Download Recovery of the resurrection plant Craterostigma wilmsii from

Journal of Experimental Botany, Vol. 53, No. 375, pp. 1805±1813, August 2002
DOI: 10.1093/jxb/erf028
Recovery of the resurrection plant Craterostigma wilmsii
from desiccation: protection versus repair
Keren Cooper1 and Jill M. Farrant2,3
1
Department of Botany, University of Cape Town, Private Bag, Rondebosch, 7701, South Africa
2
Department of Molecular and Cell Biology, University of Cape Town, Private Bag, Rondebosch, 7701, South
Africa
Received 29 October 2001; Accepted 1 May 2002
Abstract
Craterostigma wilmsii Engl. (homoiochlorophyllous)
is a resurrection species that is thought to rely
primarily on the protection of cellular components
during drying to survive desiccation. The time
taken for this protection to be instituted is thought
to preclude recovery after rapid drying. Thus the
response of C. wilmsii plants to rapid dehydration
was investigated. The effect of rapid drying on
sucrose accumulation was determined and the cellular ultrastructure was investigated during natural
(slow) and rapid dehydration and on subsequent
rehydration. The dependence of naturally and rapidly dried C. wilmsii on de novo transcription and
translation during and after rehydration was determined by examining quantum ef®ciency, changes
in photosynthetic pigments and subcellular organization of excised leaves with rehydration in water
and using the metabolic inhibitors, distamycin A
and cycloheximide. Slowly dried C. wilmsii required
no new transcription or translation during rehydration in order to recover. With rapid dehydration,
cells showed ultrastructural damage, which indicated that at least some protective mechanisms
were affected (as evidenced by a reduced accumulation of sucrose). C. wilmsii was able to limit the
damage and recover upon rehydration in water, but
rapidly dried plants did not survive if mRNA or
protein synthesis was inhibited by distamycin A or
cycloheximide, respectively. This demonstrates an
induction of repair mechanisms during rehydration,
which enables recovery from rapid drying. Thus,
although C. wilmsii does rely almost entirely on
protection during natural drying, it apparently has
3
the ability to repair if protection is inadequate and
damage is incurred.
Key
words:
Craterostigma
wilmsii,
drying
rate,
homoiochlorophyllous,
metabolic
inhibitors,
modi®ed
desiccation tolerance, transcription, translation.
Introduction
During periods of prolonged water de®cit, it may not be
possible for plants to avert the loss of turgor in the cells,
resulting in mechanical stress, concomitant membrane
damage, metabolic disruption and damage due to free
radicals (reviews by Levitt, 1980; Smirnoff, 1993;
McKersie and Leshem, 1994; Bohnert et al., 1995;
Pammenter and Berjak, 1999). Resurrection or desiccation-tolerant plants, however, have the ability to tolerate
severe water loss (>90% RWC) and resume normal
physiological functioning on rehydration (Gaff, 1989). In
general, tolerance requires either a protection against the
stresses, repair of the resulting damage or a combination of
the two (Oliver, 1996). It has been suggested that lower
orders chie¯y repair damage associated with water loss,
while angiosperms, which have been termed modi®ed
desiccation-tolerant plants since their loss of water is
slower, utilize a greater degree of protection, induced
during dehydration (Oliver, 1996). It was therefore thought
that the time taken for induction and establishment
precludes survival of rapid drying (Gaff and Loveys,
1984; Bartels et al., 1990; Oliver et al., 1998; Tuba et al.,
1998).
However, Farrant et al. (1999), in a study on angiosperm
resurrection species, have shown that Craterostigma
wilmsii Engl. (homoiochlorophyllous) recovers after
To whom correspondence should be addressed. Fax: +27 21 698 7574. E-mail: [email protected]
ã Society for Experimental Biology 2002
1806 Cooper and Farrant
rapid (6±8 h) dehydration, while Xerophyta humilis (Bak.)
Dub. & Schinz (poikilochlorophyllous) does not. They
proposed that the strategy of protection employed by X.
humilis, which involves the dismantling of the photosynthetic apparatus, is impaired by rapid drying, resulting in
excessive damage due to light stress. C. wilmsii, conversely, has a swiftly instituted protection strategy
(involving leaf curling and accumulation of anthocyanins)
to avoid light stress (Sherwin and Farrant, 1998; Farrant,
2000) which enables it to survive.
Dace et al. (1998) investigated the effect of inhibiting
transcription and translation on the recovery of slowly
dried X. humilis. The results indicated that mRNAs for
chlorophyll synthesis and recovery of electron transport in
chloroplasts are stored in a stable form in the dried leaves
and are translated upon rehydration. However, additional
evidence suggested that new genomic transcription was
necessary after approximately 18 h for the complete
recovery of photosystem (PS) II functioning (Dace et al.,
1998).
The aim of this study was to establish whether
dehydration-induced mechanisms of protection in
Craterostigma wilmsii are affected by rapid drying and
to establish, by the use of metabolic inhibitors, the extent
to which this species is dependent on de novo transcription
and translation on rehydration after slow and rapid drying.
Materials and methods
Plant material
Whole plants of C. wilmsii (Scrophulariaceae) were collected and
maintained in a greenhouse at the University of Cape Town, as
previously described (Sherwin and Farrant, 1996; Dace et al., 1998)
until the initiation of the experiments described below.
Dehydration and rehydration
Whole plants were slowly dried by withholding water, allowing the
plants to dry naturally over approximately 8 d. Rapid drying in the
order of 20 h was accomplished as described previously by Farrant
et al. (1999), namely, whole plants were removed from the soil and
suspended on nylon gauze over silica gel within a plastic container.
Dry, compressed air was passed through a tube ®lled with silica gel
(at a rate of approximately 0.13 dm3 s±1), over the plant and out
through a perforated lid. During drying, the ranges of light intensity
and temperature were 150±400 mmol m±2 s±1 and 18±33 °C,
respectively.
Detached leaves were rehydrated for 48 h in Petri dishes
containing either water, a 310 g m±3 distamycin A solution (inhibitor
of transcription) or a 220 g m±3 cycloheximide solution (inhibitor of
translation) in a plant growth room at 25 °C. The light intensity was
1200 mmol m±2 s±1 on a 16/8 h light/dark cycle. The concentrations of
the inhibitor solutions were determined by testing the incorporation
of tritiated uridine and radiolabelled amino acids (35S-met,35S-cys
mixture).
At various intervals during drying and rehydration, leaves were
sampled for the procedures described below. Relative water content
(RWC) of at least three replicate sample leaves was determined by
the standard formula RWC(%)=(water content3100)/(water content
at full turgor). Water content and dry weight were determined
gravimetrically by oven-drying at 70 °C for 48 h. Mean water
content at full turgor was determined from at least 40 replicate leaves
taken from fully hydrated plants.
Sucrose content
The sucrose content of leaves from hydrated and dry plants, as well
as at two stages during dehydration was determined for at least four
replicate samples from different (at least three) plants. The RWC
ranges for the drying stages were: drying stage (i) 70±65% RWC;
drying stage (ii) 40±25% RWC.
Leaves were ground in liquid nitrogen and mixed with cold
extraction solution consisting of 100 mM NaOH in 50% ethanol (v/
v). The volume of extraction solution added was four (for hydrated
tissue) to eight (dry tissue) times the weight of leaf material. Extracts
were heated to 100 °C in a water bath for 10 min and then rapidly
cooled on ice. The extract was neutralized by the addition of 100
mM HEPES in 100 mM acetic acid. This was centrifuged at 16 000 g
for 20 min and the supernatant retained. The remaining pellet was reextracted using the same procedure as described above and the
supernatants of the two extractions combined.
The quantitation of the sucrose was carried out using a D-glucose/
D-fructose sugar assay kit (Boehringer Mannheim, Germany) based
on the methodology of Bergmeyer and Bernt (1974). The production
of NADPH (determined by reading the absorbance at 340 nm with a
spectrophotometer (Shimadzu UV-1601, Schimadzu Scienti®c
Instruments Inc., USA)) was used to calculate the quantity of
sucrose in samples.
Cellular ultrastructure
Cellular ultrastructure was investigated using transmission electron
microscopy (TEM). Small pieces of leaf tissue (approximately
2 mm2) were excised from at least four different leaves of a
minimum of two different plants. Samples were taken from
hydrated, dry and rehydrated tissue, as well as three stages during
dehydration. These drying stages were (i) 85±70% RWC; (ii) 65±
50% RWC; (iii) 40±25% RWC. The tissue was processed according
to a method previously used for this tissue (Sherwin and Farrant,
1996). Fixation was in 2.5% glutaraldehyde in 0.1 M phosphate
buffer (pH 7.4) containing 0.5% caffeine, post-®xed in 1% osmium
in phosphate buffer. After dehydration in a graded ethanol series, the
tissue was in®ltrated with epoxy resin (Spurr, 1969) over 4 d.
Samples were embedded in epoxy resin, hardened at 60 °C for 16 h
and sectioned at a gold interference colour (95 nm) using a
microtome (Ultracut-S, Reichert, Germany). Sections were stained
with 2% uranyl acetate and 1% lead citrate (Reynolds, 1963) and
viewed with a transmission electron microscope (EM-109, Zeiss,
Germany). Observations were made of general cell structure in at
least four different sections of all leaves prepared. Use of standard
®xation techniques might have allowed slight rehydration of the
desiccated leaves. Studies using cryo®xation of dry leaves of C.
wilmsii (Vicre et al., 1999) and freeze-substitution of dry tissue from
both species used in this study (J Wesley-Smith, University of Natal
Durban, personal communication) have given similar results to those
observed in this study. However, those methods allowed the
preservation of epidermis only and one or two cell layers below
that. In order to view the subcellular changes in all leaf cells,
standard ®xation was used here. Furthermore, since all treatments in
the current study were ®xed in the same manner, comparisons among
them are valid.
Chlorophyll and carotenoid content
The photosynthetic pigment content of at least three replicates of leaf
tissue from different (at least two) plants was determined as
described by Sherwin and Farrant (1996). Pigment was extracted
overnight in 100% acetone and the absorbance of the extract was
measured at 661.6 nm, 644.8 nm and 470 nm using a spectro-
Recovery from dehydration of Craterostigma 1807
Fig. 2. Sucrose content of slowly (®lled squares) and rapidly dried
(open squares) leaves of C. wilmsii at various stages during
dehydration. The RWC range at drying stage (i) was 70±65% RWC,
while that of drying stage (ii) was 40±25% RWC. Error bars represent
standard deviation. Letters represent mean separation by ScheffeÂ's
multiple range tests (P <0.05, n=4±7).
Fig. 1. (A) Dehydration time-course of whole plants of C. wilmsii
dried slowly (®lled circles) and rapidly (open circles). (B) Rehydration
time-course of slowly dried C. wilmsii leaves rehydrated in water
(®lled circles), distamycin A (open squares) or cycloheximide (open
triangles). Error bars represent standard deviation (n=3±6).
photometer (DU 650, Beckman, USA). Chlorophyll (a+b) and
carotenoid (x+c) contents were calculated using extinction coef®cients provided by Lichtenthaler (1987). The pigment content of
hydrated material (which had not been previously dried) was used as
a control.
Quantum ef®ciency of photosystem II
The quantum ef®ciency of PSII of plants that had been either rapidly
or slowly dried was determined by chlorophyll ¯uorometry during
rehydration. Five replicate leaves from different (at least two) plants
were dark adapted for 10 min (as previously determined) using darkadaption cuvettes. FV/FM was measured using a portable, modulated
¯uorometer (OS-500, Opti-Sciences, USA) with a saturating light
intensity of 7500 mmol m±2 s±1 to obtain maximal ¯uorescence (FM).
This experiment was performed twice.
Statistical analysis
Where appropriate, statistical analyses were conducted with the
Statgraphics Plus computer program (Statistical Graphic
Corporation, 1997).
Results
Dehydration and rehydration rates
The two drying treatments resulted in markedly different
rates of water loss (Fig. 1A). Slowly or naturally dried C.
wilmsii took approximately 8 d to reach an air-dry state,
whereas rapidly dried C. wilmsii achieved similar water
contents in 12 h. In both cases the ®nal water content
achieved was similar for rapidly and slowly dried material.
Figure 1B shows the rehydration time-courses for leaves
that were rehydrated in water and with the two metabolic
inhibitors. Rehydration rates (for all treatments) of rapidly
and slowly dried plants were not signi®cantly different
(Analysis of Covariance, P <0.05, n=3±6) and, therefore,
only data for slowly dried leaves are shown. Although there
was considerable variability in RWC of leaves during the
early stages of rehydration, all three rehydration treatments
exhibited comparable rates of rehydration, reaching 100%
RWC between approximately 15 h and 20 h.
Sucrose contents during dehydration
Figure 2 shows the sucrose contents of leaves of rapidly
and slowly dried plants. This sugar showed a substantial
increase in dry leaves from both drying treatments, with the
induction of sucrose accumulation only occurring during
the late stages of dehydration (below 25% RWC). Rapidly
dried leaves, however, accumulated less sucrose than
slowly dried ones, suggesting that the protection afforded
by this molecule was incomplete in rapidly dried plants.
Cellular ultrastructure during dehydration and on
rehydration
Figures 3 and 4 depict the subcellular organization of C.
wilmsii mesophyll cells in hydrated, dry and rehydrated
states and at several stages during dehydration. Control
(hydrated) C. wilmsii cells (Fig. 3A) had a large central
vacuole, with the cytoplasm and organelles situated at the
periphery of the cell. There was little variation in cell
appearance of tissues dried (regardless of rate) to at least
70% (data not shown). At water contents below this (65±
50% RWC), the ®rst signs of turgor loss and subcellular
reorganization became evident. In general, there was little
difference in ultrastructural changes of cells from rapidly
and slowly dried tissues, although at these water contents,
some cells from rapidly dried leaves had signs of
subcellular damage (primarily rupturing of plasma and
vacuolar membranes). The most marked changes in
cellular ultrastructure were observed at water contents
1808 Cooper and Farrant
Fig. 3. (A) Mesophyll cell of hydrated (control) C. wilmsii leaf. Cells are highly vacuolate (v) and the cytoplasm and chloroplasts (c) are situated
at the cell periphery. (B) Mesophyll cell from slowly dried C. wilmsii at drying stage 3 (40±25% RWC). Note the folding of the cell wall (w) and
the numerous small vacuoles (v). Chloroplast (c). (C) Damaged mesophyll cell from rapidly dried C. wilmsii at drying stage 3 (40±25% RWC).
The membrane has been almost completely destroyed (as indicated by arrows). (D) Undamaged mesophyll cell from rapidly dried C. wilmsii at
drying stage 3 (40±25% RWC). As with slowly dried material, there is extensive cell wall (w) folding and subdivision of vacuoles (v).
Chloroplasts (c). (E) Mesophyll cell from dry, rapidly dried C. wilmsii. As with slowly dried material, there is extensive cell wall (w) folding and
subdivision of vacuoles (v). Chloroplasts (c). (F) Damaged mesophyll cell from dry, rapidly dried C. wilmsii. Note the extensive plasma
membrane withdrawal. Chloroplasts (c). (G) Mesophyll cell of rehydrated rapidly dried C. wilmsii. There is a large central vacuole (v) and
chloroplasts (c) are pressed against the cell wall.
between 40% and 25% (Fig. 3B±D). In slowly dried
tissues, considerable cell wall folding had occurred, there
was substantial subdivision of the vacuoles and some
plasma membrane withdrawal from the cell walls (Fig. 3B).
The extent of this may have been an artefact of the aqueous
processing of samples, but there was no breakage of the
membranes. In this water content range, cells of rapidly
dried leaves also had conspicuous wall folding (Fig. 3C,
Recovery from dehydration of Craterostigma 1809
Fig. 4. (A) Mesophyll cell of slowly dried C. wilmsii tissue rehydrated to 100% RWC in distamycin A. Vacuole (v); chloroplasts (c). Note the
folding of the mesophyll cell wall (w) and electron dense material in the vacuole. (B) Chloroplast of slowly dried C. wilmsii tissue rehydrated to
100% RWC in distamycin A with thylakoid membranes (t) and starch grains (s) visible within. (C) Mesophyll cell of slowly dried C. wilmsii
tissue rehydrated to 100% RWC in cycloheximide. Vacuole (v); chloroplasts (c). Note the electron-dense material in the vacuole. (D) Chloroplast
of slowly dried C. wilmsii tissue rehydrated to 100% RWC in cycloheximide. Both starch grains (s) and thylakoid membranes (t) arranged into
grana are present. (E) Mesophyll cell and chloroplasts of rapidly dried C. wilmsii rehydrated to 100% RWC in distamycin A. There is severe
plasma membrane withdrawal, vacuoles (v), where still intact, have not coalesced and chloroplasts (c) lack boundary membranes. Membrane
breaks are indicated with arrows. (F) Mesophyll cell of rapidly dried C. wilmsii rehydrated to 100% RWC in distamycin A. There is very little
subcellular organization and cell walls (w) are still folded. (G) Mesophyll cell and chloroplasts of rapidly dried C. wilmsii rehydrated to 100%
RWC in cycloheximide. There is severe plasma membrane withdrawal, vacuoles (v), where still intact, have not coalesced and chloroplasts (c)
have blistered thylakoids. (H) Mesophyll cell of rapidly dried C. wilmsii rehydrated to 100% RWC in cycloheximide. There is very little
subcellular structure left. Membrane breaks are indicated with arrows.
D), but certain cells showed membrane breakage (Fig. 3C),
while the membranes of other cells appeared intact
(Fig. 3D). In the dry state, the majority of rapidly dried
cells again showed much the same appearance as slowly
1810 Cooper and Farrant
Fig. 5. Chlorophyll (A) and carotenoid (B) content of slowly dried
(®lled squares) and rapidly dried (open squares) C. wilmsii rehydrated
for 48 h and 72 h in water, distamycin A or cycloheximide. The
control is the pigment content of hydrated material (which had not
been previously dried). Error bars represent standard deviation. Letters
represent mean separation by ScheffeÂ's multiple range tests and show
comparisons within the chlorophyll (lower case letters) and carotenoid
(upper case letters) contents separately (P <0.10, n=3±6).
dried cells, as reported in Farrant et al. (1999), with folded
cell walls, U-shaped chloroplasts and numerous vesicles
(Fig. 3E). There were, however, small pockets of cells in
rapidly dried tissue that did show damage (Fig. 3F). Upon
rehydration in water, the tissue of both slow and rapid
drying treatments reverted to the appearance of the control
leaves, with no visible damage (as exempli®ed in Fig. 3G).
The cellular organization of mesophyll cells of
slowly and rapidly leaves rehydrated in solutions of
distamycin A or cycloheximide are shown in Fig. 4.
The subcellular organization of slowly dried leaves
rehydrated in distamycin A (Fig. 4A, B) or cycloheximide (Fig. 4C, D) showed almost identical results to
those rehydrated in water (not shown). Cells were
vacuolated and the plasma membrane had become reappressed to the cell wall, having incurred no visible
damage (Fig. 4A, C). Thylakoids were intact within the
chloroplasts and starch grains were visible (Fig. 4B,
D). Possibly the only visible difference was in the cell
walls of leaves rehydrated in the metabolic inhibitors,
which exhibited slight folding as exempli®ed in
Fig. 4A, something not evident in leaves rehydrated
in water. In addition, there appeared to be osmophilic
material within the vacuole in both distamycin A- and
cycloheximide-rehydrated tissue (Fig. 4A, C).
The cellular ultrastructure of mesophyll cells from
rapidly dried C. wilmsii rehydrated in solutions of
transcriptional inhibitor or translational inhibitor is
Fig. 6. Changes in quantum ef®ciency of photosystem II for (A)
slowly dried C. wilmsii and (B) rapidly dried C. wilmsii rehydrated in
water (®lled circles), distamycin A (open squares) or cycloheximide
(open triangles). Error bars represent standard deviation (n=5).
shown in Fig. 4E±H. These cells did not show any
recovery. Both treatments exhibited considerable folding
of cell walls and neither appeared to have reconstituted the
several small vesicles into a single central vacuole (Fig. 4E,
G). In each case, cells had suffered severe plasmolysis with
numerous membrane breaks visible (Fig. 4F, H) and
blistering of the thylakoid membranes in the chloroplasts.
In addition, chloroplasts lacked boundary membranes
(Fig. 4E, G). Indeed, in some cases, there were few intact
organelles present at all (Fig. 4F, H).
Chlorophyll and carotenoid content of rehydrated
tissue
There was a loss of approximately 60% of chlorophyll and
20% of carotenoids during slow drying of C. wilmsii
(Fig. 5A, B) as has been previously reported (Farrant et al.,
1999). During rehydration of these plants, there was very
little variation among the three treatments in chlorophyll
and carotenoid content, with both pigments being
resynthesized within 48 h of rehydration. It therefore
seems that the proteins required for synthesis of photosynthetic pigments are protected during drying and activated during rehydration.
Following rapid drying, only plants rehydrated in water
were able to resynthesize photosynthetic pigments (Fig. 5A,
B). When leaves were rehydrated in either inhibitor, there
was further breakdown and total loss of chlorophyll and
carotenoids. This could be attributed to photo-bleaching
caused by free radical action. The inability to recover
pigments suggests that the components required for
pigment synthesis had not survived rapid drying.
Recovery from dehydration of Craterostigma 1811
Quantum ef®ciency of photosystem II of rehydrated
tissue
There was no statistical difference in the rate of recovery
of PSII functioning among the three rehydration treatments
of slowly dried material (Fig. 6A) (Analysis of Covariance,
P <0.05, n=5). The similarity of the curves indicates that
the inhibitor solutions did not prevent full recovery of the
leaves and FV/FM increased to control levels (>0.7) within
15 h. The variability seen in the initial stages of
rehydration corresponds with the variability of RWC
before 20 h of rehydration (Fig. 1B). Furthermore, there
was no drop in quantum ef®ciency for up to 72 h of
rehydration in either inhibitor. This suggests that the
proteins present were adequate to maintain normal
photosynthetic metabolism for a considerable period
subsequent to rehydration.
Figure 6B shows the pattern of chlorophyll ¯uorescence
during rehydration of rapidly dried material. Only those
leaves rehydrated in water survived. Although both
inhibitors prevented eventual recovery of FV/FM, leaves
rehydrated in transcriptional inhibitor (distamycin A) were
able to initiate recovery at the same rate as that of the water
controls during the ®rst 2 h of rehydration (Analysis of
Covariance, P <0.05, n=5). This might suggest that some
transcripts coding for proteins (and/or pigments) involved
in PSII activity were present and translated to allow the
initial recovery. However, for full recovery new transcription must have been necessary. Leaf explants rehydrated in
cycloheximide exhibited no recovery of PSII activity upon
rehydration (Fig. 6B), suggesting that when dried rapidly,
protein translation on rehydration is vital for recovery of C.
wilmsii.
Discussion
C. wilmsii, being an angiosperm, is classi®ed as a modi®ed
desiccation-tolerant plant (Oliver, 1996). As such, its
`strategy' in surviving desiccation is primarily to protect
the subcellular milieu during drying, so as to limit the
damage incurred (Oliver, 1996). Previous research has
indeed found that such protective mechanisms play a
signi®cant role in the survival of this resurrection species
(Farrant and Sherwin, 1998; Sherwin and Farrant, 1998;
Vicre et al., 1999; Farrant, 2000). Despite the hypothesis
that this reliance on protection necessitates slow drying (in
order to allow suf®cient time to institute the required
mechanisms), Farrant et al. (1999) have shown that C.
wilmsii does survive rapid drying in the order of 6±8 h. In
that study, slowly and rapidly dried plants showed virtually
no difference in recovery of photosynthetic functioning,
pigment contents and electrolyte leakage during dehydration. However, from the data presented here, it is evident
that rapid drying does have some effect on C. wilmsii.
Folding of the cell walls in this species has been
proposed to be a mechanism to minimize mechanical stress
(Vicre et al., 1999; Farrant, 2000). In the current study wall
folding occurred only below a RWC content of 50% and
was relatively rapidly induced, with almost complete wall
folding visible in the 40±25% RWC range. While wall
folding occurred in both rapidly and slowly dried tissues,
some cells within the rapidly dried leaves did incur some
mechanical damage, indicated by membrane tearing and
some disarrangement of cell contents. This suggests that
not all cells were able to institute suitable protective
mechanisms to maintain membrane integrity. However,
damaged cells were suf®ciently few so as not to increase
electrolyte leakage signi®cantly (Farrant et al., 1999).
Another indication that rapid drying possibly prevented
full accrual of protection mechanisms is the reduced
sucrose accumulation in leaves of rapidly dried as
compared with those of slowly dried plants. This sugar is
thought to play a number of protective functions in dry
tissues, such as by replacing water on membranes and
macromolecules (Crowe et al., 1988), vitri®cation of the
cytoplasm (Leopold and Vertucci, 1986; Vertucci and
Farrant, 1995) and ®lling and stabilization of vacuoles
(Farrant, 2000). Thus the shortfall of sucrose in rapidly
dried leaves could have allowed lesions to develop in a
number of places. In general, the quantity of sucrose was
greater than previously reported for other resurrection
plants (Ghasempour et al., 1998; Norwood et al., 1999).
This difference is probably attributable to the low light
intensities in which the experimental plants used in those
studies were maintained prior to and during desiccation.
Possibly, this resulted in lower accumulation of photosynthetic products, with fewer reserves available for
sucrose production in the leaves.
Impairment of complete protection by rapid drying has
indirectly been shown to occur in another Craterostigma
species. Alamillo and Bartels (1996) have shown that
young C. plantagineum Hochst plants have reduced
accumulation of desiccation stress proteins (dsps). They
proposed that young plants dry more rapidly than the adult
plants and therefore cannot accumulate the typical quantity
of dsps.
However, insuf®cient protection mechanisms, whatever
the nature, did not compromize the survival of rapidly
dried C. wilmsii, when rehydrated in water, these plants
survived. Since there was little ultrastructural evidence of
the subcellular damage noted during drying in rehydrated
tissues, it is likely that this damage was repaired on
rehydration.
Intriguingly these data show that slowly dried plants
rehydrated in distamycin A (inhibitor of transcription) or
cycloheximide (inhibitor of translation) recovered completely. Photosynthetic pigments lost during desiccation
were recovered, as was full functioning of PSII. C. wilmsii,
being homoiochlorophyllous, does not lose all of its
photosynthetic pigments during drying and can probably
continue photosynthesis during early recovery without the
1812 Cooper and Farrant
need for immediate resynthesis of these pigments.
However, these data show full recovery (within 48 h) of
both carotenoid and chlorophyll contents in both inhibitors. In a similar study on Xerophyta humilis, the mRNAs
required for chlorophyll and carotenoid synthesis accumulated during slow drying and were stored in the dry
state, avoiding the necessity for de novo transcription on
rehydration (Dace et al., 1998). However, translation was
required for full recovery of the plants. It is possible that in
C. wilmsii, the mRNA and enzymes required for pigment
biosynthesis are accumulated prior to or during desiccation, and are suf®ciently protected in the dry state not to
require new transcription or translation during early
rehydration. Alternatively, the photosynthetic pigments
may have been partially disassembled into colourless
intermediates (Matile et al., 1999; Takamiya et al., 2000)
needing only reconstitution into an active state on
rehydration. Again, however, this would necessitate the
presence during drying and activity upon rehydration of
enzymes involved in the reconstitution process. Added to
this, photosynthesis (as measured by PSII activity), starch
accumulation in plastids (ultrastructural observation), and
probably all other metabolism (although not measured in
this study) continue upon rehydration independently of the
de novo transcription and translation.
Although slowly dried C. wilmsii seems almost
unaffected by inhibitor treatments, an exception was the
mechanism of cell wall unfolding. The folding of cell walls
is a controlled process, which is thought to involve the
accumulation during drying of components that increase
tensile strength (Vicre et al., 1999). Upon rehydration, the
quantity of these molecules (e.g. xyloglucan and pectins)
decline and the walls `unfold' (Vicre et al., 1999). It is
conceivable that although many of the proteins needed for
the reversal of this process are synthesized before or during
desiccation, newly synthesized components (requiring
transcription and translation, since wall unfolding does
not occur in either inhibitor) are required upon rehydration
for completion. Similarly, osmophilic material remains
accumulated in vacuoles of plants rehydrated in both
inhibitors, suggesting that the components necessary to
eliminate these from vacuoles are probably synthesized de
novo during rehydration. Alternatively, these may be byproducts of metabolism, which was disrupted due to the
imbibition in metabolic inhibitors.
In general, it would appear that C. wilmsii lays down
virtually complete protection, including the components
needed for recovery on rehydration, during desiccation.
Furthermore, C. wilmsii is able to maintain photosynthetic
functioning for at least 72 h of rehydration in either
distamycin A or cycloheximide. This indicates the robust
nature of the protection of the photosynthetic metabolism
and slow turnover of the enzymes involved in photosynthesis and/or the operation of effective repair processes.
Indeed, Schneider et al. (1993) found that translation
products accumulated during drying remained for many
hours after rehydration in the tissues of C. plantagineum. It
has been proposed that the nature of protection of the
genome against desiccation- and rehydration-induced
damage is such that its reorganization into an active state
on rehydration might require more time than most other
metabolism (Dace et al., 1998). Having all the components
necessary for recovery in place, without the need for new
genome-derived information, allows for rapid recovery.
Rapidly dried plants did not survive rehydration in the
presence of the metabolic inhibitors, distamycin A or
cycloheximide, with little recovery of photosynthesis
being recorded. It was proposed above that rapidly dried
plants of C. wilmsii do not have suf®cient time to institute
the protection needed completely, resulting in injury
during desiccation. These damaged components are not
repaired nor replaced when rehydrated in inhibitors, since
this would require de novo transcription and translation,
and thus viability is lost. It is not possible at this stage to
differentiate whether rapid drying also prevents the
establishment of repair systems, or damages repair
enzymes, or whether repair is only ever initiated de novo
on rehydration. However, if protection is complete enough,
then repair probably does not need to be put into place
during drying and would only be upregulated on rehydration if needed.
In conclusion, it is proposed that the survival strategy of
C. wilmsii relies on the production, during natural (slow)
drying, of full cellular protection and of the components
required for recovery. This ensures that little repair is
required on rehydration, although repair is possible if
injury is sustained. The study's data show that while some
repair processes could be constitutively present, any
replacement of damaged components requires de novo
transcription and translation. Such a strategy would enable
an extremely rapid recovery, especially of carbon metabolism, when water is once again available. Furthermore, C.
wilmsii can institute protection relatively rapidly, a strategy probably necessitated due to its lack of xerophytic
morphological features which serve to retard water loss.
These plants occur in an environment in which they
undergo frequent cycles of drying and rehydration (Gaff,
1977) and the ability to induce protection rapidly, much of
which might even be constitutive, would be essential to its
survival. In this respect, C. wilmsii does not conform to the
character of a typical modi®ed desiccation-tolerant plant.
This is a departure from the conventional assumptions
regarding classi®cations of desiccation-tolerance and may
extend to other desiccation-tolerant species. Certainly, the
®ndings of Proctor and Smirnoff (2000) indicate that the
photosystems of `fully' desiccation-tolerant bryophytes are
kept intact during desiccation and that repair systems are
not required for initial recovery of PSII quantum ef®ciency.
Only with long-term rehydration in light conditions does
the need for repair arise, most likely due to photo-oxidative
Recovery from dehydration of Craterostigma 1813
stress. Although increased complexity of angiosperm
resurrection plants does indeed engender a greater degree
of protection, rapid drying does not necessarily constrain
these mechanisms to such a degree as to prevent survival.
In addition, the role of repair in these species may have
been underestimated. In the light of these ®ndings, it seems
appropriate to re-evaluate the general classi®cations of
modi®ed versus full desiccation-tolerance.
Acknowledgements
We would like to thank Mohamed Jaffer and the staff of the Electron
Microscope Unit at the University of Cape Town for their assistance,
as well as John and Sandie Burrows for plant collection from
Buffelskloof Private Nature Reserve. This project was funded by the
NRF.
References
Alamillo JM, Bartels D. 1996. Light and stage of development
in¯uence the expression of desiccation-induced genes in the
resurrection plant Craterostigma plantagineum. Plant, Cell and
Environment 19, 300±310.
Bartels D, Schneider K, Terstappen G, Piatkowski D, Salamini
F. 1990. Molecular cloning of abscisic acid-modulated genes
which are induced during desiccation of the resurrection plant
Craterostigma plantagineum. Planta 181, 27±34.
Bergmeyer H, Bernt E. 1974. Methods of enzymatic analysis, Vol.
2. New York: Verlag Chemie Wenheim Academic Press.
Bohnert HJ, Nelson DE, Jensen RG. 1995. Adaptations to
environmental stresses. The Plant Cell 7, 1099±1111.
Crowe JH, Crowe LM, Carpenter JF, Rudolph AS, Wistrom
CA. 1988. Interactions of sugars with membranes. Biochimica et
Biophysica Acta 947, 367±384.
Dace H, Sherwin HW, Illing N, Farrant JM. 1998. Use of
metabolic inhibitors to elucidate the mechanism of recovery from
desiccation stress in the resurrection plant Xerophyta humilis.
Plant Growth Regulation 24, 171±177.
Farrant JM. 2000. A comparison of mechanisms of desiccationtolerance among three angiosperm resurrection plant species.
Plant Ecology 151, 29±39.
Farrant JM, Sherwin HW. 1998. Mechanisms of desiccation
tolerance in seeds and resurrection plants. In: Taylor AG, Huang
X-L, eds. Progress in seed science research. New York:
Communication Services of the New York State Agricultural
Experimental Station, 109±120.
Farrant JM, Cooper KL, Kruger LA, Sherwin HW. 1999. The
effect of drying rate on the survival of three desiccation-tolerant
angiosperm species. Annals of Botany 84, 1±9.
Gaff DF. 1977. Desiccation-tolerant vegetative plants of Southern
Africa. Oecologia 31, 95±109.
Gaff DF. 1989. Responses of desiccation tolerant 'resurrection'
plants to water stress. In: Kreeb KH, Richter H, Hinckley TM,
eds. Structural and functional responses to environmental
stresses: water shortage. Berlin: XIV International Botanical
Congress, 255±268.
Gaff DF, Loveys BR. 1984. Abscisic acid content and effects
during dehydration of detached leaves of desiccation-tolerant
plants. Journal of Experimental Botany 35, 1350±1358.
Ghasempour HR, Gaff DF, Williams RPW, Gianello RD. 1998.
Contents of sugars in leaves of drying desiccation-tolerant
¯owering plants particularly grasses. Plant Growth Regulation
24, 185±191.
Leopold AC, Vertucci CW. 1986. Physical attributes of desiccated
seeds. In: Leopold AC, ed. Membranes, metabolism and dry
organisms. New York: Cornell University Press, 22±34.
Levitt J. 1980. Responses of plants to environmental stresses, Vol.
II. Water, radiation, salt and other stresses, 2nd edn. New York:
Academic Press.
Lichtenthaler HK. 1987. Chlorophylls and carotenoids, the
pigments of the photosynthetic biomembranes. Methods in
Enzymology 148, 350±382.
Matile P, HoÈrtensteiner S, Thomas H. 1999. Chlorophyll
degradation. Annual Review of Plant Physiology and Plant
Molecular Biology 50, 67±95.
McKersie BD, Leshem Y. 1994. Stress and stress coping in
cultivated plants. Netherlands: Kluwer Academic Publishers.
Norwood M, Truesdale MR, Richter A, Scott P. 1999. Metabolic
changes in leaves and roots during dehydration of the resurrection
plant Craterostigma plantagineum (Hochst). South African
Journal of Botany 65, 421±427.
Oliver MJ. 1996. Desiccation tolerance in vegetative plant cells.
Physiologia Plantarum 97, 779±787.
Oliver MJ, Woods AJ, O'Mahony P. 1998. `To dryness and
beyond'Ðpreparation for the dried state and rehydration in
vegetative desiccation-tolerant plants. Plant Growth Regulation
24, 193±210.
Pammenter NW, Berjak P. 1999. A review of recalcitrant seed
physiology in relation to desiccation-tolerance mechanisms. Seed
Science Research 9, 13±37.
Proctor MCF, Smirnoff N. 2000. Rapid recovery of photosystems
on rewetting desiccation-tolerant mosses: chlorophyll
¯uorescence and inhibitor experiments. Journal of Experimental
Botany 51, 1695±1704.
Reynolds ES. 1963. The use of lead citrate at high pH as an
electron opaque stain for electron microscopy. Journal of Cell
Biology 17, 208±212.
Schneider K, Wells B, Schmelzer E, Salamini F, Bartels D. 1993.
Desiccation leads to rapid accumulation of both cytosolic and
chloroplastic proteins in the resurrection plant Craterostigma
plantagineum Hochst. Planta 189, 120±131.
Sherwin HW, Farrant JM. 1996. Differences in rehydration of
three desiccation-tolerant angiosperm species. Annals of Botany
78, 703±710.
Sherwin HW, Farrant JM. 1998. Protection mechanisms against
excess light in the resurrection plants Craterostigma wilmsii and
Xerophyta viscosa. Plant Growth Regulation 24, 203±210.
Smirnoff N. 1993. The role of active oxygen in the response of plants
to water de®cit and desiccation. New Phytologist 125, 27±58.
Spurr AR. 1969. A low viscosity epoxy resin embedding medium
for electron microscopy. Journal of Ultrastructural Research 26,
31±46.
Takamiya K-I, Tsuchiya T, Ohta H. 2000. Degradation
pathway(s) of chlorophyll: what has gene cloning revealed?
Trends in Plant Science 5, 426±431.
Tuba Z, Proctor MCF, Csintalan Z. 1998. Ecophysiological
responses of homoiochlorophyllous and poikilochlorophyllous
desiccation-tolerant plants: a comparison and ecological
perspective. Plant Growth Regulation 24, 211±217.
Vertucci CW, Farrant JM. 1995. Acquisition and loss of
desiccation-tolerance. In: Kigel J, Galili G, eds. Seed
development and germination. New York: Marcel Dekker, 237±
271.
Vicre M, Sherwin HW, Driouich A, Jaffer MA, Farrant JM.
1999. Cell wall characteristics and structure of hydrated and dry
leaves of the resurrection plant, Craterostigma wilmsii, a
microscopical study. Journal of Plant Physiology 155, 719±726.