName
Springer-Verlag Berlin Heidelberg
Corresponding Author
Family Name
Postlethwait
Particle
Given Name
John H.
Suffix
Author
Division
Institute of Neuroscience
Organization
University of Oregon
Address
97403, Eugene, OR, USA
Email
jpostle@uoneuro.uoregon.edu
Family Name
Braasch
Particle
Given Name
Ingo
Suffix
Abstract
Division
Institute of Neuroscience
Organization
University of Oregon
Address
97403, Eugene, OR, USA
Email
ibraasch@uoneuro.uoregon.edu
Multiple rounds of whole-genome duplications (WGDs) punctuated the evolution of rayfin fish, a speciesrich group comprising about half of all vertebrates. Rayfin fish, along with lobefin vertebrates including
humans, derive from early vertebrate ancestors that evolved through two rounds of polyploidization (the first
and second rounds of vertebrate genome duplication, VGD1 and VGD2) at the dawn of the vertebrate lineage.
Furthermore, teleost fish underwent an additional round of polyploidization in their stem lineage, the teleost
genome duplication (TGD). Additional WGD events occurred independently in numerous species and higher
level taxa of teleosts and other rayfin fish, for example in salmonids, carp, and sturgeon, so that some fish
lineages experienced at least four rounds of WGD since the origin of vertebrates. This chapter provides an
overview of these polyploidization events in the fish lineage and focuses on the impact these genome
duplications (GD) had on genome evolution in selected fish taxa. We then review evidence for the TGD and
discuss its consequences for the evolution of gene content, order, and functions in the teleost lineage. We
argue that, although evidence remains sparse, the TGD may have had a profound influence on the evolutionary
success and the biodiversity of teleosts. Importantly, an in-depth understanding of the causes and
consequences of the TGD and other teleost GD events will help to inform us about the evolution of our own
paleopolyploid genome.
1
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 341/383
Chapter 17
4
Ingo Braasch and John H. Postlethwait
11
12
13
14
15
16
17
18
19
20
21
22
23
24
PR
OO
D
9
10
TE
8
EC
7
Abstract Multiple rounds of whole-genome duplications (WGDs) punctuated the
evolution of rayfin fish, a species-rich group comprising about half of all vertebrates. Rayfin fish, along with lobefin vertebrates including humans, derive from
early vertebrate ancestors that evolved through two rounds of polyploidization (the
first and second rounds of vertebrate genome duplication, VGD1 and VGD2) at the
dawn of the vertebrate lineage. Furthermore, teleost fish underwent an additional
round of polyploidization in their stem lineage, the teleost genome duplication
(TGD). Additional WGD events occurred independently in numerous species and
higher level taxa of teleosts and other rayfin fish, for example in salmonids, carp,
and sturgeon, so that some fish lineages experienced at least four rounds of WGD
since the origin of vertebrates. This chapter provides an overview of these polyploidization events in the fish lineage and focuses on the impact these genome
duplications (GD) had on genome evolution in selected fish taxa. We then review
evidence for the TGD and discuss its consequences for the evolution of gene
content, order, and functions in the teleost lineage. We argue that, although evidence remains sparse, the TGD may have had a profound influence on the evolutionary success and the biodiversity of teleosts. Importantly, an in-depth
understanding of the causes and consequences of the TGD and other teleost GD
events will help to inform us about the evolution of our own paleopolyploid
genome.
CO
RR
5
6
F
3
Polyploidy in Fish and the Teleost
Genome Duplication
2
I. Braasch J. H. Postlethwait (&)
Institute of Neuroscience, University of Oregon, Eugene, OR 97403, USA
e-mail: jpostle@uoneuro.uoregon.edu
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: 17
I. Braasch
e-mail: ibraasch@uoneuro.uoregon.edu
P. S. Soltis and D. E. Soltis (eds.), Polyploidy and Genome Evolution,
DOI: 10.1007/978-3-642-31442-1_17, Springer-Verlag Berlin Heidelberg 2012
341
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
I. Braasch and J. H. Postlethwait
The problems of this world are only truly solved in two ways:
by extinction or duplication.
Susan Sontag
25
26
27
28
29
30
31
32
33
34
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
61
62
63
64
65
66
D
38
In common usage, ‘‘fish’’ applies to any aquatic vertebrate that possesses gills and
fins (if any appendages) (Nelson 2006). Following this phenotypic definition,
‘‘fish’’ include jawless fish such as lampreys and hagfish, cartilaginous fish, such as
sharks and rays, lobefin fish including coelacanth and lungfish, and rayfin fish such
as teleosts. ‘‘Fish’’, however, are a paraphyletic assemblage because it excludes
tetrapod vertebrates, which share a more recent common ancestor with lungfish
than lungfish share with teleost fish. Until we tend to call tetrapods (including
ourselves) lobefin fish and accept that all living vertebrates are fish, the term ‘‘fish’’
should be used with caution (see Fig. 17.1).
This chapter focuses on the monophyletic group of rayfin fish, or Actinopterygii, and, in more detail, their largest subgroup, teleost fish. The jawless,
cartilaginous, and lobefin fish, however, are important for understanding evolution
of the vertebrate genome by whole-genome duplications (WGDs) at the dawn of
the vertebrate lineage and are thus featured in Chap. 12 (this volume).
Among actinopterygian fish, the teleosts are by far the largest subgroup in terms
of extant species numbers. Nelson (2006) lists 26,891 living rayfin fish species
(453 families), of which only 51 species (5 families) are not teleosts. The teleosts,
in contrast, comprise 26,840 living species (448 families), an impressive 99.8 % of
all rayfins. Even more impressive, adding all living vertebrates to the calculation
gives around 50 % of all vertebrates being teleost fish.
Draft genome assemblies of ten rayfin species (all of them clupeocephalan
teleosts) are currently publicly available (Fig. 17.1): zebrafish (Danio rerio), threespined stickleback (Gasterosteus aculeatus), medaka (Oryzias latipes) (Kasahara
et al. 2007), fugu (Takifugu rubripes) (Aparicio et al. 2002), spotted green pufferfish (Tetraodon nigroviridis) (Jaillon et al. 2004), Atlantic cod (Gadus morhua)
(Star et al. 2011), and four species of East African cichlids, including tilapia
(Oreochromis niloticus). More species such as platyfish (Xiphophorus maculatus),
Atlantic salmon (Salmo salar), rainbow trout (Oncorhynchus mykiss), and the nonteleost spotted gar (Lepisosteus oculatus; see below) are soon to be added to this
list (Fig. 17.1), and numerous additional rayfins will be sequenced as part of the
GENOME 10K Project (Haussler et al. 2009).
TE
37
EC
36
17.1 A Brief Introduction to ‘‘Fish’’
CO
RR
35
PR
OO
F
In their great numbers and degree of anatomical diversity, the
modern ray-finned fishes may be considered the most
successful of all vertebrates.
Robert L. Carroll
UN
Editor Proof
342
Book ISBN: 978-3-642-31441-4
Page: 342/383
Layout: T1 Standard SC
Chapter No.: 17
Book ISBN: 978-3-642-31441-4
Page: 343/383
Polyploidy in Fish and the Teleost Genome Duplication
Editor Proof
17
Book ID: 272454_1_En
Date: 16-8-2012
343
Spotted green pufferfish
Tetraodon nigroviridis
Medaka
Oryzias latipes
Platyfish
Xiphophorus maculatus
Atlantic cod
Gadus morhua
SaGD
Salmonids
Zebrafish
Danio rerio
TGD
F
rayfin fish
PR
OO
East African cichlids
clupeocephala
teleosts
Three-spined stickleback
Gasterosteus aculeatus
percomorpha
Japanese pufferfish
Takifugu rubripes
Carps + Goldfish
CGD
D
Goldeye
Hiodon alosoides
TE
Bowfin
Amia calva
Spotted gar
Lepisosteus oculatus
AGDs
VGD1
700
CO
RR
VGD2
600
500
400
300
200
Bichirs
Tetrapods
Lungfish
lobefin fish
EC
Sturgeons
Coelacanth
Cartilaginous fish
Jawless fish
100
0 MYA
UN
Fig. 17.1 A phylogeny of all kinds of ‘‘fish’’. Tree topology and timing of divergence events
follows data from Setiamarga et al. (2009) and www.timetree.org. The figure shows the timing of
genome duplication (GD) events (VGD1/VGD2 the vertebrate GDs, TGD the teleost GD, AGDs
acipenserid GDs, SaGD the salmonid GD, CGD the carp GD). Whether the second vertebrate
genome duplication (VGD2) occurred before or after the divergence of jawless fish remains
controversial. MYA million years ago
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
I. Braasch and J. H. Postlethwait
71
The species richness of rayfin and teleost fish poses the question of the secret of
their evolutionary success. A recurring pattern of genome evolution in rayfin fish is
their propensity to polyploidization. In line with this idea, a positive correlation of
genome size and species richness has been reported among actinopterygians as
well as for specific teleost clades (Mank and Avise 2006a).
72
17.2 Duplications in the Crown: Polyploidy in Rayfin Fish
67
68
69
PR
OO
F
70
91
17.2.1 Twigs and Leaves: Polyploidization Events in Fish
79
80
81
82
83
84
85
86
87
88
89
92
93
94
95
96
97
98
99
100
101
102
103
TE
77
78
EC
75
76
CO
RR
74
D
90
In comparison to plants, polyploidizations are rare in the animal kingdom (Muller
1925), yet the reasons for this genomic discrepancy between fauna and flora
remain controversial (Mable 2004). In the vertebrate lineage, polyploids are most
often found among amphibians and rayfin fish (Mable et al. 2011; Otto 2007;
Comai 2005). Factors that may favor the proclivity of rayfin fish to generate
surviving polyploid lineages compared to other groups of vertebrates (e.g., high
production rate of unreduced gametes, their propensity for hybridization, genomic
flexibility, etc.) have been extensively discussed recently elsewhere, yet a clear
explanation remains elusive (Mable et al. 2011). Given that polyploid events also
appear to have occurred in some jawless, cartilaginous, and lobefin fish (see
Leggatt and Iwama 2003 and references therein), it may be worth considering that
a paucity of genome duplication (GD) is an amniote vertebrate-specific phenomenon. Several works provide comprehensive lists of known cases of polyploidy in
fish (Otto and Whitton 2000; Leggatt and Iwama 2003; Le Comber and Smith
2004; Mable et al. 2011).
Here, we give an overview of the few general trends that can be inferred from
the phylogenetic distribution of polyploidization events and will then discuss
several specific cases in more detail.
73
Despite large variations in genome size, rayfin fish have surprisingly conserved
karyotypes, with the majority of diploid genomes having 48 or 50 chromosomes
(Ohno et al. 1968; Mank and Avise 2006b); thus, chromosome counts have been
used as indicators of polyploidizations in fish. A study based on chromosome
counts of 615 rayfin species concluded that at least 7–20 polyploidization events
occurred in the evolution of extant rayfins (Mank and Avise 2006b); this estimate
is likely an underestimate because less than 3 % of all living rayfin species were
included in this study.
Polyploidization events in fish are phylogenetically restricted, i.e., are unevenly
distributed across the actinopterygian tree. Some groups, especially those occupying early diverging branches in rayfin phylogeny, and among teleosts particularly the Ostariophysi (including carps and suckers and many others), seem to be
UN
Editor Proof
344
Book ISBN: 978-3-642-31441-4
Page: 344/383
Layout: T1 Standard SC
Chapter No.: 17
110
111
112
113
114
115
116
117
118
119
120
121
122
123
124
125
126
127
128
F
109
PR
OO
107
108
D
106
particularly prone to polyploidization in their lineage, while other groups,
including more derived lineages such as the percomorphs (perch and relatives, like
pufferfish, stickleback, cichlids and many more), have few or no GDs after the
teleost genome duplication (TGD) (Leggatt and Iwama 2003; Mable et al. 2011).
Polyploidizations have occurred in many individual fish genera, species, and
populations (i.e., the ‘‘leaves’’ of the rayfin tree), as for example in multiple species
of barbs (genus Barbus, family Cyprinidae) (Chenuil et al. 1999), in the cyprinid
Squalius alburnoides species complex (Alves et al. 2001), in the pond loach
(Misgurnus anguillicaudatus) (Li et al. 2011) and other loach species (Cobitidae,
order Cypriniformes) (Ferris and Whitt 1977a), and in the stinging catfish
(Heteropneustes fossilis, order Siluriformes) (Pandian and Koteeswaran 1999). On
the other hand, piscine GD events have also occurred in the last common ancestor
of higher level taxa (some of the tree’s ‘‘twigs’’). All members are polyploid in the
three families Salmonidae (salmonids) (Allendorf and Thorgaard 1984), Callichthyidae (armored catfish) (Oliveira et al. 1992), and Catostomidae (suckers) (Ferris
1984; Uyeno and Smith 1972). Common carp and goldfish (subfamily Cyprininae)
also arise from a shared tetraploid origin (Ohno et al. 1967; David et al. 2003;
Schultz 1980; Larhammar and Risinger 1994). Among rayfins diverging before
teleosts, the Acipenseridae (sturgeons) are famous for their multiple ploidy levels,
up to at least hexadecaploid (Birstein et al. 1997; Ludwig et al. 2001). In addition,
artificial polyploids, particularly triploids, are routinely produced for multiple
commercially important fish species to induce sterility and thereby improve somatic
growth rate (Piferrer et al. 2009).
TE
105
345
EC
104
Book ISBN: 978-3-642-31441-4
Page: 345/383
Polyploidy in Fish and the Teleost Genome Duplication
17.2.2 Piscine Polyploidy and the Evolution of Genome Function:
Salmon, Carp, and Calandinos
133
17.2.2.1 Salmonids
130
131
134
135
136
137
138
139
140
CO
RR
132
Traditional studies of polyploidy in fish generally involve the analysis of karyotypes and allozymes. Only recently, with the advent of genome sequencing
techniques, we have begun to understand some of the evolutionary footprints left
by polyploidization events in fish genomes.
129
Salmonids (salmon, trout, whitefish, grayling) are among the fish of highest economic importance (Davidson et al. 2010). Salmonids have been suspected to be
polyploids since the 1940s (Svärdson 1945; Kupka 1948), but it was several
decades until it was generally accepted that all living salmonids are of autotetraploid origin due to a GD event that occurred in their stem lineage 25–100 million
years ago (Ohno et al. 1968; Allendorf and Thorgaard 1984). This WGD is
unlikely to have occurred by segmental duplications because all segments of each
UN
Editor Proof
17
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
148
149
150
151
152
153
154
155
156
157
158
159
160
161
162
163
164
165
166
167
168
169
170
171
172
173
174
175
176
177
178
179
180
181
182
183
184
185
F
146
147
PR
OO
145
D
144
TE
143
chromosome appear to be in duplicate (although of course not every gene in each
segment is still present in duplicate); if segmental duplications had been responsible, then some chromosome segments would be present in three or five copies.
Salmonids still show multivalent chromosomes during meiosis as well as tetrasomic inheritance of some loci (Allendorf and Thorgaard 1984; Phillips and Rab
2001) and are thus considered pseudotetraploids, i.e., the process of diploidization
has not yet concluded in these fish (Danzmann et al. 2008).
The autotetraplodization in salmonids is sometimes referred to as ‘‘4R’’,
because it is the fourth round of WGD in this lineage since the rise of the vertebrate lineage (see later this chapter and Chap. 16, this volume). This numerating
terminology, however, imposes difficulties to distinguish independent GD events
in various lineages of rayfins as discussed above. Therefore, we suggest using the
term salmonid genome duplication (SaGD). Recent progress in salmonid genomics
(Davidson et al. 2010; Miller et al. 2011) has provided early insights into the
evolutionary dynamics of the salmonid genome after the SaGD.
In line with tetraploidization in a salmonid ancestor, the diploid genomes of
most salmonids contain 96–104 chromosome arms. The Atlantic salmon is a
special case among salmonids, because its more derived karyotype has been
secondarily reduced to 72–74 chromosome arms by chromosome fusions (Phillips
and Rab 2001; Phillips et al. 2009). Conserved synteny analysis comparing genetic
linkage maps of Atlantic salmon and rainbow trout (Danzmann et al. 2008; Lien
et al. 2011) and analyzing the genomic distribution of conserved non-coding
elements (CNEs) (Moghadam et al. 2009) show first that gene order in salmonids
generally reflects the inferred ancestral rayfin karyotypes (Nakatani et al. 2007;
Kasahara et al. 2007), and second that two salmonid homeologous chromosomes
share conserved synteny with one teleost outgroup chromosome (e.g., medaka,
zebrafish, and stickleback), as would be expected from an additional round of GD.
The duplication of hox gene clusters, generally a good indicator for GDs in
animals (see Chap. 16, this volume), has been studied in detail in salmonids.
Salmonid genomes contain at least 13 hox clusters, nearly twice as many as most
diploid teleost genomes (Moghadam et al. 2005a, b; Mungpakdee et al. 2008a).
Given that gene loss (nonfunctionalization, or pseudogenization) is the most frequent fate of gene duplicates (Lynch and Conery 2000), it is surprising that the
salmonid hox cluster repertoire, as judged by collapsing duplicated clusters, shows
no loss of hox paralogy groups between the ancestral teleost and today’s Atlantic
salmon genome, which has retained more hox paralogy groups than any other
studied teleost (Mungpakdee et al. 2008a). After the SaGD, several hox paralogs
evolved asymmetrically, as manifested by pseudogenization, elevated rates of
molecular evolution, and/or divergence of non-coding regions in one of the two
hox gene duplicates. This asymmetry is also apparent over entire hox clusters
(Mungpakdee et al. 2008a). Overall, however, expression domains of duplicated
Atlantic salmon hox genes are conserved with their unduplicated orthologs in other
teleosts, although salmon hox paralogs show evidence for quantitative subfunctionalization (Mungpakdee et al. 2008b), a predicted type of functional evolution
of duplicate genes (Force et al. 1999).
EC
142
CO
RR
141
I. Braasch and J. H. Postlethwait
UN
Editor Proof
346
Book ISBN: 978-3-642-31441-4
Page: 346/383
Layout: T1 Standard SC
Chapter No.: 17
193
194
195
196
197
198
199
200
201
202
203
204
205
206
207
208
209
210
211
212
213
214
215
216
217
218
219
220
221
222
223
224
225
226
227
F
192
PR
OO
190
191
17.2.2.2 Carp and Goldfish: Facilitated Domestication by Genome
Duplication?
An allotetraploidization occurred in a shared ancestor of common carp (Cyprinus
carpus; 2n = 100) and goldfish (Cyprinus carpio; 2n = 100). The carp genome
duplication (CaGD) occurred between 11 and 21 million years ago (Risinger and
Larhammar 1993; Ohno et al. 1967; Larhammar and Risinger 1994; David et al.
2003; Yuan et al. 2010). Evidence for the hybrid (i.e., allotetraploid) origin of
these polyploids comes from the absence of quadrivalents (Ohno et al. 1967) and
the disomic inheritance of genetic markers in view of the short time period since
the GD event (David et al. 2003). Independent chromosomal rearrangements took
place in goldfish and carp after the allotetraploidization (Ohno et al. 1967). Around
52–60 % of genomic loci of the common carp are still duplicated (David et al.
2003; Ferris and Whitt 1977b), but duplicate retention seems to be considerably
lower in goldfish (Woods and Buth 1984). As in salmonids, several duplicates of
hox cluster genes have become pseudogenes after the CaGD (Luo et al. 2007;
Yuan et al. 2010).
Like salmonids, the common carp and its relatives are important economic
species in aquaculture as food sources and as ornamental species. Common carp
and goldfish are the oldest domesticated fish species, bred in central Europe for
*2,000 years and in China for *1,000 years, respectively (reviewed in Balon
2004). During domestication of carp and goldfish, significant morphological
change has been fixed in populations compared to their wild ancestors in a short
period of time, as exemplified by the multiple, independent loss of scales in the
carp (Balon 2004), the color morphs of koi carp (David et al. 2004), and the
selection for monstrosities such as telescope eyes, lion-head, or fin loss in ornamental goldfish (Komiyama et al. 2009).
The molecular basis of the strong response to artificial selection in carp and
goldfish has rarely been explored. Based on the multigenic inheritance of coloration
in ornamental koi carp, it has been suggested that the propensity to generate the
many color morphs may be related to the tetraploidization of carps (David et al.
2004). Is it possible that allopolyploidization provided carp and goldfish with a high
degree of genomic flexibility similar to domestic polyploid plants (see Chaps. 7 and
10, this volume)—made them particularly responsive to morphological selection
D
189
TE
188
347
Overall, around 50 % of SaGD-duplicated loci have been retained in salmonids
(Allendorf and Thorgaard 1984). Although purifying selection appears to be the
prevalent mode of salmonid gene duplicate evolution, asymmetric divergence of
salmonid gene duplicates was also observed in an analysis of EST sequences when
compared to the diploid Northern pike (Esox lucius) as outgroup (Koop et al. 2008;
Leong et al. 2010). The asymmetric pattern of molecular evolution among paralogs, observed particularly often for DNA binding proteins, is thought to be caused
by relaxed functional constraints on one of the two paralogs after the SaGD (Leong
et al. 2010; Mungpakdee et al. 2008a) as for GDs in general.
EC
187
CO
RR
186
Book ISBN: 978-3-642-31441-4
Page: 347/383
Polyploidy in Fish and the Teleost Genome Duplication
UN
Editor Proof
17
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
I. Braasch and J. H. Postlethwait
234
17.2.2.3 Gene Expression Regulation in Triploid Calandino
230
231
232
PR
OO
229
F
233
and domestication? A recent study of the common carp suggests that the duplication
of the fgfr1a gene during the course of the CaGD may have permitted the breeding
of ‘‘mirror’’ forms that lack almost all scales (Rohner et al. 2009). Importantly, two
different domesticated forms of mirror carp have been bred by selection of two
different loss-of-function mutations in the same paralog, fgfr1a1 (Rohner et al.
2009).
228
261
17.3 The Doubled Trunk: The Teleost Genome Duplication
239
240
241
242
243
244
245
246
247
248
249
250
251
252
253
254
255
256
257
258
259
262
263
264
265
TE
238
EC
237
CO
RR
236
D
260
Although polyploidization events in salmonids and carp are fairly recent events on
the evolutionary timescale, they are nevertheless too old to study changes in gene
expression pattern following soon after polyploidization and hybridization in fish
and the interplay of these two factors. The relationship of polyploidization and
hybridization, however, has been analyzed in naturally occurring populations of an
Iberian cyprinid, the calandino (S. alburnoides).
The allopolyploid S. alburnoides species complex consists of di-, tri-, and
tetraploid fish of different genomic compositions derived from interspecific
hybridizations of a paternal ancestor (the A genome) and different, geographically
separated, maternal genome contributors, the southern S. pyraneicus (P genome)
or the northern S. caroliterii (C genome) (Alves et al. 2001; Pala et al. 2010).
Expression studies of seven genes in different adult tissues revealed that dosage
compensation occurs in polyploids, so that overall gene expression is reduced to
the diploid level (Pala et al. 2008, 2010). In S. alburnoides, gene dosage compensation is accomplished by a complex pattern of gene copy silencing in a genespecific and tissue-specific manner: for example, in southern triploids of the PAA
genome composition, some genes are expressed only from the A genome, others
from P and A genomes; some tissues express only A genes, while others express
both P and A genes; some genes are expressed from A in one tissue, but from P and
A in another tissue (Pala et al. 2008, 2010).
The gene expression patterns also depend on genomic composition: while the
A genome allele is dominant in southern individuals, C and A alleles are
co-dominant in northern polyploids (CAA, CCA, CCAA, or C-A-) (Pala et al. 2010).
Although the exact molecular mechanism leading to dosage compensation in the
calandino remains elusive at this point, these studies illustrate the potential these
animals provide for the study of gene regulation in diverging polyploid populations.
235
UN
Editor Proof
348
Book ISBN: 978-3-642-31441-4
Page: 348/383
Although recent polyploidization events appear to be comparatively common in
rayfin fish compared to other vertebrate clades, the most important polyploidization event for genome evolution in fish took place on the branch leading to the
teleosts, the largest clade of rayfin fish. Initial arguments for and against the
Layout: T1 Standard SC
Chapter No.: 17
Book ISBN: 978-3-642-31441-4
Page: 349/383
Polyploidy in Fish and the Teleost Genome Duplication
349
269
17.3.1 Evidence for the Teleost Genome Duplication
270
17.3.1.1 Expansion of Vertebrate Gene Families in Rayfin Fish
PR
OO
267
F
268
occurrence of such a third round of vertebrate WGD (alias 3R, fish-specific genome duplication, FSGD, or teleost genome duplication, TGD) have been overcome
by sequencing the genomes of several teleost species.
266
287
17.3.1.2 Teleost Hox Gene Clusters
276
277
278
279
280
281
282
283
284
285
288
289
290
291
292
293
294
295
296
297
298
299
300
301
302
TE
275
EC
273
274
CO
RR
272
D
286
Initially, the branch leading to the teleosts did not raise suspicion of polyploidization because the ancestral vertebrate karyotype, like that of teleosts, was initially
inferred to be 48 chromosomes (Ohno et al. 1968).
A general observation since the rise of allozyme data, however, was that many
enzyme loci with a single locus in tetrapods appeared to have multiple copies in
fish (see Morizot 1990 and references therein). With the advent of the zebrafish
and other teleosts as model organisms for developmental and genetic studies in the
1990s and the accompanying cloning of teleost DNA sequences, this impression of
‘‘more genes in fish’’ (Wittbrodt et al. 1998) was substantially reinforced.
Three alternative explanations were initially considered to explain these
observations: (1) higher frequencies of individual or local or tandem gene duplication in teleosts than in tetrapods; (2) higher retention of gene duplicates from the
ancestral vertebrate genome duplications (VGD) in teleosts than in tetrapods; or
(3) a WGD in the rayfin fish lineage after the divergence from tetrapods (Morizot
1990; Postlethwait et al. 1998; Wittbrodt et al. 1998; Meyer 1998; RobinsonRechavi et al. 2001).
271
Sequencing the hox gene clusters, which had already suggested the earlier VGDs
(see Chap. 16, this volume), settled the argument toward the teleost-specific GD
hypothesis: since the cloning of the first teleost hox genes in the late 1980s (Eiken
et al. 1987; Njolstad et al. 1988), it became apparent that hox genes were no
exception to the generalization of ‘‘more genes in fish’’ and additional paralogs of
hox genes were found in different teleosts (e.g., Misof and Wagner 1996; Aparicio
et al. 1997; Prince et al. 1998). The genomes of tetrapods and other non-teleost
vertebrates generally have four hox gene clusters (Graham et al. 1989). The discovery that hox genes are actually organized in seven gene clusters in the zebrafish, that two copies of the hoxA cluster were present in fugu, and that each
duplicated hox cluster in fugu was orthologous to a single duplicated copy in
zebrafish provided the first significant support that a GD had occurred in the
lineage leading to zebrafish and that the event was shared by fugu (Amores et al.
1998; Aparicio et al. 1997). Importantly, the hox clusters and genes closely linked
to the hox clusters were found to be distributed over eight different chromosomes
UN
Editor Proof
17
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
303
304
305
306
in zebrafish, as would be expected if a WGD increased the hox cluster number
from four to eight, followed by the loss of one of the seven hox complements
(Amores et al. 1998). Seven or eight hox gene clusters have since been identified in
several other teleosts (see e.g. Prohaska and Stadler 2004; Hoegg and Meyer 2005;
Hoegg et al. 2007, and references therein).
308
309
PR
OO
F
307
I. Braasch and J. H. Postlethwait
17.3.1.3 More than just Hox: Duplication of Other Multigene
Families in Fish
337
17.3.1.4 Global Evidence from Teleost Genome Projects
316
317
318
319
320
321
322
323
324
325
326
327
328
329
330
331
332
333
334
335
338
339
340
341
TE
315
EC
314
CO
RR
312
313
D
336
Although the genomic location of hox gene clusters and linked gene families could be
best explained by a WGD in the lineage leading to teleosts (Meyer and Schartl 1999),
several alternative possibilities were discussed, including: (1) that the seven hox
clusters in zebrafish were due to polyploidization specific to the zebrafish lineage
(Stellwag 1999); (2) that hox cluster-containing chromosomal segments had been
amplified in teleosts by local duplications (Elgar et al. 1999); and/or (3) that teleosts
had an unusually high gene duplication rate (Robinson-Rechavi et al. 2001).
Phylogenetic analyses of numerous duplicated zebrafish genes, however,
including those not related or linked to the hox genes and those distributed all over
the genome, provided evidence for their origin after the divergence of zebrafish
and tetrapod lineages (Taylor et al. 2001; Van de Peer et al. 2001). Additional
work on specific gene families was also in line with this hypothesis [e.g. mitf
(Altschmied et al. 2002), midkines (Winkler et al. 2003), sox9 (Cresko et al. 2003),
egfr (Gomez et al. 2004), pomc (de Souza et al. 2005), and receptor tyrosine
kinases (Braasch et al. 2006)].
At the same time, putting more and more genes on the genetic maps of zebrafish
(Gates et al. 1999; Postlethwait et al. 2000; Woods et al. 2000, 2005) and medaka
(Naruse et al. 2004) provided additional evidence for a teleost WGD beyond just
the duplication of hox cluster-bearing chromosomes.
Importantly, the inclusion of gene sequences from the Japanese pufferfish
(T. rubripes) revealed that these fish-specific gene duplications within multigene
families usually date back to a point during rayfin fish evolution before the
divergence of zebrafish and pufferfish (Amores et al. 1998; Taylor et al. 2003).
Therefore, it became apparent that the GD shared by these teleost fish most likely
occurred somewhere along the branch to the teleost fish; but did it occur after the
origin of teleosts? Did it include non-teleost rayfin fish? Or did it include all
teleosts and only teleosts?
310
311
UN
Editor Proof
350
Book ISBN: 978-3-642-31441-4
Page: 350/383
The sequence of the fugu (T. rubripes) genome was the second vertebrate and the
first teleost fish draft genome assembly (Aparicio et al. 2002). The authors were
cautious about making conclusions from the Takifugu genome assembly with
respect to the TGD hypothesis, stating that the distribution of gene duplicates in
Layout: T1 Standard SC
Chapter No.: 17
Book ISBN: 978-3-642-31441-4
Page: 351/383
Polyploidy in Fish and the Teleost Genome Duplication
351
371
17.3.1.5 Phylogenetic Timing of the Teleost Genome Duplication
349
350
351
352
353
354
355
356
357
358
359
360
361
362
363
364
365
366
367
368
369
372
373
374
375
376
377
378
379
380
381
382
PR
OO
348
D
346
347
TE
345
EC
344
CO
RR
343
F
370
the Takifugu genome indicated segmental or large-scale, but not tandem, duplications (Aparicio et al. 2002).
Two follow-up large-scale analyses, however, came to the conclusion that the
distribution and age of gene duplicates in the Takifugu genome were indeed
consistent with the TGD hypothesis (Vandepoele et al. 2004; Christoffels et al.
2004). Both studies found that the majority of gene duplicates in Takifugu were
located in paralogons, i.e., chromosomal blocks that share paralogous syntenies
within the genome. Also, molecular clock-based analyses of gene families containing fish-specific gene duplicates helped to date the duplication event and hinted
at their occurrence around the origin of the teleost lineage.
In contrast to the genome assembly of Takifugu, the genome assembly of the
spotted green pufferfish (T. nigroviridis) was anchored onto chromosomes, which
made possible a more detailed analysis of conserved synteny blocks within a
teleost genome (Jaillon et al. 2004). The publication of the Tetraodon genome
assembly put the TGD beyond doubt, by showing that internal paralogy and
chromosomal blocks of doubled conserved synteny extended genome-wide within
this pufferfish genome. Doubled conserved synteny refers to a human chromosomal region sharing conserved synteny with two pufferfish paralogons (see Sect.
17.3.2.2). These patterns of conserved synteny also led to reconstructions of
ancestral karyotypes (Jaillon et al. 2004).
With the publication of the medaka genome assembly, patterns of conserved
synteny within and between the genomes of medaka, pufferfish, and zebrafish,
using human as an outgroup, allowed more refined reconstruction of ancestral, preWGD karyotypes (see Sect. 17.3.2.5) (Kasahara et al. 2007).
Formal publications of the zebrafish and stickleback genome assemblies are
pending, but analyses of these two species (as exemplified in Sect. 17.3.2) overall
confirm observations made from the genomes of pufferfish and medaka: the
occurrence of a WGD in the lineage leading to the teleosts is the most parsimonious explanation of all available genome data.
342
Genome analyses of zebrafish, pufferfish, medaka, and stickleback pointed to a GD
event somewhere in the rayfin fish lineage prior to the divergence of these teleosts,
but when exactly did the event occur? Is it, for example, a trait shared only by
clupeocephalan teleosts, to which the aforementioned sequenced species belong
(Fig. 17.1)? Or did it happen in an ancestor of all teleosts? Or even earlier, before
the rise of the teleost lineage and as such represents a trait shared with earlier
diverging rayfin branches, such as those leading to bowfin, gar, sturgeon, or bichir?
Of course, the phylogenetic timing of the TGD is essential for understanding any
possible causal relationships between the TGD and the teleost radiation. This
question is further complicated by difficulties in establishing historical relationships of various rayfin fish lineages diverging basal to the teleosts as well as of the
UN
Editor Proof
17
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
I. Braasch and J. H. Postlethwait
418
17.3.2 The TGD and Gen(om)e Evolution in Teleosts
390
391
392
393
394
395
396
397
398
399
400
401
402
403
404
405
406
407
408
409
410
411
412
413
414
415
416
419
420
421
422
PR
OO
389
D
388
TE
387
EC
385
386
CO
RR
384
F
417
basal relationships within teleosts themselves based on morphological and
molecular data.
The sequencing of four hox clusters from the bichir (Polypterus senegalus), a
representative of the most basally branching extant lineage of rayfin fish, the
polypteriforms, indicated that the GD did not occur in an ancestor of all rayfins
(Chiu et al. 2004; Raincrow et al. 2011). Subsequently, several studies cloned and
sequenced homeobox genes and several other nuclear markers from different
species representing major phylogenetic lineages of rayfins (Hoegg et al. 2004;
Crow et al. 2006; Mulley et al. 2006; Hurley et al. 2007). These studies did not find
any evidence for the presence of paralogs derived from the TGD in the genomes of
extant nonteleosts, i.e., Lepisosteiformes (gars), Amiiformes (bowfin), and Acipenseriformes (sturgeons and paddlefish, which have, however, experienced their
own lineage-specific polyploidizations; see above). In contrast, paralogs were
found in all teleost lineages analyzed, including representatives of the early
branching teleost lineages, i.e., Osteoglossomorpha (bonytongues and mooneyes),
Elopomorpha (tarpons and eels), and Clupeomorpha (herrings and shads) (see
Fig. 17.1). Finally, Amores et al. (2011) recently presented the first genome-wide
synteny comparison between a non-teleost and teleost rayfins through the generation of a high-density genetic map for the spotted gar (L. oculatus). A clear
pattern of double conserved synteny became apparent between the gar and teleost
genomes showing that the gar lineage diverged from the teleost lineage before the
GD shared among teleosts (Amores et al. 2011).
To summarize, the GD initially detected in zebrafish, medaka, and pufferfish
was an event shared by all teleosts but not by non-teleost rayfin fish, and it
occurred (depending on the use of different molecular clocks) around 226–350
million years ago (Vandepoele et al. 2004; Christoffels et al. 2004; Hoegg et al.
2004; Hurley et al. 2007) (Fig. 17.1). It thus seems better to call this tetraploidization event the TGD (The often-used term Fish-Specific Genome Duplication, or
FSGD, is ambiguous due to the many GD events in multiple fish lineages, such as
salmonids, sturgeons, and cyprinids, and the paraphyletic term ‘‘fish’’.).
Reviewing morphological characters defining the teleost lineage, de Pinna
(1996) articulated: Intriguingly, molecular data have yet to provide consistent
support for teleostean monophyly. Fifteen years later, apparently it is the TGD that
provides the best synapomorphy, a molecular character supporting the monophyly
of teleosts and probably the best we will ever obtain from extant molecules.
383
UN
Editor Proof
352
Book ISBN: 978-3-642-31441-4
Page: 352/383
17.3.2.1 The TGD and Conserved Syntenies
A pair of genes that are syntenic are on the same chromosome in one species.
Conserved syntenies are situations in which a pair of genes that are syntenic in one
species have orthologs that are syntenic in another species. People often use the
Layout: T1 Standard SC
Chapter No.: 17
Polyploidy in Fish and the Teleost Genome Duplication
(b) 1
hbbe1
Dre Chromosomes
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
Hsa16
cdh1
hbbe2
si:dkey-30c15.12
cen
HBA
0Mb
10Mb
20Mb
30Mb
CDH1
40Mb
50Mb
60Mb
70Mb
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
Hsa17
hoxba
hoxbb
slc43a2a
SLC43A2
10Mb
(d) XY
cen
6Mb
12Mb
18Mb
24Mb
hoxbb
30Mb
cen
TRAF4
20Mb
30Mb
36Mb
Dre12 Orthologs
D
Hsa Chromosomes
HOXB
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
Dre3
TE
0Mb
slc16a5b
HOXB
40Mb
50Mb
SLC16A5
60Mb
70Mb
Hsa17 Orthologs
(c) X
Y
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
Dre12
traf4a
traf4b
0Mb
80Mb
slc16a5a
slc43a2b
Hsa16 Orthologs
Hsa Chromosomes
353
F
(a)
Dre Chromosomes
Book ISBN: 978-3-642-31441-4
Page: 353/383
PR
OO
Editor Proof
17
Book ID: 272454_1_En
Date: 16-8-2012
42Mb
0Mb
HOXB
cen
hoxba
10Mb
20Mb
30Mb
40Mb
50Mb
60Mb
Dre3 Orthologs
424
425
426
427
428
429
430
431
432
433
434
435
term ‘syntenic’ erroneously to mean ‘conserved synteny’. Conserved syntenies give
strong evidence for the TGD. The Synteny Database (http://teleost.cs.uoregon.edu/
synteny_db/) uses BLAST scores to identify groups of paralogous genes in a
genome rather than a single pair of genes showing the ‘‘best hit’’; the algorithm then
anchors paralogy groups to an ortholog in another genome, and finally plots
orthologs and paralogs along chromosomes (Catchen et al. 2009, 2011). Results
show that segments of human chromosomes generally have orthologs and coorthologs in paralogons on two zebrafish chromosomes. For example, the horizontal
axis of Fig. 17.2 displays genes ordered along human chromosome 16 (Hsa16) and
on the vertical axis shows the zebrafish orthologs and co-orthologs of each Hsa16
gene plotted on the appropriate linkage groups directly above the human gene. The
short arm of Hsa16 (Hsa16p) has orthologs and co-orthologs mainly on zebrafish
chromosomes Dre3 and Dre12, including duplicated hemoglobin loci, and the long
UN
423
CO
RR
EC
Fig. 17.2 Conserved syntenies are as predicted by GD. a Genes on human chromosome 16 are
aligned along the horizontal axis and their zebrafish orthologs and co-orthologs are marked
directly above each human gene on the zebrafish chromosome on which it resides. Results reveal
duplicate ohnologons on Dre3 and Dre12 for Hsa16p and Dre7 and Dre18 for Hsa16q. A gray
circle marks the location of the centromere on Hsa16. b Zebrafish ohnologons for Hsa17 are
mainly on Dre3 and Dre12. c Human orthologs of Dre12 genes occupy single portions of human
chromosomes, mainly Hsa10 and Hsa17. d Human orthologs of Dre12 are widely scattered on
mostly Hsa16 and Hsa17. Individual genes are located on each plot
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
I. Braasch and J. H. Postlethwait
456
17.3.2.2 Doubled Conserved Synteny
440
441
442
443
444
445
446
447
448
449
450
451
452
453
454
PR
OO
439
D
438
TE
437
F
455
arm (Hsa16q) has orthologs and co-orthologs on Dre7 and Dre18, including
co-orthologs of CDH1 and many other genes. Other zebrafish chromosomes also
have strings of genes that appear on the plot, including Dre24 and Dre25, but most
of these genes are more distantly related paralogs resulting from the VGDs.
The content of other human chromosomes appears to have involved more
rearrangements than Hsa16 since the divergence of the human and zebrafish lineages, either in the rayfin or the lobefin lineage or both. For example, in zebrafish,
the genetic content now on Hsa17 occupies duplicates of at least seven distinct
large chromosome segments, mostly on Dre3 and Dre12 (Fig. 17.2b), which
indicates several major translocations with respect to human, some of which
occurred before the TGD because they are shared by both copies, and some after
the TGD, because they differ between the two paralogons (or ohnologons, paralogons arising from GD).
Comparisons in the other direction, zebrafish to human, generally display a
one-to-one relationship with a substantial number of translocations and inversions
(Fig. 17.2c). For example, about half of the human orthologs of Dre12 genes
are located on Hsa17, and about half on Hsa10, each with several rearrangements;
minor numbers are located on four other zebrafish chromosomes. Other
chromosomes, like Dre3, appear to have experienced an even greater number of
chromosome rearrangements (Fig. 17.2d).
436
467
17.3.2.3 An Example: The Hox Clusters
459
460
461
462
463
464
465
468
469
470
471
472
473
CO
RR
458
EC
466
The most frequent fate of a pair of ohnologs after WGD is nonfunctionalization
(Lynch and Conery 2000); hence, ohnologons often experience reciprocal gene
loss, like the regions containing two zebrafish co-orthologs of SLC16A 5 on Dre3
and Dre12 (Figs. 17.2b and 17.3). Of the string of four genes on Hsa17 including
human SLC16A 5 , different sets are missing from each duplicated zebrafish
segment, so that the content of both zebrafish ohnologons must be summed to get
the orthologous human gene content in this region (Fig. 17.3). In the extreme, all
WGD duplicates will have resolved to singletons, and so conserved syntenies will
be maintained but with no co-orthologs to anchor the segments in a doubled
conserved synteny.
457
Circle plots can display conserved paralogs on a genome-wide scale. For example,
the Synteny Database shows that paralogs of genes within 10 Mb of the four
human HOX clusters preferentially occupy regions near each of the other clusters
(Fig. 17.4a) (For nomenclature, see https://wiki.zfin.org/display/general/ZFIN+
Zebrafish+Nomenclature+Guidelines.). Note stronger links between the pairs
HOXA/HOXB and HOXC/HOXD, which may reflect the order of their origin, with
UN
Editor Proof
354
Book ISBN: 978-3-642-31441-4
Page: 354/383
Layout: T1 Standard SC
Chapter No.: 17
17
Book ID: 272454_1_En
Date: 16-8-2012
Polyploidy in Fish and the Teleost Genome Duplication
Fig. 17.3 Doubled
conserved synteny. The
region at about 73 Mb on
Hsa17 including SLC16A5
shares syntenies with small
regions of Dre3 and Dre12.
Zebrafish has two coorthologs of SLC16A5
(boxed), but some of its
neighbors were reciprocally
lost from one or the other
ohnologon. In addition, an
inversion on Dre12 reversed
gene orders and caused other
genes to disrupt the ancestral
group. This pattern of
doubled conserved synteny
with disrupting inversions is
quite common
355
LOC794371
Dre3
F
atp5h
PR
OO
ARMC7
SLC16A5
KCTD2
Hsa17
ATP5H
Dre12
zgc:153947
LOC565514
zgc:153278
TE
D
Editor Proof
Book ISBN: 978-3-642-31441-4
Page: 355/383
486
17.3.2.4 Symmetry of Gene Loss Between Members of Ohnologons
478
479
480
481
482
483
484
487
488
489
490
491
492
493
494
CO
RR
476
477
Immediately after GD, both ohnologons have identical gene compositions. As
nonfunctionalization events occur, ohnologons begin to diverge from each other.
Gene loss events raise the question: Do both members of a pair of ohnologons tend
to lose genes at the same rate? Or do genes tend to disappear more frequently from
one duplicated chromosome segment than the other? According to one hypothesis,
gene losses would be purely stochastic. According to an alternative hypothesis,
expression of a number of genes in the same neighborhood might be regulated by a
central element, for example, an element that helped to regulate local chromosome
UN
475
EC
485
VGD1 giving rise to HOXA/B and HOXC/D clusters (Amores et al. 1998) and
VGD2 providing all four clusters. A circle plot of orthologs and paralogs of genes
surrounding zebrafish hox clusters (Fig. 17.4b) shows that zebrafish has eight hox
cluster-containing ohnologons even though only seven of them contain proteincoding hox genes (Amores et al. 1998). While zebrafish has two copies of the
hoxa, hoxb, and hoxc clusters, it has a single hoxd cluster that contains proteincoding genes. The hoxdb cluster, while lacking protein-coding genes, retains a
microRNA-10 paralog, which is embedded in Hox clusters from flies to fish to
humans (Woltering and Durston 2006), as well as paralogs of many surrounding
genes easily detected in the plot (Fig. 17.4b). Likewise, stickleback lacks a second
hoxc cluster, but the location where it ‘should’ go is apparent from the circle plot
(Fig. 17.4c).
474
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 356/383
Editor Proof
356
(a)
I. Braasch and J. H. Postlethwait
HOXC
(b)
HOXB
(c)
hoxaa
hoxda
hoxab
hoxca
hoxca
(hoxcb)
hoxba
hoxcb
hoxba
hoxda
PR
OO
hoxdb
hoxbb
F
hoxbb
HOXA
hoxab
hoxaa
HOXD
hoxdb
Fig. 17.4 Hox cluster ohnologons. a Paralogs within 10 Mb of human HOX clusters mapped on
the human genome. b Paralogs within 5 Mb of zebrafish hox clusters mapped on the zebrafish
genome. Although only a microRNA gene remains in the hoxdb cluster, the paralogy of this
segment and its relationship to hoxda is evident. c Paralogs within 3 Mb of stickleback hox
clusters mapped on the stickleback genome
501
502
503
504
505
506
507
508
509
510
511
512
513
514
515
516
517
518
519
520
521
522
523
524
D
500
TE
498
499
EC
497
structure, and loss of that element would then lead to rapid loss of the several
genes that element controls and hence asymmetric gene loss. Analysis of a few
individual chromosome regions suggests that asymmetries often occur in teleosts
(Canestro et al. 2009; Braasch et al. 2006; Siegel et al. 2007), but this question has
not been explored sufficiently on a genome-wide scale.
To study this question, we used the Synteny Database to plot the zebrafish and
stickleback chromosomes on which orthologs, co-orthologs, and paralogs of each
human gene lie for all human chromosomes. Figure 17.5 displays results for two
human chromosomes (Hsa7 and Hsa18) for both zebrafish and stickleback. Figure 17.5a shows genes on Hsa7 as gray dots along the bottom with the zebrafish
orthologs and paralogs directly above; note that gene order in these plots reflects
gene arrangements in the human genome, not the fish genome.
Results revealed several principles (Fig. 17.5a–d). (1) Many regions of
ohnology are clear, as marked by rectangles. For example, the region from 18 to
38 Mb on Hsa7 has two clear ohnologons in both zebrafish and stickleback, with
orthologous chromosome segments marked by color (Fig. 17.5a, b). (2) In contrast, many regions have ambiguous ohnologons, such as the region from the left
telomere of Hsa7 to about 18 Mb for both zebrafish and stickleback. (3) Some
ohnolog pairs are clear in one species but ambiguous in the other; for example,
from 38 to 64 Mb of Hsa7, where three or four zebrafish chromosomes have
strings of ‘hits’ compared to two clear regions in stickleback. Regions like these
(also note for stickleback region 101–142 Mb the ‘ghost’ paralogons on groupXII
and groupXVIII) are often due to paralogs arising in the VGD1 and VGD2 events
rather than the TGD. (4) Most chromosome translocations occurred before the
divergence of the zebrafish and stickleback lineages. Evidence supporting this
conclusion is that, with respect to the human chromosome, orthologous blocks
tend to have the same termini in both fish species. And (5), a minority of translocations occurred in one or the other fish lineage after divergence; for example,
the region from 77 to 88 Mb and the region from 101 to 142 Mb are syntenic (on
the same chromosome) in human (Hsa7) and in both ohnologons in stickleback
CO
RR
496
UN
495
Layout: T1 Standard SC
Chapter No.: 17
Book ISBN: 978-3-642-31441-4
Page: 357/383
Polyploidy in Fish and the Teleost Genome Duplication
(a)
357
2
4
6
Dre Chromosomes
8
10
12
F
14
16
18
PR
OO
20
22
24
Hsa7
(b)
groupII
Gac Chromosomes
groupIV
groupVI
groupVIII
groupX
groupXII
groupXIV
groupXVI
groupXVIII
groupXX
Hsa7
20Mb
40Mb
60Mb
80Mb
100Mb
D
0Mb
120Mb
140Mb
(c)
TE
Hsa7 Orthologs
2
4
6
Dre Chromosomes
8
10
12
EC
14
16
18
20
22
24
(d)
CO
RR
Hsa18
groupII
groupIV
Gac Chromosomes
groupVI
groupVIII
groupX
groupXII
groupXIV
groupXVI
groupXVIII
groupXX
Hsa18
0Mb
UN
Editor Proof
17
Book ID: 272454_1_En
Date: 16-8-2012
10Mb
20Mb
30Mb
40Mb
50Mb
60Mb
70Mb
Hsa18 Orthologs
Fig. 17.5 Ohnologons in zebrafish (a, c) and stickleback (b, d) for Hsa7 (a, b) and Hsa18 (c, d).
These plots show doubled conserved synteny as well as the pattern of chromosome translocations
in various lineages
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
532
533
534
535
536
537
538
539
540
541
542
543
544
545
546
547
548
549
550
551
552
553
554
555
556
557
558
559
560
561
562
563
564
565
566
567
F
531
PR
OO
530
D
528
529
TE
527
(groupIV and groupXX)—thus representing the ancestral condition—but for
zebrafish, these two regions are syntenic for only one of the copies (Dre4) but not
the other (Dre18 and Dre25). These results demonstrate a translocation in the
zebrafish lineage after it diverged from the stickleback lineage (Fig. 17.5a, b).
Note that the effects of inversions in the fish lineage are invisible in these plots
because gene orders in the fish genomes are displayed according to their order in
the human genome.
Having identified ohnologons, we can now pose the question of gene loss
asymmetries. Over the human genome, we identified 49 unambiguous ohnologon
pairs in zebrafish or stickleback or both, and counted the number of fish orthologs
and paralogs of human genes occupying each segment. For zebrafish, 7 of 39
(17.9 %), and for stickleback, 13 of 44 (29.5 %) ohnologon pairs had significantly
different number of genes [v2 test, 1 df, p \ 0.01; 10/39 (25.6 %), which increased
to 18/44 (40.9 %) if p \ 0.05]. We conclude: (1) that most ohnologons appear to
lose genes approximately randomly between the two duplicated copies, but (2) that
about 20–40 % of ohnologons in these fish species have resolved TGD ohnologs
asymmetrically between the two duplicated chromosome segments, with one
member of the pair retaining significantly more genes than the other. (3) Of 13
segments with significance at the 0.01 level in one or the other fish, 6 are significant at 0.01 or 0.05 level in both species, which diverged deeply in teleost
phylogeny (Fig. 17.1), and for all of those pairs, chromosome segments retaining
the most genes in zebrafish are orthologs of those retaining the most in stickleback.
This result is consistent with the idea that the basis for most asymmetric gene
losses occurred before the divergence of zebrafish and stickleback lineages.
A question that remains is the mechanism that led to asymmetric resolution of
gene duplicates after the TGD. One hypothesis is that, if a long-range regulatory
function that controls the expression of many genes in a neighborhood disappears
from one ohnologon, then nonfunctionalization of the genes that the regulatory
function controls can follow without penalty given the maintenance of that longrange function in the sister ohnologon. That hypothesis is yet to be seriously
investigated. An example of resolution asymmetries is shown in Fig. 17.6 for
Hsa15 in zebrafish and stickleback, along with chromosome diagrams comparing
genes in their order along a segment of Hsa15 and their fish orthologs and coorthologs in zebrafish (52 ortholog pairs in one ohnologon vs. 4 ortholog pairs in
the other) and stickleback (54 ortholog pairs vs. 16 ortholog pairs) (Fig. 17.6c, d).
A related question is whether ohnologs residing in ohnologons that suffer
asymmetric rates of gene loss also experience asymmetric rates of gene evolution.
Asymmetric rates of molecular evolution have been found for a substantial number
of TGD paralogs (Van de Peer et al. 2001; Steinke et al. 2006; Brunet et al. 2006),
sometimes correlating with asymmetric loss of genes from ohnologons (Braasch
et al. 2006; Siegel et al. 2007). Available evidence from other WGDs, however,
suggests that gene evolutionary rate asymmetry does not strongly depend on the
conservation of syntenies (Bu et al. 2011).
EC
526
CO
RR
525
I. Braasch and J. H. Postlethwait
UN
Editor Proof
358
Book ISBN: 978-3-642-31441-4
Page: 358/383
Layout: T1 Standard SC
Chapter No.: 17
Editor Proof
17
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 359/383
Polyploidy in Fish and the Teleost Genome Duplication
(a)
359
2
4
6
Dre Chromosomes
8
10
12
F
14
16
18
PR
OO
20
22
24
Hsa15
(b)
groupII
Gac Chromosomes
groupIV
groupVI
groupVIII
groupX
groupXII
groupXIV
groupXVI
groupXVIII
groupXX
Hsa15
0Mb
20Mb
40Mb
(c)
PAK6(1of2)
sb:cb730
srp14
zgc:136872
FMN1
GJD2 MEIS2 SRP14
C15orf57
C15orf24 ACTC1
PLCB2
AQR
C15orf52
ATPBD4
EXD1
EHD4 LRRC57
RPAP1 PLA2G4F
CDAN1
VPS39
TTBK2
ZFP106 UBR1
spint1b
cx35
zgc:86709
meis2.1
SCG5
ENSGACG00000007697
ENSGACG00000007661
ENSGACG00000005610
GJD2(2of2)
CASC5
GacgroupXVIII
54 pairs
im:7147183
80Mb
GREM1
SCG5CHRM5
ARHGAP11A RYR3
Hsa15
16 pairs
GacgroupXV
FMN1
AC012652.1
EIF2AK4
CASC5
CHAC1
MAPKBP1
FAM98B
RPUSD2 SPINT1
TYRO3
SPRED1
BUB1B DISP2
ZFYVE19 NUSAP1
JMJD7
C15orf41
THBS1 PAK6
IVD GCHFR DLL4
RTF1 MGA
GJD2
MEIS2
FSIP1 PLCB2
DNAJC17
C15orf57
AVEN
ACTC1 RASGRP1
GPR176
C15orf24
AQR
SRP14
C15orf29
ATPBD4
MAPKBP1(2of2)
ENSGACG00000003916
TYRO3
LRRC57 JMJD7
100Mb
NDUFAF1
RPAP1
CHAC1(1of2)
DLL4(2of2)
LOC100334937
vps39
LOC100331590
si:ch211-283f6.8
zgc:63611
zgc:154068
zgc:110758
meis2.2
MEIS2(2of2)
fam98b
spred1
zgc:154061
CAPN3(2of2)
VPS39
CDAN1
TMEM87A
TTBK2
ZFP106
UBR1
LRRC57
TMEM87A
ZFP106
capn3
LOC100331516
ubr1
ENSGACG00000009824
FMN1(2of2)
CHRM5(2of2)
THBS1(1of2)
RYR3(2of2)
STARD9
HAUS2
SNAP23
CAPN3
GANC
TMEM62
CAPN3(1of2)
VPS39
GANC
ENSGACG00000005683
ENSDARG00000091548
LOC565314
zgc:77419
PAK6(1of2)
FMN1(1of2) GCHFR
BUB1B
FSIP1 EIF2AK4
SRP14
RASGRP1 CHRM5(1of2) DNAJC17 ZFYVE19
THBS1(2of2)
SPINT1(1of2)
RYR3(1of2)
PLCB2
GREM1
ENSGACG00000006109
C15orf57
DISP2
UBR1
RPUSD2 IVD
RPAP1
FAM98B
SPRED1
C15orf41
GJD2(1of2) MEIS2
ENSGACG00000010421
AQR
ATPBD4
TMEM62
SNAP23
ENSGACG00000011127
CDAN1
TTBK2(1of2)
RTF1
NUSAP1
ENSGACG00000012183
NDUFAF1
EC
HAUS2
C15orf24
C15orf29
GPR176
AVEN
zgc:123218
zgc:56072
LOC565088 ivd
LOC564712
RPAP1
si:dkey-170l10.1
TE
4 pairs
Dre20
NP_001185682.1
disp2 itpka
MAPKBP1
jmjd7
mga
tyro3
JMJD7
STARD9
RPUSD2
MAPKBP1
HAUS2
FAM98B PAK6 BAHD1
MGA
SNAP23
SPRED1 BUB1B
IVD
SPINT1 TYRO3
CAPN3
C15orf41 THBS1
DISP2 ZFYVE19
ITPKA
GANC TMEM62
CHRM5
GREM1
ARHGAP11A
Hsa15
LOC565386
ENSDARG00000056535
lrrc57
ENSDARG00000079066
actc1b
PLCB2
chrm5a
spint1a
LOC100005728 zgc:92683
arhgap11a
LOC796966
52 pairs
D
ENSDARG00000088538
THBS1(2of2)
zgc:195050
Dre17
(d)
60Mb
Hsa15 Orthologs
569
570
571
572
573
574
575
576
577
578
579
580
581
17.3.2.5 Reconstruction of the Ancestral Karyotype and Teleost Chromosome
Rearrangements
As we discussed earlier, the ancestral haploid teleost (post-TGD) genome was
inferred to contain 24 chromosomes based on karyotype data (Ohno et al. 1968;
Mank and Avise 2006b). Using several teleost genetic maps as well as the genome
assemblies of Tetraodon and medaka, several attempts have been made to
reconstruct the ancestral pre-TGD protokaryotype (Naruse et al. 2004; Jaillon et al.
2004; Woods et al. 2005; Kohn et al. 2006; Kasahara et al. 2007; Nakatani et al.
2007). All studies largely agree that the teleost karyotype is derived from 11 to 12
pre-TGD protochromosomes, which is in agreement with the inferred number of
24 post-TGD chromosomes (Mank and Avise 2006b). There are, however, some
differences in the assignment of chromosomal blocks from different extant teleosts
to the protochromosomes (Woods et al. 2000; Kasahara et al. 2007). The reconstruction of the ancestral teleost karyotype has so far relied on using tetrapod
UN
568
CO
RR
Fig. 17.6 Asymmetrically resolved ohnologons for Hsa15 in zebrafish (a) and stickleback (b).
Chromosome contents displayed with proper gene orders for zebrafish duplicates (c, 52 vs. 4
ortholog pairs) and stickleback (d, 54 vs. 16 pairs)
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
588
589
590
591
592
593
594
595
596
597
598
599
600
601
602
603
604
605
606
607
608
609
610
611
612
613
614
615
616
617
618
619
620
621
622
623
624
625
626
F
587
PR
OO
586
D
585
TE
584
genomes as outgroups, but the genome sequence of gar or bowfin, rayfin fish that
can serve as outgroup to the TGD, will help to provide a more detailed picture of
the ancestral teleost genome.
Nevertheless, the current state of karyotype reconstruction is helpful to compare
and cross-refer different types of data on duplicated genes in teleosts. As an
example, for zebrafish and stickleback we downloaded all pairs of predicted
duplicated genes and their genomic locations from the EnsemblCompara GeneTree set (Vilella et al. 2009) and parsed them for the ‘Clupeocephala’ duplication
node, which indicates a duplication on the branch leading to teleosts, after
divergence from tetrapods, but before the divergence of teleosts. We then plotted
the location of these pairs as Oxford grids for zebrafish and stickleback genomes
(Fig. 17.7) and overlaid them with the 12 ancestral pre-TGD protochromosomes
a–m inferred from the analysis of the medaka genome (Kasahara et al. 2007). This
analysis clearly shows that paralogs are nonrandomly distributed over the genome;
for example, of the 228 genes in the analysis that reside on zebrafish chromosome
Dre3 and have TGD duplicates, the vast majority, 105 genes, are on Dre12 and a
sizable minority on Dre1, and of the 167 genes on stickleback groupXIV that are
duplicated and assigned to chromosomes, 152 are on groupXIII. These results
show that the vast majority of ‘Clupeocephala-duplicated’ genes obtained from the
tree-only method used by Ensembl supports a TGD origin of the paralogs and is in
line with conserved synteny data and ancestral karyotype reconstructions.
Teleost genomes are rearranged with respect to tetrapod genomes, which may
be due to chromosome rearrangements that were facilitated by the TGD, for
example through illegitimate recombination between homeologous (paralogous)
chromosomes. This hypothesis, however, is controversial (Comai 2005; Semon
and Wolfe 2007a; Hufton et al. 2008), because chromosome restructuring may
have likewise occurred before the TGD, on the long branch separating teleosts and
tetrapods after the divergence of rayfin fish from lobefin fish.
Using the genetic map of the spotted gar, which contains nearly 1,000 coding
markers, Amores et al. (2011) recently showed unexpectedly high conservation of
synteny between human and gar when compared to zebrafish and stickleback. This
suggests that chromosome rearrangements and the loss of ancestral syntenies
accelerated after the TGD, but before the divergence of stickleback from zebrafish,
and supports the hypothesis that WGD can facilitate syntenic rearrangements.
The zebrafish genome appears to be more rearranged than the genome of
percomorph teleosts (stickleback, medaka, pufferfish). Chromosomal blocks of the
inferred ancestral pre-TGD karyotype (Kasahara et al. 2007; Nakatani et al. 2007)
are generally distributed over many more chromosomes in zebrafish than in percomorphs. For example, as shown in Fig. 17.7, chromosomal blocks of the
ancestral pre-TGD chromosome m are found on four stickleback chromosomes
(GacI, GacIII, GacVIII, GacXXI), but on six chromosomes in zebrafish (Dre2,
Dre6, Dre8, Dre11, Dre22, Dre24); pieces of ancestral chromosome j are found on
two chromosomes in stickleback (GacII, GacXIX), but on three in zebrafish (Dre7,
Dre18, Dre25), and so forth. A genome sequence of another representative of the
ostariophysians, which, after percomorphs, is the next most species-rich group of
EC
583
CO
RR
582
I. Braasch and J. H. Postlethwait
UN
Editor Proof
360
Book ISBN: 978-3-642-31441-4
Page: 360/383
Layout: T1 Standard SC
Chapter No.: 17
Polyploidy in Fish and the Teleost Genome Duplication
chr
1
2
3
4
5
6
7
8
9
10
11
12
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
Un
8
1
51
0
0
2
18
1
49
0
0
0
25
47
0
0
7
0
1
4
2
1
15
0
1
1
10
0
0
0
30
22
17
0
0
24
9
0
0
20
0
0
6
0
25
0
48
7
86
0
3
8
0
0
29
0
0
0
0
4
105
0
0
0
1
0
0
1
0
0
7
0
17
1
4
4
0
2
0
2
0
1
0
0
0
0
0
7
3
26
0
1
0
0
0
0
44
1
48
1
7
14
1
48
3
3
0
18
36
1
0
3
0
0
36
0
2
7
0
3
12
0
7
67
0
22
1
4
0
0
0
0
0
0
0
0
2
60
1
0
2
8
0
1
13
1
0
1
2
0
0
0
41
0
0
18
0
0
0
70
2
16
0
45
33
0
1
0
0
0
0
0
0
0
25
13
14
0
0
2
12
1
8
2
0
2
1
1
0
0
1
0
0
17
0
0
0
2
2
0
0
0
14
54
0
0
2
0
0
1
0
0
4
2
1
2
0
6
1
0
0
1
0
2
0
0
0
58
0
1
3
I
II
III
IV
V
VI
VII
VIII
IX
X
XI
10
1
1
49
0
35
0
0
0
0
2
7
0
1
0
77
1
0
6
6
63
0
0
1
0
125
0
3
0
1
0
0
0
1
2
0
6
13
14
15
16
6
49
1
0
0
27
0
3
0
0
0
0
0
1
3
4
0 0
0
0
0
1
1
1
8
0
18
0
0
36
8
0
0
1
0
0
3
0
2
1
8
0
0
2
11
0
36
0
0
0
0
0
14
3
0
0
0
174 4
4 132
0
1
0
2
3
2
1
0
0
1
2
2
XII
XIII
XV
XVI XVII XVIII XIX
2
8
(b)
2
0
0
48
0
1
0
1
0
2
9
0
2
0
0
0
9
32
1
28
0
0
0
1
1
2
0
1
0
0
0
7
19
8
1
0
1
0
0
0
17
0
0
0
0
0
1
2
17
18
19
20
21
22
23
24
XIV
2
0
0
8
1
0
0
24
1
2
1
0
0
1
0
2
1
4
0
0
0
1
0
4
10
0
0
1
0
3
9
7
0
0
2
XX
XXI
Un
28
0
0
2
5
1
5
0
0
6
107 0
0
0
0
0
0
0
0
14
0
0
0
0
0
4
0
0
1 147
1
0
22
2
4
1
1
0
0
0
0
0
0
0
0
30
25
Un
48
0
6
0 152
0
0
0
0
103 10
0
0
1
0
0
0
0
0
70 12
0
0
0
0
1
0
0
0
14
2
3
0
97
0
1
2
10
0
0
0
0
0
0
14
8
0
0
0
0
40
4
8
8
ancestral
chromosomes:
D
6
1
2
16
0
4
0
2
1
0
0
0
0
0
1
123 0
8
3
2 150
1
2
0
1
0
0
1
0
0
0
0
0
0
0
0
0
1
0
3
1
0
74 20 11
0
0
0
4
11
0
3 126 1
2
2
0
0
0
56
9
1
33
TE
I
II
III
IV
V
VI
VII
VIII
IX
X
XI
XII
XIII
XIV
XV
XVI
XVII
XVIII
XIX
XX
XXI
Un
EC
group
361
F
(a)
Book ISBN: 978-3-642-31441-4
Page: 361/383
PR
OO
Editor Proof
17
Book ID: 272454_1_En
Date: 16-8-2012
2
0
14
0
8
8
0
0
19
a
b
c
d
e
f
g
h
i
j
k
l
m
0
0
23
2
7
51
628
629
630
631
632
633
teleosts and includes zebrafish and carps (Nelson 2006), will be necessary to
clarify whether this enhanced rearrangement rate is specific to the zebrafish genome or a more general phenomenon on this major branch of teleost diversity.
UN
627
CO
RR
Fig. 17.7 Oxford grid of duplicated genes in zebrafish (a) and stickleback (b). Chromosomal
location of paralogs determined by EnsemblCompara GeneTrees (Vilella et al. 2009) to be
generated at the ‘Clupeocephala’ node. Each box in the grid shows the number of duplicated
genes shared between the indicated chromosomes. The distribution of duplicated genes is
nonrandom as expected from the TGD. Most clusters of duplicated genes can be traced back to
the ancestral protochromosome (Kasahara et al. 2007), from which they are derived (see color
code)
17.3.2.6 Global Patterns of Gene Retention and Loss After the TGD
The TGD obviously had a major impact on the evolution of genome structure in
teleosts. We still do not know, however, how many paralogs from the TGD have
escaped nonfunctionalization and have been retained in duplicate in extant
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
641
642
643
644
645
646
647
648
649
650
651
652
653
654
655
656
657
658
659
660
661
662
663
664
665
666
667
668
669
670
671
672
673
F
640
PR
OO
639
D
637
638
TE
636
teleosts. Genome-wide estimates range between 12 and 24 % depending on the
approach and dataset (Postlethwait et al. 2000; Jaillon et al. 2004; Woods et al.
2005; Brunet et al. 2006; Kassahn et al. 2009) and do not differ significantly
among sequenced teleost genomes (Kassahn et al. 2009). Between *1,800 and
*2,200 TGD-derived pairs of gene duplicates can be identified in the five
sequenced teleost genomes (I. Braasch, ‘‘unpublished data’’). Loss of paralogs
after the TGD appears to be time-dependent, with most losses occurring soon after
the GD event (Braasch et al. 2009a; Sato et al. 2009). A similar trend can be
observed when comparing paralog loss after different GD events in teleosts, from
the CaGD *11–21 million years ago (mya) with a paralog retention rate of
*60 % (David et al. 2003), over the sucker genome duplication (SuGD) and
SaGD, which occurred *50 mya and 25–100 mya, respectively, and which both
have estimated paralog retention rates of *50 % (Ferris 1984; Allendorf and
Thorgaard 1984), to the TGD that happened *226–350 mya and that has an
estimated paralog retention of *12–24 % (see above).
Retained paralogs from the TGD are enriched for transcription factors, developmental genes, and cell communication proteins (Brunet et al. 2006; Kassahn
et al. 2009), similar to results for the earlier rounds of WGD in vertebrates (see
Putnam et al. 2008; Huminiecki and Heldin 2010; Chap. 16, this volume). Some
gene classes appear to have a high rate of TGD paralog retention, such as genes
encoding components of pigmentation pathways (Braasch et al. 2007, 2009a) and
of signaling pathways involved in long-term potentiation of synaptic transmission,
and olfactory and taste transduction (Sato et al. 2009). However, a more detailed
analysis of gene family evolution after the TGD awaits the genome assembly of a
rayfin outgroup to the TGD, the spotted gar (see below).
EC
635
17.3.3 Functional Divergence of Gene Duplicates: Examples
in Fish
CO
RR
634
I. Braasch and J. H. Postlethwait
After a polyploidization event, ohnolog pairs are initially identical, both in regulatory elements and in protein-coding areas, so they are fully redundant. Ohnologs,
however, soon begin to accumulate mutations that become fixed in populations
and distinguish the a copy from the b copy; these mutations can occur in both
regulatory and protein coding regions.
Some mutations can cause one copy of a duplicated pair of genes to become
nonfunctional, either because they inactivate the protein by premature stop
mutation or destructive amino acid substitution, or because they eliminate gene
expression. Nonfunctionalization is the most frequent fate of gene duplicates
(Lynch and Conery 2000). For genes that are not dosage-sensitive, nonfunctionalization of one copy is likely to carry little selective penalty (and may provide in
some cases an advantage), reflecting the fact that most mutations are recessive in a
diploid.
UN
Editor Proof
362
Book ISBN: 978-3-642-31441-4
Page: 362/383
Layout: T1 Standard SC
Chapter No.: 17
Book ISBN: 978-3-642-31441-4
Page: 363/383
Polyploidy in Fish and the Teleost Genome Duplication
363
684
17.3.3.1 Subfunctionalization
680
681
682
685
686
687
688
689
690
691
692
693
694
695
696
697
698
699
700
701
702
703
704
705
706
707
708
709
710
711
712
713
714
715
PR
OO
679
The classic example that led to the idea of subfunctionalization—duplicate gene
preservation by the reciprocal loss of ancestral gene subfunctions—is the zebrafish
pair of engrailed-1 co-orthologs (Force et al. 1999). The mouse En1 gene is
expressed in both the pectoral appendage bud and in a specific set of hindbrain
interneurons (Joyner and Martin 1987; Davis et al. 1991; Gardner and Barald
1992). In contrast, the zebrafish co-orthologs eng1a and eng1b partitioned these
two expression domains between them, with eng1a expressed in the pectoral
appendage bud and, reciprocally, eng1b expressed in the hindbrain interneurons, a
case of spatial subfunctionalization (Force et al. 1999). The reciprocal loss of
ancestral regulatory domains like this, which can occur purely by neutral evolution, makes both copies essential, with eng1a essential to provide Engrailed
function to the pectoral fin bud and eng1b necessary to deliver Engrailed protein to
interneurons.
Many genes evolved like eng1 genes to show spatial subfunctionalization, but
some genes evolved by temporal subfunctionalization, duplicate gene expression
in the same tissue but at multiple developmental stages. For example, Sox1 is
expressed in tetrapods at several developmental stages in the lens, but after the
TGD, sox1a came to be expressed several hours earlier in the lens than is
sox1b (Okuda et al. 2006). Subfunctionalization is pervasive: a global analysis of
TGD ohnologs showed that nearly all that were examined had diverged in their
patterns of spatial and/or temporal expression during embryogenesis and that such
regulatory changes were more frequent than changes at the level of protein
function (Kassahn et al. 2009). Tetrapod enhancers, which represent a type of
regulatory subfunction, and CNEs, many of which may be enhancers, evolved
divergent sequences in teleosts by altering a few specific bases that underlie
divergent functions, as for medaka and fugu ohnologs derived from Hoxa2
(Tumpel et al. 2006) or zebrafish and stickleback Fgf8 co-orthologs (Canestro
et al. 2007); this result would be expected to occur by subfunctionalization. Cases
of preserved ohnologs that have no documented differences in embryonic
expression domains may display differences at developmental stages or tissues that
have not yet been examined.
D
678
TE
677
EC
676
CO
RR
675
F
683
Besides nonfunctionalization of one member of a duplicate pair, reciprocal
changes to both members of the pair cause both ohnologs to become essential.
Reciprocal changes can be of two main sorts, either reciprocal loss of ancestral
gene subfunctions that occur purely by neutral processes (subfunctionalization), or
by mutation in one or both copies that result in both ohnologs being positively
selected for new functions (neofunctionalization) (Force et al. 1999). Mutations
that lead to the partitioning of subfunctions between ohnologs do not preclude the
later origin of novel subfunctions [called subneofunctionalization (He and Zhang
2005; Rastogi and Liberles 2005)] and vice versa. Here, we note a few examples
for each of several fates of TGD ohnologs.
674
UN
Editor Proof
17
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
723
724
725
726
727
728
729
730
731
732
733
734
735
736
737
738
739
740
741
742
743
744
745
746
747
748
749
750
751
752
753
754
755
756
757
758
759
760
F
722
PR
OO
721
D
719
720
TE
718
Besides spatial or temporal differences in expression domains, some ohnologs
may achieve quantitative subfunctionalization, the accumulation of activityreducing mutations in both ohnologs, so that neither gene by itself can provide
sufficient product to achieve normal function (Force et al. 1999; Stoltzfus 1999).
Mutations that partially reduce gene function should be relatively common, and if
a population fixes a partial loss-of-function allele at one copy, then it is unlikely to
fix a null allele in the gene’s ohnolog. Although quantitative subfunctionalization
may be common after WGD in yeast (Scannell and Wolfe 2008), few examples
exist from the TGD, probably because of the lack of appropriate analyses. We do
know that quantitative subfunctionalization occurred in many of the hox gene
duplicates after the SaGD (Mungpakdee et al. 2008b). The finding that most pairs
of CNEs in teleost ohnologs experience substantial degeneration in element length
might also reflect cases of quantitative subfunctionalization (Woolfe and Elgar
2007). The evolutionary significance of quantitative subfunctionalization may be
that it preserves both duplicates long enough to increase the opportunity for later
mutations that might secondarily increase gene activity, as appears to have happened for the medaka copy of aldh1a2, a gene originating in VGD2 (Canestro
et al. 2009).
Protein structure evolution: Individual tetrapod genes can encode proteins with
multiple functions. In some cases, a single protein sequence performs several
functions; in other cases, a single gene encodes different protein sequences, often
with different functions, due to alternative splicing or start sites.
Amino acid substitutions can cause proteins encoded by a pair of ohnologs to
diverge in sequence in functionally important ways. For example, the chemokine
Cxcl12 guides gastrulation and the migration of primordial germ cells (PGCs) in
several vertebrates (Nair and Schilling 2008; Richardson and Lehmann 2010).
Zebrafish has co-orthologs of Cxcl12 that appear to have arisen in the TGD
(Doitsidou et al. 2002). Cxcl12 functions by binding Cxcr4 receptor, which is also
duplicated in zebrafish. Cxcl12a, but not Cxcl12b, guides PGCs to their target, the
gonad, and a single amino acid substitution causes the relative affinity of Cxcl12
ligands to switch from one of the duplicated Cxcr4 receptors to the other, thereby
controlling whether the Cxcl12:Cxcr4 ligand:receptor system controls PGC
migration or gastrulation (Boldajipour et al. 2011). These experiments are a particularly deeply studied case study supporting the concept of protein
subfunctionalization.
Alternative splicing can generate different proteins with variant functions. For
example, the human STAT1 gene, which mediates interferon signaling, makes two
different proteins by alternative splicing: STAT1-alpha, which contains a TAZ2binding domain, and STAT1-beta, which lacks that domain and may act as a
negative regulator of STAT1-alpha (Bromberg et al. 1996). The TGD provided
teleosts with duplicate stat1 genes, and, due to reciprocal mutations that affect
transcript splicing, the zebrafish stat1a gene came to encode a protein that contains
the TAZ2-binding domain like the human STAT1-alpha isoform, while
stat1b evolved to encode a protein that lacks this domain like STAT1-beta (Song
et al. 2011). Thus, due to subfunctionalization of splicing signals, two genes in
EC
717
CO
RR
716
I. Braasch and J. H. Postlethwait
UN
Editor Proof
364
Book ISBN: 978-3-642-31441-4
Page: 364/383
Layout: T1 Standard SC
Chapter No.: 17
Book ISBN: 978-3-642-31441-4
Page: 365/383
Polyploidy in Fish and the Teleost Genome Duplication
365
772
17.3.3.2 Neofunctionalization
768
769
770
773
774
775
776
777
778
779
780
781
782
783
784
785
786
787
788
789
790
791
792
793
794
795
796
797
798
799
800
801
802
PR
OO
767
In the race between nonfunctionalization, subfunctionalization, and neofunctionalization, nonfunctionalization generally rides the fastest horse. Subfunctionalization
appears to place second, and neofunctionalization—the preservation of ohnologs by
the origin of new functions (Ohno 1970)—shows in third. As with subfunctionalization, neofunctionalization can occur at the level of either gene regulation or protein
structure. Preservation by neofunctionalization differs from subfunctionalization in
two main ways: first, it involves positive selection rather than neutral events, and
second, it involves the origin of gene subfunctions that did not exist in the ancestral
gene rather than the partitioning of ancestral subfunctions. Genome-wide studies
suggest that the acquisition of novel protein domains might have occurred in a quarter
of duplicate pairs after the TGD and happened more frequently in duplicates than in
single-copy genes (Kassahn et al. 2009). Although several specific cases of
neofunctionalization after the TGD have been suggested (Yao and Ge 2010; Sha et al.
2008; Howarth et al. 2008), conclusions are often difficult due to assumptions about the
ancestral state, which is usually based on tetrapods, species that are far removed from
the teleost lineage.
To distinguish neofunctionalization from subfunctionalization, one must
accurately identify the ancestral state. Inferences about the ancestral state can be
difficult with WGD but are often easier with tandem duplications, which have led
to neofunctionalization of vitellogenin and aquaporin genes in teleost egg hydration (Finn and Kristoffersen 2007; Zapater et al. 2011; Cerda 2009), the evolution
of antifreeze proteins by Antarctic teleosts (Deng et al. 2010), and the preservation
of fatty acid-binding protein genes (Karanth et al. 2009).
Several examples of neofunctionalization after the TGD have been suggested.
(1) Phosphoglucose isomerase (PGI) ohnologs from the TGD, after the partitioning
of tissue-specific regulatory elements for expression, diverged so that the copy
expressed in organs of the body cavity became more negative in charge while the
muscle isoform became more positively charged, although the functional significance of these changes is not yet well understood (Sato and Nishida 2007). (2)
After the TGD, cyprinid fish, like zebrafish, retained only one copy of the
D
766
TE
765
EC
763
764
CO
RR
762
F
771
zebrafish encode two protein variants that are encoded by a single gene in human.
Another example is the synapsin gene in fugu (Yu et al. 2003).
Alternative start sites can also result in alternative exon usage. The human
MITF gene controls pigment production both in melanocytes and in the pigmented
retina, with different transcription start sites employed by the two different
cell types (Udono et al. 2000). Zebrafish and other teleosts have two MITF
co-orthologs, with mitfa expressed in melanophores and mitfb in the pigmented
retina; correspondingly, evolution of the two ohnologs resulted in mitfa using one
transcription start site and mitfb using the other (Lister et al. 2001; Altschmied
et al. 2002). Again, two different tissue-specific proteins encoded by one gene in
mammals are formed from sister ohnologs in teleosts due to subfunctionalization.
761
UN
Editor Proof
17
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
810
811
812
813
814
815
816
817
818
819
820
821
822
823
824
825
826
827
828
829
830
831
832
833
834
835
836
837
838
839
840
841
842
843
844
F
809
PR
OO
807
808
D
806
TE
805
androgen receptor gene (ar), but percomorph fish, like pufferfish and medaka,
retained both ohnologs, one of which (arb) experienced substantial sequence
alterations that are consistent with neofunctionalization; further, these changes
occurred after the divergence of basal teleost lineages, including eels, but before
the radiation of percomorphs (Douard et al. 2008). (3) Some TGD duplicates show
elevated evolutionary rates, like csf1rb and its neighbor pdgfrbb in cichlids, which
could be due either to neofunctionalization or to relaxed selection; it has been
suggested that such behavior could be a causal factor in the evolution of the
spectacular coloration patterns of some teleost fish (Braasch et al. 2006). To learn
whether neofunctionalization is at play in these examples and many others, the
duplicates must be compared to each other by function in addition to sequence and
compared to the structure and function of the ancestral genes in a rayfin fish that
diverged before the TGD.
In two cases of neofunctionalization—a hox gene and a sodium channel gene—
TGD duplicates have been investigated functionally in a phylogenetic context that
leaves little doubt about the functions of the preduplication gene. Zebrafish has coorthologs of Hoxa13 derived from the TGD (Amores et al. 1998), and, as determined
by a broad phylogenetic study, these ohnologs began to diverge asymmetrically long
after their initial preservation subsequent to the TGD (Crow et al. 2009). Ohnolog
knockdown experiments in zebrafish, coupled with rescue experiments, showed that
hoxa13a, the faster evolving ohnolog, contains many derived a clade-specific amino
acid replacements that are necessary for development of the yolk sac extension (the
hind yolk), a derived feature in zebrafish and its relatives (Crow et al. 2009).
A great example of neofunctionalization after the TGD occurred in electric fish.
The distantly related African elephantfish (Mormyridae; Osteoglossiformes) and
South American knifefish (Gymnotideae; Gymnotiformes) independently evolved a
muscle-derived electric organ, which facilitates electrical communication and
utilizes the voltage-gated sodium channel subunit Scn4a. Teleosts have two
scn4a genes from the TGD, both expressed in skeletal muscle in most teleosts; in
two lineages of electric fish, however, scn4aa independently lost its expression in
skeletal muscle and gained expression in the electric organ (Arnegard et al. 2010).
Phylogenetic analyses showed that this neofunctionalization event occurred more
than 100 million years after the TGD with a signal of positive selection (Arnegard
et al. 2010). Meanwhile, scn4ab maintained the ancestral expression in skeletal
muscle. These results for Scn4a and Hoxa13 both show that, long after the preservation of ohnologs by subfunctionalization, one copy can still provide the genetic
variation necessary to lead to the innovation of new morphologies and functions.
EC
804
CO
RR
803
I. Braasch and J. H. Postlethwait
17.3.3.3 Spotted Gar as a Rayfin Outgroup for Functional Studies of TGD
Paralogs
UN
Editor Proof
366
Book ISBN: 978-3-642-31441-4
Page: 366/383
As pointed out above, knowledge of the functions of the preduplication gene is
essential for inferring subfunctionalization versus neofunctionalization. Investigators often infer the properties of the pre-TGD gene from the phenotype of well-
Layout: T1 Standard SC
Chapter No.: 17
851
852
853
854
855
856
857
858
859
860
861
862
863
864
865
866
867
868
869
870
871
872
873
874
875
876
877
878
879
880
881
882
883
884
F
850
PR
OO
849
D
848
studied tetrapods, especially mouse. The tetrapod lineage, however, diverged from
the teleost lineage about 450 million years ago and involved the evolution of
substantial differences in morphologies from ancestral bony fish. In addition, rayfin
fish followed an independent lineage for about 200 million years before the TGD;
thus, what appears to involve neofunctionalization when comparing teleost
co-orthologs to tetrapod genes (Fig. 17.8a) could instead be due to the tetrapod
lineage having lost a subfunction originally present in the single-copy gene in the
last common ancestor of teleosts and tetrapods (Fig. 17.8b). A third alternative is
the reciprocal loss of ancestral VGD ohnologs, which could also mimic neofunctionalization, pointing out the importance of careful distinction between
orthologs and ohnologs (Fig. 17.8c). Finally, the novel function thought to have
arisen after the TGD might have actually originated in ancestral rayfin fish,
pointing out the importance of identifying a fish representing this node
(Fig. 17.8d). In addition, distinguishing among the possibilities shown in
Fig. 17.8b–d requires examination of an outgroup diverging basal to the diagrammed tree, such as a cartilaginous fish.
Recently, the spotted gar L. oculatus has been identified as an unduplicated
rayfin outgroup to the TGD (see Sect. 17.3.1.5 and Amores et al. 2011). The
spotted gar occupies a lineage of ‘ancient fish’ that includes not only lepisosteids
(gars), but also bowfin (Amia calva) and possibly acipenseriforms (sturgeons, e.g.
Acipenser transmontanus, and paddlefish, e.g. Polyodon spathula) (Inoue et al.
2003; Kikugawa et al. 2004; and Fig. 17.1). Among these ‘ancient fish’, spotted
gar is the species that has the smallest genome and can most readily be fertilized in
vitro, providing embryos that are amenable to in situ hybridization studies and that
grow to adults in laboratory aquaria with little care. Furthermore, no lineagespecific polyplodization has occurred in gar, in contrast to acipenseriforms. Thus,
the spotted gar makes an ideal outgroup for the investigation of the mechanisms of
evolution of gene function after GD.
TE
847
367
EC
846
CO
RR
845
Book ISBN: 978-3-642-31441-4
Page: 367/383
Polyploidy in Fish and the Teleost Genome Duplication
17.3.4 Mind the Gap: The TGD and the Teleost Radiation—Is
There a Connection?
The preceding paragraphs demonstrate that rayfin species experienced many WGD
events and that teleosts are the most species-rich group of vertebrates. Importantly,
many lineages experiencing WGD, such as barbs, armored catfish, and salmonids,
appear to be particularly species-rich (Le Comber and Smith 2004). These considerations raise the question of whether WGD facilitates lineage diversification in
fish. Ever since its discovery, the TGD has been suggested to have had a major
impact on the radiation and biodiversity of the teleost lineage as a whole (Amores
et al. 1998; Wittbrodt et al. 1998; Meyer and Schartl 1999; Taylor et al. 2001,
2003; Postlethwait et al. 2004; Volff 2005; Meyer and Van de Peer 2005;
Froschauer et al. 2006; Ravi and Venkatesh 2008; Volff et al. 2011; Christoffels
UN
Editor Proof
17
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 368/383
I. Braasch and J. H. Postlethwait
(a) Neofunctionalization and
(b) Subfunction loss in tetrapods;
subfunctionalization after TGD
g1
at
ele
os
t
g1
bt
ele
os
t
g1
pre
-TG
D
ray
G1
fin
tet
rap
od
s
g1
at
ele
os
t
g1
bt
ele
os
t
g1
pre
-TG
D
ray
G1
fin
tet
rap
od
s
subfunctionalization
sub
sub
TGD
loss
TGD
(c) Reciprocal ohnologs gone missing;
(d) Origin of new function in ancestral
rayfin; subfunctionalization after TGD
s
ray
tet
rap
od
-TG
D
G1
g1
D
pre
os
ele
at
g1
TGD
sub
TE
VGD
ohnologs
t
t
os
ele
rap
tet
G1
reciprocal
OGM
g1
s
od
-TG
D
pre
os
sub
g1
ele
bt
g1
g1
at
ele
os
t
t
ray
fin
fin
subfunctionalization after TGD
TGD
PR
OO
F
neo
bt
Editor Proof
368
new
886
887
888
889
890
891
et al. 2004; Van de Peer et al. 2009), but likewise, this has been questioned by
others based on the fossil record and speciation rate analyses (Donoghue and
Purnell 2005; Hurley et al. 2007; Santini et al. 2009).
UN
885
CO
RR
EC
Fig. 17.8 Functional analysis of a pre-TGD diverging rayfin fish is important to distinguish
neofunctionalization from subfunctionalization. a Preservation of one ohnolog by neofunctionalization (the function symbolized by a star) and preservation of the other by subfunctionalization. b Preservation of teleost ohnologs by subfunctionalization; the ‘star’ function was
ancestral, not newly originated after the TGD, but was lost in the tetrapod lineage, as
demonstrated by the pre-TGD-diverging rayfin outgroup. c The ‘star’ function that appears to be
newly originated after the TGD was ancestral in a VGD ohnolog that was lost in the tetrapod
lineage, while the rayfin lineage reciprocally lost the other ohnolog. d. A novel function not
present in the ancestral gene could have originated in stem rayfins and then have been lost in one
TGD ohnolog. Distinguishing among b, c and d requires an additional outgroup, such as a
cartilaginous fish
17.3.4.1 Testing Speciation Models in the Fish World
Genetic principles argue that a WGD event that is followed by population isolation, lineage-specific reciprocal nonfunctionalization events, and later hybridization would tend to promote speciation (Lynch and Force 2000; Werth and
Layout: T1 Standard SC
Chapter No.: 17
369
ancestral population AA
whole genome duplication
ohnolog divergence
Aa Aa ; Ab Ab
lineage divergence
reciprocal nonfunctionalization
Aa Aa ; A- A-
A- A- ; Ab Ab
populations hybridize
Aa A- ; Ab Ahybrids mate
9 Aa - ; Ab survives
F
Polyploidy in Fish and the Teleost Genome Duplication
Fig. 17.9 How WGD could
facilitate lineage
diversification. ‘A’ indicates
the original gene. ‘Aa’ and
‘Ab’ indicate functional but
diverged ohnologs. ‘A-’
indicates a nonfunctionalized allele. A ‘-’
indicates an allele that could
be either functional or
nonfunctional. The analysis
assumes that non-functional
alleles are recessive to
functional alleles
Editor Proof
Book ISBN: 978-3-642-31441-4
Page: 369/383
PR
OO
17
Book ID: 272454_1_En
Date: 16-8-2012
3 Aa - ; A- Asurvives
3 A- A- ; Ab survives
1 A- A- ; A- Alethal
915
17.3.4.2 TGD-Based Morphological Evolution in Teleosts
898
899
900
901
902
903
904
905
906
907
908
909
910
911
912
913
916
917
918
919
TE
897
EC
896
CO
RR
895
UN
894
D
914
Windham 1991). As Fig. 17.9 shows, the fertility of hybrids would be expected to
be somewhat reduced if even a single duplicated gene had reciprocal nonfunctionalization, but if reciprocal nonfunctionalization (or reciprocal gene loss)
occurred at least once on several or many chromosomes, which is likely, then
hybrids would have greatly reduced fertility, thus strengthening lineage
divergence.
Regarding the TGD, while one study estimated *7 % reciprocal gene loss rate
between zebrafish and Tetraodon, which would amount to *1,700 genes (Semon
and Wolfe 2007b), more recent work including more teleost species found little
support for reciprocal gene loss coinciding with speciation events in teleosts
(Kassahn et al. 2009). The latter is more in line with our observation that differences in asymmetric gene loss from paralogons are rare between zebrafish and
stickleback (see Sect. 17.3.2.4).
Similar to reciprocal nonfunctionalization, speciation after WGD may also be
promoted by reciprocal subfunctionalization that results in genomic incompatibilities in post-WGD hybrids (Lynch and Force 2000; Werth and Windham 1991).
Differential functional evolution of TGD paralogs in divergent teleost lineages is
being found in more and more cases (see below), yet associations with genetic
incompatibilities and speciation in teleosts remain to be shown.
Finally, it has also been suggested that the TGD may have caused reduced
probability of extinction in teleosts (Crow and Wagner Crow and Wagner 2006).
The extinction rate of teleost lineages, however, is apparently higher than in nonteleost rayfin lineages (Santini et al. 2009).
892
893
Besides their species richness, teleosts are also extremely diverse in terms of
morphology, physiology, behavior, ecology, and biogeography. If the TGD laid
the groundwork for divergence in such traits, one would expect to find differential
function of TGD paralogs in various teleost lineages. Testing this proposition
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
928
929
930
931
932
933
934
935
936
937
938
939
940
941
942
943
944
945
946
947
948
949
950
951
952
953
954
955
956
957
958
959
960
961
962
F
926
927
PR
OO
924
925
D
923
TE
922
involves experimentally challenging studies in different fish species, and only a
handful of examples are currently available. For teleost sox9 co-orthologs, several
ancient expression domains are differentially retained in zebrafish, stickleback, and
medaka, indicating lineage-specific subfunctionalization (Cresko et al. 2003;
Kluver et al. 2005; Yokoi et al. 2009); similar lineage-specific partitioning of
expression domains have been reported for fgf8 (Jovelin et al. 2007) and tyrp1
(Braasch et al. 2009b) co-orthologs.
A better way to compare gene functions across teleosts is to compare mutant
phenotypes. Unfortunately, few genes have yet been mutated in teleosts other than
zebrafish and medaka, and still fewer have been mutated in both species. Most
teleosts have two TGD ohnologs encoding FGF receptor-1; in zebrafish and carp,
both cyprinids, mutation of fgfr1a causes a scaleless phenotype (Rohner et al.
2009). In contrast, mutation of fgfr1a in medaka, a perciform fish, deletes the trunk
and tail (the headfish mutant) (Yokoi et al. 2007; Shimada et al. 2008). This
finding shows that subfunction partitioning in fgfr1 ohnologs occurred after the
divergence of medaka and zebrafish lineages. Such divergence in subfunction
partitioning after the TGD may provide the genomic basis for some morphological
divergence in teleosts.
And what about the TGD and morphological complexity? The identification of
the seven hox clusters in teleosts, generally not considered to be more complex
than other vertebrate groups, called into question the idea that morphological
complexity along the body axis and the number of hox clusters and genes may be
directly linked (Amores et al. 1998; Prince 2002). The high retention rate of TGD
paralogs for developmentally important genes (Brunet et al. 2006; Kassahn et al.
2009), however, argues for an involvement of at least some TGD paralogs in
teleost morphological evolution. Also, some aspects of teleost physiology and
morphology are unique among vertebrates. To these belong, for example, the
complexity and diversity of teleost body pigmentation and color patterning
(Braasch et al. 2008); some functional modules of pigment cell development and
differentiation have been retained in large part in duplicate in teleosts after the
TGD (Braasch et al. 2009a).
Another question that has so far been rarely addressed but will need to be
investigated in more detail is the possible involvement of TGD duplicates in
morphological and physiological novelties. For example, is functional divergence
of TGD paralogs causing some of the synapomorphies of the teleost lineage
(de Pinna 1996), such as the truly symmetric (homocercal) tail fin? Or are
functional shifts in TGD paralogs involved in lineage-specific key innovations?
We met one intriguing case earlier, the parallel, independent gain of electric
organs used for electrical communication in knifefish and elephantfish by
neofunctionalization of the scn4aa paralog (Arnegard et al. 2010). A more
comprehensive picture, however, can only be gathered by studying TGD paralogs
in more than just a few teleost representatives and adding an appropriate rayfin
outgroup to the analyses.
EC
921
CO
RR
920
I. Braasch and J. H. Postlethwait
UN
Editor Proof
370
Book ISBN: 978-3-642-31441-4
Page: 370/383
Layout: T1 Standard SC
Chapter No.: 17
963
Book ISBN: 978-3-642-31441-4
Page: 371/383
Polyploidy in Fish and the Teleost Genome Duplication
371
17.3.4.3 The Significance of the TGD for the Teleost Radiation
996
17.4 Outlook
971
972
973
974
975
976
977
978
979
980
981
982
983
984
985
986
987
988
989
990
991
992
993
994
997
998
999
1000
1001
1002
PR
OO
970
D
969
TE
968
EC
966
967
CO
RR
965
F
995
Based on the fossil record, a major temporal delay of about 150 million years
separated the TGD event and the major radiation of the teleost lineage in the
percomorphs (Donoghue and Purnell 2005; Hurley et al. 2007). Recent diversification rate shift analyses lead to the recognition that three incidents of accelerated
speciation rates occurred within rayfin fish: on the branches leading (1) to the
teleosts, (2) to the ostariophysians (zebrafish and relatives), and (3) to the percomorphs (perch-related fish, including medaka, stickleback, and pufferfish),
respectively (Santini et al. 2009; Alfaro et al. 2009). Diversification rate shifts
shortly after the TGD accounted for about 10 % of species diversity in teleosts
(Santini et al. 2009). As noted above, however, TGD duplicates can alter their
functions hundreds of millions of years after the GD itself. Whether individual fish
taxa with additional WGD events (salmonids, barbs, etc.) are actually more species
rich than diploid taxa has not been critically tested.
For plants, it has recently been shown that recent polyploids had lower rates of
speciation and higher rates of extinction than diploid control lineages (Mayrose
et al. 2011). In contrast to recent polyploids, the number of ancient polyploidization events that provided today’s flora is higher than expected if diversification
rates of diploids and polyploids were equal (Mayrose et al. 2011). Furthermore,
ancient WGDs appear to be associated with several major radiations in the
angiosperms (Soltis et al. 2009). In contrast, the ratio of species death rate
(extinction) to species birth rate is 1.5 times higher in teleosts than in non-teleost
rayfin fish (Santini et al. 2009). Thus, crucial questions are, when and under which
circumstances do we expect a polyploidization event to deploy its full potential for
lineage diversification and radiation? Given the deep influence of the TGD on
genome evolution in teleosts illustrated in this chapter and given the described
potential for ongoing non-, sub-, and neofunctionalization long after the TGD
event, its major impact may not be apparent immediately after the GD event.
Obviously, the TGD event was not an evolutionary dead-end, but quite the
contrary should be considered a genomic exaptation that generated countless
‘‘spandrels’’ (Gould and Lewontin 1979) in the teleost genome upon which
selection could act long after the actual polyploidization event, when ‘‘opportunity
meets preparation’’.
964
As we have seen—although recognized since early studies of vertebrate genomes—the apparent propensity for polyploidization in the rayfin fish lineage, and
among them, of cyprinid teleosts in particular, still requires an explanation. Since
the first genomic evidence was published in 1998, the TGD hypothesis has
overcome initial criticism, and the TGD is so far the best-studied GD in ‘‘fish’’. We
are confident that many questions regarding the evolution of genome structure,
UN
Editor Proof
17
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
I. Braasch and J. H. Postlethwait
CO
RR
EC
TE
D
PR
OO
F
Editor Proof
372
Book ISBN: 978-3-642-31441-4
Page: 372/383
Fig. 17.10 The cover of Susumo Ohno’s book (1970) underlines the importance of fish for
understanding our own past
1004
1005
1006
1007
1008
1009
1010
1011
1012
1013
gene families, and regulatory circuitry after the TGD can be solved by adding the
sequences of basal rayfins (such as the spotted gar) and of basal teleosts to the
genomic pond. The even harder question about the significance of the TGD for the
teleost radiation will require extensive work on gene function evolution at all
levels of teleost diversity, including numerous non-model fish species. The advent
of next-generation sequencing techniques and the application of developmental
biology studies to an increasing arsenal of fish species are promising for the
acceleration of important research opportunities in this direction. Yet other
questions, for example whether the TGD was an auto- or an allopolyploidization
event, may never be solved due to the millions of years that elapsed since the
event.
UN
1003
Layout: T1 Standard SC
Chapter No.: 17
Book ISBN: 978-3-642-31441-4
Page: 373/383
Polyploidy in Fish and the Teleost Genome Duplication
373
1024
1025
1026
1027
Acknowledgments We would like to thank Cristian Canestro for extensive discussions as well
as Irene Pala for discussions of gene expression in the calandino. This work was supported by a
grant from the Volkswagen Foundation Germany (IB) and National Institutes of Health grant R01
RR020833 (JHP).
1028
References
1029
1030
1031
1032
1033
1034
1035
1036
1037
1038
1039
1040
1041
1042
1043
1044
1045
1046
1047
1048
1049
1050
1051
1052
1053
1054
1055
1056
1057
1058
Alfaro ME, Santini F, Brock C, Alamillo H, Dornburg A, Rabosky DL, Carnevale G, Harmon LJ
(2009) Nine exceptional radiations plus high turnover explain species diversity in jawed
vertebrates. Proc Nat Acad Sci USA 106(32):13410–13414. doi:10.1073/pnas.0811087106
0811087106 [pii]
Allendorf FW, Thorgaard GH (1984) Tetraploidy and the evolution of salmonid fishes. In: Turner
BT (ed) Evolutionary genetics of fishes. Plenum Press, New York, pp 1–53
Altschmied J, Delfgaauw J, Wilde B, Duschl J, Bouneau L, Volff JN, Schartl M (2002)
Subfunctionalization of duplicate mitf genes associated with differential degeneration of
alternative exons in fish. Genetics 161(1):259–267
Alves MJ, Coelho MM, Collares-Pereira MJ (2001) Evolution in action through hybridisation and
polyploidy in an Iberian freshwater fish: a genetic review. Genetica 111(1–3):375–385
Amores A, Force A, Yan YL, Joly L, Amemiya C, Fritz A, Ho RK, Langeland J, Prince V, Wang
YL, Westerfield M, Ekker M, Postlethwait JH (1998) Zebrafish hox clusters and vertebrate
genome evolution. Science 282(5394):1711–1714
Amores A, Catchen J, Ferrara A, Fontenot Q, Postlethwait JH (2011) Genome evolution and meiotic
maps by massively parallel DNA sequencing: spotted gar, an outgroup for the teleost genome
duplication. Genetics 188(4):799–808. doi:10.1534/genetics.111.127324 188/4/799 [pii]
Aparicio S, Hawker K, Cottage A, Mikawa Y, Zuo L, Venkatesh B, Chen E, Krumlauf R,
Brenner S (1997) Organization of the Fugu rubripes hox clusters: evidence for continuing
evolution of vertebrate hox complexes. Nat Genet 16(1):79–83. doi:10.1038/ng0597-79
Aparicio S, Chapman J, Stupka E, Putnam N, Chia JM, Dehal P, Christoffels A, Rash S, Hoon S,
Smit A, Gelpke MD, Roach J, Oh T, Ho IY, Wong M, Detter C, Verhoef F, Predki P, Tay A,
Lucas S, Richardson P, Smith SF, Clark MS, Edwards YJ, Doggett N, Zharkikh A, Tavtigian
SV, Pruss D, Barnstead M, Evans C, Baden H, Powell J, Glusman G, Rowen L, Hood L, Tan
YH, Elgar G, Hawkins T, Venkatesh B, Rokhsar D, Brenner S (2002) Whole-genome shotgun
assembly and analysis of the genome of Fugu rubripes. Science 297(5585):1301–1310.
doi:10.1126/science.1072104 1072104[pii]
Arnegard ME, Zwickl DJ, Lu Y, Zakon HH (2010) Old gene duplication facilitates origin and
diversification of an innovative communication system-twice. P Nat Acad Sci USA
107(51):22172–22177. doi:10.1073/Pnas.1011803107
1021
1022
PR
OO
1019
1020
D
1018
TE
1017
EC
1016
CO
RR
1015
F
1023
Studying other, more recent GDs in rayfins, such as the salmonid and carp
polyloidizations, will help to reveal in more detail the immediate epigenetic and
genomic changes that follow a polyploidization. Luckily, the multitude and distribution of piscine GD events over evolutionary time will contribute to studies of
the gradual evolution of vertebrate animals after different types of GD.
Finally, as illustrated so marvelously by the cover of Ohno’s book (Ohno 1970)
(Fig. 17.10), the study of polyploidizations in fish will not only allow us to better
understand our own fishy heritage, but importantly, will help to inform us about
the evolutionary processes that have shaped the vertebrate genome after VGD1
and VGD2, which had such a profound influence on our own, human evolution.
1014
UN
Editor Proof
17
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
EC
TE
D
PR
OO
F
Balon EK (2004) About the oldest domesticates among fishes. J Fish Biol 65:1–27. doi:10.1111/
j.1095-8649.2004.00563.x
Birstein VJ, Hanner R, DeSalle R (1997) Phylogeny of the Acipenseriformes: cytogenetic and
molecular approaches. Environ Biol Fish 48(1–4):127–156
Boldajipour B, Doitsidou M, Tarbashevich K, Laguri C, Yu SR, Ries J, Dumstrei K, Thelen S,
Dorries J, Messerschmidt EM, Thelen M, Schwille P, Brand M, Lortat-Jacob H, Raz E (2011)
Cxcl12 evolution—subfunctionalization of a ligand through altered interaction with the
chemokine receptor. Development 138(14):2909–2914. doi:10.1242/Dev.068379
Braasch I, Salzburger W, Meyer A (2006) Asymmetric evolution in two fish-specifically
duplicated receptor tyrosine kinase paralogons involved in teleost coloration. Mol Biol Evol
23(6):1192–1202. doi:10.1093/molbev/msk003 msk003 [pii]
Braasch I, Schartl M, Volff JN (2007) Evolution of pigment synthesis pathways by gene and
genome duplication in fish. BMC Evol Biol 7:74. doi:10.1186/1471-2148-7-74 1471-2148-774 [pii]
Braasch I, Volff JN, Schartl M (2008) The evolution of teleost pigmentation and the fish-specific
genome duplication. J Fish Biol 73(8):1891–1918. doi:10.1111/J.1095-8649.2008.02011.X
Braasch I, Brunet F, Volff JN, Schartl M (2009a) Pigmentation pathway evolution after wholegenome duplication in fish. Genome Biol Evol 1:479–493. doi:10.1093/gbe/evp050
Braasch I, Liedtke D, Volff JN, Schartl M (2009b) Pigmentary function and evolution of tyrp1
gene duplicates in fish. Pigment Cell Melanoma Res 22(6):839–850. doi:10.1111/j.1755148X.2009.00614.x PCR614 [pii]
Bromberg JF, Horvath CM, Wen ZL, Schreiber RD, Darnell JE (1996) Transcriptionally active
Stat1 is required for the antiproliferative effects of both interferon alpha and interferon
gamma. P Nat Acad Sci USA 93(15):7673–7678
Brunet FG, Crollius HR, Paris M, Aury JM, Gibert P, Jaillon O, Laudet V, Robinson-Rechavi M
(2006) Gene loss and evolutionary rates following whole-genome duplication in teleost fishes.
Mol Biol Evol 23(9):1808–1816. doi:10.1093/Molbev/Mls049
Bu L, Bergthorsson U, Katju V (2011) Local synteny and codon usage contribute to asymmetric
sequence divergence of Saccharomyces cerevisiae gene duplicates. BMC Evol Biol 11:279.
doi:10.1186/1471-2148-11-279 1471-2148-11-279 [pii]
Canestro C, Yokoi H, Postlethwait JH (2007) Evolutionary developmental biology and genomics.
Nat Rev Genet 8(12):932–942. doi:10.1038/nrg2226 nrg2226 [pii]
Canestro C, Catchen JM, Rodriguez-Mari A, Yokoi H, Postlethwait JH (2009) Consequences of
lineage-specific gene loss on functional evolution of surviving paralogs: ALDH1A and
retinoic acid signaling in vertebrate genomes. PLoS Genet 5(5):e1000496. doi:10.1371/
journal.pgen.1000496
Catchen JM, Conery JS, Postlethwait JH (2009) Automated identification of conserved synteny
after whole-genome duplication. Genome Res 19(8):1497–1505. doi:10.1101/Gr.090480.108
Catchen JM, Braasch I, Postlethwait JH (2011) Conserved synteny and the zebrafish genome.
Method Cell Biol 104:259–285. doi:10.1016/B978-0-12-374814-0.00015-X
Cerda J (2009) Molecular pathways during marine fish egg hydration: the role of aquaporins.
J Fish Biol 75(9):2175–2196. doi:10.1111/j.1095-8649.2009.02397.x JFB2397 [pii]
Chenuil A, Galtier N, Berrebi P (1999) A test of the hypothesis of an autopolyploid vs.
allopolyploid origin for a tetraploid lineage: application to the genus Barbus (Cyprinidae).
Heredity (Edinb) 82(Pt 4):373–380. doi:her489 [pii]
Chiu CH, Dewar K, Wagner GP, Takahashi K, Ruddle F, Ledje C, Bartsch P, Scemama JL,
Stellwag E, Fried C, Prohaska SJ, Stadler PF, Amemiya CT (2004) Bichir HoxA cluster
sequence reveals surprising trends in ray-finned fish genomic evolution. Genome Res
14(1):11–17. doi:10.1101/gr.1712904 14/1/11[pii]
Christoffels A, Koh EG, Chia JM, Brenner S, Aparicio S, Venkatesh B (2004) Fugu genome
analysis provides evidence for a whole-genome duplication early during the evolution of rayfinned fishes. Mol Biol Evol 21(6):1146–1151. doi:10.1093/molbev/msh114 msh114 [pii]
Comai L (2005) The advantages and disadvantages of being polyploid. Nat Rev Genet 6(11):836–
846. doi:10.1038/nrg1711
CO
RR
1059
1060
1061
1062
1063
1064
1065
1066
1067
1068
1069
1070
1071
1072
1073
1074
1075
1076
1077
1078
1079
1080
1081
1082
1083
1084
1085
1086
1087
1088
1089
1090
1091
1092
1093
1094
1095
1096
1097
1098
1099
1100
1101
1102
1103
1104
1105
1106
1107
1108
1109
1110
1111
1112
I. Braasch and J. H. Postlethwait
UN
Editor Proof
374
Book ISBN: 978-3-642-31441-4
Page: 374/383
Layout: T1 Standard SC
Chapter No.: 17
375
EC
TE
D
PR
OO
F
Cresko WA, Yan YL, Baltrus DA, Amores A, Singer A, Rodriguez-Mari A, Postlethwait JH
(2003) Genome duplication, subfunction partitioning, and lineage divergence: sox9 in
stickleback and zebrafish. Dev Dyn 228(3):480–489. doi:10.1002/dvdy.10424
Crow KD, Wagner GP (2006) Proceedings of the SMBE tri-national young investigators’
workshop. What is the role of genome duplication in the evolution of complexity and
diversity? Mol Biol Evol 23(5):887–892. doi:10.1093/molbev/msj083 msj083 [pii]
Crow KD, Stadler PF, Lynch VJ, Amemiya C, Wagner GP (2006) The ‘‘fish-specific’’ Hox
cluster duplication is coincident with the origin of teleosts. Mol Biol Evol 23(1):121–136.
doi:10.1093/molbev/msj020 msj020 [pii]
Crow KD, Amemiya CT, Roth J, Wagner GP (2009) Hypermutability of HoxA13A and functional
divergence from its paralog are associated with the origin of a novel developmental feature in
zebrafish and related taxa (cypriniformes). Evolution 63(6):1574–1592. doi:10.1111/j.15585646.2009.00657.x EVO657 [pii]
Danzmann RG, Davidson EA, Ferguson MM, Gharbi K, Koop BF, Hoyheim B, Lien S,
Lubieniecki KP, Moghadam HK, Park J, Phillips RB, Davidson WS (2008) Distribution of
ancestral proto-actinopterygian chromosome arms within the genomes of 4R-derivative
salmonid fishes (rainbow trout and Atlantic salmon). BMC Genomics 9:557. doi:10.1186/
1471-2164-9-557
David L, Blum S, Feldman MW, Lavi U, Hillel J (2003) Recent duplication of the, common carp
(Cyprinus carpio L.) genome as revealed by analyses of microsatellite loci. Mol Biol Evol
20(9):1425–1434. doi:10.1093/molbev/msg173
David L, Rothbard S, Rubinstein I, Katzman H, Hulata G, Hillel J, Lavi U (2004) Aspects of red
and black color inheritance in the Japanese ornamental (Koi) carp (Cyprinus carpio L.).
Aquaculture 233(1–4):129–147. doi:10.1016/j.aquaculture.2003.10.033
Davidson WS, Koop BF, Jones SJM, Iturra P, Vidal R, Maass A, Jonassen I, Lien S, Omholt SW
(2010) Sequencing the genome of the Atlantic salmon (Salmo salar). Genome Biol 11(9):403.
doi:10.1186/gb-2010-11-9-403
Davis CA, Holmyard DP, Millen KJ, Joyner AL (1991) Examining pattern formation in mouse,
chicken and frog embryos with an en-specific antiserum. Development 111(2):287–298
de Pinna MCC (1996) Teleostean monophyly. In: Stiassny MLJ, Parenti LR, Johnson GD (eds)
Interrelationships of fishes. Academic, San Diego, pp 147–162
de Souza FS, Bumaschny VF, Low MJ, Rubinstein M (2005) Subfunctionalization of expression
and peptide domains following the ancient duplication of the proopiomelanocortin gene in
teleost fishes. Mol Biol Evol 22(12):2417–2427. doi:10.1093/molbev/msi236 msi236 [pii]
Deng C, Cheng CH, Ye H, He X, Chen L (2010) Evolution of an antifreeze protein by
neofunctionalization under escape from adaptive conflict. Proc Nat Acad Sci USA
107(50):21593–21598. doi:10.1073/pnas.1007883107 1007883107 [pii]
Doitsidou M, Reichman-Fried M, Stebler J, Koprunner M, Dorries J, Meyer D, Esguerra CV,
Leung T, Raz E (2002) Guidance of primordial germ cell migration by the chemokine SDF-1.
Cell 111(5):647–659 S0092867402011352[pii]
Donoghue PC, Purnell MA (2005) Genome duplication, extinction and vertebrate evolution. Trends
Ecol Evol 20(6):312–319. doi:10.1016/j.tree.2005.04.008 S0169-5347(05)00108-4 [pii]
Douard V, Brunet F, Boussau B, Ahrens-Fath I, Vlaeminck-Guillem V, Haendler B, Laudet V,
Guiguen Y (2008) The fate of the duplicated androgen receptor in fishes: a late
neofunctionalization event? BMC Evol Biol 8:336. doi:10.1186/1471-2148-8-336 Artn 336
Eiken HG, Njolstad PR, Molven A, Fjose A (1987) A zebrafish homeobox-containing gene with
embryonic transcription. Biochem Biophys Res Commun 149(3):1165–1171
Elgar G, Clark MS, Meek S, Smith S, Warner S, Edwards YJ, Bouchireb N, Cottage A, Yeo GS,
Umrania Y, Williams G, Brenner S (1999) Generation and analysis of 25 Mb of genomic
DNA from the pufferfish Fugu rubripes by sequence scanning. Genome Res 9(10):960–971
Ferris SD (1984) Tetraploidy and the evolution of the catostomid fishes. In: Turner BT (ed)
Evolutionary genetics of fishes. Plenus Press, New York, pp 54–93
Ferris SD, Whitt GS (1977a) Duplicate gene expression in diploid and tetraploid loaches
(cypriniformes, Cobitidae). Biochem Genet 15(11–12):1097–1112
CO
RR
1113
1114
1115
1116
1117
1118
1119
1120
1121
1122
1123
1124
1125
1126
1127
1128
1129
1130
1131
1132
1133
1134
1135
1136
1137
1138
1139
1140
1141
1142
1143
1144
1145
1146
1147
1148
1149
1150
1151
1152
1153
1154
1155
1156
1157
1158
1159
1160
1161
1162
1163
1164
1165
1166
Book ISBN: 978-3-642-31441-4
Page: 375/383
Polyploidy in Fish and the Teleost Genome Duplication
UN
Editor Proof
17
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
EC
TE
D
PR
OO
F
Ferris SD, Whitt GS (1977b) Evolution of duplicate gene-expression in carp (Cyprinus carpio).
Experientia 33(10):1299–1301
Finn RN, Kristoffersen BA (2007) Vertebrate vitellogenin gene duplication in relation to the ‘‘3R
hypothesis’’: correlation to the pelagic egg and the oceanic radiation of teleosts. PLoS One
2(1):e169. doi:10.1371/journal.pone.0000169
Force A, Lynch M, Pickett FB, Amores A, Yan YL, Postlethwait J (1999) Preservation of
duplicate genes by complementary, degenerative mutations. Genetics 151(4):1531–1545
Froschauer A, Braasch I, Volff JN (2006) Fish genomes, comparative genomics and vertebrate
evolution. Curr Genomics 7(1):43–57. doi:10.2174/138920206776389766
Gardner CA, Barald KF (1992) Expression patterns of engrailed-like proteins in the chick
embryo. Dev Dyn 193(4):370–388. doi:10.1002/aja.1001930410
Gates MA, Kim L, Egan ES, Cardozo T, Sirotkin HI, Dougan ST, Lashkari D, Abagyan R, Schier
AF, Talbot WS (1999) A genetic linkage map for zebrafish: comparative analysis and
localization of genes and expressed sequences. Genome Res 9(4):334–347
Gomez A, Volff JN, Hornung U, Schartl M, Wellbrock C (2004) Identification of a second egfr
gene in Xiphophorus uncovers an expansion of the epidermal growth factor receptor family in
fish. Mol Biol Evol 21(2):266–275. doi:10.1093/molbev/msh017 msh017[pii]
Gould SJ, Lewontin RC (1979) Spandrels of San-Marco and the Panglossian paradigm—a
critique of the adaptationist program. Proc R Soc Lond B Biol Sci 205(1161):581–598
Graham A, Papalopulu N, Krumlauf R (1989) The murine and Drosophila homeobox gene
complexes have common features of organization and expression. Cell 57(3):367–378.
doi:0092-8674(89)90912-4 [pii]
Haussler D, O’Brien SJ, Ryder OA, Barker FK, Clamp M, Crawford AJ, Hanner R, Hanotte O,
Johnson WE, McGuire JA, Miller W, Murphy RW, Murphy WJ, Sheldon FH, Sinervo B,
Venkatesh B, Wiley EO, Allendorf FW, Amato G, Baker CS, Bauer A, Beja-Pereira A,
Bermingham E, Bernardi G, Bonvicino CR, Brenner S, Burke T, Cracraft J, Diekhans M,
Edwards S, Ericson PGP, Estes J, Fjelsda J, Flesness N, Gamble T, Gaubert P, Graphodatsky
AS, Graves JAM, Green ED, Green RE, Hackett S, Hebert P, Helgen KM, Joseph L, Kessing B,
Kingsley DM, Lewin HA, Luikart G, Martelli P, Moreira MAM, Nguyen N, Orti G, Pike BL,
Rawson DM, Schuster SC, Seuanez HN, Shaffer HB, Springer MS, Stuart JM, Sumner J,
Teeling E, Vrijenhoek RC, Ward RD, Warren WC, Wayne R, Williams TM, Wolfe ND, Zhang
YP, Graph-Odatsky A, Johnson WE, Felsenfeld A, Turner S, Scientists GKC, Grp M, Grp B,
Grp AR, Grp F, Grp GP, Grp A (2009) Genome 10 K: a proposal to obtain whole-genome
sequence for 10,000 vertebrate species. J Hered 100(6):659–674. doi:10.1093/jhered/esp086
He X, Zhang J (2005) Rapid subfunctionalization accompanied by prolonged and substantial
neofunctionalization in duplicate gene evolution. Genetics 169(2):1157–1164. doi:10.1534/
genetics.104.037051 genetics.104.037051 [pii]
Hoegg S, Meyer A (2005) Hox clusters as models for vertebrate genome evolution. Trends Genet
21(8):421–424. doi:10.1016/j.tig.2005.06.004 S0168-9525(05)00165-4 [pii]
Hoegg S, Brinkmann H, Taylor JS, Meyer A (2004) Phylogenetic timing of the fish-specific
genome duplication correlates with the diversification of teleost fish. J Mol Evol 59(2):190–
203. doi:10.1007/s00239-004-2613-z
Hoegg S, Boore JL, Kuehl JV, Meyer A (2007) Comparative phylogenomic analyses of teleost
fish hox gene clusters: lessons from the cichlid fish Astatotilapia burtoni. BMC Genomics
8:317. doi:10.1186/1471-2164-8-317 1471-2164-8-317 [pii]
Howarth DL, Law SHW, Barnes B, Hall JM, Hinton DE, Moore L, Maglich JM, Moore JT,
Kullman SW (2008) Paralogous vitamin D receptors in teleosts: transition of nuclear receptor
function. Endocrinology 149(5):2411–2422. doi:10.1210/En.2007-1256
Hufton AL, Groth D, Vingron M, Lehrach H, Poustka AJ, Panopoulou G (2008) Early vertebrate
whole genome duplications were predated by a period of intense genome rearrangement.
Genome Res 18(10):1582–1591. doi:10.1101/gr.080119.108 gr.080119.108 [pii]
Huminiecki L, Heldin CH (2010) 2R and remodeling of vertebrate signal transduction engine.
BMC Biol 8:146. doi:10.1186/1741-7007-8-146 1741-7007-8-146 [pii]
CO
RR
1167
1168
1169
1170
1171
1172
1173
1174
1175
1176
1177
1178
1179
1180
1181
1182
1183
1184
1185
1186
1187
1188
1189
1190
1191
1192
1193
1194
1195
1196
1197
1198
1199
1200
1201
1202
1203
1204
1205
1206
1207
1208
1209
1210
1211
1212
1213
1214
1215
1216
1217
1218
1219
I. Braasch and J. H. Postlethwait
UN
Editor Proof
376
Book ISBN: 978-3-642-31441-4
Page: 376/383
Layout: T1 Standard SC
Chapter No.: 17
377
EC
TE
D
PR
OO
F
Hurley IA, Mueller RL, Dunn KA, Schmidt EJ, Friedman M, Ho RK, Prince VE, Yang Z,
Thomas MG, Coates MI (2007) A new time-scale for ray-finned fish evolution. Proc Biol Sci
274(1609):489–498
Inoue JG, Miya M, Tsukamoto K, Nishida M (2003) Basal actinopterygian relationships: a
mitogenomic perspective on the phylogeny of the ‘‘ancient fish’’. Mol Phylogenet Evol
26(1):110–120
Jaillon O, Aury JM, Brunet F, Petit JL, Stange-Thomann N, Mauceli E, Bouneau L, Fischer C,
Ozouf-Costaz C, Bernot A, Nicaud S, Jaffe D, Fisher S, Lutfalla G, Dossat C, Segurens B,
Dasilva C, Salanoubat M, Levy M, Boudet N, Castellano S, Anthouard V, Jubin C, Castelli V,
Katinka M, Vacherie B, Biemont C, Skalli Z, Cattolico L, Poulain J, De Berardinis V, Cruaud
C, Duprat S, Brottier P, Coutanceau JP, Gouzy J, Parra G, Lardier G, Chapple C, McKernan
KJ, McEwan P, Bosak S, Kellis M, Volff JN, Guigo R, Zody MC, Mesirov J, Lindblad-Toh K,
Birren B, Nusbaum C, Kahn D, Robinson-Rechavi M, Laudet V, Schachter V, Quetier F,
Saurin W, Scarpelli C, Wincker P, Lander ES, Weissenbach J, Roest Crollius H (2004)
Genome duplication in the teleost fish Tetraodon nigroviridis reveals the early vertebrate
proto-karyotype. Nature 431(7011):946–957. doi:10.1038/nature03025 nature03025 [pii]
Jovelin R, He X, Amores A, Yan YL, Shi R, Qin B, Roe B, Cresko WA, Postlethwait JH (2007)
Duplication and divergence of fgf8 functions in teleost development and evolution. J Exp Zool
B Mol Dev Evol 308(6):730–743. doi:10.1002/jez.b.21193
Joyner AL, Martin GR (1987) En-1 and En-2, two mouse genes with sequence homology to the
Drosophila engrailed gene: expression during embryogenesis. Genes Dev 1(1):29–38
Karanth S, Lall SP, Denovan-Wright EM, Wright JM (2009) Differential transcriptional
modulation of duplicated fatty acid-binding protein genes by dietary fatty acids in zebrafish
(Danio rerio): evidence for subfunctionalization or neofunctionalization of duplicated genes.
BMC Evol Biol 9:219. doi:10.1186/1471-2148-9-219 1471-2148-9-219 [pii]
Kasahara M, Naruse K, Sasaki S, Nakatani Y, Qu W, Ahsan B, Yamada T, Nagayasu Y, Doi K,
Kasai Y, Jindo T, Kobayashi D, Shimada A, Toyoda A, Kuroki Y, Fujiyama A, Sasaki T,
Shimizu A, Asakawa S, Shimizu N, Hashimoto S, Yang J, Lee Y, Matsushima K, Sugano S,
Sakaizumi M, Narita T, Ohishi K, Haga S, Ohta F, Nomoto H, Nogata K, Morishita T, Endo
T, Shin IT, Takeda H, Morishita S, Kohara Y (2007) The medaka draft genome and insights
into vertebrate genome evolution. Nature 447(7145):714–719. doi:10.1038/nature05846 nature05846 [pii]
Kassahn KS, Dang VT, Wilkins SJ, Perkins AC, Ragan MA (2009) Evolution of gene function
and regulatory control after whole-genome duplication: comparative analyses in vertebrates.
Genome Res 19(8):1404–1418. doi:10.1101/Gr.086827.108
Kikugawa K, Katoh K, Kuraku S, Sakurai H, Ishida O, Iwabe N, Miyata T (2004) Basal jawed
vertebrate phylogeny inferred from multiple nuclear DNA-coded genes. BMC Biol 2:3.
doi:10.1186/1741-7007-2-3 1741-7007-2-3 [pii]
Kluver N, Kondo M, Herpin A, Mitani H, Schartl M (2005) Divergent expression patterns of Sox9
duplicates in teleosts indicate a lineage specific subfunctionalization. Dev Genes Evol
215(6):297–305. doi:10.1007/s00427-005-0477-x
Kohn M, Hogel J, Vogel W, Minich P, Kehrer-Sawatzki H, Graves JA, Hameister H (2006)
Reconstruction of a 450-My-old ancestral vertebrate protokaryotype. Trends Genet
22(4):203–210. doi:10.1016/j.tig.2006.02.008 S0168-9525(06)00063-1 [pii]
Komiyama T, Kobayashi H, Tateno Y, Inoko H, Gojobori T, Ikeo K (2009) An evolutionary
origin and selection process of goldfish. Gene 430(1–2):5–11. doi:10.1016/j.gene.2008.10.019
S0378-1119(08)00543-X [pii]
Koop BF, von Schalburg KR, Leong J, Walker N, Lieph R, Cooper GA, Robb A, Beetz-Sargent
M, Holt RA, Moore R, Brahmbhatt S, Rosner J, Rexroad CE, McGowan CR, Davidson WS
(2008) A salmonid EST genomic study: genes, duplications, phylogeny and microarrays.
BMC Genomics 9:545. doi:10.1186/1471-2164-9-545
Kupka E (1948) Chromosomale Verschiedenheiten bei schweizerischen Coregonen (Felchen).
Rev Suisse Zool 55:293–295
CO
RR
1220
1221
1222
1223
1224
1225
1226
1227
1228
1229
1230
1231
1232
1233
1234
1235
1236
1237
1238
1239
1240
1241
1242
1243
1244
1245
1246
1247
1248
1249
1250
1251
1252
1253
1254
1255
1256
1257
1258
1259
1260
1261
1262
1263
1264
1265
1266
1267
1268
1269
1270
1271
1272
Book ISBN: 978-3-642-31441-4
Page: 377/383
Polyploidy in Fish and the Teleost Genome Duplication
UN
Editor Proof
17
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
EC
TE
D
PR
OO
F
Larhammar D, Risinger C (1994) Molecular genetic aspects of tetraploidy in the common carp
Cyprinus carpio. Mol Phylogenet Evol 3(1):59–68. doi:10.1006/mpev.1994.1007 S10557903(84)71007-4 [pii]
Le Comber SC, Smith C (2004) Polyploidy in fishes: patterns and processes. Biol J Linn Soc
82(4):431–442
Leggatt RA, Iwama GK (2003) Occurrence of polyploidy in the fishes. Rev Fish Biol Fisher
13(3):237–246
Leong JS, Jantzen SG, von Schalburg KR, Cooper GA, Messmer AM, Liao NY, Munro S, Moore
R, Holt RA, Jones SJM, Davidson WS, Koop BF (2010) Salmo salar and Esox lucius fulllength cDNA sequences reveal changes in evolutionary pressures on a post-tetraploidization
genome. BMC Genomics 11:279. doi:10.1186/1471-2164-11-279
Li YJ, Yu Z, Zhang MZ, Qian C, Abe S, Arai K (2011) The origin of natural tetraploid loach
Misgurnus anguillicaudatus (Teleostei: Cobitidae) inferred from meiotic chromosome
configurations. Genetica 139(6):805–811. doi:10.1007/s10709-011-9585-x
Lien S, Gidskehaug L, Moen T, Hayes BJ, Berg PR, Davidson WS, Omholt SW, Kent MP (2011)
A dense SNP-based linkage map for Atlantic salmon (Salmo salar) reveals extended
chromosome homeologies and striking differences in sex-specific recombination patterns.
BMC Genomics 12(1):615. doi:10.1186/1471-2164-12-615 1471-2164-12-615 [pii]
Lister JA, Close J, Raible DW (2001) Duplicate mitf genes in zebrafish: complementary
expression and conservation of melanogenic potential. Dev Biol 237(2):333–344
Ludwig A, Belfiore NM, Pitra C, Svirsky V, Jenneckens I (2001) Genome duplication events and
functional reduction of ploidy levels in sturgeon (Acipenser, Huso and Scaphirhynchus).
Genetics 158(3):1203–1215
Luo J, Stadler PF, He S, Meyer A (2007) PCR survey of Hox genes in the goldfish Carassius
auratus auratus. J Exp Zool B Mol Dev Evol 308(3):250–258. doi:10.1002/jez.b.21144
Lynch M, Conery JS (2000) The evolutionary fate and consequences of duplicate genes. Science
290(5494):1151–1155 8976 [pii]
Lynch M, Force AG (2000) The origin of interspecific genomic incompatibility via gene
duplication. Am Nat 156(6):590–605
Mable BK (2004) ‘Why polyploidy is rarer in animals than in plants’: myths and mechanisms.
Biol J Linn Soc 82(4):453–466
Mable BK, Alexandrou MA, Taylor MI (2011) Genome duplication in amphibians and fish: an
extended synthesis. J Zool 284(3):151–182. doi:10.1111/j.1469-7998.2011.00829.x
Mank JE, Avise JC (2006a) Cladogenetic correlates of genomic expansions in the recent
evolution of actinopterygiian fishes. Proc R Soc Lond B Biol Sci 273(1582):33–38.
doi:10.1098/rspb.2005.3295
Mank JE, Avise JC (2006b) Phylogenetic conservation of chromosome numbers in actinopterygiian fishes. Genetica 127(1–3):321–327. doi:10.1007/s10709-005-5248-0
Mayrose I, Zhan SH, Rothfels CJ, Magnuson-Ford K, Barker MS, Rieseberg LH, Otto SP (2011)
Recently formed polyploid plants diversify at lower rates. Science 333(6047):1257.
doi:10.1126/Science.1207205
Meyer A (1998) Hox gene variation and evolution. Nature 391 (6664):225, 227–228.
doi:10.1038/34530
Meyer A, Schartl M (1999) Gene and genome duplications in vertebrates: the one-to-four (-toeight in fish) rule and the evolution of novel gene functions. Curr Opin Cell Biol 11(6):699–
704. doi:S0955-0674(99)00039-3 [pii]
Meyer A, Van de Peer Y (2005) From 2R to 3R: evidence for a fish-specific genome duplication
(FSGD). BioEssays 27(9):937–945. doi:10.1002/bies.20293
Miller MR, Brunelli JP, Wheeler PA, Liu S, Rexroad CE 3rd, Palti Y, Doe CQ, Thorgaard GH
(2011) A conserved haplotype controls parallel adaptation in geographically distant salmonid
populations. Mol Ecol. doi:10.1111/j.1365-294X.2011.05305.x
Misof BY, Wagner GP (1996) Evidence for four hox clusters in the killifish Fundulus heteroclitus
(teleostei). Mol Phylogenet Evol 5(2):309–322. doi:10.1006/mpev.1996.0026 S10557903(96)90026-3 [pii]
CO
RR
1273
1274
1275
1276
1277
1278
1279
1280
1281
1282
1283
1284
1285
1286
1287
1288
1289
1290
1291
1292
1293
1294
1295
1296
1297
1298
1299
1300
1301
1302
1303
1304
1305
1306
1307
1308
1309
1310
1311
1312
1313
1314
1315
1316
1317
1318
1319
1320
1321
1322
1323
1324
1325
1326
I. Braasch and J. H. Postlethwait
UN
Editor Proof
378
Book ISBN: 978-3-642-31441-4
Page: 378/383
Layout: T1 Standard SC
Chapter No.: 17
379
EC
TE
D
PR
OO
F
Moghadam HK, Ferguson MM, Danzmann RG (2005a) Evidence for Hox gene duplication in
rainbow trout (Oncorhynchus mykiss): a tetraploid model species. J Mol Evol 61(6):804–818.
doi:10.1007/s00239-004-0230-5
Moghadam HK, Ferguson MM, Danzmann RG (2005b) Evolution of Hox clusters in Salmonidae:
a comparative analysis between Atlantic salmon (Salmo salar) and rainbow trout
(Oncorhynchus mykiss). J Mol Evol 61(5):636–649. doi:10.1007/s00239-004-0338-7
Moghadam HK, Ferguson MM, Danzmann RG (2009) Comparative genomics and evolution of
conserved noncoding elements (CNE) in rainbow trout. BMC Genomics 10:278. doi:10.1186/
1471-2164-10-278 Artn 278
Morizot DC (1990) Use of fish gene maps to predict ancestral vertebrate genome organization. In:
Ogita Z-I, Markert CL (eds) Isozymes: structure, function, and use in biology, and medicine.
Wiley-Liss, Inc., New York, pp 207–234
Muller HJ (1925) Why polyploidy is rarer in animals than in plants. Am Nat 59(663):346–353
Mulley JF, Chiu CH, Holland PW (2006) Breakup of a homeobox cluster after genome
duplication in teleosts. Proc Nat Acad Sci USA 103(27):10369–10372. doi:10.1073/
pnas.0600341103 0600341103 [pii]
Mungpakdee S, Seo HC, Angotzi AR, Dong XJ, Akalin A, Chourrout D (2008a) Differential
evolution of the 13 Atlantic salmon Hox clusters. Mol Biol Evol 25(7):1333–1343.
doi:10.1093/molbev/msn097
Mungpakdee S, Seo HC, Chourrout D (2008b) Spatio-temporal expression patterns of anterior
Hox genes in Atlantic salmon (Salmo salar). Gene Expr Patterns 8(7–8):508–514.
doi:10.1016/j.gep.2008.06.004
Nair S, Schilling TF (2008) Chemokine signaling controls endodermal migration during zebrafish
gastrulation. Science 322(5898):89–92. doi:10.1126/science.1160038 1160038 [pii]
Nakatani Y, Takeda H, Kohara Y, Morishita S (2007) Reconstruction of the vertebrate ancestral
genome reveals dynamic genome reorganization in early vertebrates. Genome Res
17(9):1254–1265. doi:10.1101/gr.6316407
Naruse K, Tanaka M, Mita K, Shima A, Postlethwait J, Mitani H (2004) A medaka gene map: the
trace of ancestral vertebrate proto-chromosomes revealed by comparative gene mapping.
Genome Res 14(5):820–828. doi:10.1101/gr.2004004 2004004 [pii]
Nelson JS (2006) Fishes of the world, 4th edn. Wiley, Hoboken
Njolstad PR, Molven A, Hordvik I, Apold J, Fjose A (1988) Primary structure, developmentally
regulated expression and potential duplication of the zebrafish homeobox gene ZF-21.
Nucleic Acids Res 16(19):9097–9111
Ohno S (1970) Evolution by gene duplication. Springer, Berlin
Ohno S, Muramoto J, Christia L, Atkin NB (1967) Diploid-tetraploid relationship among oldworld members of fish family Cyprinidae. Chromosoma 23(1):1
Ohno S, Wolf U, Atkin NB (1968) Evolution from fish to mammals by gene duplication.
Hereditas-Genetisk A 59(1):169 &
Okuda Y, Yoda H, Uchikawa M, Furutani-Seiki M, Takeda H, Kondoh H, Kamachi Y (2006)
Comparative genomic and expression analysis of group B1 sox genes in zebrafish indicates
their diversification during vertebrate evolution. Dev Dyn 235(3):811–825. doi:10.1002/
dvdy.20678
Oliveira C, Almeidatoledo LF, Mori L, Toledofilho SA (1992) Extensive chromosomal
rearrangements and nuclear-DNA content changes in the evolution of the armored catfishes
genus Corydoras (pisces, siluriformes, callichthyidae). J Fish Biol 40(3):419–431
Otto SP (2007) The evolutionary consequences of polyploidy. Cell 131(3):452–462. doi:10.1016/
j.cell.2007.10.022
Otto SP, Whitton J (2000) Polyploid incidence and evolution. Annu Rev Genet 34:401–437.
doi:10.1146/annurev.genet.34.1.401 34/1/401 [pii]
Pala I, Coelho MM, Schartl M (2008) Dosage compensation by gene-copy silencing in a triploid
hybrid fish. Curr Biol 18(17):1344–1348. doi:10.1016/j.cub.2008.07.096
CO
RR
1327
1328
1329
1330
1331
1332
1333
1334
1335
1336
1337
1338
1339
1340
1341
1342
1343
1344
1345
1346
1347
1348
1349
1350
1351
1352
1353
1354
1355
1356
1357
1358
1359
1360
1361
1362
1363
1364
1365
1366
1367
1368
1369
1370
1371
1372
1373
1374
1375
1376
1377
1378
Book ISBN: 978-3-642-31441-4
Page: 379/383
Polyploidy in Fish and the Teleost Genome Duplication
UN
Editor Proof
17
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
EC
TE
D
PR
OO
F
Pala I, Schartl M, Brito M, Vacas JM, Coelho MM (2010) Gene expression regulation and lineage
evolution: the north and south tale of the hybrid polyploid Squalius alburnoides complex.
Proc R Soc B Biol Sci 277(1699):3519–3525. doi:10.1098/rspb.2010.1071
Pandian TJ, Koteeswaran R (1999) Natural occurrence of monoploids and polyploids in the
Indian catfish, Heteropneustes fossilis. Curr Sci India 76(8):1134–1137
Phillips R, Rab P (2001) Chromosome evolution in the Salmonidae (pisces): an update. Biol Rev
Camb Philos Soc 76(1):1–25
Phillips RB, Keatley KA, Morasch MR, Ventura AB, Lubieniecki KP, Koop BF, Danzmann RG,
Davidson WS (2009) Assignment of Atlantic salmon (Salmo salar) linkage groups to specific
chromosomes: conservation of large syntenic blocks corresponding to whole chromosome arms
in rainbow trout (Oncorhynchus mykiss). BMC Genet 10:46. doi:10.1186/1471-2156-10-46
Piferrer F, Beaumont A, Falguiere JC, Flajshans M, Haffray P, Colombo L (2009) Polyploid fish
and shellfish: production, biology and applications to aquaculture for performance improvement and genetic containment. Aquaculture 293(3–4):125–156. doi:10.1016/j.aquaculture.
2009.04.036
Postlethwait JH, Yan YL, Gates MA, Horne S, Amores A, Brownlie A, Donovan A, Egan ES,
Force A, Gong Z, Goutel C, Fritz A, Kelsh R, Knapik E, Liao E, Paw B, Ransom D, Singer A,
Thomson M, Abduljabbar TS, Yelick P, Beier D, Joly JS, Larhammar D, Rosa F, Westerfield
M, Zon LI, Johnson SL, Talbot WS (1998) Vertebrate genome evolution and the zebrafish
gene map. Nat Genet 18(4):345–349. doi:10.1038/ng0498-345
Postlethwait JH, Woods IG, Ngo-Hazelett P, Yan YL, Kelly PD, Chu F, Huang H, Hill-Force A,
Talbot WS (2000) Zebrafish comparative genomics and the origins of vertebrate chromosomes. Genome Res 10(12):1890–1902
Postlethwait J, Amores A, Cresko W, Singer A, Yan YL (2004) Subfunction partitioning, the
teleost radiation and the annotation of the human genome. Trends Genet 20(10):481–490.
doi:10.1016/j.tig.2004.08.001 S0168-9525(04)00213-6 [pii]
Prince VE (2002) The Hox paradox: more complex(es) than imagined. Dev Biol 249(1):1–15.
doi:10.1006/Dbio.2002.0745
Prince VE, Joly L, Ekker M, Ho RK (1998) Zebrafish Hox genes: genomic organization and
modified colinear expression patterns in the trunk. Development 125(3):407–420
Prohaska SJ, Stadler PF (2004) The duplication of the Hox gene clusters in teleost fishes. Theory
Biosci 123(1):89–110. doi:10.1016/j.thbio.2004.03.004
Putnam NH, Butts T, Ferrier DE, Furlong RF, Hellsten U, Kawashima T, Robinson-Rechavi M,
Shoguchi E, Terry A, Yu JK, Benito-Gutierrez EL, Dubchak I, Garcia-Fernandez J, GibsonBrown JJ, Grigoriev IV, Horton AC, de Jong PJ, Jurka J, Kapitonov VV, Kohara Y, Kuroki Y,
Lindquist E, Lucas S, Osoegawa K, Pennacchio LA, Salamov AA, Satou Y, Sauka-Spengler
T, Schmutz J, Shin IT, Toyoda A, Bronner-Fraser M, Fujiyama A, Holland LZ, Holland PW,
Satoh N, Rokhsar DS (2008) The amphioxus genome and the evolution of the chordate
karyotype. Nature 453(7198):1064–1071. doi:10.1038/nature06967 nature06967 [pii]
Raincrow JD, Dewar K, Stocsits C, Prohaska SJ, Amemiya CT, Stadler PF, Chiu CH (2011) Hox
clusters of the bichir (Actinopterygii, Polypterus senegalus) highlight unique patterns of
sequence evolution in gnathostome phylogeny. J Exp Zool B Mol Dev Evol 316(6):451–464.
doi:10.1002/jez.b.21420
Rastogi S, Liberles DA (2005) Subfunctionalization of duplicated genes as a transition state to
neofunctionalization. BMC Evol Biol 5:28. doi:10.1186/1471-2148-5-28 1471-2148-5-28 [pii]
Ravi V, Venkatesh B (2008) Rapidly evolving fish genomes and teleost diversity. Curr Opin
Genet Dev 18(6):544–550. doi:10.1016/j.gde.2008.11.001 S0959-437X(08)00151-2 [pii]
Richardson BE, Lehmann R (2010) Mechanisms guiding primordial germ cell migration:
strategies from different organisms. Nat Rev Mol Cell Biol 11(1):37–49. doi:10.1038/
nrm2815 nrm2815 [pii]
Risinger C, Larhammar D (1993) Multiple loci for synapse protein SNAP-25 in the tetraploid
goldfish. Proc Nat Acad Sci USA 90(22):10598–10602
CO
RR
1379
1380
1381
1382
1383
1384
1385
1386
1387
1388
1389
1390
1391
1392
1393
1394
1395
1396
1397
1398
1399
1400
1401
1402
1403
1404
1405
1406
1407
1408
1409
1410
1411
1412
1413
1414
1415
1416
1417
1418
1419
1420
1421
1422
1423
1424
1425
1426
1427
1428
1429
1430
I. Braasch and J. H. Postlethwait
UN
Editor Proof
380
Book ISBN: 978-3-642-31441-4
Page: 380/383
Layout: T1 Standard SC
Chapter No.: 17
381
EC
TE
D
PR
OO
F
Robinson-Rechavi M, Marchand O, Escriva H, Bardet PL, Zelus D, Hughes S, Laudet V (2001)
Euteleost fish genomes are characterized by expansion of gene families. Genome Res
11(5):781–788. doi:10.1101/gr.165601
Rohner N, Bercsenyi M, Orban L, Kolanczyk ME, Linke D, Brand M, Nusslein-Volhard C,
Harris MP (2009) Duplication of fgfr1 permits Fgf signaling to serve as a target for selection
during domestication. Curr Biol 19(19):1642–1647. doi:10.1016/j.cub.2009.07.065 S09609822(09)01542-5 [pii]
Santini F, Harmon LJ, Carnevale G, Alfaro ME (2009) Did genome duplication drive the origin
of teleosts? A comparative study of diversification in ray-finned fishes. BMC Evol Biol 9:194.
doi:10.1186/1471-2148-9-194 1471-2148-9-194 [pii]
Sato Y, Nishida M (2007) Post-duplication charge evolution of phosphoglucose isomerases in
teleost fishes through weak selection on many amino acid sites. BMC Evol Biol 7:204.
doi:10.1186/1471-2148-7-204 Artn 204
Sato Y, Hashiguchi Y, Nishida M (2009) Temporal pattern of loss/persistence of duplicate genes
involved in signal transduction and metabolic pathways after teleost-specific genome
duplication. BMC Evol Biol 9:127. doi:10.1186/1471-2148-9-127 1471-2148-9-127 [pii]
Scannell DR, Wolfe KH (2008) A burst of protein sequence evolution and a prolonged period of
asymmetric evolution follow gene duplication in yeast. Genome Res 18(1):137–147.
doi:10.1101/gr.6341207 gr.6341207 [pii]
Schultz RJ (1980) Role of polyploidy in the evolution of fishes. In: Lewis WH (ed) Polyploidy—
biological relevance. Basic life science, vol 13. Plenum Press, New York, pp 313–340
Semon M, Wolfe KH (2007a) Rearrangement rate following the whole-genome duplication in
teleosts. Mol Biol Evol 24(3):860–867. doi:10.1093/molbev/msm003 msm003 [pii]
Semon M, Wolfe KH (2007b) Reciprocal gene loss between Tetraodon and zebrafish after whole
genome duplication in their ancestor. Trends Genet 23(3):108–112. doi:10.1016/
j.tig.2007.01.003 S0168-9525(07)00021-2 [pii]
Setiamarga DH, Miya M, Yamanoue Y, Azuma Y, Inoue JG, Ishiguro NB, Mabuchi K, Nishida
M (2009) Divergence time of the two regional medaka populations in Japan as a new time
scale for comparative genomics of vertebrates. Biol Lett 5(6):812–816. doi:10.1098/
rsbl.2009.0419 rsbl.2009.0419 [pii]
Sha Z, Yu P, Takano T, Liu H, Terhune J, Liu Z (2008) The warm temperature acclimation
protein Wap65 as an immune response gene: its duplicates are differentially regulated by
temperature and bacterial infections. Mol Immunol 45(5):1458–1469. doi:10.1016/
J.Molimm.2007.08.012
Shimada A, Yabusaki M, Niwa H, Yokoi H, Hatta K, Kobayashi D, Takeda H (2008) Maternalzygotic medaka mutants for fgfr1 reveal its essential role in the migration of the axial
mesoderm but not the lateral mesoderm. Development 135(2):281–290. doi:10.1242/
dev.011494 135/2/281 [pii]
Siegel N, Hoegg S, Salzburger W, Braasch I, Meyer A (2007) Comparative genomics of ParaHox
clusters of teleost fishes: gene cluster breakup and the retention of gene sets following whole
genome duplications. BMC Genomics 8:312. doi:10.1186/1471-2164-8-312 1471-2164-8-312
[pii]
Soltis DE, Albert VA, Leebens-Mack J et al (2009) Polyploidy and angiosperm diversification.
Am J Bot 96(1):336–348
Song H, Yan YL, Titus T, He XJ, Postlethwait JH (2011) The role of stat1b in zebrafish
hematopoiesis. Mech Develop 128(7–10):442–456. doi:10.1016/J.Mod.2011.08.004
Star B, Nederbragt AJ, Jentoft S, Grimholt U, Malmstrom M, Gregers TF, Rounge TB, Paulsen J,
Solbakken MH, Sharma A, Wetten OF, Lanzen A, Winer R, Knight J, Vogel JH, Aken B,
Andersen O, Lagesen K, Tooming-Klunderud A, Edvardsen RB, Tina KG, Espelund M,
Nepal C, Previti C, Karlsen BO, Moum T, Skage M, Berg PR, Gjoen T, Kuhl H, Thorsen J,
Malde K, Reinhardt R, Du L, Johansen SD, Searle S, Lien S, Nilsen F, Jonassen I, Omholt
SW, Stenseth NC, Jakobsen KS (2011) The genome sequence of Atlantic cod reveals a unique
immune system. Nature 477(7363):207–210. doi:10.1038/nature10342 nature10342 [pii]
CO
RR
1431
1432
1433
1434
1435
1436
1437
1438
1439
1440
1441
1442
1443
1444
1445
1446
1447
1448
1449
1450
1451
1452
1453
1454
1455
1456
1457
1458
1459
1460
1461
1462
1463
1464
1465
1466
1467
1468
1469
1470
1471
1472
1473
1474
1475
1476
1477
1478
1479
1480
1481
1482
1483
Book ISBN: 978-3-642-31441-4
Page: 381/383
Polyploidy in Fish and the Teleost Genome Duplication
UN
Editor Proof
17
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 17
Book ID: 272454_1_En
Date: 16-8-2012
EC
TE
D
PR
OO
F
Steinke D, Salzburger W, Braasch I, Meyer A (2006) Many genes in fish have species-specific
asymmetric rates of molecular evolution. BMC Genomics 7:20. doi:10.1186/1471-2164-7-20
1471-2164-7-20 [pii]
Stellwag EJ (1999) Hox gene duplication in fish. Semin Cell Dev Biol 10(5):531–540.
doi:10.1006/scdb.1999.0334 S1084-9521(99)90334-8 [pii]
Stoltzfus A (1999) On the possibility of constructive neutral evolution. J Mol Evol 49(2):169–
181. doi:JME1907 [pii]
Svärdson G (1945) Chromosome studies on Salmonidae. Rep Swed State Inst Fresh Fish Res
23:1–151
Taylor JS, Van de Peer Y, Braasch I, Meyer A (2001) Comparative genomics provides evidence
for an ancient genome duplication event in fish. Philos Trans R Soc Lond B Biol Sci
356(1414):1661–1679. doi:10.1098/rstb.2001.0975
Taylor JS, Braasch I, Frickey T, Meyer A, Van de Peer Y (2003) Genome duplication, a trait
shared by 22,000 species of ray-finned fish. Genome Res 13(3):382–390. doi:10.1101/
gr.640303
Tumpel S, Cambronero F, Wiedemann LM, Krumlauf R (2006) Evolution of cis elements in the
differential expression of two Hoxa2 coparalogous genes in pufferfish (Takifugu rubripes).
Proc Nat Acad Sci USA 103(14):5419–5424. doi:10.1073/pnas.0600993103 0600993103 [pii]
Udono T, Yasumoto K, Takeda K, Amae S, Watanabe K, Saito H, Fuse N, Tachibana M,
Takahashi K, Tamai M, Shibahara S (2000) Structural organization of the human
microphthalmia-associated transcription factor gene containing four alternative promoters.
Biochim Biophys Acta 1491(1–3):205–219
Uyeno T, Smith GR (1972) Tetraploid origin of karyotype of catostomid fishes. Science
175(4022):644
Van de Peer Y, Taylor JS, Braasch I, Meyer A (2001) The ghost of selection past: rates of
evolution and functional divergence of anciently duplicated genes. J Mol Evol 53(4–5):436–
446. doi:10.1007/s002390010233
Van de Peer Y, Maere S, Meyer A (2009) The evolutionary significance of ancient genome
duplications. Nat Rev Genet 10(10):725–732. doi:10.1038/nrg2600 nrg2600 [pii]
Vandepoele K, De Vos W, Taylor JS, Meyer A, Van de Peer Y (2004) Major events in the
genome evolution of vertebrates: paranome age and size differ considerably between rayfinned fishes and land vertebrates. Proc Nat Acad Sci USA 101(6):1638–1643. doi:10.1073/
pnas.0307968100 0307968100[pii]
Vilella AJ, Severin J, Ureta-Vidal A, Heng L, Durbin R, Birney E (2009) EnsemblCompara
GeneTrees: complete, duplication-aware phylogenetic trees in vertebrates. Genome Res
19(2):327–335. doi:10.1101/Gr.073585.107
Volff JN (2005) Genome evolution and biodiversity in teleost fish. Heredity (Edinb) 94(3):280–
294. doi:10.1038/sj.hdy.6800635 6800635 [pii]
Volff JN, Brunet F, Böhne A, Galiana-Arnoux D (2011) Evolution of fish genomes. In: Farrell
AP, Cech JJ, Richards JG, Stevens ED (eds) Encyclopedia of fish physiology: from genome to
environment. Elsevier Inc., San Diego
Werth CR, Windham MD (1991) A model for divergent, allopatric speciation of polyploid
pteridophytes resulting from silencing of duplicate-gene expression. Am Nat 137(4):515–526
Winkler C, Schafer M, Duschl J, Schartl M, Volff JN (2003) Functional divergence of two
zebrafish midkine growth factors following fish-specific gene duplication. Genome Res
13(6A):1067–1081. doi:10.1101/gr.1097503 GR-10975R [pii]
Wittbrodt J, Meyer A, Schartl M (1998) More genes in fish? BioEssays 20(6):511–515.
doi:10.1002/(sici)1521-1878(199806)20:6\511:aid-bies10[3.0.co;2-3
Woltering JM, Durston AJ (2006) The zebrafish hoxDb cluster has been reduced a single
microRNA. Nat Genet 38(6):601–602. doi:10.1038/Ng0606-601
Woods TD, Buth DG (1984) High-level of gene silencing in the tetraploid goldfish. Biochem Syst
Ecol 12(4):415–421
Woods IG, Kelly PD, Chu F, Ngo-Hazelett P, Yan YL, Huang H, Postlethwait JH, Talbot WS
(2000) A comparative map of the zebrafish genome. Genome Res 10(12):1903–1914
CO
RR
1484
1485
1486
1487
1488
1489
1490
1491
1492
1493
1494
1495
1496
1497
1498
1499
1500
1501
1502
1503
1504
1505
1506
1507
1508
1509
1510
1511
1512
1513
1514
1515
1516
1517
1518
1519
1520
1521
1522
1523
1524
1525
1526
1527
1528
1529
1530
1531
1532
1533
1534
1535
1536
1537
I. Braasch and J. H. Postlethwait
UN
Editor Proof
382
Book ISBN: 978-3-642-31441-4
Page: 382/383
Layout: T1 Standard SC
Chapter No.: 17
383
EC
TE
D
PR
OO
F
Woods IG, Wilson C, Friedlander B, Chang P, Reyes DK, Nix R, Kelly PD, Chu F, Postlethwait
JH, Talbot WS (2005) The zebrafish gene map defines ancestral vertebrate chromosomes.
Genome Res 15(9):1307–1314. doi:10.1101/gr.4134305 gr.4134305 [pii]
Woolfe A, Elgar G (2007) Comparative genomics using fugu reveals insights into regulatory
subfunctionalization. Genome Biol 8(4):R53. doi:10.1186/gb-2007-8-4-r53 gb-2007-8-4-r53
[pii]
Yao K, Ge W (2010) Kit system in the zebrafish ovary: evidence for functional divergence of two
isoforms of kit (kita and kitb) and kit ligand (kitlga and kitlgb) during folliculogenesis. Biol
Reprod 82(6):1216–1226. doi:10.1095/biolreprod.109.082644 biolreprod.109.082644 [pii]
Yokoi H, Shimada A, Carl M, Takashima S, Kobayashi D, Narita T, Jindo T, Kimura T,
Kitagawa T, Kage T, Sawada A, Naruse K, Asakawa S, Shimizu N, Mitani H, Shima A,
Tsutsumi M, Hori H, Wittbrodt J, Saga Y, Ishikawa Y, Araki K, Takeda H (2007) Mutant
analyses reveal different functions of fgfr1 in medaka and zebrafish despite conserved ligandreceptor relationships. Dev Biol 304(1):326–337. doi:10.1016/j.ydbio.2006.12.043 S00121606(06)01502-8 [pii]
Yokoi H, Yan YL, Miller MR, BreMiller RA, Catchen JM, Johnson EA, Postlethwait JH
(2009) Expression profiling of zebrafish sox9 mutants reveals that sox9 is required for
retinal differentiation. Dev Biol 329(1):1–15. doi:10.1016/j.ydbio.2009.01.002 S00121606(09)00020-7 [pii]
Yu WP, Brenner S, Venkatesh B (2003) Duplication, degeneration and subfunctionalization of the
nested synapsin-timp genes in fugu. Trends Genet 19(4):180–183 S0168952503000489[pii]
Yuan J, He Z, Yuan X, Jiang X, Sun X, Zou S (2010) Speciation of polyploid Cyprinidae fish of
common carp, crucian carp, and silver crucian carp derived from duplicated Hox genes. J Exp
Zool B Mol Dev Evol 314(6):445–456. doi:10.1002/jez.b.21350
Zapater C, Chauvigne F, Norberg B, Finn RN, Cerda J (2011) Dual neofunctionalization of a
rapidly evolving aquaporin-1 paralog resulted in constrained and relaxed traits controlling
channel function during meiosis resumption in teleosts. Mol Biol Evol 28(11):3151–3169.
doi:10.1093/molbev/msr146 msr146 [pii]
CO
RR
1538
1539
1540
1541
1542
1543
1544
1545
1546
1547
1548
1549
1550
1551
1552
1553
1554
1555
1556
1557
1558
1559
1560
1561
1562
1563
1564
1565
Book ISBN: 978-3-642-31441-4
Page: 383/383
Polyploidy in Fish and the Teleost Genome Duplication
UN
Editor Proof
17
Book ID: 272454_1_En
Date: 16-8-2012
Metadata of the chapter that will be visualized in
SpringerLink
Book Title
Polyploidy and Genome Evolution
Series Title
Chapter Title
Polyploidization and Sex Chromosome Evolution in Amphibians
Copyright Year
2012
Copyright HolderName
Springer-Verlag Berlin Heidelberg
Corresponding Author
Family Name
Evans
Particle
Given Name
Ben J.
Suffix
Author
Division
Department of Biology
Organization
McMaster University
Address
Life Sciences Building Room 328, 1280 Main Street West, L8S 4K1,
Hamilton, ON, Canada
Email
evansb@mcmaster.ca
Family Name
Alexander Pyron
Particle
Given Name
R.
Suffix
Division
Department of Biological Sciences
Organization
The George Washington University
Address
2023 G St. NW, 20052, Washington, DC, USA
Email
Author
Family Name
Wiens
Particle
Given Name
John J.
Suffix
Division
Department of Ecology and Evolution
Organization
Stony Brook University
Address
11794-5245, Stony Brook, NY, USA
Email
Abstract
Genome duplication, including polyploid speciation and spontaneous polyploidy in diploid species, occurs
more frequently in amphibians than mammals. One possible explanation is that some amphibians, unlike
almost all mammals, have young sex chromosomes that carry a similar suite of genes (apart from the genetic
trigger for sex determination). These species potentially can experience genome duplication without
disrupting dosage stoichiometry between interacting proteins encoded by genes on the sex chromosomes and
autosomal chromosomes. To explore this possibility, we performed a permutation aimed at testing whether
amphibian species that experienced polyploid speciation or spontaneous polyploidy have younger sex
chromosomes than other amphibians. While the most conservative permutation was not significant, the frog
genera Xenopus and Leiopelma provide anecdotal support for a negative correlation between the age of sex
chromosomes and a species’ propensity to undergo genome duplication. This study also points to more
frequent turnover of sex chromosomes than previously proposed, and suggests a lack of statistical support
for male versus female heterogamy in the most recent common ancestors of frogs, salamanders, and
amphibians in general. Future advances in genomics undoubtedly will further illuminate the relationship
between amphibian sex chromosome degeneration and genome duplication.
Book ISBN: 978-3-642-31441-4
Page: 385/410
Chapter 18
4
Ben J. Evans, R. Alexander Pyron and John J. Wiens
11
12
13
14
15
16
17
18
19
20
21
22
23
D
9
10
TE
8
EC
7
Abstract Genome duplication, including polyploid speciation and spontaneous
polyploidy in diploid species, occurs more frequently in amphibians than mammals.
One possible explanation is that some amphibians, unlike almost all mammals, have
young sex chromosomes that carry a similar suite of genes (apart from the genetic
trigger for sex determination). These species potentially can experience genome
duplication without disrupting dosage stoichiometry between interacting proteins
encoded by genes on the sex chromosomes and autosomal chromosomes. To explore
this possibility, we performed a permutation aimed at testing whether amphibian
species that experienced polyploid speciation or spontaneous polyploidy have
younger sex chromosomes than other amphibians. While the most conservative
permutation was not significant, the frog genera Xenopus and Leiopelma provide
anecdotal support for a negative correlation between the age of sex chromosomes
and a species’ propensity to undergo genome duplication. This study also points to
more frequent turnover of sex chromosomes than previously proposed, and suggests
a lack of statistical support for male versus female heterogamy in the most recent
common ancestors of frogs, salamanders, and amphibians in general. Future
advances in genomics undoubtedly will further illuminate the relationship between
amphibian sex chromosome degeneration and genome duplication.
CO
RR
5
6
PR
OO
3
Polyploidization and Sex Chromosome
Evolution in Amphibians
2
F
1
Book ID: 272454_1_En
Date: 16-8-2012
B. J. Evans (&)
Department of Biology, McMaster University, Life Sciences Building Room 328,
1280 Main Street West, Hamilton, ON L8S 4K1, Canada
e-mail: evansb@mcmaster.ca
R. Alexander Pyron
Department of Biological Sciences, The George Washington University,
2023 G St. NW, Washington, DC 20052, USA
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: 18
J. J. Wiens
Department of Ecology and Evolution, Stony Brook University,
Stony Brook, NY 11794-5245, USA
P. S. Soltis and D. E. Soltis (eds.), Polyploidy and Genome Evolution,
DOI: 10.1007/978-3-642-31442-1_18, Springer-Verlag Berlin Heidelberg 2012
385
Layout: T1 Standard SC
Chapter No.: 18
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 386/410
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
61
62
63
64
65
66
F
PR
OO
28
29
D
27
Why polyploidization is more common in plants than in animals is a central
question in biology (Mable 2004; Muller 1925; Orr 1990), and multiple explanations have been put forward (reviewed in Gregory and Mable 2005; Mable 2004;
Orr 1990; Otto and Whitton 2000). One possibility is that the propensity for a
species to undergo polyploidization is related to the extent of sex chromosome
degeneration. Sex chromosome degeneration is the evolution of differences in gene
content that goes beyond the fundamental difference in the presence or absence of
a genetic trigger for sex determination. Degenerate sex chromosomes could
present problems during ‘‘diploidization’’ of a polyploid genome. Diploidization
refers to the phenomenon by which a polyploid species transitions to a mode of
chromosomal inheritance that is similar to a diploid species. A key feature of
diploidization is the switch from polysomic inheritance, where multivalents form
during cell division, to disomic inheritance, where only bivalents form (Wolfe
2001). This phenomenon is probably achieved via divergence between duplicated
pairs of homologous chromosomes. Diploidization therefore could be instantaneous when polyploidization occurs via allopolyploidization (genome duplication
associated with hybridization among diverged species) because duplicated
homologous chromosome pairs already diverged from one another in the ancestral
species. When a polyploid genome with duplicated sex chromosomes becomes
diploidized, one pair of sex chromosomes presumably begins to segregate autosomally. With a degenerate Y-chromosome, for instance, the nascent autosomal
pair that was previously a pair of sex chromosomes would initially have three
possible genotypes: AXAX, AX0, and 00 where AX refers to an autosomal allele
derived from an ancestral X-chromosome and 0 refers to a missing allele that was
lost on the ancestral Y-chromosome. If the 00 genotype is deleterious or lethal,
there would be reproductive incompatibilities in the early stages of diploidization
until the degenerate chromosome is lost. This fitness cost could be mitigated if a
functional paralogous allele were still present on the sex chromosomes, as would
be expected in an autopolyploid (formed from genome duplication within a species). In allopolyploids, however, sex chromosomes could degenerate in unique
ways in each ancestral species, giving rise to diverged gene content, so homozygous null genotypes would be a bigger problem. In both types of polyploids,
dosage balance requirements—natural selection favoring a specific relative
expression level of interacting genes (Papp et al. 2003; Qian and Zhang 2008)—
could impose a fitness cost on a homozygous or heterozygous null autosomal
genotype in a polyploid.
Polyploidization could also present challenges to species with degenerate sex
chromosomes that have also evolved mechanisms for dosage compensation (Orr
1990). Dosage compensation equilibrates expression levels of genes that have one
allele in one sex and two alleles in the other sex (for example, X-linked genes have
one allele in XY males but two alleles in XX females). In this way the stoichiometry
of expression of X-linked and autosomal genes is constant, or ‘‘balanced’’, in males
TE
26
EC
25
18.1 Introduction
CO
RR
24
B. J. Evans et al.
UN
Editor Proof
386
Layout: T1 Standard SC
Chapter No.: 18
Book ISBN: 978-3-642-31441-4
Page: 387/410
Polyploidization and Sex Chromosome Evolution in Amphibians
387
101
18.1.1 Sex Chromosome Evolution
74
75
76
77
78
79
80
81
82
83
84
85
86
87
88
89
90
91
92
93
94
95
96
97
98
99
102
103
104
105
106
107
PR
OO
73
D
72
TE
71
EC
70
CO
RR
69
F
100
and females. Orr (1990) argued that a polyploid lineage would initially be established via backcrossing a new polyploid individual to diploid individuals, and that
this would disrupt this balance.
The evolution of differences in gene content is a combined consequence of the
migration of genes, especially genes with sex-specific function, to one or the other
sex chromosome, and also the loss of genes from the region of suppressed recombination on the sex-specific sex chromosome (for example, the Y-chromosome).
The disparity in gene content between the sex chromosomes is thought to increase
over time as a consequence of natural selection (Bergero and Charlesworth 2009;
Charlesworth et al. 2005). Substantial disparity in gene content between the sex
chromosomes could be coupled with selective pressure favoring the evolution of
mechanisms of dosage compensation. For this reason, the proposal that degenerate
sex chromosomes deter polyploidization (including species that lack dosage compensation) is not independent of Orr’s (1990) proposal that dosage compensation
deters polyploidization. In either case, if sex chromosome degeneration acts as a
barrier to polyploidization, this would predict that polyploid species or species with
polymorphism in ploidy levels would have relatively ‘‘young’’, minimally degenerate sex chromosomes as compared with other species.
In this chapter we briefly review polyploidization in frogs and salamanders and
general features of sex chromosome evolution. Using previously published
information, we then use a maximum likelihood approach to analyze the evolution
of new sex-determining mechanisms in frogs and salamanders in a phylogenetic
context, where new mechanisms are inferred either from a change in which
chromosomes are the sex chromosomes, the evolution of a new trigger for sex
determination, or from observed polymorphism in sex-determining mechanisms.
Following this, we explore whether polyploidization occurs more frequently soon
after a new sex-determining mechanism evolves using a permutation test that
accommodates uncertainty in ancestral reconstruction. We conclude that novel
sex-determining mechanisms have evolved in amphibians even more frequently
than previously proposed, and that amphibians with young sex chromosomes may
be more likely to experience genome duplication, resulting either in polyploid
speciation or in spontaneous polyploidy of individuals of an otherwise diploid
species. A major caveat to the latter result is that information on the age of sexdetermining mechanisms of most polyploid amphibians is lacking.
67
68
Sex chromosomes originate from autosomes (Ohno 1967) but differ in carrying
genetic information that (a) differs between the sexes and (b) triggers or represses
sex-specific gonadal differentiation. The ‘‘heterogametic’’ sex produces two types
of gametes, each type with a different sex chromosome, and the ‘‘homogametic’’
sex produces only one type of gamete with respect to the sex chromosomes. The sex
chromosomes of species with male heterogamy are called ‘‘X’’ and ‘‘Y’’ (females
UN
Editor Proof
18
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 18
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 388/410
114
115
116
117
118
119
120
121
122
123
124
125
126
127
128
129
130
131
132
133
134
135
136
137
138
139
140
141
142
143
144
145
146
147
148
149
150
151
152
F
113
PR
OO
112
D
111
TE
110
have two X chromosomes and males have an X and a Y), and the sex chromosomes
of species with female heterogamy are called ‘‘Z’’ and ‘‘W’’ (males have two Z
chromosomes, and females have a Z and a W). In species with genetic sex determination, gonadal differentiation—also known as primary sex determination—is
achieved either using a sex chromosome-specific genetic trigger or by gene dosage,
where the homogametic sex carries two doses of an activator of that sex, or a
repressor of the heterogametic sex.
The age of sex chromosomes influences important aspects of their evolution
and divergence, including divergence in gene content, and the origin of dosage
compensation. For example, the sex chromosomes of therian (placental and
marsupial) mammals are extremely old ([180 million years; Graves 2008), and
the Y chromosome is much smaller than the X and carries fewer genes. This
disparity in size and gene content arose due to Y chromosome ‘‘degeneration’’ as a
consequence of suppressed recombination with the X chromosome (Charlesworth
and Charlesworth 2000). Suppressed recombination ensures that male progeny
inherit an intact copy of the genetic trigger for testis formation, which in therians is
the SRY gene. But this also permits deleterious mutations to accumulate in
Y-linked genes (Muller 1964), leading to loss of function and deletion. ‘‘Muller’s
ratchet’’, the stochastic loss of the least deleterious allele in a population
(Felsenstein 1974), leads to a decline in fitness. This decline occurs more quickly
in regions of the genome that do not recombine. Hill-Robertson effects, background selection, and hitchhiking of deleterious alleles also contribute to fitness
declines of non-recombining portions of the genome (reviewed in Charlesworth
and Charlesworth 2000). Degeneration of the therian Y-chromosome occurred in a
stepwise fashion as the region of suppressed recombination expanded in large
increments (Skaletsky et al. 2003). In therians the disparity in gene content
increased over time after the origin of SRY and associated suppression of
recombination between the X and Y chromosomes. Most angiosperm plants that
have separate sexes (dioecy), in contrast, have comparatively young sex chromosomes that are not substantially differentiated, although exceptions exist
(Bergero and Charlesworth 2011; Charlesworth 2002). Polyploid species are
prevalent in angiosperms (Otto and Whitton 2000) but absent in therians (Svartman et al. 2005), and these observations thus provide anecdotal support for the
contention that the extent of sex chromosome degeneration is negatively correlated
with the incidence of polyploid speciation. Amphibians offer an interesting focal
group with which to further evaluate this hypothesis because some features of
amphibian genome evolution resemble plants more than other animal groups such
as therian mammals. In particular, unlike therian mammals, sex chromosomes in
many amphibians are relatively young, chromosome degeneration is modest or
absent, and polyploidization is fairly common.
Species that determine sex exclusively using environmental triggers do not
have genomic differences between the sexes and therefore have no sex chromosomes. In amphibians, sex determination is genetic so all species are expected to
have sex chromosomes. In addition, temperature has been reported to influence
offspring sex ratios of various species of the salamander genera Pleurodeles and
EC
109
CO
RR
108
B. J. Evans et al.
UN
Editor Proof
388
Layout: T1 Standard SC
Chapter No.: 18
159
160
161
162
163
164
165
166
167
168
169
170
171
172
173
174
175
176
177
178
179
180
181
182
183
184
185
186
187
188
189
190
191
192
193
F
158
PR
OO
157
D
156
Hynobius and the frog genera Bufo, Rana, and laboratory-generated polyploids of
the genus Xenopus (Hayes 1998; Kobel 1996; Schmid and Steinlein 2001). Sex
chromosomes are cytologically distinct in some amphibian species (Schmid et al.
2010). Differences in gene content between the sex chromosomes, which is suggested by cytologically distinct sex chromosomes, led to the independent evolution
of dosage compensation mechanisms in placental mammals, birds, and other
groups such as Drosophila and Caenorhabditis (Arnold et al. 2008; Straub and
Becker 2007). However, in amphibians evidence of dosage compensation has not
been found (Hayes 1998; Schmid et al. 1986; Schmid and Steinlein 2001). One
possible reason for this is that we do not yet know the identity of any amphibian
genes that are hemizygous in the heterogametic sex, so a rigorous test for dosage
compensation in amphibians is not yet possible. These genes would be restricted to
the portion of the X or Z chromosome that does not recombine with the Y or W
chromosome, respectively.
A recent study of European tree frogs identified one way that amphibians
circumvent sex chromosome degeneration (Stöck et al. 2011). In three species, no
recombination occurred between the sex chromosomes in males generated from
intraspecific crosses, yet no intraspecific sex chromosome divergence was observed
(Stöck et al. 2011). This suggests that Muller’s ratchet is periodically reset in these
species by infrequent recombination between the sex chromosomes. Sex chromosome degeneration can also be circumvented by genomic translocation of the sexdetermining locus to another chromosomal pair, or by re-assignment of the sexdetermining function to a gene located elsewhere in the genome. Both of these
phenomena result in a change in which chromosomes are the sex chromosomes
(hereafter ‘‘sex chromosome turnover’’). Ancient examples of sex chromosome
turnover are evinced in amniotes by homology between the sex chromosomes of
platypuses and those of birds but not those of therian mammals (Graves 2008). In
amphibians, sex chromosome turnover is common and is suggested by variation
among and within species in male versus female heterogamy (Ezaz et al. 2006; Hillis
and Green 1990). Using maximum parsimony, Hillis and Green (1990) analyzed
variation in male and female heterogamy in amphibians in a phylogenetic context
and concluded that sex chromosome turnover occurred at least seven times.
TE
155
389
EC
154
CO
RR
153
Book ISBN: 978-3-642-31441-4
Page: 389/410
Polyploidization and Sex Chromosome Evolution in Amphibians
18.1.2 How Many Frog and Salamander Species
are Polyploid?
Comprehensive reviews of polyploidization in amphibians are available in Bogart
(1980), Kawamura (1984), Duellman and Trueb (1994), Beçak and Beçak (1998),
Otto and Whitton (2000), Gregory and Mable (2005), Schmid et al. (2010), and
Mable et al. (2011). The two most recent of these reviews have up-to-date lists of
known polyploid species and associated citations that document polyploidy.
Schmid et al. (2010) also summarize male and female heterogamy in frogs and
salamanders, including information on species with cytologically detectable sex
UN
Editor Proof
18
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 18
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 390/410
201
202
203
204
205
206
207
208
209
210
211
212
213
214
215
216
217
218
219
220
221
222
223
224
225
226
227
228
229
230
231
232
233
234
235
236
237
238
F
PR
OO
200
D
198
199
TE
197
EC
196
chromosome divergence (their Table 8, pp 160–161). In their Supplementary
Information, Mable et al. (2011) provide data on confirmed diploid species that are
closely related to the polyploids. A key difference between these two reviews is
that Mable et al. (2011) include only bisexually reproducing polyploids whereas
Schmid et al. (2010) also include unisexual polyploids. We have attempted to
compile this information as inclusively as possible in Table 18.1, including some
minor corrections and a few additional species and associated citations. Thus, not
all polyploids listed in this table are bisexual, and some are diploid species in
which polyploid individuals occur spontaneously or by induction due to laboratory
manipulation.
Fifty polyploid frog species have been described, including seven triploids, 30
tetraploids, 11 octoploids, and two dodecaploids derived from 15 families and 20
genera (Table 18.1). Three tetraploids and two dodecaploids have been reported
from the genus Xenopus but not yet formally described as species (Evans 2007,
2008; Evans et al. 2004a, 2005a; Tymowska 1991). Stable triploids are known from
three frog genera (Bufo, Eupsophus, and Rana), tetraploids from 16 (Aphantophryne, Astylosternus, Bufo, Chiasmocleis, Dicroglossus, Eleuthrodactylus, Hyla,
Neobatrachus, Odontophrynus, Phyllomedusa, Pleurodema, Pyxicephalus,
Scaphiophryne Silurana, Tomopterna, and Xenopus), octoploids from three
(Ceratophrys, Pleurodema, and Xenopus), and dodecaploids only from Xenopus.
Spontaneous or experimentally induced polyploidy has been reported in at least five
frog species. Six polyploid species of salamander, including four triploids and two
tetraploids, are known from only two genera (Ambystoma and Siren) from two
families (Table 18.1). Spontaneous or experimentally induced triploidy or tetraploidy has been reported in eight salamander species.
The origin of polyploidy necessarily is preceded by the existence of one or more
diploid ancestors. Interestingly, a number of polyploid frog species are inferred to
have originated from ancestral diploid species that do not have known extant diploid
descendants. In Xenopus and Silurana, for example, three currently unknown
diploid species contributed their genomes to extant tetraploid species (reviewed in
Evans 2008). There are also three currently unknown tetraploid species that contributed their genomes to extant octoploid and dodecaploid Xenopus species
(reviewed in Evans 2008). The tetraploid Hyla versicolor is thought to be derived
from multiple independent allopolyploidization events between three diploid species, two of which are currently unknown, and probably extinct given that the region
in which they occur (temperate North America) is well studied (Holloway et al.
2006). In Ceratophrys, there are no known tetraploid species even though three
species in this genus are octoploid (Table 18.1). Similarly, the tetraploid species
Bufo pewzowi is thought to be derived from the diploid B. turanensis and another
unidentified diploid (Stöck et al. 2009), and various tetraploid species of Neobatrachus are derived from diploid ancestors whose diploid descendants are currently
unknown (Mable and Roberts 1997). It is tempting to speculate from these
observations that polyploidization contributed to the long-term survival of these
lineages, given that the diploid ancestors of extant polyploids seem to have gone
extinct in many cases. However, we lack information on how frequently
CO
RR
194
195
B. J. Evans et al.
UN
Editor Proof
390
Layout: T1 Standard SC
Chapter No.: 18
Book ISBN: 978-3-642-31441-4
Page: 391/410
Polyploidization and Sex Chromosome Evolution in Amphibians
391
Table 18.1 A list of known polyploid amphibians compiled primarily from Schmid et al. (2010)
, Mable et al. (2011) and citations therein
Family and species
Ploidy
Family and species
Ploidy
(Frogs)
(Frogs continued)
Tetraploid
Tetraploid
Tetraploid
Tetraploid
F
Myobatrachidae
Neobatrachus aquilonius
Neobatrachus centralis
Arthroleptidae
Neobatrachus sudelli
Astylosternus diadematus Tetraploid Neobatrachus kunapalari
Microhylidae
Scaphiophryne gottlebei
Bufonidaea
Bufo asmaerae /
Tetraploid Aphantophryne(Cophixalus)
Amietophrynus
pansa
asmaerae
Bufo baturae
Triploid
Chiasmocleis leucosticta
Bufo kerinyagae
Tetraploid Odontophrynidae
Bufo oblongus and
Tetraploid Odontophrynus americanus
subspecies (synonym
of ‘‘B. danatensis’’)
Bufo pewzowi and
Tetraploid Pipidae
subspecies (also a
synonym of
‘‘B. danatensis’’)
Bufo poweri
Triploid
Silurana epitropicalis
Triploid
‘‘Silurana new tetraploid 1’’
Bufo pseudoraddeia
(‘‘Silurana sp. Nov VII’’,
‘‘Silurana paratropicalis’’)b,c
Bufo viridis
Triploid
‘‘Silurana new tetraploid 2’’b
a
Triploid
Xenopus borealis
Bufo zugmayeri
Xenopus clivii
Ceratophryidae
Xenopus fraseri
Tetraploid Xenopus gilli
Craugastoridae
Eleutherodactylus
(Haddadus) binotatus
Ceratophrys dorsata /
Octoploid Xenopus laevisd
Ceratophrys aurita
Ceratophrys joazeirensis Octoploid Xenopus muelleri
Ceratophrys ornata
Octoploid Xenopus pygmaeus
Xenopus largeni (‘‘Xenopus
sp. Nov. III’’)e
Triploid
PR
OO
Alsodidae
Eupsophus vertebralis
Tetraploid
Tetraploid
Tetraploid
Tetraploid
D
TE
EC
CO
RR
UN
Editor Proof
18
Book ID: 272454_1_En
Date: 16-8-2012
Tetraploid
Tetraploid
Tetraploid
Tetraploid
Tetraploid
Tetraploid
Tetraploid
Tetraploid
Tetraploid
Tetraploid
Tetraploid
(continued)
Layout: T1 Standard SC
Chapter No.: 18
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 392/410
B. J. Evans et al.
Family and species
(Frogs continued)
Dicroglossus
(Hoplobotrachus)
occipitalis
Tetraploid
‘‘Xenopus new tetraploid 1’’ (‘‘Xenopus
sp. Nov. VI’’,’’ Xenopus muelleri
west’’)e,f
Xenopus amieti
Tetraploid
Tetraploid
Xenopus andrei
Xenopus boumbaensis
Xenopus itombwensisg
Xenopus lenduensish
Xenopus vestitus
Xenopus wittei
‘‘Xenopus sp. nov. X’’e,i
Hylidae
Ploidy
Octoploid
Octoploid
Octoploid
Octoploid
Octoploid
Octoploid
Octoploid
Octoploid
Dodecaploid
Dodecaploid
Dodecaploid
Dodecaploid
Tetraploid
Tetraploid
Ranidae
Rana esculenta
Rana japonica*k
Triploid
Triploid
Triploid /
tetraploid
Rana nigromaculata**
Triploid /
Triploid
Triploid
Triploid
Rana pipiens*l
Rana rugosa*m
Triploid
Triploid
Triploid
CO
RR
Leiopelmatidae
Leiopelma
hochstetteri**
Family and species
(Salamanders)
Ambystomatidae
Ambystoma
jeffersonianum
Ambystoma
mexicanum**
tetraploid
Ambystoma nothagenesn
Ambystoma platineumn
Ambystoma tremblayin
Plethodontidae
Eurycea bislineata**
Xenopus longipes
Xenopus ruwenzoriensis
Tetraploid ‘‘X. cf. boumbaensis’’j
Octoploid ‘‘Xenopus sp. Nov. VIII’’e,i
Tetraploid Pyxicephalidae
Pyxicephalus (Tomopterna) delalandii
Tomopterna tandyi
Triploid
D
Leptodactylidae
Pleurodema bibroni
Pleurodema cordobae
Pleurodema kriegi
EC
Hyla versicolor
Phyllomedusa
tetraploidea
Tetraploid
PR
OO
Dicroglossidae
Ploidy
F
Ploidy
TE
Table 18.1 (continued)
Family and species
(Frogs)
UN
Editor Proof
392
Triploid /
tetraploid
(continued)
Layout: T1 Standard SC
Chapter No.: 18
18
Book ISBN: 978-3-642-31441-4
Page: 393/410
Polyploidization and Sex Chromosome Evolution in Amphibians
Table 18.1 (continued)
Family and species
(Frogs)
Editor Proof
Book ID: 272454_1_En
Date: 16-8-2012
Ploidy
Family and species
(Frogs continued)
393
Ploidy
Salamandridae
F
Triploid
Triploid
Triploid
Triploid
PR
OO
Pleurodeles waltl*
Ichthyosaura alpestris*
Cynops pyrrhogaster**o
Notophtalamus
viridescens**p
Lissotriton vulgaris**q
Sirenidaer
Siren intermedia
Siren lacertina
Triploid
Tetraploid
Tetraploid
CO
RR
EC
TE
D
Additional citations are provided for examples not included in these references, and for unnamed
species
* Experimentally induced
** Spontaneously observed, in some cases also experimentally induced
a
Following taxonomy of Stöck et al. (2009). Further work is needed to confirm stable triploid
ploidy of B. pseudoraddei and B. zugmayeri (Stöck et al. 2009)
b
Evans et al. (2004b)
c
‘‘Silurana paratropicalis’’ is a nomen nudem; see Blackburn DC and Beier M (2011)
d
Here we consider as X. laevis all diverged populations within this clade as identified by Evans
et al. (2004). This includes Xenopus sp. Nov. IX (a.k.a. X. congo 3) from Tymowska (1991), X.
petersi, and X. victorianus
e
Tymowska (1991)
f
Kobel et al. (1996)
g
Table 13 of Schmid et al. (2010) incorrectly lists Xenopus itombwensis as a dodecaploid
h
Evans et al. (2011)
i
Species status requires further investigation
j
Evans (2007)
k
Kawamura and Tokunaga (1952)
l
Briggs (1947)
m
Kashiwagi (1993)
n
These names refer to individuals that are not species in the sense of being reproductively
isolated lineages that persist through time. They are unisexual progeny resulting from hybridization of other species
o
Fankhauser et al. (1942)
p
Fankhauser (1941); Fankhauser and Watson (1942)
q
Litvinchuk et al. (1998)
r
Morescalchi and Olmo (1974) found Pseudobranchus striatus to be polyploid but this result
was not supported by the analysis of Moler and Kezer (1993)
240
241
242
243
polyploidization occurs and how frequently diploids outcompete polyploids, so it is
difficult to test this. It is also plausible, for example, that variation in ploidy level is a
neutral phenomenon influenced by stochastic survival and extinction of polyploids
and diploids, or by variation among lineages, including polyploids (Mayrose et al.
2011), in their ability to speciate by polyploidization.
UN
239
Layout: T1 Standard SC
Chapter No.: 18
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 394/410
250
251
252
253
254
255
256
257
258
259
260
261
262
263
264
265
266
267
268
269
270
271
272
273
274
275
276
277
278
279
280
281
282
283
284
285
286
F
249
PR
OO
248
About one third of the described polyploid frog species belong to the genus
Xenopus. At least six independent instances of genome duplication gave rise to the
ploidy levels seen among extant species in this group, including multiple episodes
that generated the highest ploidy level of any vertebrate—dodecaploidy (reviewed
in Evans 2008). Tetraploid Xenopus evolved at least once, octoploid Xenopus
evolved independently at least three times, and dodecaploid Xenopus evolved
independently at least two times (and possibly more depending on the species
status of Xenopus cf. boumbaensis and of Xenopus sp. nov. VIII; Table 1, Evans
2007, 2008; Evans et al. 2008a, 2011, 2005a; Tymowska 1991). Tetraploidy also
occurred independently in Silurana (Evans 2007; Evans et al. 2005a).
With respect to genome duplication, something is clearly special about Xenopus—
but what? One possible clue emerges from the recent discovery of the first known
genetic trigger of sex determination in amphibians by Yoshimoto et al. (2008). These
researchers identified a female-specific gene called DMW in the tetraploid species
Xenopus laevis. DMW is a W-chromosome linked gene that evolved via gene duplication from another important regulator of sexual differentiation called DMRT1
(Yoshimoto et al. 2008). DMW may function by blocking DMRT1 induction of testis
differentiation (Yoshimoto et al. 2010, 2008). Potentially relevant to the high incidence
of polyploidization in Xenopus is the discovery that DMW originated extremely
recently in amphibian evolution—after divergence of Silurana and Xenopus, but
before diversification of most or all extant species of Xenopus (Bewick et al. 2011). Not
surprisingly, the sex chromosomes of Xenopus are not cytologically distinct (Tymowska 1991). Gene contents of the W and Z chromosomes of Xenopus are therefore
probably very similar, and Xenopus species presumably lack mechanisms of dosage
compensation operating over most sex-linked genes because both sexes have two
alleles at most loci on the sex chromosomes. The preponderance of polyploids in
Xenopus is therefore consistent with the proposal that polyploidization is more likely to
occur in lineages with young, minimally degenerate sex chromosomes.
Another possible link between sex chromosome evolution and polyploidization is provided by Leiopelma hochstetteri. This species has intraspecific variation in the presence of a recently evolved univalent W chromosome that
governs sex determination in females (Green 1988). Leiopelma hochstetteri is
diploid but also has spontaneous triploidy (that is, polyploidy without speciation;
Green et al. 1984). It is not clear whether novel mechanisms for sex determination are more likely to evolve and persist in species that have nondegenerate
sex chromosomes, but this seems plausible under the same reasoning discussed
above with respect to the propensity for lineages to experience polyploidization.
More specifically, if a new pair of sex chromosomes appears in a population then
the old ones would segregate as a newly established autosomal pair. For this
reason, ancestral sex chromosomes with similar gene content would lack or have
few null alleles when they segregate autosomally. While the observation of
D
247
TE
246
EC
245
18.1.3 Examples of polyploidy in species with demonstrably
young sex chromosomes
CO
RR
244
B. J. Evans et al.
UN
Editor Proof
394
Layout: T1 Standard SC
Chapter No.: 18
Book ISBN: 978-3-642-31441-4
Page: 395/410
Polyploidization and Sex Chromosome Evolution in Amphibians
395
291
18.2 Evolution of Sex Determination Systems in Amphibians
292
18.2.1 Methods
296
297
298
299
300
301
302
303
304
305
306
307
308
309
310
311
312
313
314
315
316
317
318
319
320
321
322
323
324
PR
OO
295
Changes in the heterogametic sex, evolution of new triggers for sex determination,
and polymorphism in sex chromosomes mark the origin of novel features in
genetic pathways for sex determination. In order to quantify how many times this
has happened in amphibians, we began with the large amphibian phylogeny
reported by Pyron and Wiens (2011). We trimmed from this tree all species except
those for which we had information on either heterogamy or polyploidy, or both,
and retained the original maximum likelihood branch lengths among the retained
species. For illustrative purposes, we also retained diploid species (confirmed or
presumed) from the phylogeny of Pyron and Wiens (2011) that are closely related
to polyploid species. In many cases diploidy has been confirmed in these species or
other closely related species (see Supplementary Information of Mable et al.
2011). We had heterogamy information for Physalaemus (Engystomops) petersi,
but this species was not included in the phylogeny. Therefore, we used a closely
related species (Physalaemus cuvieri) that is included in the phylogeny to represent Physalaemus petersi. Physalaemus petersi was the only species from this
genus that was analyzed, so this substitution should be uncontroversial (note that
placing some Physalaemus in Engystomops makes no difference as Engystomops
and Phsalaemus are sister taxa). To better illustrate the phylogenetic distribution
of polyploid species in Fig. 18.1, we also substituted Chiasmocleis hudsoni, which
was present in the phylogeny of Pyron and Wiens (2011), with the tetraploid
species C. leucosticta, which was not present in the phylogeny of Pyron and Wiens
(2011). However, C. leucosticta was not included in any of the analyses described
below because we lack data on heterogamy for this species.
A total of 143 species (97 frogs, 45 salamanders, and one caecilian as an
outgroup) were included. We then converted this tree to a chronogram (a timecalibrated phylogeny) using the penalized likelihood approach (Sanderson 2002),
implemented in r8s version 1.71 (Sanderson 2003). We used the calibration points
detailed in Wiens (2011) but some had to be excluded given the more limited
taxon sampling used here, and some were added or modified given the differences
in taxon sampling (e.g., we added two calibration points within the genus Hyla,
given our more extensive sampling of species in that genus relative to Wiens
(2011). We used the following 17 calibration points. The first 16 were treated as
D
294
TE
293
EC
289
CO
RR
288
F
290
spontaneous triploidy suggests a tolerance of polyploidy, L. hochstetteri is not a
polyploid species, so a direct link between the age of the sex chromosomes and
polyploid speciation (as opposed to the toleration of polyploidy) is not established by this species.
287
UN
Editor Proof
18
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 18
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 396/410
B. J. Evans et al.
Necturus punctatus
Necturus alabamensis
Necturus maculosus
Necturus beyeri
Necturus lewisi
Aneides ferreus
Hydromantes flavus
Hydromantes supramontis
Hydromantes imperialis
Hydromantes italicus
Hydromantes ambrosii
Eurycea junaluska
Eurycea bislineata
Nototriton abscondens
Nototriton picadoi
Nototriton richardi
Oedipina parvipes
Oedipina pseudouniformis
Oedipina cyclocauda
Oedipina poelzi
Cryptotriton veraepacis
Dendrotriton rabbi
Chiropterotriton dimidiatus
Thorius dubitus
Pleurodeles waltl
Pleurodeles poireti
Triturus marmoratus
Triturus pygmaeus
Triturus dobrogicus
Triturus karelinii
Triturus cristatus
Triturus carnifex
Notophthalmus perstriatus
Notophthalmus viridescens
Ambystoma laterale
Ambystoma mexicanum
Ambystoma dumerilii
Ambystoma tigrinum
Pseudobranchus striatus
Pseudobranchus axanthus
Siren intermedia
Siren lacertina
Hynobius quelpaertensis
Hynobius tokyoensis
Hynobius hidamontanus
Leiopelma hamiltoni
Leiopelma pakeka
Leiopelma archeyi
Leiopelma hochstetteri
Bombina orientalis
Discoglossus pictus
Xenopus gilli
Xenopus laevis
Silurana epitropicalis
Silurana tropicalis
Hymenochirus boettgeri
Pelodytes punctatus
Rana clamitans
Rana catesbeiana
Rana pipiens
Rana sphenocephala
Rana blairi
Rana berlandieri
Rana tagoi
Rana japonica
Rana temporaria
Rana tsushimensis
Rana rugosa
Rana esculenta
Rana lessonae
Rana ridibunda
Rana plancyi
Rana nigromaculata
Buergeria buergeri
Hoplobatrachus crassus
Hoplobatrachus occipitalis
Tomopterna cryptotis
Tomopterna tandyi
Tomopterna delalandii
Pyxicephalus adspersus
Pyxicephalus edulis
Aphantophryne pansa
Cophixalus humicola
Scaphiophryne spinosa
Scaphiophryne gottlebei
Chiasmocleis leucosticta
Chiasmocleis shudikarensis
Astylosternus batesi
Astylosternus diadematus
Strabomantis biporcatus
Pristimantis pulvinatus
Pristimantis shrevei
Pristimantis euphronides
Eleutherodactylus johnstonei
Eleutherodactylus glamyrus
Eleutherodactylus cavernicola
Eleutherodactylus turquinensis
Eleutherodactylus cuneatus
Eleutherodactylus casparii
Eleutherodactylus emiliae
Eleutherodactylus albipes
Physalaemus petersi
Pleurodema thaul
Pleurodema bibroni
Pseudopaludicola falcipes
Vitreorana antisthenesi
Bufo boulengeri
Bufo viridis
Bufo balearicus
Bufo oblongus
Bufo pewzowi
Bufo bufo
Bufo regularis
Bufo poweri
Eupsophus calcaratus
Eupsophus roseus
Eupsophus insularis
Eupsophus migueli
Eupsophus vertebralis
Proceratophrys boiei
Odontophrynus cultripes
Odontophrynus americanus
Ceratophrys cranwelli
Ceratophrys ornata
Phyllomedusa bahiana
Phyllomedusa burmeisteri
Phyllomedusa tetraploidea
Phyllomedusa distincta
Hyla femoralis
Hyla chrysoscelis
Hyla versicolor
Hyla arborea
Hyla intermedia
Hyla sarda
Hyla meridionalis
Pseudis tocantins
Gastrotheca riobambae
Gastrotheca pseustes
Gastrotheca walkeri
Crinia signifera
Neobatrachus pictus
Neobatrachus sudelli
PR
OO
F
Editor Proof
396
Salamanders
CO
RR
EC
TE
D
Frogs
UN
Fig. 18.1 Heterogamy and polyploidy in salamanders and frogs. The phylogeny is from Pyron
and Wiens (2011) with branch lengths proportional to time based on a relaxed molecular clock
and calibration points described in the text. A black scale bar indicates 40 million years of
evolution. Red and blue rectangles on tips indicate male or female heterogamy respectively;
green, orange, and yellow rectangles indicate de novo sex determining systems; missing data on
heterogamy have no rectangles in this column. In the right column, black rectangles indicate
polyploid species; other species are either confirmed or assumed diploid. Some polyploid species
listed in Table 18.1 are not included in this figure due to a lack of phylogenetic information. Pie
charts on nodes indicate ancestral reconstructions of heterogametic state
Layout: T1 Standard SC
Chapter No.: 18
331
332
333
334
335
336
337
338
339
340
341
342
343
344
345
346
347
348
349
350
351
352
353
354
355
356
357
358
359
360
361
362
363
364
365
366
367
F
330
PR
OO
329
(1) Most recent common ancestor (MRCA) of extant salamanders, at least 150.8
Mya (Millions of years ago), based on the fossil Iridotriton hechti of the
Kimmeridgian/Early Tithonian (Late Jurassic), which is considered to be a
crown-group caudate (Evans et al. 2005b).
(2) MRCA of Salamandroidea (all salamanders exclusive of cryptobranchids,
hynobiids, and sirenids), at least 125.0 Mya (early Barremian, Cretaceous),
based on Galverpeton and Valdotriton (Evans and Milner 1996).
(3) MRCA of plethodontids and proteids, at least 65.5 Mya. The oldest known
amphiumid fossil (Proamphiuma cretacea) is late Maastrichtian or early
Paleocene, and thus from approximately 65.5 Mya (Gardner 2003). The split
between Plethodontidae and Amphiumidae must be at least this old. We do
not have amphiumids included here, but the clade of plethodontids and
proteids must be at least this old given the well-supported clade consisting of
proteids, rhyacotritonids, amphiumids, and plethodontids; see Pyron, Wiens
(2011) and earlier studies.
(4) MRCA of Aneides and Hydromantes, at least 19 Mya. Given the presence of
an Aneides vertebra in the Arikareean period (Tihen and Wake 1981), the
MRCA of the clade containing modern Aneides must be at least 19 Myo
(Millions of years old).
(5) MRCA of Triturus and Notophthalmus at least 33.9 Mya, based on fossils of
Triturus from the Eocene of Europe (33.9–55.8 Mya; Milner 2000)
(6) MRCA of Ambystomatidae and Salamandridae, at least 56.8 Mya, based on a
fossil dicamptodontid (Paleocene; Tiffanian; 60.2–56.8 Mya; Naylor and Fox
1993), and given that the ambystomatidae is the sister group to the Dicamptodontidae (so the sister group to Ambystomatidae ? Dicamptodontidae
must be at least this old).
(7) MRCA of frogs and salamanders, at least 245 Mya, based on a fossil anuran
(Triadobatrachus) from the Early Triassic (251–245 Mya) of Madagascar
(Carroll 1988; Rage and Rocek 1989)
(8) MRCA of pipoids and all other frogs, at least 145.5 Mya, given Rhadinosteus
parvus, ostensibly a rhinophrynid and clearly a pipoid, from the Late Jurassic
(Tithonian, 145.5–150.8 Mya; Rocek 2000).
(9) MRCA of Hymenochirus and Xenopus, at least 83.5 Mya, given the pipid Pachybatrachus taqueti from the Upper Cretaceous (Coniacian-Santonian, 83.5–89.3
Mya), which is thought to be closely related to Hymenochirus (Rocek 2000).
(10) MRCA of Myobatrachidae (represented here by the limnodynastine Neobatrachus and the myobatrachine Crinia) at least 54.6 Mya, given fossils
assigned to the limnodynastine genus Lechriodus (Evans et al., 2008b;
Sanchiz, 1998).
(11) MRCA of Bufonidae ? Leptodactylidae ? Centrolenidae, at least 55.8 Mya,
given putative fossil Bufo from the late Paleocene (55.8–58.7 Mya; Baéz 2000).
D
328
TE
327
397
constraints on the minimum age of each clade, and the final calibration point was a
fixed age for the root of the tree.
EC
326
CO
RR
325
Book ISBN: 978-3-642-31441-4
Page: 397/410
Polyploidization and Sex Chromosome Evolution in Amphibians
UN
Editor Proof
18
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 18
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 398/410
375
376
377
378
379
380
381
382
383
384
385
386
387
388
389
390
391
392
393
394
395
396
397
398
399
400
401
402
403
404
405
406
408
407
409
410
411
412
F
374
PR
OO
373
D
372
TE
371
EC
370
(12) MRCA of Ranidae (sensu Wiens et al. 2009) at least 33.9 Mya, given fossil
Rana from the Late Eocene (37.2–33.9 Mya; Rocek and Rage 2000).
(13) Crown group of Terrarana (the clade including the families Brachycephalidae,
Ceuthomantidae, Craugastoridae, Eleutherodactylidae, and Strabomantidae,
or more simply, the clade including Eleutherodactylus and related genera) at
least 35 Mya, based on an Eleutherodactylus fossil in amber from the La Toca
formation (Dominican Republic) estimated to be *35 Myo (Poinar and
Cannatella 1987).
(14) Stem group of Ceratophryidae (the clade including the genera Ceratophrys,
Chacophrys, and Lepidobatrachus) at least 65.5 Mya based on the late Cretaceous fossil genera Beelzebufo and Baurubatrachus (Evans et al. 2008b). Evans
et al. (2008b) considered the Madagascan taxon Beelzebufo to be a ceratophryine. This taxon is of Maastrichtian (Late Cretaceous) age (65.5–70.6
Mya). The South American genus Baurubatrachus is also considered to be a
ceratophryine (Evans et al. 2008b; Rocek 2000). Although the exact relationships of these taxa are somewhat uncertain, the presence of seemingly ceratophryine fossils in South America suggests that the stem group age of
Ceratophryidae is at least 65.5 Mya. The relationships of ceratophryids are
uncertain, but in this molecular analysis, they appear as the sister group to a
clade including Odontophrynidae and Alsodidae (Eupsophus).
(15) Crown-group age of North American and European Hyla clade, at least 16
Myo; given fossil Hyla similar to extant H. arborea and H. meridionalis in
the Lower Miocene of Austria (*16 Myo; Sanchiz 1998). We assume that
these Hyla are closely related to Hyla presently extant in Europe. However,
we cannot assume that these fossils are younger than the crown-group age
of the extant European species. We assume instead that the crown group of
the clade of Hyla is at least 16 Myo based on these European fossils.
(16) MRCA of H. avivoca-H. chrysocelis-H. versicolor clade; H. miocenica is
thought to be closely related to H. chrysocelis and H. versicolor and occurs
in the Barstovian of the Middle Miocene (14–16 Myo; Holman 2003). In
our phylogeny, H. avivoca, H. chrysocelis, and H. versicolor form a clade.
We assume that the stem group age of these three species is at least 14 Myo.
(17) We fixed the root age of the tree using the estimated age from Wiens (2011)
for the MRCA of lissamphibians (frogs, salamanders, caecilians) of 368.3
Mya, using penalized likelihood. Although the use of a fixed calibration
point (rather than a minimum constraint) may seem controversial, it should
be noted that at least one node must be given a fixed age. Furthermore, our
focus here is not on re-estimating these ages, but providing relative
assessments of clade ages (see below).
CO
RR
368
369
B. J. Evans et al.
UN
Editor Proof
398
Construction of a chronogram using r8s requires a cross-validation step that
identifies a best-fitting value for the ‘‘smoothing parameter’’, which specifies the cost
of differing rates of evolution between neighboring branches (Sanderson 2002).
Cross-validation considered smoothing parameter values from 100 –105.5 in exponential increments of 0.5. These cross-validation analyses failed until a species with a
Layout: T1 Standard SC
Chapter No.: 18
419
420
421
422
423
424
425
426
427
428
429
430
431
432
433
434
435
436
437
438
439
440
441
442
443
444
445
446
447
448
449
450
451
452
453
454
455
456
F
418
PR
OO
417
D
416
zero-length branch (Bufo pewzoi) was removed. After removing this species, the
cross-validation analyses showed that a value of 101 gave the lowest Chi-squared
error. We then generated a chronogram for the 142 remaining species using this
smoothing parameter. We then performed a second analysis using this same
smoothing parameter but including all 143 species. This second analysis gave
identical divergence-date estimates throughout the tree as the first analysis with 142
species. The resulting chronogram (Fig. 18.1) shows species for which we have
information on phylogenetic relationships and either heterogamy and/or ploidy.
Using the R package ‘‘ape’’ (Paradis et al. 2004), which is a software package for
phylogenetic analysis, we tested the fit of alternative models for the evolution of new
sex determination systems for a total of 90 species (55 frogs and 35 salamanders) for
which we had data on heterogamy or sex chromosome polymorphism. This was done
to select an appropriate model for ancestral reconstruction and for use in a permutation test described below. We coded all species as having either male heterogamy
(0), female heterogamy (1), or, for each of three species (Xenopus laevis, Leiopelma
hochstetteri, and Rana rugosa), a unique ‘‘de novo’’ sex-determining mechanism (2,
3, and 4), in which categories we include species with polymorphisms in mechanisms
for sex determination. Xenopus laevis was assigned a unique heterogametic state in
order to account for the finding that this species evolved its W-linked sex-determining gene after the split from S. tropicalis (Bewick et al. 2011), which also has
female heterogamy. Leiopelma hochstetteri was assigned a unique heterogametic
state in order to account for the finding that this species recently evolved a derived
W0 sex-determining system that is unique to this lineage (Green et al. 1993; Sharbel
et al. 1998). Rana rugosa was assigned a unique heterogamy state in order to
accommodate evidence for recent and possibly repeated instances of sex chromosome turnover (Ogata et al. 2008). The three de novo states were coded as separate
character states in order to ensure that known instances of novel sex determination
mechanisms were included in the analysis even though they did not necessarily
involve a change in heterogamy. We note that the newly evolved sex chromosomes
of L. hochstetteri and R. rugosa are polymorphisms, and it is not clear whether these
new polymorphisms will eventually fix in each species, and thus actually constitute a
sex chromosome turnover. However, in both of these examples, at least one of the
polymorphic systems for sex determination is species-specific and therefore new.
Using the ‘‘ace’’ function of the ‘‘ape’’ package, we then evaluated the
following models for evolution of the five heterogamy states:
TE
415
399
EC
414
CO
RR
413
Book ISBN: 978-3-642-31441-4
Page: 399/410
Polyploidization and Sex Chromosome Evolution in Amphibians
(1) All rates equal (one rate for all possible transitions between states, one parameter).
(2) One reversible rate between XY and ZW, and one reversible rate to and from
any of the de novo states (two rates in total, two parameters). By reversible, we
mean, for example, that the rate of change from XY to ZW is equal to the rate
of change from ZW to XY.
(3) One rate for XY to ZW, another rate for ZW to XY, and one reversible rate to
and from any of the de novo states (three rates in total, three parameters).
(4) Rates between each of the five heterogamy states are reversible and unique
(ten rates in total, ten parameters).
UN
Editor Proof
18
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 18
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 400/410
B. J. Evans et al.
(5) All rates unique (twenty rates in total, twenty parameters).
458
18.2.2 Results and Discussion
464
465
466
467
468
469
470
471
472
473
474
475
476
477
478
479
480
481
482
483
484
485
486
487
488
489
490
491
492
493
494
PR
OO
463
D
462
TE
461
The likelihoods of each of these models were compared using the Akaike Information Criterion (Akaike 1974) calculated as 2k–2ln(L) where k is the number of
parameters in the model and L is the maximum value of the likelihood function of the
model. A P value was generated with a hierarchical likelihood ratio test (hLRT) with
degrees of freedom equal to the difference in free parameters of the models under the
assumption of a Chi-squared distribution. For the hLRT we evaluated whether
adding complexity to the models resulted in a significant increase in model fit. The
likelihoods of Models 1, 2, 3, 4 and 5 were -70.74572 (AIC = 143.5), -53.82688
(AIC = 111.7), -53.81533 (AIC = 113.6), -52.31287 (AIC = 124.6), and
-49.97701 (AIC = 140.0) respectively. Model 2 thus was favored by the Akaike
Information Criterion. According to the hLRT, Model 2 was also preferred over
Model 1 (P\0.0001), but Model 3 was not preferred over Model 2 (P = 0.879).
Model 4 was not preferred over Model 2 (P = 0.932) or Model 3 (P = 0.885), and
Model 5 was not preferred over Model 2 (P = 0.982), or Model 3 (P = 0.999), or
Model 4 (P = 0.912). These results suggests that the transition rate from ZW to XY
is not significantly higher than the transition rate from XY to ZW. Model 2 was
therefore used to reconstruct ancestral heterogamy states and also used for simulations in our permutation test described below. The model used in the ancestral
reconstructions differs slightly from the model used in the permutation test in that the
rate of reversal from the de novo heterogamy states to other heterogamy states was set
to zero for the ancestral reconstructions. This was not possible with the permutation
test, which requires a reversible model. We present results from this slightly different
version for the ancestral reconstructions for illustrative purposes because with this
model there is zero likelihood for all of the de novo states in all of the ancestral
reconstructions. Other inferences, such as the likelihood of male and female heterogamy in the most recent common ancestor of frogs, of salamanders, and of frogs and
salamanders discussed below, are identical with both of these models.
The ancestral state reconstructions estimated from the analysis with 90 species
but plotted on the chronogram with 143 species (Fig. 18.1) suggest that there is not
strong statistical support to distinguish whether the ancestral hetrogamy state was
female or male heterogamy (that is, ZW females and ZZ males) in frogs or
salamanders. The marginal likelihood of female heterogamy (ZW) for the most
recent common ancestor of frogs is 0.544, for the most recent common ancestor of
salamanders is 0.499, and for the most recent common ancestor of frogs and
salamanders is 0.513. Thus, the support for ZW versus XY heterogamy as the
ancestral state in each group is effectively equivocal.
EC
460
CO
RR
459
F
457
UN
Editor Proof
400
Layout: T1 Standard SC
Chapter No.: 18
Book ISBN: 978-3-642-31441-4
Page: 401/410
Polyploidization and Sex Chromosome Evolution in Amphibians
401
497
18.3.1 Methods
503
504
505
506
507
508
509
510
511
512
513
514
515
516
517
518
519
520
521
522
523
524
525
526
527
528
529
530
531
532
533
534
PR
OO
502
D
501
TE
500
In our analysis, we consider the sex-determining system to have changed every
time that a change in heterogamy occurred from XY to ZW, from ZW to XY, or
from ZW or XY to one of the three de novo sex-determining systems. To quantify
the number of times that the system for sex determination changed in amphibians,
we used the stochastic character mapping approach proposed by Nielsen (2002) as
implemented by the R package ‘‘phytools’’ (Revell 2011). This approach simulates
character evolution on a phylogeny, conditioning on the observed character states
of the terminals. In this way, one can estimate the number of character-state
transitions that occurred and also evaluate where in the phylogeny changes are
likely to have occurred. The stochastic mapping simulations were performed using
a reversible version of Model 2 described above.
As discussed earlier, recent discoveries implicate sex chromosome turnover in
facilitating polyploid speciation or the tolerance of polyploidization, at least in
Xenopus and Leiopelma. These observations raise the question: is this a general
phenomenon in amphibians? Since a change in heterogamy necessarily involves a
change in the sex-determining system, we predicted that the time since a change in
heterogamy (XY to ZW or ZW to XY) would be lower in species that are polyploid or
that tolerate polyploidy than expected by chance, if the same number of polyploid
species (or polyploid-tolerant species) were to evolve randomly on the phylogeny.
We note that this hypothesis does not involve correlation between polyploidy and a
particular heterogamy state, so standard approaches to test for phylogenetic correlation among traits cannot be used to test this prediction. Instead, we developed a
novel permutation test that accommodates uncertainty in when and in which lineages
sex chromosome turnover occurred during amphibian evolution.
From the set of 90 species for which heterogamy information was available, we
identified five phylogenetically independent instances of stable or spontaneous
polyploidy. We emphasize that this is an underestimate of the number of independent polyplodization events, and we were able to use only five instances of
polyploidization because we lack heterogamy and/or phylogenetic information for
the other examples listed in Table 18.1. Diploidy has either been confirmed for the
other species for which heterogamy information was available (Mable et al. 2011)
or was assumed. The five examples of independent polyploidization or tolerance of
polyploidization are:
EC
499
CO
RR
498
F
496
18.3 Is Polyploidy Tolerated to a Greater Degree in Species
with Young Sex Chromosomes?
495
(1) Siren intermedia, a tetraploid, which may have descended from a tetraploid
ancestor that also gave rise to S. lacertina (Morescalchi and Olmo 1974). We
note that the polyploid status of the family Sirenidae has not been confirmed
by additional studies (Mable et al. 2011).
UN
Editor Proof
18
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 18
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 402/410
542
543
544
546
545
547
548
549
550
551
552
553
554
555
556
557
558
559
560
561
562
563
564
565
566
567
568
569
570
571
572
573
574
575
576
577
578
579
F
PR
OO
540
541
For this permutation test, we used as a test statistic the mean time since the origin
of the current heterogamy state for the five polyploid lineages. This mean was
calculated from 1,000 simulations that are conditioned on the observed heterogamy
states, using the stochastic character mapping approach described by Nielsen (2002)
and implemented by the R package ‘‘phytools’’ (Revell 2011). Each stochastic
mapping simulation provides one possible evolutionary scenario that is consistent
with the data. It was necessary to perform many (1,000) stochastic mapping
simulations for the observed data in order to accommodate uncertainty in these
evolutionary scenarios (that is, to accommodate uncertainty in the ancestral
reconstruction of the evolution of heterogamy).
If polyploidy tends to occur soon after a change in heterogamy, then the
observed test statistic should be smaller than the distribution of statistics calculated
after repeatedly randomly selecting five species across the tree to be polyploid, and
performing 1,000 stochastic character mapping simulations for each of the randomizations. The observed test statistic was therefore compared to a distribution of
statistics generated from 100 randomizations where, in each randomization, five
species are selected to be polyploid, with 1,000 stochastic character mapping
simulations that were conditioned on the observed heterogamy states performed
for each of the randomizations. The difference between the test statistic and the
randomizations therefore is that the mean path length (that is, for each polyploid,
the path length between the most recent change in heterogamy and the branch tip)
was calculated respectively either from real polyploid species (for the test statistic), or from five species selected at random from the 90 species for which we have
heterogamy data (for each of the randomizations).
This test is conservative in the sense that it does not consider subsequent, phylogenetically independent instances of polyploidization that occurred in Xenopus (three
additional independent instances of octoploidization and at least two additional independent instances of dodecaploidization). More specifically, because Xenopus has a
relatively young sex-determining system, a test statistic generated by counting each of the
independent polyploidization events in Xenopus is even lower than the test statistic that
counts polyploidization of Xenopus only once (see below).
One limitation of this analysis is that the frequency of transitions in amphibian
sex-determining systems is undoubtedly underestimated. It is possible, for example,
that changes in heterogamy occurred in other species that were excluded in the
D
539
TE
537
538
(2) Ambystoma mexicanum, a species with spontaneous triploidy (Humphrey 1963)
that is closely related to the unisexual triploids A. jeffersonianum, A. platineum,
and A. tremblayi (see notes on species status of unisexuals in Table 18.1).
(3) Xenopus laevis, a tetraploid species (Tymowska 1991). We excluded X. gilli
from this analysis even though we have heterogamy data for this species
because it shares a polyploid ancestor with X. laevis.
(4) Rana esculenta, a naturally occurring diploid/triploid hybridogenic species
formed from hybridization of R. lessonae and R. ridibunda (Uzzell et al. 1975).
(5) Leiopelma hochstetteri, a diploid species with spontaneous triploidy (Green
et al. 1984).
EC
536
CO
RR
535
B. J. Evans et al.
UN
Editor Proof
402
Layout: T1 Standard SC
Chapter No.: 18
Book ISBN: 978-3-642-31441-4
Page: 403/410
Polyploidization and Sex Chromosome Evolution in Amphibians
403
604
18.3.2 Results and Discussion
589
590
591
592
593
594
595
596
597
598
599
600
601
602
605
606
607
608
609
610
611
612
613
614
615
616
617
618
619
PR
OO
587
588
D
586
TE
584
585
EC
582
583
Stochastic mapping of heterogamy state, including the independent evolution of
three de novo sex chromosomes provides an average estimated number of times
that the sex chromosomes turned over of 32 (95% confidence interval: 25–41)
based on 1,000 simulations that were conditioned on the observed heterogamy
states. This is much higher than the maximum parsimony inference of only seven
changes by Hillis and Green (1990). In the example simulation depicted in
Fig. 18.2a, for instance, there are 28 changes in heterogamy.
These simulations also suggest that more changes occurred from male heterogamy to female heterogamy than the reverse, even though our model comparison
suggested that the rate of change in each direction was not significantly different.
Out of 1,000 stochastic mapping simulations, the mean number of changes from
male to female heterogamy (XY to ZW) was 22 (95 % confidence interval: 13–28)
and the mean number of changes from female to male heterogamy (ZW to XY)
was 7 (95 % confidence interval: 2–14). The example simulation depicted in
Fig. 18.2a is typical of the other simulations in the sense that there are 19 changes
CO
RR
581
F
603
analysis because differences between the sex chromosomes were not cytologically
detectable. It is also possible that some species experienced a change in the sexdetermining system that did not involve a change in heterogamy. Gastrotheca
pseustes, for instance, is known to have polymorphism in the morphology of the Y
chromosome (Schmid et al. 1990), but this species was coded as XY because these
size variants may involve homologous chromosomes and no change in the sexdetermining system. Intraspecific polymorphism in sex-determining mechanisms
has also been observed in Rana narina, Eleutherodactylus maussi, Rana japonica
and R. narina (Eggert 2005), but phylogenetic information was lacking from these
species in the phylogeny of Pyron and Wiens (2011). Furthermore, adding more taxa
might influence the inferred timing of the transitions to heterogamy, even if the
phylogeny remains the same (e.g., added taxa could subdivide long branches and
help clarify where on a given branch the heterogamy transition occurred).
Another limitation of this analysis is that we use the total time that the polyploid species have been in the observed heterogamy states for our test statistic,
rather than the difference between these times and the age of each polyploid
species. The latter difference would be a better metric for this test because it
focuses on events prior to polyploidization. However, it is difficult to estimate the
age of each polyploid (or of a randomly selected species in the permutation)
because in some cases the diploid ancestor of the polyploid is unknown or extinct
(see above), or other cases because we lack phylogenetic information from the
sister taxon. We note that results are contingent on the phylogeny and evolutionary
model, and that this analysis does not accommodate uncertainty in divergence
times and phylogenetic relationships.
580
UN
Editor Proof
18
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 18
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 404/410
B. J. Evans et al.
(a)
(b)
Salamanders
1
Frogs
PR
OO
5
2
F
Editor Proof
404
XY
ZW
de novo 1
de novo 2
de novo 3
4
3
622
623
624
625
626
627
628
629
630
631
632
633
634
635
636
from male to female heterogamy but only 6 changes from female to male
heterogamy.
The observed average path length to a change in the sex-determining system for
the five polyploids averaged over 1,000 stochastic mapping simulations, was
89.0 million years. This is not to suggest that no sex chromosome degeneration
occurred within this period. In fact, if sex chromosome degeneration in amphibians
occurred at a similar rate as it did in therian mammals (4.6 genes per million years;
Graves 2004), the ancestors of these polyploids probably did have somewhat
degenerate sex chromosomes. Rather, this result suggests that the amount of sex
chromosome degeneration that typically occurs within this period of time was not
of sufficient magnitude to prevent polyploidization or the tolerance of spontaneous
polyploidization. The permutation test indicates that this test statistic is not significantly lower than the distribution of statistics calculated when polyploids
evolved five times on random branches in this phylogeny (P = 0.059; average
path length in permutations was 141.2 million years and the standard deviation
was 32.7 million years), although the P value is close to 0.05. If we include six
independent polyploidizations in Xenopus (one tetraploidization, three
CO
RR
621
UN
620
EC
TE
D
Fig. 18.2 Simulations that stochastically map evolution of heterogamy form the basis of the
permutation test. Shown here is (a) an example of a stochastic mapping simulation for the
evolution of the observed heterogamy states and (b) highlighted simulated paths to each of the
five observed polyploid species. Species with missing heterogamy data from Fig. 18.1 have been
removed for these analyses and species names are omitted for clarity. In (b) simulated ages of the
observed heterogamy states for Ambystoma mexicanum, Siren intermedia, Leiopelma hochstetteri, Xenopus laevis, and Rana esculentia are labeled 1, 2, 3, 4, and 5 respectively. The test
statistic is the average age of the observed heterogamy state for these five polyploid species,
averaged over 1,000 stochastic mapping simulations. This test statistic is compared to analogous
calculations from 100 permutations where five species were randomly selected to be polyploid
and 1,000 stochastic mapping simulations were performed for each randomization
Layout: T1 Standard SC
Chapter No.: 18
643
644
645
646
647
648
649
650
651
652
653
654
655
656
657
658
659
660
661
662
663
664
665
666
667
668
669
670
671
672
673
674
675
676
677
678
679
680
681
F
642
PR
OO
641
D
640
TE
639
405
octoploidizations, and at least two dodecaploidizations; reviewed in Evans 2008)
in the observed test statistic, in addition to the four other examples of polyploidization itemized above, the average observed path length to a change in the sexdetermining system for the ten independent polyploid lineages is 68.1 million
years. This test statistic is significantly smaller than statistics calculated from 100
permutations where one of the five randomly selected polyploids is also assumed
to undergo six independent polyploidizations (P = 0.020; average path length in
permutations was 169.8 million years and the standard deviation was 55.3 million
years). Although there are at least six independent polyploidizations in Xenopus,
this test suffers from pseudoreplication in that these polyploid lineages may share
the same system for sex determination (i.e., DMW). Additional data on whether
other polyploid species of amphibians have male or female heterogamy would
clearly help illuminate the question of whether species with young sex chromosomes are more tolerant of polyploidization. It is surprising how little is known
about heterogamy of polyploid amphibians given that karyotypes of essentially all
of these species were inspected in order to identify polyploidy in the first place.
One possible reason for this dearth of information on heterogamy of polyploid
species is that many of these species may lack morphologically distinct sex
chromosomes. This proposal, if accurate, would be consistent with the contention
that polyploidization is better tolerated by species with minimally degenerate sex
chromosomes.
Additional insights into the influence of sex chromosome evolution and polyploidization may be gained from studies of laboratory-generated polyploid
Xenopus. Laboratory allopolyploidization in Xenopus duplicates autosomal chromosomes but generates female polyploid individuals with 3 Z chromosomes and 1
W chromosome and male polyploid individuals with 4 Z chromosomes (Kobel and
Du Pasquier 1986). Thus, the W chromosome of one of the ancestral diploids is not
inherited by Xenopus allopolyploids, and therefore never gets a chance to segregate as an autosome. In this way, the mechanism of Xenopus allopolyploidization
in nature may circumvent autosomal segregation of a W chromosome (whether
degenerate or not) (reviewed in Evans 2008), and this could account for the
unusually high incidence of polyploidization in this genus.
Analyses presented here show that changes in the system for sex chromosome
turnover were much more common in amphibians than previously proposed [*32
versus 7 as proposed by Hillis and Green (1990)], and that there is not strong
support for female versus male heterogamy in the ancestor of salamanders, frogs,
or the most recent common ancestor of salamanders and frogs. These changes need
not involve the evolution of completely novel systems for sex determination, and
some of these inferred changes may be reversals to an ancestral system. We also
found that changes from male to female heterogamy occurred more frequently
than changes from female to male heterogamy, although the transition rates
between each state were not significantly different. This result also contradicts the
conclusions of Hillis and Green (1990), who stated that there was a bias in
evolution from female heterogamy to male heterogamy. One reason for these
differences is that our analysis included new data on heterogamy (reviewed in
EC
638
CO
RR
637
Book ISBN: 978-3-642-31441-4
Page: 405/410
Polyploidization and Sex Chromosome Evolution in Amphibians
UN
Editor Proof
18
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 18
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 406/410
B. J. Evans et al.
691
18.4 Conclusions
685
686
687
688
689
PR
OO
684
F
690
Schmid et al. 2010), more species, and a more comprehensive phylogeny
(Pyron and Wiens 2011). Another reason for our higher estimate in the number of
changes is that we considered three de novo changes in sex determination as a new
heterogamy state, thereby forcing a sex chromosome turnover in these lineages
(but this is only 3 out of *32 changes). Differences in the resolution, relationships,
and branch lengths of the phylogenies used in each study are also likely to have
played a role in these differing conclusions. Finally, and importantly, the different
analytical approaches may have influenced the results (that is, the use of maximum
parsimony by Hillis and Green (1990) and maximum likelihood here).
682
683
703
704
705
706
Acknowledgements We are particularly grateful to Barbara Mable who provided a comprehensive critical assessment of an earlier version of this chapter. We also thank Liam Revell for
advice and assistance with the R package ‘‘phytools’’ and Jim Bogart and Ben Bolker for
comments.
707
References
708
709
710
711
712
713
714
715
716
717
718
719
Akaike H (1974) A new look at the statistical model identification. IEEE Trans Autom Contr
19:716–723
Arnold AP, Itoh Y, Melamed E (2008) A bird’s-eye view of sex chromosome dosage
compensation. Annu Rev Genomics Hum Genet 9:109–127
Baéz AM (2000) Tertiary anurans from South America. In: Heatwole H, Carroll RL (eds)
Amphibian biology. Surrey Beatty, Chipping Norton, Australia, pp 1388–1401
Beçak ML, Beçak W (1998) Evolution by polyploidy in amphibia: new insights. Cytogenet Cell
Genet 80:28–33
Bergero R, Charlesworth D (2009) The evolution of restricted recombination in sex
chromosomes. Trends Ecol Evol 24:94–102
Bergero R, Charlesworth D (2011) Preservation of the Y transcriptome in a 10 million-year-old
plant sex chromosome system. Curr Biol 21:1470–1474
696
697
698
699
700
701
TE
695
EC
694
CO
RR
693
D
702
Polyploidization generates new species and duplicates genes; the resulting genetic
redundancy has the capacity to degrade or to undergo innovation. The question of
why some lineages frequently undergo polyploidization whereas others do not thus
has important implications for evolution and adaptation. Eventually we will have a
much more comprehensive understanding of genetic variation in (a) the triggers of
sex determination in amphibians, (b) the extent of suppressed recombination that
surrounds these genetic triggers, (c) the extent of sex chromosome degeneration that
exists in amphibians, and (d) whether or not other lineages of polyploid amphibians
have minimally degenerate sex chromosomes. Future discoveries in these areas can
undoubtedly be leveraged to provide exciting new insights into the role of sex
chromosome degeneration in the propensity of species to tolerate polyploidization.
692
UN
Editor Proof
406
Layout: T1 Standard SC
Chapter No.: 18
407
EC
TE
D
PR
OO
F
Bewick AJ, Anderson DW, Evans BJ (2011) Evolution of the closely related, sex-related genes
DM-W and DMRT1 in African clawed frogs (Xenopus). Evolution 65:698–712
Blackburn DC and Beier M (2011) ‘‘Xenopus paratropicalis’’ is not a valid name. Zootaxa
3035:57–58
Briggs R (1947) The experimental production and development of triploid frog embryos. J Exp
Zool 106:237–266
Bogart JP (1980) Evolutionary significance of polyploidy in amphibians and reptiles. In: Lewis
WH (ed) Polyploidy, biological relavance. Basic life sciences, New York, pp 341–378
Carroll RL (1988) Vertebrate paleontology and evolution W. H. Freeman and Company, New York
Charlesworth B, Charlesworth D (2000) The degeneration of Y chromosomes. Philos Trans R
Soc London B 355:1563–1572
Charlesworth D (2002) Plant sex determination and sex chromosomes. Heredity 88:94–101
Charlesworth D, Charlesworth B, Mariais G (2005) Steps in the evolution of heteromorphic sex
chromosomes. Heredity 95:118–128
Duellman WE, Trueb L (1994) Biology of amphibians. The Johns Hopkins University Press, Baltimore
Eggert C (2005) Sex determination: the amphibian models. Reprod Nutr Dev 44:539–549
Evans BJ (2007) Ancestry influences the fate of duplicated genes millions of years after
duplication in allopolyploid clawed frogs (Xenopus). Genetics 176:1119–1130
Evans BJ (2008) Genome evolution and speciation genetics of allopolyploid clawed frogs
(Xenopus and Silurana). Front Biosci 13:4687–4706
Evans BJ, Cannatella DC, Melnick DJ (2004a) Understanding the origins of areas of endemism in
phylogeographic analyses: a reply to Bridle et al. Evolution 58:1397–1400
Evans BJ, Kelley DB, Tinsley RC, Melnick DJ, Cannatella DC (2004b) A mitochondrial DNA
phylogeny of clawed frogs: phylogeography on sub-Saharan Africa and implications for
polyploid evolution. Mol Phylogenet Evol 33:197–213
Evans BJ, Carter TF, Hanner R et al (2008a) A new species of clawed frog (genus Xenopus), from
the Itombwe Plateau, democratic republic of the congo: implications for DNA barcodes and
biodiversity conservation. Zootaxa 1780:55–68
Evans BJ, Greenbaum E, Kusamba C et al (2011) Description of a new octoploid frog species
(Anura: Pipidae: Xenopus) from the democratic republic of the congo, with a discussion of the
biogeography of African clawed frogs in the Albertine Rift. J Zool London 283:276–290
Evans BJ, Kelley DB, Melnick DJ, Cannatella DC (2005a) Evolution of RAG-1 in polyploid
clawed frogs. Mol Biol Evol 22:1193–1207
Evans SE, Jones MEH, Krause DW (2008b) A giant frog with South American affinities from the
Late Cretaceous of Madagascar. Proc Nat Acad Sci 105:2951–2956
Evans SE, Lally C, Chure DC, Elder A, Maisano JA (2005b) A new fully metamorphosed
salamander from the Late Jurassic of North America. Zool J Linn Soc 143
Evans SE, Milner AR (1996) A metamorphosed salamander from the early Cretaceous of Las
Hoyas, Spain. Philos Trans R Soc London B 351:627–646
Ezaz T, Stiglec R, Veyrunes F, Graves JAM (2006) Relationships between vertebrate ZW and
XY sex chromosome systems. Curr Biol 16:R736–R743
Felsenstein J (1974) The evolutionary advantage of recombination. Genetics 78:737–756
Fankhauser G, Crotta R, Perrot M (1942) Spontaneous and cold-induced triploidy in the Japanese
newt Triturus pyrrhogaster. J Exp Zool 89:167–181
Fankhauser G (1941) The frequency of polyploidy and other spontaneous aberrations of chromosome
number among larvae of the newt Triturus viridescens. Proc Nat Acad Sci 27(11):507–512
Fankhauser G, Watson RC (1942) Heat-indiced triploidy in the newt, Triturus viridescens. Proc
Nat Acad Sci 28:436–440
Gardner JD (2003) The fossil salamander Proamphiuma cretacea Estes (Caudata: Amphiumidae)
and relationships within the Amphiumidae. J Vertebr Paleontol 23:769–782
Graves JAM (2004) The degenerate Y chromosome—can conversion save it? Reprod Fertil Dev
16:527–534
Graves JAM (2008) Weird animal genomes and the evolution of vertebrate sex and sex
chromosomes. Annu Rev Genet 42:565–586
CO
RR
720
721
722
723
724
725
726
727
728
729
730
731
732
733
734
735
736
737
738
739
740
741
742
743
744
745
746
747
748
749
750
751
752
753
754
755
756
757
758
759
760
761
762
763
764
765
766
767
768
769
770
771
772
773
Book ISBN: 978-3-642-31441-4
Page: 407/410
Polyploidization and Sex Chromosome Evolution in Amphibians
UN
Editor Proof
18
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 18
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 408/410
EC
TE
D
PR
OO
F
Green DM (1988) Cytogenetics of the endemic New Zealand frog, Leiopelma hochstetteri:
extraordinary supernumerary chromosome variation and a unqie sex-chromosome system.
Chromosoma 97:55–70
Green DM, Kezer J, Nussbaum RA (1984) Triploidy in Hochstetter’s frog, Leiopelma
hochstetteri, from New Zealand. New Zealand J Zool 11:457–460
Green DM, Zeyl CW, Sharbel TF (1993) The evolution of hypervariable sex and supernumerary (B)
chromosomes in the relict New Zealand frog, Leiopelma hochstetteri. J Evol Biol 6:417–441
Gregory TR, Mable BK (2005) Polyploidy in animals. In: Gregory TR (ed) The Evolution of the
Genome. Elsevier Academic Press, Burlington, pp 428–517
Hayes TB (1998) Sex determination and primary sex differentiation in amphibians: genetic and
developmental mechanisms. J Exp Zool 281:373–399
Hillis DM, Green DM (1990) Evolutionary changes of heterogametic sex in the phylogenetic
history of amphibians. J Evol Biol 3:49–64
Holloway AK, Cannatella DC, Gerhardt HC, Hillis DM (2006) Polyploids with different origins
and ancestors form a single sexual polyploid species. Am Nat 167:E88–E101
Holman JA (2003) Fossil frogs and toads of North America Indiana University Press.
Bloomington and Indianapolis, IN
Kashiwagi K (1993) Production of triploids and their reproductive capacity in Rana rugosa. Sci
Rep Lab Amphibian Biol Hiroshima Univ 12:23–36
Kawamura T, Tokunaga C (1952) The sex of triploid frogs, Rana japonica Günther. J Sci
Hiroshima Univ, Ser B, Div 1 (Zoology) 13
Humphrey RR (1963) Polyploidy in the Mexican axolotl (Ambystoma mexicanum) resulting from
multinucleate ova. Proc Nat Acad Sci 50:1122–1127
Kobel HR, Loumont C, Tinsley RC (1996) The extant species. In: Tinsley RC, Kobel HR (eds)
The Biology of Xenopus. Clarendon Press, Oxford, pp 9–33
Kawamura T (1984) Polyploidy in amphibians. Zool Sci 1:1–15
Kobel HR (1996) Allopolyploid speciation. In: Tinsley RC, Kobel HR (eds) The Biology of
Xenopus. Clarendon Press, Oxford, pp 391–401
Kobel HR, Du Pasquier L (1986) Genetics of polyploid Xenopus. Trends Genet 2:310–315
Litvinchuk SN, Rosanov JM, Borkin LJ (1998) A case of natural triploidy in a smooth newt Triturus
vulgaris (Linneaus, 1958), from Russia (Caudata: Salamandridae). Herpetozoa 11:93–95
Mable BK (2004) ‘Why polyploidy is rarer in animals than in plants’: myths and mechanisms.
Biol J Linn Soc 82:453–466
Mable BK, Alexandrou MA, Taylor MI (2011) Genome duplication in amphibians and fish: an
extended synthesis. J Zool 284:151–182
Mable BK, Roberts JD (1997) Mitochondrial DNA evolution in the genus Neobatrachus (Anura:
Myobatrachidae). Copeia 1997:680–689
Mayrose I, Zhan SH, Rothfels CJ et al (2011) Recently formed polyploid plants diversify at lower
rates. Science 333:1257
Milner AR (2000) Mesozoic and Tertiary Caudata and Albanerpetontidae. In: Heatwole H, Carrol
RL (eds) Amphibian Biology. Surrey Beatty, Chipping Norton, Australia, pp 31–108
Morescalchi A, Olmo E (1974) Sirenids: a family of polyploid urodeles? Experientia 30:491–492
Moler PE, Kezer J (1993) Karyology and systematics of the salamander genus Pseudobranchus
(Sirenidae). Copeia 1993:39–47
Muller HJ (1925) Why polyploidy is rarer in animals in plants. Am Nat 59:346–353
Muller HJ (1964) The relation of recombination to mutational advance. Mutat Res 106:2–9
Naylor BG, Fox RC (1993) A new ambystomatid salamander Dicamptodon antiquus n. sp. from
the Paleocene of Alberta. Can J Earth Sci 30:814–818
Nielsen R (2002) Mapping mutations on phylogenies. Syst Biol 51:729–739
Ogata M, Hasegawa Y, Ohtani H, Mineyama M, Miura I (2008) The ZZ/ZW sex-determining
mechanism originated twice and independently during evolution of the frog, Rana rugosa.
Heredity 100:92–99
Ohno S (1967) Sex chromosomes and sex-linked genes. Springer, Berlin
Orr HA (1990) ‘Why polyploidy is rarer in animals than in plants’ revisited. Am Nat 136:759–770
CO
RR
774
775
776
777
778
779
780
781
782
783
784
785
786
787
788
789
790
791
792
793
794
795
796
797
798
799
800
801
802
803
804
805
806
807
808
809
810
811
812
813
814
815
816
817
818
819
820
821
822
823
824
825
826
827
B. J. Evans et al.
UN
Editor Proof
408
Layout: T1 Standard SC
Chapter No.: 18
409
EC
TE
D
PR
OO
F
Otto SP, Whitton J (2000) Polyploid incidence and evolution. Annu Rev Genet 34:401–437
Papp B, Pál C, Hurst LD (2003) Dosage sensitivity and the evolution of gene families in yeast.
Nature 424:194–197
Paradis E, Claude J, Strimmer K (2004) APE: analysis of phylogenetics and evolution in R
language. Bioinf 20:289–290
Poinar GO, Cannatella DC (1987) An upper Eocene frog from the Dominican Republic and its
implication for Caribbean biogeography. Science 237:1215–1216
Pyron RA, Wiens JJ (2011) A large-scale phylogeny of amphibia including over 2800 species,
and a revised classification of extant frogs, salamanders, and caecilians. Mol Phylogenet Evol
61:543–583
Qian W, Zhang J (2008) Gene dosage and gene duplicability. Genetics 179:2319–2324
Rage JC, Rocek Z (1989) Redescription of Triadobatrachus massinoti (Piveteau, 1936) an anuran
amphibian from the early Triassic. Palaeontographica Paleontologica 206:1–16
Revell JJ (2011) Phytools: phylogenetic tools for comparative biology (and other things). (R Package)
Rocek Z (2000) Mesozoic anurans. In: Heatwole H, Carrol RL (eds) Amphibian biology. Surrey
Beatty, Chipping Norton, Australia, pp 1295–1331
Rocek Z, Rage J-C (2000) Tertiary Anura of Europe, Africa, Asia, North America, and Australia.
In: Heatwole H, Carrol RL (eds) Amphibian Biology. Surrey Beatty, Chipping Norton,
Australia, pp 1332–1387
Sanchiz FB (1998) Salienta. In: Wellnhofer P (ed) Encyclopedia of paleoherpetology, Part 4,
Salienta. Verlag, Pfeil, Munich, pp 1–276
Sanderson MJ (2002) Estimating absolute rates of molecular evolution and divergence times: a
penalized likelihood approach. Mol Biol Evol 19:1218–1231
Sanderson MJ (2003) r8s: inferring absolute rates of evolution and divergence times in the
absence of a molecular clock. Bioinformatics 19:301–302
Schmid M, Sims SH, Haaf T, Macgregor HC (1986) Chromosome banding in amphibia X. 18S
and 28S ribosomal RNA genes, nucleolus organizers and nucleoli in Gastrotheca riobambae.
Chromosoma 94:139–145
Schmid M, Steinlein C (2001) Sex chromosomes, sex-linked genes, and sex determination in the
vertebrate class Amphibia. In: Scherer G, Schmid M (eds) Genes and mechanisms in
vertebrate sex determination. Verlag, Basel, pp 143–176
Schmid M, Steinlein C, Bogart JP et al (2010) The chromosomes of terraranan frogs. Cytogenetic
Genome Res 130–131:1–568
Schmid M, Steinlein C, Friedl R et al (1990) Chromosome banding in Amphibia. XV. Two types
of Y chromosomes and heterochromatin hypervariability in Gastrotheca pseustes (Anura,
Hylidae). Chromosoma 99:413–423
Sharbel TF, Green DM, Houben A (1998) B-chromosome origin in the endemic New Zealand
frog Leiopelma hochstetteri through sex chromosome devolution. Genome 41:14–22
Skaletsky H, Kuroda-Kawaguchi T, Minx PJ et al (2003) The male-specific region of the human
Y chromosome is a mosaic of discrete sequence classes. Nature 432:823–837
Stöck M, Horn A, Grossen C et al (2011) Ever-young sex chromosomes in European tree frogs.
PLoS Biol 9:e1001062
Stöck M, Ustinova J, Lamatsch DK et al (2009) A vertebrate reproductive system involving three
ploidy levels: Hybrid origin of triploids in a contact zone of diploid and tetraploid Paleartic
green toads (Bufo viridis subgroup). Evolution 64:944–959
Straub T, Becker PB (2007) Dosage compensation: the beginning and end of a generalization. Nat
Rev Genet 8:47–57
Svartman M, Stone G, Stanyon R (2005) Molecular cytogenetics discards polyploidy in
mammals. Genomics 85:425–430
Tihen JA, Wake DB (1981) Vertebrae of plethodontid salamanders from the Lower Miocene of
Montana. J Herpetology 15:35–40
Tymowska J (1991) Polyploidy and cytogenetic variation in frogs of the genus Xenopus.
In: Green DS, Sessions SK (eds) Amphibian cytogenetics and evolution. Academic Press, San
Diego, pp 259–297
CO
RR
828
829
830
831
832
833
834
835
836
837
838
839
840
841
842
843
844
845
846
847
848
849
850
851
852
853
854
855
856
857
858
859
860
861
862
863
864
865
866
867
868
869
870
871
872
873
874
875
876
877
878
879
880
881
Book ISBN: 978-3-642-31441-4
Page: 409/410
Polyploidization and Sex Chromosome Evolution in Amphibians
UN
Editor Proof
18
Book ID: 272454_1_En
Date: 16-8-2012
Layout: T1 Standard SC
Chapter No.: 18
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 410/410
EC
TE
D
PR
OO
F
Uzzell T, Berger L, Günther R (1975) Diploid and triploid progeny from a diploid female of Rana
esculenta (Amphibia Salientia). Proc Acad Nat Sci Philadelphia 127:81–91
Wiens JJ (2011) Re-evolution of lost mandibular teeth in frogs after more than 200 million years,
and re-evaluating Dollo’s Law. Evolution 65:1283–1296
Wiens JJ, Sukumaran J, Pyron RA, Brown RM (2009) Evolutionary and biogeographic origins of
high tropical diversity in Old World frogs (Ranidae). Evolution 63:1217–1231
Wolfe KH (2001) Yesterdays’s polyploids and the mystery of diploidization. Nat Rev Genet
2:333–341
Yoshimoto S, Ikeda K, Izutsu Y et al (2010) Opposite roles of DMRT1 and its W-linked paralog,
DM-W, in sexual dimorphism of Xenopus laevis: implications of a ZZ/ZW-type sexdetermining system. Development 137:2519–2526
Yoshimoto S, Okada E, Umemoto H et al (2008) A W-linked DM-domain gene, DM-W, participates
in primary ovary development in Xenopus laevis. Proc Nat Acad Sci 105:2469–2474
CO
RR
882
883
884
885
886
887
888
889
890
891
892
893
894
B. J. Evans et al.
UN
Editor Proof
410
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 411/415
Index
PR
OO
F
1
2
19
20
21
22
23
24
25
26
27
28
29
B
Background
selection, 388
Bichir, 352
Bivalent formation/bivalent, 218, 219
Bivalent(s), 37–41, 43–45
Bottom-up approach
(to genome alignment), 97
Bowfin, 351, 360, 367
Brachypodium, 96
Brassica napus, 283, 285
30
31
32
33
34
35
Cephalochordate, 309, 310, 313, 316, 325
Chiasma, 218, 219
Chromosome, 272, 280, 283, 284, 287
non-reduction, 99
number, 78–80
Cod, 343
Comparative expression
profiling, 202
Compensating
aneuploidy, 283
Concerted evolution, 103, 193
Conserved synteny, 346, 351–355, 360
Cotton, 183, 273, 282, 285
Crossover (co), 34, 39–41
Cytomixis, 42
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
D
Detecting whole genome duplication, 77–82,
84, 85, 87, 88
Diakinesis, 34, 42
Diploidization, 386
Diplotene, 34
Disomic, 43
Disomic inheritance, 218–220
Divergent resolution/reciprocal gene loss after
polyploidization, 9, 10, 12
DNA replication, 34, 35
Dosage
balance, 293, 302
compensation, 386, 387
sensitivity, 281
Dosage/dosage balance and
duplicate genes, 2, 5, 14
Double reduction, 43, 44, 46
Double strand breaks (DSB), 34, 39, 40
Duplicate gene expression, 192, 195
51
52
53
54
55
56
57
58
59
60
61
62
63
64
65
66
67
68
69
70
D
A
Allopolyploidy, 7, 8, 9, 22, 183, 184, 245–247,
250, 252, 254, 255, 257, 259–263,
272, 282
Amphibians, 385, 387–389, 394, 395, 401,
404–406
Amplification fragment length polymorphism
(AFLP), 245, 252, 257, 262
Anaphase, 34, 41, 43, 46
Aneuploidy, 26, 218, 272, 283, 285
Apomixis, 47, 48
Arabidopsis suecica, 285
Arabidopsis thaliana, 285
Autonomous pairing sites (APS), 37
Autopolyploids, 7, 8, 23, 98
Axial element (AE), 35
CO
RR
EC
TE
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
UN
Editor Proof
Layout: T1 Standard SC
Chapter No.: BM
C
Calandinos, 345
Carp, 341, 345, 347, 348
Catfish, 345, 367
Centromere, 34, 35, 39, 41, 43
Centromeric divergence, 100
P. S. Soltis and D. E. Soltis (eds.), Polyploidy and Genome Evolution,
DOI: 10.1007/978-3-642-31442-1, Springer-Verlag Berlin Heidelberg 2012
411
Layout: T1 Standard SC
Chapter No.: BM
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 412/415
Index
E
Ecological and adaptive consequences of
hybridization and polyploidy in
spartina, 231
Epigenetic, 216–218, 245, 246, 249,
257, 262–264
Expression, 272, 280–282, 285–287
Expression stoichiometry, 386
79
80
81
82
83
84
85
86
87
88
89
90
F
Fertility, 210, 217–219
Fiber, 202
Fish, 341–344, 349, 350, 352
Fluorescence in situ hybridization (fish), 213,
283, 286, 287
Fractionation
Biased, 141
Mechanism, 143
Fugu, 342, 349, 350, 363, 365
Functional
coherence, 104
91
92
93
94
95
96
97
98
99
100
101
102
103
104
105
106
107
108
109
110
111
112
113
114
115
116
117
118
119
120
G
Gamete(s), gametogenesis, 33, 34, 42, 43,
46–48
Gametophyte, 42, 47
Gar, 342, 351, 352, 360, 366, 367
Gene
balance hypothesis, 281
conversion, 194
copy number, 80, 81
duplication and loss, 85–87
Chordate, 309, 310, 313, 315, 322, 324,
325, 327, 328
Vertebrate, 309, 310, 313, 315, 316,
319, 321, 327, 328
dosage, 283
expression, 27, 245, 252, 256, 257, 261,
262, 264
function evolution, 310, 321, 322
loss, 101, 191
Gene tree
parsimony, 82, 83–85
reconciliation, 81–87
Genome
dominance, 141
doubling, 127
duplication, 293, 300, 341, 343, 344, 349,
351, 353
duplication and evolution, 309, 310, 313,
316, 323
evolution, 112, 117, 127
evolution following hybridization and
allopolyploid speciation in spartina,
233
groups, 187
rearrangement, 139, 140
sequence(s), 94
size, 189
Genomic balance, 27
Genomic dominance, 141, 198
Genomic downsizing, 189
Genomic in situ hybridization (GISH), 211,
283, 284, 286, 287
Genomic plasticity, 109, 127
Ghost duplication, 98
Goldfish, 345, 347
Gossypium, 188, 271, 284
barbadense, 188
hirsutum, 187
121
122
123
124
125
126
127
128
129
130
131
132
133
134
135
136
137
138
H
Heterodimerization of duplicate genes, 4
Heterogametic;Heterogamy, 387–389, 400,
403, 405
Heterosis, 25
Hexaploids, 28
Hierarchical clustering approach
(to genome alignment), 97
Hill–Robertson effect, 388
Hitchhiking, 388
Homeolog, 195, 278–282, 284–286
Homoelogous (nonhomologous
chromosomes), 38, 39
Homoeolog expression, 197–200
Homoeologous gene expression, 197
Homoeologous pairing, 188
Homoeologue, 257, 261
Homogametic;Homogamy, 387
Homogenization/intergenomic homogenization, 211, 217, 218
Homologous (chromosomes)
homology, 34–40, 45
Hox, 313, 314, 316, 318, 320, 346, 347, 349,
350, 352, 354, 355, 363, 364, 366, 370
Hybridization, 38, 40–43, 46, 47, 245–247,
250, 252–257, 260, 261, 264
139
140
141
142
143
144
145
146
147
148
149
150
151
152
153
154
155
156
157
158
159
160
161
162
163
164
I
Illegitimate recombination, 190
Introduction, 226, 227, 230, 232
Inversion(s)/chromosomal inversion(s), 217,
219
ITS
165
166
167
168
169
170
AQ1
CO
RR
EC
TE
D
PR
OO
F
71
72
73
74
75
76
77
78
UN
Editor Proof
412
Layout: T1 Standard SC
Chapter No.: BM
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 413/415
413
K
Karyotype, 272, 283, 284
evolution, 100
restructuring, 210, 217, 218, 220
175
176
L
Leptotene, 34, 35
177
178
179
180
181
182
183
184
185
186
187
188
189
190
M
Medaka, 342, 350–352, 359, 360, 363, 366,
370, 371
Meiosis, 33
Metaphase, 33, 34
Methylation-sensitive amplification
polymorphism (msap), 257, 260–262
Microarray, 197, 252–257, 260, 261, 264
Monosomy, 272, 283–286
Muller’s ratchet, 388
Multivalent(s), 36–39, 41, 43, 44, 46, 47
formation, 7, 8
formation after
polyploidization, 211, 218, 219
191
192
193
194
195
196
197
198
199
200
201
202
203
204
205
206
207
208
209
210
N
Neofunctionalization, 363, 365–367,
370, 371
of duplicate genes, 3, 11, 14
of pathways, 6
Next-generation sequencing (NGS), 211,
213–215
Nobilization, 99
Non-crossover, 40
Nonfunctionalization, 346, 354, 355, 358,
361–363, 365, 369
Nonfunctionalization
of duplicate genes, 2
Nonreciprocal
exchange, 194
homoeologous recombination, 191
Nuclear-cytoplasmic interaction
hypothesis, 215
Null allele, 394
Nullisomy, 272, 283, 284, 286
211
212
213
214
215
216
P
Pachytene, 33, 34, 36, 42
Pairing (of chromosomes), 33
Pairing-partner switch(es) (PPS), 37–39, 41
Paleopolyploidy, 95
Ph1 locus, 38, 39
Placode, 309, 313, 318, 327–329
Platyfish, 342
Poaceae (gramineae), 94
Polyploidization, 341, 344, 345, 348,
350, 362, 371
Polyploidy, 129
Polysomic, 45
Presence absence variation, 141, 142
Prophase, 34, 41
Psuedogenization, 191, 193
Pufferfish, 345, 350–352, 366
Punnett square, 43
Putative ancestral region (PAR), 97
217
218
219
220
221
222
223
224
225
226
227
228
229
230
231
232
233
234
235
Q
Quadrivalent(s), 43–45, 218
236
237
D
TE
EC
CO
RR
O
Octoploids, 28
Ohnolog, 309, 311, 312, 314, 321,
323–325, 329, 354, 356, 358,
362–367, 370
Ohnologon, 354–356, 358
PR
OO
F
171
172
173
174
UN
Editor Proof
Index
R
Rabl-configuration, 35, 36
Random-end pairing model, 37
Rates
of gene duplication, 1, 2
of speciation after polyploidization, 13
Ray flower/ray floret, 262, 263, 264
rDNA, 277, 278
Recent polyploidy, 273, 274, 285, 286
Recombination, 33, 34, 38–41, 43, 44, 46, 47
Recombination nodules (RNS), 39
Recurrent reticulate evolution and polyploidy
in spartina, 226
Repeated polyploidy, 273, 274, 276
Retention rates of duplicate genes, 2, 5, 15
Retroelements/retrotransposons, 210, 211,
212, 214, 215
Rho, 96
Ribosomal DNA (rDNA), 210, 211, 213,
215, 216
Rice, 96
Rice chromosomes 11 and 12, 102
238
239
240
241
242
243
244
245
246
247
248
249
250
251
252
253
254
255
256
257
258
259
S
Saccharomyces cerevisiae, 294
Salmon, 342, 345, 346
Satellite repeat(s), 211
Secondary association(s), 96
260
261
262
263
264
Layout: T1 Standard SC
Chapter No.: BM
Book ID: 272454_1_En
Date: 16-8-2012
Book ISBN: 978-3-642-31441-4
Page: 414/415
Index
F
T
Tandem repeat(s), 211, 212
Teleost, 341, 349–352, 359, 360, 366, 369
Telomere, 35, 36, 39
Telomere bouquet (formation), 35, 36
Telophase, 34
Tetraodon, 351, 359, 369
Tetrasomic (inheritance), 43
Tetrasomy, 272, 283–286
Tilapia, 342
PR
OO
305
306
307
308
309
310
311
312
313
314
Tissue-specific silencing, 281, 282
Tobacco, 210, 214, 215
Tragopogon, 271, 273–288
castellanus, 274, 275
mirus, 271–275, 277–279, 282, 283
miscellus, 271–274, 276–285
x mirabilis, 287
Transcriptome shock, 252, 253, 255, 262
Transcriptomic shock, 272, 282, 286, 287
Transgressive, 255
Translocation(s)/intergenomic
translocation(s), 211, 217, 218, 219
Transposable elements, 123, 189
Transposon, 322
Transposon/transposable
element, 252, 253, 260
Trichomes, 182, 198, 202
Tripartite brain, 329
Triploids, 28
Trisomy, 272, 283–286
Triticum, 110, 111, 112, 115, 119, 121, 125,
284, 285
Trout, 342, 346
CO
RR
EC
TE
Segmental allopolyploids, 25
Segregation, 33, 34, 40, 41, 43–45, 47, 48
Self-incompatibility, 248, 263
Senecio
cambrensis, 245–266, 273, 285, 286
eboracensis, 246, 247, 249, 250, 264
squalidus, 245–251, 253, 254, 257, 258,
262, 263, 266
vulgaris, 248–251, 253–255, 257, 258, 261,
264–266
x baxteri, 245, 250, 252–257, 260–262
Sequenom, 280, 282, 284
Sex chromosome, 385–389, 394, 399, 401,
404, 405
degeneration, 386, 387, 404
turnover, 389, 405
Sigma, 97
Silencing, 192, 196–199
Sister chromatid, 34, 35, 39, 41, 43, 46
Spartina anglica, 273, 285, 286
Speciation, 245–247
Speciation after polyploidization, 9, 11–13
Squalius alburnoides, 345, 348
Stickleback, 342, 345, 351, 355, 356, 358–360,
370, 371
Subfunctionalization, 282, 285, 363–366,
369, 370
Subfunctionalization of duplicate genes, 3, 4,
11, 13, 15
Synapsis, 34, 35, 37–39
Synaptonemal complex (SC), 34, 39
Synonymous substitution, 182, 192
Synteny
blocks, 97
conservation, 314
Synthetic nicotiana tabacum, 211, 214–216,
219, 220
Synthetic polyploids, 271, 273–275, 278,
282, 285, 287
D
S (cont.)
265
266
267
268
269
270
271
272
273
274
275
276
277
278
279
280
281
282
283
284
285
286
287
288
289
290
291
292
293
294
295
296
297
298
299
300
301
302
303
304
UN
Editor Proof
414
315
316
317
318
319
320
321
322
323
324
325
326
327
328
329
330
331
332
333
334
335
336
337
U
Unequal crossing over, 194
Unreduced gametes, 33, 42, 46–48
Urochordate evolution, 329
338
339
340
341
V
Vertebrate, 341, 342, 344, 348–350, 362, 364,
367, 370, 372
Vertebrate Origin and Innovations
Neural Crest, 309, 313, 327, 328
342
343
344
345
346
W
Wheat, 109–123, 125–129, 273, 285
Whole-genome duplication, 94, 271, 272, 277
347
348
349
X
Xenopus, 391, 393, 394, 399, 401, 402, 404
350
351
Z
Zebrafish, 353, 355, 364–366
Zygotene, 34–36
352
353
354
AQ2
356
F
Chapter No.:
PR
OO
357
358
363
361
362
360
364
365
Query Refs.
Details Required
370
368
367
369
371
AQ1
Please provide appropriate page number for the index
term ‘ITS’.
376
373
374
375
AQ2
Please provide index terms with appropriate page
numbers for chapters 4 and 9.
Author’s Response
CO
RR
EC
TE
D
377
UN
Editor Proof
Author Queries
355
MARKED PROOF
Please correct and return this set
Please use the proof correction marks shown below for all alterations and corrections. If you
wish to return your proof by fax you should ensure that all amendments are written clearly
in dark ink and are made well within the page margins.
Instruction to printer
Leave unchanged
Insert in text the matter
indicated in the margin
Delete
Textual mark
under matter to remain
New matter followed by
or
through single character, rule or underline
or
through all characters to be deleted
Substitute character or
substitute part of one or
more word(s)
Change to italics
Change to capitals
Change to small capitals
Change to bold type
Change to bold italic
Change to lower case
Change italic to upright type
under matter to be changed
under matter to be changed
under matter to be changed
under matter to be changed
under matter to be changed
Encircle matter to be changed
(As above)
Change bold to non-bold type
(As above)
Insert ‘superior’ character
Marginal mark
through letter or
through characters
through character or
where required
or
new character or
new characters
or
under character
e.g.
Insert ‘inferior’ character
(As above)
Insert full stop
Insert comma
(As above)
Insert single quotation marks
(As above)
Insert double quotation marks
(As above)
over character
e.g.
(As above)
or
or
(As above)
Transpose
Close up
Insert or substitute space
between characters or words
Reduce space between
characters or words
linking
and/or
or
or
Insert hyphen
Start new paragraph
No new paragraph
or
characters
through character or
where required
between characters or
words affected
and/or