Journal of Experimental Botany, Vol. 62, No. 3, pp. 895–905, 2011
doi:10.1093/jxb/erq317 Advance Access publication 18 October, 2010
This paper is available online free of all access charges (see http://jxb.oxfordjournals.org/open_access.html for further details)
RESEARCH PAPER
P. Pérez1,*, G. Rabnecz2, Z. Laufer3, D. Gutiérrez1, Z. Tuba2,3,† and R. Martı́nez-Carrasco1
1
Institute of Natural Resources and Agrobiology of Salamanca, CSIC, Apartado 257, 37071 Salamanca, Spain
Institute of Botany and Ecophysiology, Faculty of Agriculture and Environmental Sciences, Szent István University, Páter K. 1., 2103
}, Hungary
Gödöllo
3
‘Plant Ecology’ Departmental Research Group of the Hungarian Academy of Sciences, Szent István University, Páter K. 1., 2103
}, Hungary
Gödöllo
2
y
Deceased.
* To whom correspondence should be addressed. E-mail: pilar.perez@irnasa.csic.es
Received 9 March 2010; Revised 17 September 2010; Accepted 21 September 2010
Abstract
Recovery of photosynthesis in rehydrating desiccated leaves of the poikilochlorophyllous desiccation-tolerant plant
Xerophyta scabrida was investigated. Detached leaves were remoistened under 12 h light/dark cycles for 96 h.
Water, chlorophyll (Chl), and protein contents, Chl fluorescence, photosynthesis–CO2 concentration response, and
the amount and activity of Rubisco were measured at intervals during the rehydration period. Leaf relative water
contents reached 87% in 12 h and full turgor in 96 h. Chl synthesis was slower before than after 24 h, and Chla:Chlb
ratios changed from 0.13 to 2.6 in 48 h. The maximum quantum efficiency recovered faster during rehydration than
the photosystem II operating efficiency and the efficiency factor, which is known to depend mainly on the use of the
electron transport chain products. From 24 h to 96 h of rehydration, net carbon fixation was Rubisco limited, rather
than electron transport limited. Total Rubisco activity increased during rehydration more than the Rubisco protein
content. Desiccated leaves contained, in a close to functional state, more than half the amount of the Rubisco
protein present in rehydrated leaves. The results suggest that in X. scabrida leaves Rubisco adopts a special,
protective conformation and recovers its activity during rehydration through modifications in redox status.
Key words: Chlorophyll fluorescence, desiccation tolerance, non-photochemical quenching, photosynthesis, poikilochlorophylly,
relative water content, Rubisco, Xerophyta scabrida.
Introduction
Desiccation-tolerant (DT) plants can withstand the loss of
up to 90–95% of the water of their vegetative tissues and
revive when humidity is available, in contrast to the
majority of plants (Proctor and Tuba, 2002). Desiccation
tolerance entails cellular, biochemical, and molecular
changes during dehydration (Vicré et al., 2004), including
the accumulation of carbohydrates (Whittaker et al., 2001;
Toldi et al., 2009), late embryogenesis-abundant (LEA)
proteins (Ingram and Bartels, 1996), and antioxidants
(Kranner et al., 2002; Mowla et al., 2002; Vicré et al.,
2004), as well as altered expression of target genes and
transcription factors (Frank et al., 1998; Ramanjulu and
Bartels, 2002). Recovery from the desiccated state is much
faster in homoiochlorophyllous DT (HDT) plants such as
Abbreviations: Chl, chlorophyll; DT, desiccation tolerant; HDT, homoiochlorophyllous DT; PDT, poikilochlorophyllous DT; RuBP, ribulose-1,5-bisphosphate; RWC,
relative water content.
ª 2010 The Author(s).
This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/bync/2.5), which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.
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Restoration of photosystem II photochemistry and carbon
assimilation and related changes in chlorophyll and protein
contents during the rehydration of desiccated Xerophyta
scabrida leaves
896 | Pérez et al.
hydrated HDT leaves were reported (Peeva and Cornic,
2009), but information concerning the fate of ribulose1,5-bisphosphate carboxylase oxygenase (Rubisco) and the
relative capacities of Rubisco carboxylation and electron
transport in rehydrating PDT plants is scarce. In earlier
studies, desiccated fronds (Harten and Eickmeier, 1986) and
leaves (Daniel and Gaff, 1980) of DT plants conserved from
40% to 62% of the control Rubisco activity. A decrease in
Rubisco content was observed during dehydration of the C4
DT plant Sporobolus stapfianus (Martinelli et al., 2007),
whereas Rubisco (fraction I) protein did not appear to
decrease relative to hydrated X. viscosa leaves (Daniel and
Gaff, 1980). Consistent with this, it was surmised that the
carboxylating enzymes in X. scabrida would only be
inactivated, but not degraded, during desiccation (Tuba
et al., 1998b). In contrast, Rubisco activity was undetectable
below 51% RWC (12 h rehydration) in R. serbica leaves
(Degl’Innocenti et al., 2008). Drying-induced disruption of
the electron transport chain causes oxidative stress (Vicré
et al., 2004), which can induce aggregation and polymerization, membrane association, and the degradation of
Rubisco (Marı́n-Navarro and Moreno, 2006). On the other
hand, in stressed Lemna minor fronds Rubisco was not
degraded but gradually became polymerized to inactive
aggregates, accompanied by a reduction in the number of
sulphydryl groups (Ferreira and Shaw, 1989). The Benson–
Calvin cycle enzymes have a tendency to form soluble and
membrane-bound multienzyme complexes (Sainis and Harris,
1986; Gontero et al., 1988, 1993; Sainis et al., 1989; Persson
and Johansson, 1989; Hermoso et al., 1992; Anderson et al.,
1995; Agarwal et al., 2009) with higher catalytic efficiency
and less susceptibility to auto-oxidation and proteolysis than
free enzymes (Gontero et al., 1988, 1993).
The aim of this study was to determine to what extent the
recovery from desiccation of X. scabrida photosynthesis is
dependent on photochemical and carboxylation capacities.
The hypothesis was that restoration of Rubisco activity
limits the attainment of photosynthetic competence of
rehydrated PDT plants. To test this hypothesis, Chl
fluorescence and photosynthesis–CO2 response curves were
determined while turgor was being regained. To assess the
carboxylation capacity, the free or aggregated state, as well
as the amount and activity of Rubisco were determined in
desiccated and rehydrating leaves.
Materials and methods
A description of X. scabrida morphology has been provided in an
earlier article (Tuba et al., 1993b). Briefly, it is a 40–90 cm high,
branched pseudoshrub with perennial leaves. Dry leaves are
usually 24–30 cm long, 5–6 mm wide, and folded over along the
midrib. In July 2004, desiccated X. scabrida (Pax) Th. Dur. et
Schinz branches were collected in Tanzania (Mindu Hill, WSW of
Morogoro town, 6°50.78’S, 37°36.76’E) and were kept in paper
bags at room temperature until rehydration and analysis. Central
sections of desiccated leaves having a purple-black or blackishgreen coloration were selected for this study. As representative of
time zero inmediately prior to watering, triplicate desiccated leaves
were briefly immersed in water in a vacuum desiccator to saturate
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Haberlea rhodopensis (Georgieva et al., 2005, 2007) than in
poikilochlorophyllous DT (PDT) plants such as Xerophyta
scabrida (Tuba et al., 1993a, 1994). The former retain their
chlorophyll (Chl), preserve their photosynthetic apparatus,
and undergo morphological changes during drying that
protect their tissues against oxidative stress (Vicré et al.,
2004). In contrast, the latter lose all of their Chl and
dismantle their photosynthetic apparatus during drying,
and they resynthesize these molecules after rehydration
(Tuba et al., 1994, 1998a; Sherwin and Farrant, 1996).
Xerophyta scabrida preserves most of its Chl when dried in
the dark, so most of the loss seems to result from
photooxidative degradation (Tuba et al., 1997). The PDT
strategy evolved in plants that are anatomically complex
and that include the largest in size of all DT species, and it
can be seen as the younger strategy in evolutionary terms
(Proctor and Tuba, 2002).
DT plants regain their water content within a time span
ranging from minutes in bryophytes and pteridophytes
(Csintalan et al., 1999) to days in angiosperms (Tuba et al.,
1994; Proctor and Tuba, 2002; Georgieva et al., 2005;
Degl’Innocenti et al., 2008). As could be expected for HDT
plants, the Chl contents, the Chla:Chlb ratio, and the
relative amounts of the Chl–protein complexes remain
mostly unchanged in control, desiccated, and rehydrated
leaves (Georgieva et al., 2005, 2007). In remoistened PDT
plants, Chl resynthesis begins after 6–12 h of rehydration
(Tuba et al., 1993a, 1994) at 36% relative water contecnt
(RWC) (Degl’Innocenti et al., 2008), and is completed by
48–72 h at 84% RWC. Synthesis of the photosystem II
(PSII) reaction centre and antenna proteins correlates with
the recovery and increase in photosynthetic capacity (Ingle
et al., 2008).
Non-radiative energy dissipation can play an important
protective role during both desiccation and rehydration. In
several mosses (Csintalan et al., 1999) and in Ramonda
serbica (Augusti et al., 2001; Degl’Innocenti et al., 2008),
non-photochemical quenching (NPQ) shows a transient
increase upon remoistening. Maximum quantum efficiency,
Fv/Fm, is completely recovered at 48 h (Degl’Innocenti
et al., 2008) or 72 h (Tuba et al., 1994) after rewatering.
Faster increases during rehydration were recorded in Fv/Fm
than in PSII operating efficiency, Fq#/Fm# (also termed
UPSII; Csintalan et al., 1999). Nonetheless, the involvement
in the non-photochemical energy dissipation of basal, nonradiative decays and of the regulated non-photochemical
energy loss (Baker et al., 2007; Klughammer and Schreiber,
2008) during the rehydration of PDT plants has not been
investigated.
Full recovery of photochemical activity in PDT plants
requires the assimilation of CO2 as an acceptor of the
products of photosynthetic electron transport. In the moss
Polytrichum formosum, carbon fixation is completely restored 3 h after rewetting (Proctor et al., 2007) but is
resumed at 12 h (51% RWC), and is not fully re-established
at 48 h (84% RWC) after rehydration in higher plant DT
species (Tuba et al., 1994; Degl’Innocenti et al., 2008).
Recently, photosynthesis–CO2 concentration responses in
Restoration of photosynthesis in rehydrating Xerophyta scabrida leaves | 897
Water contents
Triplicate samples of desiccated leaves were weighed before and
after drying in an oven for 48 h at 60 °C; the second of these
recordings was taken as the dry weight. This was preferred to the
oven-dry weight after full rehydration, which may be affected by
losses during rewatering due to respiration or release of soluble
compounds. Additional leaves (in triplicate) that were submerged
in water were blotted and their fresh weight was determined at 12,
24, 48, 72, 96, and 120 h after the start of rehydration. No further
weight gain was recorded after 96 h and this was considered as the
turgid weight. The RWCs at successive times in the rehydration
period were determined as (fresh weight–dry weight)3100/(turgid
weight–dry weight). Chl and protein contents, and Rubisco
activity were determined in other leaves sequentially sampled
during rehydration (see below) and were expressed on a turgid
weight basis. The latter was estimated from the fresh weight of
these leaf samples and the water contents measured in the samples
used for RWC measurements.
Chl fluorescence and gas exchange measurements
For Chl fluorescence and CO2 assimilation measurements, triplicate leaf samples kept in water were collected between 3 h and 8 h
after the start of the photoperiod at the times indicated above and
placed in the fluorometer leaf clip or the infrared gas analyser
(IRGA) leaf chamber (see below) with both ends of the leaves
wrapped in moistened filter paper. After measurements, the leaf
samples were returned to the water container in the growth room
for a 30 min adaptation period prior to harvesting for leaf analyses
(see below). Chl fluorescence was measured with a modulated
fluorometer (PAM-2000, Walz, Effeltrich, Germany). Leaf sections
were kept in the dark for 20 min with leaf clips (Gutiérrez et al.,
2009), after which dark-adapted state fluorescence parameters were
measured. Fo was recorded and a saturating flash of light (;8000
lmol m2 s1) was applied for 0.8 s to determine Fm. Fo and Fm,
respectively, represent the minimal and maximal fluorescence in
the dark-adapted state, and Fv/Fm [(Fm–Fo)/Fm] represents the
maximum quantum efficiency. Light-adapted leaves were illuminated with the red actinic light source of the fluorometer to obtain
an irradiance of 1500 lmol m2 s1. Saturating light pulses were
given every 20 s until steady-state Chl fluorescence parameter
values were obtained, the fluorescence values being recorded
immediately before (F#, steady-state fluorescence) and after (Fm#,
maximal fluorescence in the light) each pulse. Then, the leaf was
covered with a black cloth, the actinic light was switched off, and
an infrared light was switched on for 3 s to quickly reoxidize the
PSII centres and measure Fo#, the minimal fluorescence with an
NPQ similar to that found in the steady-state under light. The
equipment determines Fq#/Fm# [(Fm#–F#)/Fm#], which is the PSII
operating efficiency (also termed UPSII) (Baker et al., 2007). The
PSII efficiency factor Fq#/Fv# (also termed qP) [(Fm#–F#)/ (Fm#–Fo#)]
and Fv#/Fm# [(Fm#–Fo#)/Fm#], the PSII maximum efficiency under
light, were calculated. The fraction of PSII centres in the open
state, qL, equates to (Fq#/Fv#) (Fo#/F#). The quantum yield of basal,
non-radiative decays, UNO, is 1/[NPQ+1+qL(Fm/Fo–1)], where
NPQ is (Fm/Fm#)–1, and the quantum yield of non-photochemical
quenching, UNPQ, is 1–(Fq#/Fm#)–UNO (Kramer et al., 2004).
Light-saturated photosynthesis–CO2 response curves of leaves
were recorded at the same times and with the same sampling
scheme as Chl fluorescence. Measurements were carried out with
an air flow rate of 300 ml min1, 1500 lmol m2 s1 irradiance,
and a 1.660.23 kPa vapour pressure deficit, using a 1.7 cm2
window leaf chamber connected to a portable IRGA (CIRAS-2,
PP Systems, Hitchin, Herts, UK) with differential operation in an
open system. Temperature was kept at 25 °C with the Peltier
system of the IRGA. The air CO2 concentration was decreased in
four steps from 34 Pa to 6 Pa and then increased from 34 Pa to
180 Pa in six steps. Chloroplast CO2 concentration, Cc, the
maximum carboxylation rate allowed by Rubisco, Vcmax, and the
rate of photosynthetic electron transport, J, were determined from
the photosynthesis responses to CO2 with the Rubisco kinetic
parameters and the Excel utility of Sharkey et al. (2007).
Rubisco activity assay
Triplicate leaf samples that had been equilibrated in aerated water
in the growth chamber after Chl fluorescence and gas exchange
measurements were blotted dry, rapidly transferred in situ to liquid
nitrogen, and stored at –80 °C until analysed. Rubisco activity was
assayed on the basis of the procedure described by Lilley and
Walker (1974), modified by Ward and Keys (1989) and Sharkey
et al. (1991). Aliquots (80 mg) of frozen leaves were ground in
a mortar with liquid nitrogen, extracted with 4 ml of 100 mM
N,N-bis(2-hydroxyethyl)glycine (Bicine)-NaOH (pH 7.8), 10 mM
MgCl2, 0.5 mM dithiothreitol (DTT), 1 mM ethylenediaminetetraacetic acid (EDTA), 1 mM ethylene glycol tetraacetic acid
(EGTA), 1% (v/v) Triton X-100, 0.25% (w/v) bovine serum
albumin (BSA), 20% (v/v) glycerol, 1 mM benzamidine, 1 mM
e-aminocaproic acid, 10 lM leupeptin, and 1 mM phenylmethylsulphonyl fluoride (PMSF), and then centrifuged at 13 000 g. The
total time from extraction to the assay of initial Rubisco activity
was <2.5 min. Activity was assayed by adding extract (40 ll) to
a mixture of 100 mM Bicine (pH 8.2), 20 mM MgCl2, 10 mM
NaHCO3, 18 mM KCl, 0.6 mM ribulose-1,5-bisphosphate
(RuBP), 0.2 mM NADH, 1 mM ATP, 5 mM creatine phosphate,
25 U ml1 phosphocreatine kinase, 47U ml1 phosphoglycerate
kinase, 47 U ml1 glyceraldehyde 3-phosphate dehydrogenase,
10 mM DTT, 1 mM EDTA, 0.02% (w/v) BSA (800 ll total
volume) and recording the decrease in absorbance at 340 nm
minus 400 nm for 40–60 s, at a stoichiometry of 2:1 between
NADH oxidation and RuBP carboxylation. The spectrophotometer cell compartment was thermostated with a circulating water
bath. To assay total Rubisco activity, an aliquot of the extract was
incubated with NaHCO3 and MgCl2 for 10 min at room temperature before the addition of coupling enzymes and NADH; the
reaction was started by adding RuBP. The activation state was
estimated as initial activity, as a percentage of total activity.
Activation and assays were performed either at room temperature
or at 35 °C. Commercial coupling enzymes suspended in ammonium sulphate were precipitated by centrifugation and dissolved in
20% glycerol (Sharkey et al., 1991). With the assay buffer
described, the initial lag in the reaction reported by others (Ward
and Keys, 1989; Sharkey et al., 1991) was not observed.
Chlorophyll and protein analysis
Total Chl, Chla, and Chlb in 80% acetone extracts of frozen
triplicate subsamples were determined according to Arnon (1949),
who used the extinction coefficents for Chla (16.75 l g1 cm1 and
82.04 l g1 cm1 at 645 nm and 663 nm, respectively) and Chlb
(45.6 l g1 cm1 and 9.27 l g1 cm1 at 645 nm and 663 nm,
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the intercellular air spaces with water. Subsequently the leaf
material was blotted, weighed, frozen in liquid nitrogen, and
stored at –80 °C for Chl, protein, and Rubisco activity measurements (see below). Previous experience (Tuba et al., 1993b) has
shown that placing whole plants with their roots in water does not
result in a recovery response in X. scabrida, because the roots are
dry and unable to transport water to the leaves; only a direct
rewatering of the leaves by immersion in water led to regreening.
Moreover, in the natural habitat, new root development and water
uptake were preceded by the rehydration and regreening of the
leaves. Consequently, in the present experiments, additional leaf
samples were rehydrated by submerging them in a 10.0 l glass tank
filled with tap water and aerated with a pump (Tuba et al., 1993a,
1994). The container was placed in a growth chamber with
a 21 °C/15 °C day/night temperature, under a 340 lmol m2 s1
photon flux density in a 12 h photoperiod (modified after Tuba
et al., 1994). The water was changed daily.
898 | Pérez et al.
20% methanol, and 0.1% SDS (pH 8.3) using an electrotransfer cell
(Mini Trans-Blot, Bio-Rad, Madrid, Spain) at 400 mA. The blots
were blocked immediately following transfer in 2% ECL Advance
blocking reagent (GE Healthcare, Barcelona, Spain) in 20 mM
TRIS, 137 mM NaCl (pH 7.6) with 0.1% (v/v) Tween-20 (TBS-T)
for 1 h at room temperature with shaking. Blots were briefly rinsed
twice in TBS-T, then probed with 1:50 000 diluted polyclonal
antibody specific for the large Rubisco subunit (Rubisco quantitation kit, Agrisera, Vännäs, Sweden) for 1 h at room temperature
with shaking. The antibody solution was decanted and the blot
was briefly rinsed twice, and then washed once for 15 min and
three times for 5 min in TBS-T at room temperature with shaking.
Next, the blots were incubated in secondary antibody (anti-chicken
Ig Y peroxidase conjugate, Sigma, Spain) diluted at 1:160 000 in
2% ECL Advance blocking solution for 1 h at room temperature
with shaking. The blots were then washed as above and developed
for 5 min with ECL Advance detection reagent according to the
manufacturer’s instructions. Images of the blots were obtained
using a CCD imager (Fluor-S Multilmager, Bio-Rad).
Statistical analyses
One-way analyses of variance with sampling time as a factor were
carried out with the GenStat 6.2 (VSN International Ltd, Hemel
Hempstead, UK) statistical software. From these analyses, the
standard errors of the differences (SEDs), and the least significant
differences of means (three replicates) at P <0.05 probability were
derived. The latter were used for the inspection of differences
among values for each sampling time. The homogeneity of
variance and the significance of the analysis were not modified
appreciably by using arcsine-square root transformation of percentage variables (Rubisco activation and percentage in soluble
protein). Therefore, the untransformed data were used.
Results
Water and Chl contents
Leaf RWC (Table 1) increased sharply from 3.4% to 87.0%
during the first 12 h of rehydration, and additional water
Table 1. Relative water contents (RWC), Chl contents, and Chl fluorescence parameters in leaves of Xerophyta scabrida during
rehydration
Chl fluorescence parameters in the light were measured at 1500 lmol m2 s1 light intensity.
Parameter
RWC (%)
Chla (mg g1 t. wt)
Chlb (mg g1 t. wt)
Chla:Chlb
Chl a+b (mg g1 t. wt)
Fo
Fm
Fv/Fm
Fq’/Fm’
Fv’/Fm’
Fq’/Fv’
qL
UNPQ
UNO
Time of rehydration (h)
0
12
24
48
72
96
3.4 a
0.010 a
0.039 a
0.13 a
0.049 a
87 b
0.014 a
0.049 a
0.18 a
0.063 a
0.015 a
0.020 a
0.21 a
92 c
0.047 a
0.043 a
1.1 b
0.090 a
0.13 c
0.25 b
0.44 b
0a
0.28 a
0a
0a
0.60 a
0.40 b
96 d
0.19 b
0.070 a,b
2.7 c
0.26 b
0.090 b
0.36 c
0.75 c
0.047 b
0.50 b
0.10 b
0.060 a
0.67 a
0.28 a
97 e
0.29 b
0.11b
2.6 c
0.41 b
0.095 b
0.42 c
0.77 c
0.086 c
0.53 b
0.16 b
0.081 a
0.69 a
0.23 a
100 f
0.43 c
0.17 c
2.5 c
0.60 c
0.091 b
0.45 c
0.80 c
0.099 c
0.56 b
0.18 b
0.088 a
0.67 a
0.24 a
P
SED
<0.001
<0.001
<0.001
<0.001
<0.001
<0.001
<0.001
<0.001
0.004
0.041
0.01
0.064
0.26
0.017
0.04
0.054
0.023
0.24
0.074
0.012
0.037
0.085
0.017
0.072
0.034
0.025
0.034
0.037
P, probability in the analysis of variance; SED, standard error of the difference among means (n¼3); within each row, values with the same letter
are not significanty different; t. wt, turgid weight.
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respectively) given by MacKinney (1941). The soluble proteins were
extracted by grinding frozen leaf subsamples to a fine powder in
50 mM N-[tri(hydroxymethyl)methyl] glycine (Tricine) buffer (pH
8.0), 2 mM EDTA, 10 mM NaCl, 5 mM MgCl2, 75 mM sucrose,
5 mM e-aminocaproic acid, 2 mM benzamidine, 8 mM b-mercaptoethanol (+bme), and 2 mM PMSF for 5 min on ice. This was
followed by centrifugation at 12 500 g at 4 °C for 30 min. Protein
concentrations were measured in the decanted supernatant
(Bradford, 1976), and 5 vols of cold acetone were added to an
aliquot containing 200 mg of protein, which was left overnight in
the freezer. The samples were then centrifuged at 12 000 g at 4 °C
for 15 min. The acetone was allowed to evaporate off. The
precipitates were dissolved in 65 mM TRIS-HCl (pH 6.8), 3 M
sucrose, 0.6 M bme, 5% sodium dodecylsulphate (SDS, w/v), and
0.01% bromophenol blue at 96 °C for 7 min. The samples were then
cooled to room temperature and aliquots of the SDS-dissociated
extracts, containing 15 lg of protein, were loaded onto a 12.5%
SDS–polyacrylamide gel (Mini-Protean 3 Cell, Bio Rad). This
protein amount was within the range of linear response of optical
density to the concentration of BSA standard (66 kDa), according
to previous calibration measurements. The solubilized proteins were
separated by SDS–PAGE (Laemmli, 1970) using a 0.75 mm thick
gel (12.5% resolving, 4% stacking). Electrophoresis was carried out
at room temperature at a constant 200 V. The gels were fixed in
500:150:75 (v/v/v) water–methanol–acetic acid mixture for 75 min,
stained in EZ Blue Gel Staining (Sigma) solution for 2 h, and
subsequently rinsed in water to remove excess stain. Finally, the gels
were scanned with a high-resolution scanner (Scanjet G4050,
Hewlett Packard, Spain) and the amount of Rubisco subunits was
determined by densitometry with image analysis software (Image
Quant, Molecular Dynamics, GE Healthcare, Spain). Alternatively,
when electrophoresis with non-reducing (–bme) gels was performed,
the frozen leaf samples were extracted in buffer without bme
containing 10 mM iodoacetamide to prevent the formation of
disulphide bonds, before the addition of SDS loading buffer without
bme and boiling. To a separate aliquot, bme was added to a final
concentration of 0.6 M, as a control of the same samples under
reducing (+bme) conditions (Marı́n-Navarro and Moreno, 2006).
Following electrophoresis, additional gels were blotted for
75 min to PVDF membranes (Bio-Rad, Madrid, Spain) pre-wetted
in methanol and equilibrated in 25 mM TRIS, 192 mM glycine,
Restoration of photosynthesis in rehydrating Xerophyta scabrida leaves | 899
was gained until full turgor was reached at 96 h after the
start of rehydration. Concomitant changes in specific leaf
area, with a maximum of ;0.17 cm2 mg1 dry weight by
12 h and little further change, have been reported previously (Tuba et al., 1993b). Chl contents (Table 1) were low
in desiccated leaves, Chlb being relatively more abundant
than Chla. Chl contents rose during the rehydration period,
with a faster increase after 24 h. Chla accumulated to
a greater extent than Chlb, such that the Chla:Chlb ratios,
which were initially <1, reached a value >2.5—within the
range found in other plants—by 48 h.
The maximum quantum efficiency of PSII photochemistry
(Fv/Fm, Table 1) increased from 12 h to 48 h of rehydration and changed little thereafter. This change was
a consequence of a small increase in Fo and a large increase
in Fm. The lack of variable fluorescence in the lightadapted state prevented the determination of the fluorescence parameters until 24 h of rehydration. In illuminated
leaves (1500 lmol m2 s1), the PSII operating efficiency
and the efficiency factor (Fq#/Fm# and Fq#/Fv#, respectively;
Table 1) underwent increases from 24 h to 72 h of rehydration, while Fv#/Fm# rose from 24 h to 48 h with little
further change, as was the case for Fv/Fm. The pattern of
change with time in the fraction of open PSII centres (qL)
was similar to that in Fq#/Fv#, but with lower absolute
values. The quantum yield of non-photochemical quenching (UNPQ, Table 1) underwent little change during the
rehydration period, while the quantum yield of nonradiative decay (UNO) decreased by 43% from 24 h to
72 h. By comparison, a rise in NPQ was observed in this
interval (data not shown).
CO2 fixation
Regardless of the CO2 concentration used in measurements,
there was no CO2 uptake at 24 h of rehydration (Fig. 1A)
or before. Photosynthesis increased in the following 3 d
rehydration period and was Rubisco limited up to very high
chloroplast CO2 concentrations at 48 h and 72 h (Fig. 1B,
C). The transition from Rubisco-limited to RuBP regeneration-limited photosynthesis decreased to 33 Pa CO2 partial
pressure at 96 h (Fig. 1D), which is still high in comparison
with other plants. Vcmax and J were calculated (Fig. 2) from
the photosynthesis–CO2 response curves. Except for a drop
in Vcmax at 72 h, which can be attributed to a variation
between samples, both J and Vcmax increased from 24 h to
96 h, without reaching a plateau. There were relatively
higher increases in J than in Vcmax.
Rubisco activity and contents
No Rubisco activity was detected in assays carried out at
room temperature, but activity was recorded when the
enzyme activation and assays were performed at 35 °C
(Fig. 3A). Both the initial and total Rubisco activities
increased during the 96 h hydration period, with a faster
Fig. 1. Photosynthetic responses of Xerophyta scabrida leaves to
the CO2 concentration inside the chloroplast during rehydration.
Observed data (filled circles); Rubisco-limited photosynthesis
(solid line); RuBP regeneration-limited photosynthesis (dotted line).
Measurements were performed under 1500 lmol m2 s1
irradiance, at 25 °C and 1.660.23 kPa vapour pressure deficit.
The statistical significance of the parameters derived from this
figure is shown in Fig. 2.
Fig. 2. Maximum carboxylation rate allowed by Rubisco, Vcmax
(filled circles), and the rate of photosynthetic electron transport, J
(filled squares), in Xerophyta scabrida leaves during rehydration.
Vertical bars represent least significant differences between means
(n¼3).
rise—from 12% to 75% final, total Rubisco activity—during
the first 12 h. The activation of the enzyme was 44–64% in
the first 48 h and increased to 74–83 % in the last 2 d.
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Chl fluorescence
900 | Pérez et al.
Fig. 3. Changes during rehydration of Xerophyta scabrida leaves
in (A) initial (filled circles) and total (filled squares) Rubisco activities
and Rubisco activation state (filled triangles) measured at 35 °C;
and (B) Rubisco (filled squares) and total soluble proteins (filled
circles) and amount of Rubisco as a percentage of soluble protein
(filled triangles). Rubisco was quantified by densitometric analysis
of +bme SDS–PAGE. Values are expressed on a turgid weight
basis. Vertical bars represent least significant differences between
means (n¼3).
iodoacetamide with either +bme or –bme showed few
electrophoretic differences, the two Rubisco subunits undergoing normal migration. The examination of the iodoacetamide –bme and +bme gel electrophoresis (Fig. 6)
revealed that in both desiccated and rehydrating X. scabrida
leaves Rubisco formed high molecular weight aggregates. It
was therefore investigated whether the reversible aggregation of Rubisco might account for the differences between
the gas exchange measurements and Rubisco contents.
Leaves harvested at each sampling time were analysed
immediately after illumination with bright light in moistened air, rather than after an equilibration period in water
after the photosynthesis measurements (see Materials and
methods). In these leaves, Rubisco remained in an aggregated state (data not shown).
Discussion
The rehydration of leaves and the recovery of Chl contents
in this experiment were in general agreement with earlier
reports on poikilochlorophyllous resurrection plants (Tuba
et al., 1994; Proctor and Tuba, 2002; Degl’Innocenti et al.,
2008). It may be remarked that Chlb was relatively more
abundant than Chla in desiccated leaves, but by 48 h the
Chla:Chlb ratio was >2.0. This suggests a rise during
rehydration in the PSII reaction centres with respect to the
antenna complexes (Habash et al., 1995; Bailey et al., 2001),
which is consistent with the reported up-regulation of PSII
genes (Ingle et al., 2007, 2008).
The PSII operating efficiency (Fq#/Fm#), in contrast to Fv/
Fm and Fv#/Fm#, was still relatively low at 48 h after the
start of rehydration and continued to increase up to the
following day, as did the PSII efficiency factor. Fq#/Fv# is
generally much more affected by the ability to utilize the
products of the electron transport chain than by changes in
NPQ (Baker et al., 2007). This points to the capacity for
carbon assimilation as the factor that limits the efficiency at
which the light absorbed by PSII is used for photochemistry
in rehydrating X. scabrida leaves. A similar conclusion was
reached for the HDT H. rodopensis during dehydration
(Georgieva et al., 2005). Fq#/Fv# is non-linearly related to
Fig. 4. Effect of rehydration of Xerophyta scabrida leaves on Vcmax
(filled circles), in vitro total Rubisco activity (filled squares), and
Rubisco protein (filled triangles), as a percentage of values at 96 h.
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Rubisco protein amounts (Fig. 3B) were quantified by
SDS–PAGE densitometric analysis of samples extracted
with +bme (see Materials and methods). There was little
change in the amount of Rubisco protein during the first
48 h of rehydration, although this was followed by an
increase of ;56%. Similarly, total soluble protein remained
unchanged for 48 h and then increased by ;18% (Fig. 3B).
As a fraction of soluble protein, Rubisco was relatively low
initially and increased (from 19% to 27%) in the last 2 d of
rehydration. When Rubisco protein contents and in vitro
and in vivo estimated (Vcmax) total Rubisco activities were
compared (Fig. 4), it was found that the continued increase
in the latter was accompanied by relatively smaller changes
in total in vitro Rubisco activity after 12 h of rehydration
and by an increase in Rubisco protein only after 48 h.
To examine further the disparity between Vcmax and
Rubisco contents, the electrophoresis of samples extracted
either with +bme or with iodoacetamide and –bme to block
the sulphydryl groups was compared. Whereas with the
reducing agent the two Rubisco subunits migrated according to their molecular weights (Fig. 5), with blocked
sulphydryl groups the Rubisco remained at the top of the
gel. In separate aliquots of the same samples to which bme
had been added following iodoacetamide treatment, the
large Rubisco subunit showed normal migration in the gels,
but the small Rubisco subunit was not observed. In
comparison, wheat samples extracted with bme or with
Restoration of photosynthesis in rehydrating Xerophyta scabrida leaves | 901
the proportion of PSII centres that are in the open state
(with the primary quinone electron acceptor of PSII, QA,
oxidized), as estimated by qL (Baker et al., 2007). Changes
during rehydration in this parameter and Fq#/Fv# were
parallel and showed an increasing fraction of open PSII
centres as the carboxylation capacity increased. In contrast
to previous reports on the engagement of non-photochemical energy dissipation upon the remoistening of DT leaves
(Csintalan et al., 1999; Augusti et al., 2001; Degl’Innocenti
et al., 2008), the lack of significant changes in UNPQ suggests
that the down-regulation of PSII was not a major cause of
the changes observed during rehydration in the maximum
efficiency of PSII in the light. The significant decline in UNO
probably reflects the reconstitution of functional PSII
antennae and reaction centres.
The capacity for carbon assimilation was also recovered
during rehydration, although the assimilation rates could
differ from those of intact plant leaves due to signalling and
metabolic interactions with other organs. In agreement with
the results concerning Chl fluorescence, the responses of
carbon assimilation to the CO2 concentration in the
chloroplast (Figs 1, 2) indicated that the recovery of
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Fig. 5. Soluble proteins in desiccated Xerophyta scabrida (X) and fresh wheat (W) leaves. (A) SDS–PAGE and (B) immunoblotting of
samples extracted in buffer containing 10 mM iodoacetamide without b-mercaptoethanol (i-bme), containing iodoacetamide to which
bme had been added prior to electrophoresis (i+bme), or with bme without iodoacetamide (+bme). Molecular markers were loaded onto
lanes 1 and 10 and BSA standard onto lanes 5 and 9. Rubisco aggregates (Ag) and large (L) and small (S) subunits are indicated on the
left.
902 | Pérez et al.
photosynthetic capacity during rehydration was relatively
more limited by carboxylation than by the rate of electron
transport. Rubisco activity may therefore be of paramount
importance for the photosynthetic competence of rehydrated desiccation-tolerant plants. The present results show
that a significant fraction of the Rubisco protein found in
rehydrated leaves is present in desiccated leaves (Fig. 3),
and that new synthesis occurs later in the process of
rehydration. Notably, in both desiccated and rehydrated
X. scabrida leaves—in contrast to wheat—Rubisco was in
an aggregated state (Fig. 5), as in L. minor fronds under
osmotic stress (Ferreira and Shaw, 1989). Our gas exchange
measurements and Rubisco activity assays revealed that free
or membrane-bound Rubisco aggregates in X. scabrida were
inactive in desiccated leaves and in the early rehydration
stages. Treatments such as high light intensity in gas
exchange analysis or mild warming under reducing conditions in activity assays rendered Rubisco progressively
more active. This suggests that the integrity of Rubisco was
preserved in the aggregates, but that a modification required
for the enzyme to become functional was facilitated by
rehydration. In dormant Retama raetam tissues, Rubisco
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Fig. 6. Soluble proteins in Xerophyta scabrida leaves after different rehydration periods. (A) SDS–PAGE and (B) immunoblotting of
samples extracted in buffer containing 10 mM iodoacetamide without b-mercaptoethanol (i-bme), or containing iodoacetamide to which
bme had been added prior to electrophoresis (i+bme). Leaves desiccated (0) or rehydrated for 24 h (24), 48 h (48), or 96 h (96).
Molecular markers were loaded onto lanes 5 and 10, and BSA standard onto lane 1. Rubisco aggregates (Ag) and large (L) and small (S)
subunits are indicated on the left.
Restoration of photosynthesis in rehydrating Xerophyta scabrida leaves | 903
Acknowledgements
This work has been funded by the Spanish Ministry of
Science and Innovation and the Hungarian Science and
Technology Office (Integrated Action HH2006-0019, ESP41/2006), and by the Spanish National Research and
Development Programme-European Regional Development
Fund ERDF (Project AGL2006-13541-C02-02/AGR). DG
was the recipient of a Junta de Castilla y León fellowship.
This paper is dedicated to the memory of Zoltán Tuba, who
passed away while this research was in progress.
References
Agarwal R, Ortlebb S, Sainis JK, Melzer M. 2009. Immunoelectron
microscopy for locating Calvin cycle enzymes in the thylakoids of
Synechocystis 6803. Molecular Plant 2, 32–42.
Anderson LE, Goldhaber-Gordo IM, Li D, Tang XY, Xiang M,
Prakash N. 1995. Enzyme–enzyme interaction in the chloroplast,
glyceradehyde-3-phosphate dehydrogenase, triose phosphate
isomerase and aldolase. Planta 196, 245–255.
Ramonda serbica during dehydration and rehydration. Photosynthesis
Research 67, 79–88.
Bailey S, Walters RG, Jansson S, Horton P. 2001. Acclimation of
Arabidopsis thaliana to the light environment: the existence of separate
low light and high light responses. Planta 213, 794–801.
Baker NR, Harbinson J, Kramer DM. 2007. Determining the
limitations and regulation of photosynthetic energy transduction in
leaves. Plant, Cell and Environment 30, 1107–1125.
Bradford M. 1976. A rapid and sensitive method for the quantitation
of microgram quantities of protein utilizing the principle of protein–dye
binding. Analytical Biochemistry 72, 248–254.
Cho JH, Hwang H, Cho MH, Kwon YK, Jeon JS, Bhoo SH,
Hahn TR. 2008. The effect of DTT in protein preparations for
proteomic analysis: removal of a highly abundant plant enzyme,
ribulose bisphosphate carboxylase/oxygenase. Journal of Plant
Biology 51, 297–301.
Csintalan Zs, Proctor MCF, Tuba Z. 1999. Chlorophyll
fluorescence during drying and rehydration in the mosses
Rhytidiadelphus loreus (Hedw.) Warnst., Anomodon viticulosus
(Hedw.) Hook. & Tayl. and Grimmia pulvinata (Hedw.) Sm. Annals of
Botany 84, 235–244.
Daniel V, Gaff DF. 1980. Sulfhydryl and disulphide levels in protein
fractions from hydrated and dry leaves of resurrection plants. Annals of
Botany 45, 163–171.
Degl’Innocenti E, Guidi L, Stevanovic B, Navari F. 2008. CO2
fixation and chlorophyll a fluorescence in leaves of Ramonda serbica
during a dehydration–rehydration cycle. Journal of Plant Physiology
165, 723–733.
Ferreira RB, Shaw NM. 1989. Effect of osmotic stress on protein
turnover in Lemna minor fronds. Planta 179, 456–465.
Frank W, Phillips J, Salamini F, Bartels D. 1998. Two dehydrationinducible transcripts from the resurrection plant Craterostigma
plantagineum encode interacting homeodomain leucine zipper
proteins. The Plant Journal 15, 413–421.
Georgieva K, Maslenkova L, Peeva V, Markovska Y, Stefanov D,
Tuba Z. 2005. Comparative study on the changes in photosynthetic
activity of the homoiochlorophyllous desiccation-tolerant Haberlea
rhodopensis and desiccation-sensitive spinach leaves during
desiccation and rehydration. Photosynthesis Research 85, 191–203.
Georgieva K, Szigeti Z, Sarvari E, Gaspar L, Maslenkova L,
Peeva V, Peli E, Tuba Z. 2007. Photosynthetic activity of
homoiochlorophyllous desiccation tolerant plant Haberlea rhodopensis
during dehydration and rehydration. Planta 225, 955–964.
Gontero B, Cardenas M, Ricard J. 1988. A functional five enzyme
complex of chloroplasts involved in the Calvin cycle. European Journal
of Biochemistry 173, 437–443.
Arnon DI. 1949. Copper enzymes in isolated chloroplasts. Polyphenol
oxidase in Beta vulgaris. Plant Physiology 24, 1–15.
Gontero B, Mulliert G, Rault M, Giudico-Orticoni MT, Ricard J.
1993. Structural and functional properties of a multi-enzyme complex
from spinach chloroplasts: modulation of the kinetic properties of
enzymes in the aggregated state. European Journal of Biochemistry
217, 1075–1082.
Augusti A, Scartazza A, Navari-Izzo F, Sgherri CLM,
Stevanovic B, Brugnoli E. 2001. Photosystem II photochemical
efficiency, zeaxanthin and antioxidant contents in the poikilohydric
Gutiérrez D, Gutiérrez E, Pérez P, Morcuende R, Verdejo AL,
Martinez-Carrasco R. 2009. Acclimation to future atmospheric CO2
increases photochemical efficiency and mitigates photochemistry
Downloaded from jxb.oxfordjournals.org at Centro de Información y Documentación Científica on January 18, 2011
and other proteins also appear to be present as high
molecular weight complexes (Pnueli et al., 2002). These
complexes precipitated during extraction with reducing
buffers, a result that was observed for the small Rubisco
subunit only when bme was added to extracts containing
iodoacetamide. Pnueli et al. (2002) suggested that the
dilution of reducing equivalents upon rehydration releases
proteins from the aggregates into their soluble, active form.
However, some of the DTT concentrations used by Pnueli
et al. (2002) in the protein extraction buffer have been
shown to cause Rubisco aggregation and precipitation (Cho
et al., 2008). Moreover, the present results suggested that
the increase in Rubisco activity during rehydration was not
associated with protein release from the aggregates. The
lower oxidation states of thiol groups (disulphide and
sulphenic acid) may easily be reverted again to the
sulphydryl state by disulphide exchange with free thiols, by
DTT (in vitro) or by thioredoxins and glutaredoxins
(Marcus et al., 2003; Moreno et al., 2008). It is possible
that oxidative conditions during desiccation could induce
the formation of disulphides in the Rubisco molecule, and
that the recovery of photochemical activity could lead to an
increasingly reduced stroma, favouring the reductive activation of Rubisco. While upon desiccation of X. scabrida, and
indeed of all poikilochlorophyllous plant species, Chl and
the photosynthetic apparatus are lost, it is concluded that
Rubisco is preserved in large amounts in a close to
functional state. Rubisco aggregation may be a part of the
poikilochlorophylly strategy.
904 | Pérez et al.
inhibition by warm temperatures in wheat under field chambers.
Physiologia Plantarum 137, 86–100.
plant Sporobolus stapfianus during dehydration stress. Journal of
Experimental Botany 58, 3929–3939.
Habash DZ, Matthew JP, Parry MAJ, Keys AJ, Lawlor DW. 1995.
Increased capacity for photosynthesis in wheat grown at elevated
CO2: the relationship between electron transport and carbon
metabolism. Planta 197, 482–489.
Moreno J, Garcı́a-Murria MJ, Marı́n-Navarro J. 2008. Redox
modulation of Rubisco conformation and activity through its cysteine
residues. Journal of Experimental Botany 59, 1605–1614.
Harten JB, Eickmeier WG. 1986. Enzyme dynamics of the
resurrection plant Selaginella lepidophylla (Hook. & Grev.) Spring
during rehydration. Plant Physiology 82, 61–64.
Ingle RA, Collett H, Cooper K, Takahashi Y, Farrant JM, Illing N.
2008. Chloroplast biogenesis during rehydration of the resurrection
plant Xerophyta humilis: parallels to the etioplast–chloroplast transition.
Plant, Cell and Environment 31, 1813–1824.
Ingle RA, Schmidt UG, Farrant JM, Thomson JA, Mundree SG.
2007. Proteomic analysis of leaf proteins during dehydration of the
resurrection plant Xerophyta viscosa. Plant, Cell and Environment
30, 435–446.
Ingram J, Bartels D. 1996. The molecular basis of dehydration
tolerance in plants. Annual Review of Plant Physiology and Plant
Molecular Biology 47, 377–403.
Peeva V, Cornic G. 2009. Leaf photosynthesis of Haberlea
rhodopensis before and during drought. Environmental and
Experimental Botany 65, 310–318.
Persson O, Johansson G. 1989. Studies of protein–protein
interaction using countercurrent distribution in aqueous two phase
systems. Biochemical Journal 259, 863–870.
Pnueli L, Hallak-Herr E, Rozenberg M, Cohen M, Goloubinoff P,
Kaplan A, Mittler R. 2002. Molecular and biochemical mechanisms
associated with dormancy and drought tolerance in the desert legume
Retama raetam. The Plant Journal 31, 319–330.
Proctor MCF, Ligrone R, Duckett JG. 2007. Desiccation tolerance
in the moss Polytrichum formosum: physiological and fine-structural
changes during desiccation and recovery. Annals of Botany
99, 75–93.
Proctor MCF, Tuba Z. 2002. Poikilohydry and homoihydry: antithesis
or spectrum of possibilities? New Phytologist 156, 327–349.
Klughammer C, Schreiber U. 2008. Complementary PS II quantum
yields calculated from simple fluorescence parameters measured by
PAM fluorometry and the saturation pulse method. PAM Application
Notes 1, 27–35.
Ramanjulu S, Bartels D. 2002. Drought- and desiccation-induced
modulation of gene expression in plants. Plant, Cell and Environment
25, 141–151.
Kramer DM, Johnson G, Kiirats O, Edwards GE. 2004.
New fluorescence parameters for determination of QA redox state
and excitation energy fluxes. Photosynthesis Research
79, 209–218.
Sainis JK, Harris GC. 1986. The association of d-ribulose-1,
5-bisphosphate carboxylase with phosphoriboisomerase and
phosphoribulokinase. Biochemical and Biophysical Research
Communications 139, 947–954.
Kranner I, Beckett RP, Wornik S, Zorn M, Pfeifhofer HW. 2002.
Revival of a resurrection plant correlates with its antioxidant status.
The Plant Journal 31, 13–24.
Sainis JK, Merriam K, Harris GC. 1989. The association of
ribulose-1,5-bisphosphate carboxylase/oxygenase with
phosphoribulokinase. Plant Physiology 89, 368–374.
Laemmli UK. 1970. Cleavage of structural proteins during the
assembly of the head of bacteriophage T4. Nature 227, 680–685.
Sharkey TD, Bernacchi CJ, Farquhar GD, Singsaas EL. 2007.
Fitting photosynthetic carbon dioxide response curves for C3 leaves.
Plant, Cell and Environment 30, 1035–1040.
Lilley R McC, Walker DA. 1974. An improved spectrophotometric
assay for ribulose bisphosphate carboxylase. Biochimica et Biophysica
Acta 358, 226–229.
MacKinney G. 1941. Absorption of light by chlorophyll solutions.
Journal of Biological Chemistry 140, 315–322.
Marcus Y, Altman-Gueta H, Finkler A, Gurevitz M. 2003. Dual role
of cysteine 172 in redox regulation of ribulose 1,5-bisphosphate
carboxylase/oxygenase activity and degradation. Journal of
Bacteriology 185, 1509–1517.
Marı́n-Navarro J, Moreno J. 2006. Cysteines 449 and 459 modulate
the reduction–oxidation conformational changes of ribulose
1,5-bisphosphate carboxylase/oxygenase and the translocation of the
enzyme to membranes during stress. Plant, Cell and Environment
29, 898–908.
Martinelli T, Whittaker A, Masclaux-Daubresse C, Farrant JM,
Brilli F, Loreto F, Vazzana C. 2007. Evidence for the presence of
photorespiration in desiccation-sensitive leaves of the C4‘resurrection’
Sharkey TD, Savitch LV, Butz ND. 1991. Photometric method for
routine determination of kcat and carbamylation of rubisco.
Photosynthesis Research 28, 41–48.
Sherwin HW, Farrant JM. 1996. Differences in rehydration of three
desiccation-tolerant angiosperm species. Annals of Botany
78, 703–710.
Toldi O, Tuba Z, Scott P. 2009. Vegetative desiccation tolerance:
is it a goldmine for bioengineering crops? Plant Science
176, 187–199.
Tuba Z, Zs Csintalan, Szente K, Nagy Z, Grace J. 1998B. Carbon
gains by desiccation tolerant plants at elevated CO2. Functional
Ecology 12, 39–44.
Tuba Z, Hartmut K, Maroti I, Zs Csintalan, Pócs T. 1993b.
Regreening of dessicated leaves of the poikilochlorophyllous
Xerophyta scabrida upon rehydration. Journal of Plant Physiology
142, 103–108.
Downloaded from jxb.oxfordjournals.org at Centro de Información y Documentación Científica on January 18, 2011
Hermoso R, Fonollá J, de Felipe MR, Vivo MA, Chueca A,
Lázaro J, Lopez Gorgé J. 1992. Double immunogold localization of
thioredoxin f and photosynthetic fructose-1,6-bisphosphatase in
spinach leaves. Plant Physiology and Biochemistry 30, 39–46.
Mowla SB, Thomson JA, Farrant JM, Mundree SG. 2002. A novel
stress-inducible antioxidant enzyme identified from the resurrection
plant Xerophyta viscosa Baker. Planta 215, 716–726.
Restoration of photosynthesis in rehydrating Xerophyta scabrida leaves | 905
Tuba Z, Lichtenthaler HK, Csintalan Z, Nagy Z, Szente K. 1994.
Reconstitution of chlorophylls and photosynthetic CO2 assimilation
upon rehydration of the desiccated poikilochlorophyllous plant
Xerophyta scabrida (Pax) Th. Dur. et Schinz. Planta 192, 414–420.
Tuba Z, Lichtenthaler HK, Maroti I, Csintalan Zs. 1993a.
Resynthesis of thylakoids and functional chloroplasts in the desiccated
leaves of the poikilochlorophyllous plant Xerophyta scabrida upon
rehydration. Journal of Plant Physiology 142, 742–748.
Tuba Z, Proctor MCF, Csintalan Zs. 1998a. Ecophysiological
responses of homoichlorophyllous and poikilochlorophyllous
desiccation tolerant plants: a comparison and an ecological
perspective. Plant Growth Regulation 24, 211–217.
Vicré M, Farrant JM, Driouich A. 2004. Insights into the cellular
mechanisms of desiccation tolerance among angiosperm
resurrection plant species. Plant, Cell and Environment
27, 1329–1340.
Ward DA, Keys AJ. 1989. A comparison between the coupled
spectrophotometric and uncoupled radiometric assay for RuBP
carboxylase. Photosynthesis Research 22, 167–171.
Whittaker A, Bochicchio A, Vazzana C, Lindsey G, Farrant JM.
2001. Changes in leaf hexokinase activity and metabolite levels in
response to drying in the desiccation-tolerant species Sporobolus
stapfianus and Xerophyta viscosa. Journal of Experimental Botany
52, 961–969.
Downloaded from jxb.oxfordjournals.org at Centro de Información y Documentación Científica on January 18, 2011
Tuba Z, Smirnoff N, Zs Csintalan, Nagy Z, Szente K. 1997.
Respiration during slow desiccation of the poikilochlorophyllous
desiccation tolerant plant Xerophyta scabrida at present-day CO2
concentrations. Plant Physiology and Biochemistry 35, 381–386.