Download Mechanical stabilization of desiccated vegetative tissues of the

Journal of Experimental Botany, Vol. 55, No. 397, pp. 651±661, March 2004
DOI: 10.1093/jxb/erh089 Advance Access publication 13 February, 2004
RESEARCH PAPER
Mechanical stabilization of desiccated vegetative tissues
of the resurrection grass Eragrostis nindensis: does a TIP
3;1 and/or compartmentalization of subcellular
components and metabolites play a role?
Clare Vander Willigen1, N. W. Pammenter2, Sagadevan G. Mundree1 and Jill M. Farrant1,*
1
Department of Molecular and Cellular Biology, University of Cape Town, Private Bag, Rondebosch, 7001,
South Africa
2
School of Life and Environmental Sciences, University of Natal, Durban, 4041, South Africa
Received 14 October 2003; Accepted 15 December 2003
Abstract
During dehydration, numerous metabolites accumulate in vegetative desiccation-tolerant tissues. This is
thought to be important in mechanically stabilizing
the cells and membranes in the desiccated state.
Non-aqueous fractionation of desiccated leaf tissues
of the resurrection grass Eragrostis nindensis
(Ficalho and Hiern) provided an insight into the subcellular localization of the metabolites (because of
the assumptions necessary in the calculations the
data must be treated with some caution). During
dehydration of the desiccant-tolerant leaves,
abundant small vacuoles are formed in the bundle
sheath cells, while cell wall folding occurs in the
thin-walled mesophyll and epidermal cells, leading to
a considerable reduction in the cross-sectional area
of these cells. During dehydration, proline, protein,
and sucrose accumulate in similar proportions in the
small vacuoles in the bundle sheath cells. In the
mesophyll cells high amounts of sucrose accumulate
in the cytoplasm, with proline and proteins being
present in both the cytoplasm and the large central
vacuole. In addition to the replacement of water by
compatible solutes, high permeability of membranes
to water may be critical to reduce the mechanical
strain associated with the in¯ux of water on rehydration. The immunolocalization of a possible TIP 3;1 to
the small vacuoles in the bundle sheath cells may be
important in both increased water permeability as
well as in the mobilization of solutes from the small
vacuoles on rehydration. This is the ®rst report of a
possible TIP 3;1 in vegetative tissues (previously
only reported in orthodox seeds).
Key words: Aquaporin, desiccation-tolerance, non-aqueous
fractionation, water stress.
Introduction
Many plants are subject to water-de®cit stress at some
stage during their life and, consequently, a diverse array of
strategies to survive this stress has developed. A few
unusual so-called resurrection plants are able to survive
extreme loss of water (desiccation) from their vegetative
tissues (Gaff, 1971). Mechanical damage is proposed to be
one of the major causes of irreversible desiccation-induced
damage in plants (Iljin, 1957). Extreme loss of water
results in a loss of cell volume as tissue water content
declines. Water plays an important structural role in plant
cells and it has been proposed that cells cannot contract
beyond a certain minimum `critical volume' without
irreparable damage to membranes (Meryman, 1974).
Vegetative tissues which are able to survive desiccation
to an air-dry state have mechanisms that cope with this
potentially lethal mechanical stress. Angiosperm desiccation-tolerant plants, relying predominantly on protective
mechanisms during dehydration, use two different
approaches to minimize this stress: water replacement or
a reduction in cellular volume (Gaff, 1971).
The desiccated cells of some resurrection plants become
®lled with vacuoles containing non-aqueous contents,
* To whom correspondence should be addressed. Fax: +27 21 689 7573. E-mail: [email protected]
Downloaded from http://jxb.oxfordjournals.org at Main Library L610 Lawrence Livermore Nat Lab on 4 November 2009
652 Vander Willigen et al.
replacing the volume occupied by water in the hydrated
state. This water replacement strategy has been proposed
to reduce the mechanical stress on the cellular substructure
(Farrant, 2000). The presence of numerous small vacuoles,
similar in appearance to the protein storage vacuoles found
in orthodox seeds, has been reported in a number of
resurrection species (Dalla Vecchia et al., 1998; Farrant,
2000; Vander Willigen et al., 2001a). Neither the formation of these small vacuoles nor their contents have been
investigated and hence their importance in reducing
mechanical stress is unclear.
An alternative strategy to prevent lethal mechanical
stress on cellular desiccation is to reduce cellular volume.
Extensive cell wall folding in some desiccation-tolerant
species has been recorded to cause as much as a 78%
reduction in cellular volume on dehydration (Farrant,
2000; Vander Willigen et al., 2001a). Unlike cytorrhysis
(irreversible cell wall collapse) observed in many desiccation-sensitive tissues, cell wall folding is associated with
changes in the biomechanical properties of the cell walls
brought about by changes in the biochemical properties of
the wall structure on dehydration (Vicre et al., 1999).
Vander Willigen et al. (2001a) proposed that this
controlled reduction in cellular volume on dehydration
was associated with unusual water relations in these
tissues. This possibly prevented the development of
negative turgor and, thereby, reduced the potential for
irreparable mechanical damage in these resurrection
plants.
Resurrection plants which are able to minimize
mechanical stress during dehydration must also have
mechanisms to minimize similar stresses associated with
the in¯ux of water into the cells on rehydration. Since
aquaporins play a crucial role in water movement, their
behaviour under water stress conditions is starting to
receive attention (Yamada et al., 1995, 1997; Kirsh et al.,
2000; Mitra et al., 2001). There are now some reports of
changes in the expression of aquaporins under stress
conditions which in¯uence not only the hydraulic conductivity of membranes but also water potential via
aquaporin phosphorylation (Liu et al., 1994; Johansson
et al., 1996, 1998; Sarda et al., 1997; Barrieu et al., 1999).
Not long after the discovery of water channel proteins in
plants, their role in desiccation tolerance was suggested
(Hartung et al., 1998). Five aquaporins have been isolated
from resurrection angiosperms: three PIPs (plasma membrane intrinsic proteins) and a TIP (tonoplast intrinsic
protein) from Craterostigma plantagineum (Mariaux et al.,
1998) and one TIP 1;3 from Sporobolus stap®anus (Neale
et al., 2000). Changes in the expression of these ®ve genes
during desiccation were reported; however no protein
studies on these TIPs or their role in desiccation tolerance
have been undertaken. Most of the studies on the role of
aquaporins in water stress have focused on the roles of
PIPs; very few studies have considered the function of
TIPs in water-limited environments.
The focus of this paper was to investigate the processes
that may be involved in dealing with mechanical stresses
associated with desiccation in the resurrection grass,
Eragrostis nindensis. The accumulation of small vacuoles
and metabolites on dehydration was investigated, and the
compartmentalization of the metabolites was characterized
using non-aqueous fractionation. In addition, the protein
expression and immunolocalization of a proposed TIP 3;1
(formerly a-TIP) were undertaken. The possible roles of
these phenomena in reducing mechanical stress are
discussed.
Materials and methods
Plant material
Mature Eragrostis nindensis (Ficalho and Hiern) plants from an
inselberg (granitic outcrop) located in a north-western region of
South Africa were potted in the sandy soil from the site and
translocated to a greenhouse at the University of Cape Town. All
plants remained hydrated for at least one month prior to any
experimentation.
The second leaf on a tiller from at least ®ve different E. nindensis
plants was harvested at intervals during desiccation (withholding of
watering) from plants which were initially hydrated to full turgor.
Prior investigation (Vander Willigen et al., 2001b) showed that the
outermost leaf on a tiller is sensitive to desiccation and so these
leaves have not been included in this study. After 2 weeks in the dry
state, plants were rewatered and the soil was held at ®eld capacity
during regular sampling over a rehydration period. Relative water
contents (RWC) were determined gravimetrically at each point, as
described previously (Vander Willigen et al., 2001b). The mean
RWC at each sampling point was calculated from leaf segments
either side of that used in each assay. Because diurnal changes in the
expression of TIPs and metabolites have been reported (Henzler
et al., 1999; Norwood et al., 2000), tissue was always harvested at
09.00 h, immediately plunged into liquid nitrogen, and stored at
±80 °C. Since the sampling intervals during rehydration were less
than 24 h, sampling times are indicated in the relevant legends.
TIP antibodies
The TIP antibodies used in this study were kindly provided by Dr JC
Rogers, (Institute of Biological Chemistry, Washington State
University, USA). The TIP 3;1 was isolated from Phaseolus vulgaris
seed. Antibodies were raised in rabbits against polypeptides
synthesized according to the following sequence: HQPLAPEDY,
by Jauh et al. (1998). This polypeptide forms part of a conserved
domain from the carboxyl-terminal, cytoplasmic tails of this TIP
isoform and has been found to be conserved across many species
(Jauh et al., 1998). In addition, antibodies raised against the entire
TIP 3;1 isolated from P. vulgaris (Johnson et al., 1989) were also
used. Preliminary experiments con®rmed that these antibodies only
cross-reacted with proteins of an expected size (c. 26 kDa) in
E. nindensis.
Transmission electron microscopy
Hydrated and desiccated leaves were prepared using conventional
and freeze-substitution techniques, respectively. Conventional preparation methods are described in Sherwin and Farrant (1996).
For preparation of material by freeze-substitution, leaf segments
(c. 0.5 mm2) were ®rst cryo®xed by immersion in dry, precooled
Downloaded from http://jxb.oxfordjournals.org at Main Library L610 Lawrence Livermore Nat Lab on 4 November 2009
Mechanical stabilization in desiccated tissues 653
(±80 °C) acetone using a Reichert KF8 cyro®xation instrument
(Leica, Vienna, Austria). Thereafter they were transferred to a Leica
EM AFS automatic freeze-substitution system (Leica, Vienna,
Austria). The samples were then kept at ±80 °C for 24 h in a
precooled solution of 0.1% tannic acid in anhydrous acetone. After
rinsing with anhydrous acetone, the samples were left in a precooled
freeze-substitution solution containing 2% (w/v) anhydrous glutaraldehyde (Electron Microscopy Sciences, Fort Washington, USA)
and 2% (w/v) uranyl acetate in dry acetone, for 48 h at ±80 °C. The
temperature was then set to increase by 1 °C h±1 up to ±20 °C. At this
temperature 2% (w/v) osmium tetroxide in dry acetone was added to
the freeze-substitution solution. After 24 h in this medium, the
samples were transferred into fresh osmium-free freeze-substitution
solution and rewarming by 1 °C h±1 was resumed. Samples were then
washed with acetone at room temperature and detached from the
nickel grids before in®ltrating with increasing concentrations of low
viscosity epoxy resin (Spurr, 1969) over at least 5 d. The in®ltrated
specimens were embedded in the resin and polymerized at 60 °C for
48 h. Ultrathin sections (120±150 nm) from the outermost parts of
the embedded tissue were cut on a Reichert Ultracut S (Leica,
Vienna, Austria) and mounted on Formvar-coated copper grids (300
mesh). Sections were brie¯y (less than 1 min) stained with 2% uranyl
acetate and lead citrate (Reynolds, 1963) and viewed with a JEM200CX transmission electron microscope (JEOL, Tokyo, Japan).
Protein extraction
Leaf segments (from 0.5±4 cm distal to the leaf sheath) of
E. nindensis and whole seeds P. vulgaris were ground in liquid
nitrogen and homogenized in Trizol (Life Technologies, GIBCO
BRL) for 5 min at room temperature with vortexing (c. 0.1 g tissue in
1 ml Trizol per microcentrifuge tube) according to the methods of
Chomzynski and Sacchi (1987). The protein extraction method was
modi®ed slightly by washing the pellets three times with cold 0.1 M
ammonium acetate and once with cold acetone (centrifugation at
12 000 g for 10 min at 4 °C and 5 min incubation periods between
washes). Air-dried protein samples were stored at ±20 °C.
Protein quanti®cation
Resuspended proteins were quanti®ed according to the method of
Bradford (1976) using a protein assay kit (Pierce, Rockford USA).
The absorbance of 40 ml protein samples that had been incubated
with 2 ml of the diluted Bradford reagent for 2 min, was read at
595 nm (Beckman DU 530 spectrophotometer, Fullerton, CA, USA).
Protein concentrations were calculated from a standard curve,
constructed using bovine serum albumin (BSA).
Western blot analysis
Proteins (15 mg per lane) were incubated at 37 °C for 10 min and
electrophoresed on 20% SDS-PAGE gels containing 0.1% bis
acrylamide. The gel, PVDF membrane, Whatmann 3MM paper, and
foam pads (all cut to the same size as the gel) were soaked in transfer
buffer (192 mM glycine, 25 mM TRIS, 3.5 mM SDS, 20% (v/v)
methanol). The proteins were transferred to the PVDF western
blotting membrane (Boehringer Mannheim) for 1 h at 30 V in a
precooled western blotting tank (Hoefer, San Francisco, USA). The
membrane was rinsed with TBS (50 mM TRIS, 150 mM NaCl,
pH 7.5) and non-speci®c binding sites were blocked with 0.5% (w/v)
skimmed milk powder in TBS for 1 h at room temperature with
shaking. The membrane was then incubated overnight with the
primary antibody (1:2000 dilution (polypeptide); 1: 3000 dilution
(entire protein)) diluted with 0.5% (w/v) skimmed milk powder at
4 °C with shaking. After two washes with TBS containing 0.1%
Tween 20 for 10 min, the membrane was incubated with the
secondary antibody (goat anti-rabbit IgG conjugated to horseradish
peroxidase) at 1:1000 dilution for 1 h at room temperature with
shaking. The membrane was thoroughly washed before detection
using chemiluminescence according to the manufacturer's instructions (Amersham Pharmacia, Buckinghamshire, UK). Blots were
developed on high performance chemiluminescence ®lm
(Amersham Pharmacia, Buckinghamshire, UK), scanned and
analysed using Biorad Quantity One 4.4 software (Biorad,
California, USA). Duplicate gels were routinely stained with
Coomassie Brilliant Blue R-250 (0.25% (w/v) in 50% (v/v)
methanol, 7% acetic acid (v/v)) to con®rm equal protein loading.
All blots were repeated at least tree times on separately extracted
protein samples from different plants.
Immuno¯uorescence studies
Leaf segments (c. 5 mm2 pieces) from 2 cm distal to the leaf sheath,
collected during numerous stages of dehydration and rehydration,
were ®xed in 4% (w/v) paraformaldehyde in 0.1 M Sùrensen's
phosphate buffer (pH 7.2) overnight at 4 °C. Specimens were then
dehydrated through an alcohol series (50%, 70%, 95%, and 100%
ethanol, 100% isopropanol, 100% butanol) and embedded in paraf®n
wax at 60 °C overnight. Thick wax-embedded transverse sections
(10 mm) were cut on a Leitz rotary microtome (Vienna, Austria) and
attached to APTES- (3-aminopropyltriethoxysilane) coated glass
microscope slides by heating to 60 °C for 1 h. After dewaxing with
xylol and rehydrating through an ethanol gradient, the sections were
washed with PBS (50 mM NaH2PO4, 150 mM NaCl, pH 7.4) and
blocked with 1% BSA in PBS for 1 h in a moist chamber at room
temperature. Sections were incubated with the TIP 3;1 polypeptide
antibody (1:100 dilution) or 1% BSA (control) in PBS overnight at
4 °C in a moist chamber. Thereafter, the sections were jet-washed
with 0.1% Triton X-100 in PBS and then rinsed for a further 15 min
in the buffer before incubating in the ¯uorochrome tagged secondary
goat anti-rabbit IgG antibody (Alexa ¯uor 568, Molecular Probes,
Oregon USA: dilution 1:1000) for 2 h in a moist chamber at room
temperature in the dark. The slides were then thoroughly washed
with PBS containing Triton X-100. Sections were mounted with
ProLong Antifade (Molecular Probes, Oregon, USA) and viewed
with an inverted ¯uorescent microscope (Nikon, Tokyo, Japan)
under oil emersion with a B2 DM510 epi-¯uorescence ®lter (Zeiss,
Tokyo, Japan). The approximate absorption and ¯uorescence
emission maxima for Alexa ¯uor 568 are 578 nm and 603 nm,
respectively (Molecular Probes Inc., 2000). A confocal laser
scanning microscopy (550 Confocal Microscope, Leica,
Cambridge) was used to con®rm the results, using a narrower
range of wavelengths (600±610 nm). Images were captured with an
Axiocam digital camera (Zeiss, Hallbergmoos, Germany) using
AxioVision 2.05 software (Carl Zeiss Vision GmbH, Hallbergmoos,
Germany). Care was taken to use the identical exposure times and
images were not adjusted thereafter for comparative purposes.
Non-aqueous fractionation of dry leaf tissues
The subcellular compartments of dry leaf tissues (4.260.6% RWC)
were separated using slight modi®cations of the non-aqueous
fractionation method described by Stitt et al. (1989). Dry leaves
(c. 2 g) were ground in liquid nitrogen and dried in a lyophilizer for
24 h. Care was taken at all stages after lyophilization to avoid any
condensation forming in the samples. Each sample was resuspended
in a precooled tetrachloroethylene:heptane solution (66:34 (v/v),
density: 1.3 g cm±3) which had been dried over molecular sieve
beads. The samples were ultrasonicated on ice (Sonorex TK 52H,
Bandelin, Berlin) for a total of 60 s in 10 s pulses to prevent
overheating. The homogenate, diluted 3-fold with dry, precooled
heptane, was then passsed through a glass micro®bre ®lter and
centrifuged for 10 min at 2200 g (Sigma 3K10 centrifuge, Germany).
The supernatant was discarded while the sediment was resuspended
in 3 ml of the 66:34 tetrachloroethylene:heptane solution. Aliquots
Downloaded from http://jxb.oxfordjournals.org at Main Library L610 Lawrence Livermore Nat Lab on 4 November 2009
654 Vander Willigen et al.
(of the suspension) (200 ml) were kept aside for measurement of
enzyme activity and metabolite assays in the unfractionated material
and the remainder was layered onto a prepared linear density
gradient (12 ml between 1.3 and 1.62 g cm±3). The gradient was
prepared in polyallomer ultracentrifuge tubes immediately prior to
use from dried solutions of heptane and tetrachloroethylene using a
gradient former connected to a peristaltic pump (Miniplus 3, Gilson,
France). The sample with the preformed gradient was centrifuged for
2 h at 15 000 g at 5 °C (swing-out rotor SW28, ultracentrifuge
(Beckman L5-65, Fullerton, USA)). The content of the centrifuge
tube was then collected in eight fractions and each fraction separated
into two portions; 1/3 for marker enzyme assays and 2/3 for
metabolite assays. All aliquots were diluted 3-fold with heptane and
centrifuged for 10 min at 2200 g. The supernatant was discarded and
the pellets were dried in a desiccator over silica gel for 24 h and then
extracted for the various assays.
Marker enzyme assays
Extracts were prepared by adding 500 ml of a solution containing
0.25 mM potassium phosphate (pH 7.5), 0.5 mM dithiothreitol
(DTT), and 0.5 mM EDTA to the dried pellets. After vigorous
shaking, another 500 ml of the same solution was added. The enzyme
activities were measured in the supernatant of the solution after
centrifugation for 2 min in a bench-top centrifuge. NADPglyceraldehyde-3-phosphate dehydrogenase (EC 1.2.1.13) was
measured by the oxidation of NADPH at 340 nm using the method
described by Stitt et al. (1989). This assay was used as a
cytosolic marker. Soluble acid invertase was used as a lytic (central)
vacuole marker, assayed as the hydrolysis of sucrose to glucose
according to the methods of Kingston-Smith et al. (1998). The TIP
3;1 was found to be associated with the small vacuoles (see below)
and so this subcellular compartment was identi®ed by blotting the
fraction extracts onto nitrocellulose membranes which were then
incubated with TIP 3;1 antibodies as described previously. Care was
taken to blot equal amounts of extract and spot intensity was
recorded using Biorad Quantity One 4.4 software (Biorad,
California, USA). All samples were loaded onto the same ®ve
replicate membranes.
Metabolite assays
The following assays were performed on bulk leaf tissue samples at
various stages of dehydration and rehydration as well as on aliquots
of the dry leaf samples after non-aqueous fractionation. In both cases
(bulk leaf samples and fractionated samples) at least three replicate
assays from different plants were performed, with two internal
replicates for each assay.
Sugars: Ground frozen leaf samples (and the fractionated samples)
were extracted in cold 100 mM NaOH (50% (v/v) ethanol/water).
Chloroform (15%, v/v) was added and the samples incubated on ice
for 10 min. Thereafter the pH was adjusted to 7.5 with 100 mM
HEPES in 100 mM glacial acetic acid. After centrifugation at 28 000
g for 20 min at 4 °C, the supernatant was removed. A repeat
extraction was performed on the pellets. The supernatants were
pooled and then centrifuged. Sucrose, glucose, and fructose in the
supernatant were calculated from the spectrophotometric measurement of NADPH production using a D-glucose/D-fructose sugar
assay kit (Boehringer Mannheim, Germany).
Free proline: Free proline was measured spectrophotometrically
according to the methods of Magne and Larher (1992) for samples
with high sugar contents. Frozen ground leaf samples (and the
fractionated samples) were extracted in 100% toluene. A 1%
ninhydrin solution in 60:40 (v/v) glacial acetic acid:water was then
added and the samples heated in a boiling water bath for 1 h. Twice
the sample volume of toluene was added to the cooled samples. After
vigorous shaking, the upper phase was read at 520 nm.
Concentrations were calculated from a standard curve derived
from known concentrations of proline.
Protein: Bradford (1976) assays, as described previously, were
performed on the fractions.
Determination of cellular and subcellular volumes
Desiccated leaves were hand-sectioned dry and viewed immediately
in an iso-osmotic solution of PEG 6000 using a light microscope
(Ascopscope; Zeiss, Hallbergmoos, Germany). Preliminary experiments were undertaken to ensure that no swelling of the tissues
occurred prior to photographing the sections. Hydrated leaves were
sectioned and viewed in deionized water. Cellular cross-sectional
areas of more than 40 digital images from at least eight individual
plants from both the hydrated and dehydrated leaves were measured
using an image analysis program (AnalySIS, Soft Imaging
Software), and it was assumed that cross-sectional area is an
unbiased estimate of volume (Winter et al., 1993). Volumes of
subcellular compartments in dry leaves were estimated from crosssectional areas measured from digital images of transmission
electron micrographs of the dry leaves. At least 15 grid blocks
from each of four different sections from at least two different
specimens per sample were analysed.
Preparation of these images was as previously described (see
`Transmission electron microscopy').
Results
Formation of vacuoles during desiccation
During dehydration of the desiccation-tolerant inner leaves
of E. nindensis, numerous small vacuoles formed in the
bundle sheath cells (BSC) of these leaves (Fig. 1A). Unlike
the mesophyll cells in which extensive cell wall folding
was observed (Fig. 1B), the thicker-walled BSC maintained most of their volume and shape in the desiccated
state, comprising 37% and 33% of the cross-sectional area
of the leaves in the hydrated and dehydrated states,
respectively. The folding of the thin walls of the mesophyll
cells led to a considerable reduction in volume on
dehydration, such that, in the dry state, they comprised
only 22% of the leaf cross-sectional area compared with
49% in the hydrated state.
The characteristics of the small vacuoles in the BSC
during dehydration are summarized in Table 1. As
dehydration occurred, the large central vacuole became
reduced in size and small vacuoles started to form from the
periphery of the cell. In the air-dry state no central vacuole
was apparent; the cells were packed with numerous small
vacuoles. It is unclear whether this change in size and
number of vacuoles was due to fragmentation of the central
vacuole or de novo synthesis of the smaller ones. However,
although total vacuolar volume of the BSC did not change
during desiccation, the length of tonoplast visible on the
micrographs increased (Table 1). After a 6 h rehydration
period, to a RWC of 15%, the number of vacuoles had been
reduced by 66% compared with the number in the dry state
(Table 1).
Downloaded from http://jxb.oxfordjournals.org at Main Library L610 Lawrence Livermore Nat Lab on 4 November 2009
Mechanical stabilization in desiccated tissues 655
Fig. 1. Transmission electron micrograph of a bundle sheath cell (A) and mesophyll and epidermal tissues (B) of dry inner E. nindensis leaves
prepared by freeze-substitution. Numerous small vacuoles (v) pack the bundle sheath cell and the walls of these cells (bcw) are not folded. Note
the extensive mesophyll cell wall (mcw) folding and absence of small vacuoles in the mesophyll and epidermal cells. The magni®cation bars
represent 2 mm.
Table 1. Characteristics of vacuoles of the bundle sheath cells of the desiccation-tolerant inner leaves of E. nindensis during
dehydration and rehydration
Standard deviations are given in parentheses. Letters represent the mean separation by Scheffe's multiple range tests, showing comparisons in
each parameter between treatments (P <0.05, n=20±25, at least eight individual plants).
Dehydration
Number of vacuoles/cell
Ratio of no. of large:small vacuoles
Total vacuolar cross-sectional area (mm2/cell)
Total length of tonoplast visible on micrograph (mm/cell)
Rehydration
98% RWC
26% RWC
4% RWC
15% RWC
1 (0) a
1:0
38 (3) a
74 (5) a
17 (2) b
1: 15
37 (3) a
83 (6) a
67 (3) c
0: 67
34 (3) a
125 (13) b
23 (6) b
1: 22
35 (4) a
95 (9) ab
Tonoplast intrinsic protein
Western blots using polypeptide antibodies to TIP 3;1
suggested that the protein was present only (or recognized
the antigen only) in the desiccated leaves of E. nindensis
(Fig. 2). It thus seemed that a TIP 3;1 appeared very late
during dehydration, once the leaves had less than
0.4360.22 g H2O g±1 dry mass (11% RWC) and that it
was degraded rapidly on rehydration. This result was
con®rmed using the TIP 3;1 antibodies raised against the
entire protein (Johnson et al., 1989; data not shown). Like
all other vegetative tissues studied to date (Jauh et al.,
1998), TIP 3;1 was not found in the desiccation-sensitive
outer leaves of E. nindensis (data not shown).
Immuno¯uorescence studies suggested that TIP 3;1 was
present only in dry inner leaves of E. nindensis (Fig. 3).
The localization of the ¯uorescent probe to the abundant
small vacuoles present in the BSC (Fig. 1) of this tissue in
the dry state suggests the presence of a TIP 3;1 in these
small vacuole tonoplasts in E. nindensis. Unfortunately the
wavelength and intensity of the background ¯uorescence
changed with tissue hydration state and thus could not be
eliminated completely. This is likely to be a consequence
of the changes in pigment composition and concentration
with desiccation (Vander Willigen et al., 2001b).
Subcellular distribution of metabolites
During dehydration of the inner leaves, an accumulation of
proline, sucrose, and protein was recorded (Fig. 4A1, B1,
C1). Proline and sucrose contents decreased early on
rehydration (Fig. 4A2, B2), whereas a higher concentration
of proteins was maintained during the rehydration process
(Fig. 4C2). Non-aqueous fractionation of the dry leaves
enabled an analysis of the subcellular compartmentation of
the accumulated metabolites without any rehydration of
the tissue, although the analysis is complicated by the
presence of two tissue types, BSCs and mesophyll/
epidermal cells. Only vacuolar and cytoplasmic (which
included mitochondria, chloroplasts, and nuclear compartments) fractions were analysed separately.
Figure 5 shows the distribution of enzyme markers
(reaction to TIP 3;1 antibody for small vacuoles) and hence
the distribution of subcellular compartments (cytoplasm,
large central vacuole, small vacuoles) in the dry leaf tissue
fraction from the non-aqueous density gradient. From
Downloaded from http://jxb.oxfordjournals.org at Main Library L610 Lawrence Livermore Nat Lab on 4 November 2009
656 Vander Willigen et al.
Fig. 2. Western blot of desiccation-tolerant inner leaves of
E. nindensis probed with TIP 3;1 polypeptide antibodies. Proteins
(15 mg lane±1) were extracted from tissue at a range of water contents
during dehydration and rehydration (absolute water content (AWC)
and relative water content (RWC) denoted on top of the gel). The
dehydration and rehydration time is also indicated.
these data the proportional distribution of metabolites
among the subcellular compartments was calculated using
a method similar to that described by Riens et al. (1991)
(Table 2). This table also shows the initial concentration of
the metabolites in the dry leaf tissue. From the distribution
of metabolites among compartments and the initial concentration of metabolites in whole leaf tissue, the mass
percentage of each metabolite in each compartment was
calculated (Table 2). To allocate the metabolites to
subcellular compartments in the two tissue types (BSCs
and mesophyll/epidermal cells) requires that the contribution of each compartment in each tissue type to total leaf
volume be known. This was achieved by multiplying the
volume contribution of each compartment to each tissue
type (Table 3) by the proportion that each tissue type
contributed to the whole dried leaf (3364% for the BSCs
and 2263% for mesophyll/epidermal cells). The results of
these calculations are also shown in Table 3 (the volumes
of vascular tissues or other cellular constituents not
analysed by non-aqueous fractionation were not measured
directly). The mass percentage of a metabolite in a
compartment (Table 2) was then allocated to a tissue
type according to the ratio of that compartment in each
tissue type (for example, 6.4 g protein in the cytoplasm per
100 g whole leaf tissue is distributed in the ratio 1:9
between BSC cytoplasm and mesophyll/epidermal cytoplasm). The resultant distribution of metabolites among
compartments in the tissue types is given in Table 4.
Allocating metabolites amongst compartments in this
manner requires two assumptions: (i) the concentration
of a metabolite in a compartment is the same in both tissue
types, and (ii) all the metabolites measured in bulk leaf
tissue are accounted for in the analysis of the subcellular
compartments (i.e. concentrations of protein, proline, and
sugars in structural material and vascular tissue are low). In
addition, combining proportions based on mass (Table 2)
Fig. 3. Immunolocalization of TIP 3;1 in desiccation-tolerant inner
leaves of E. nindensis in the dehydrated state (A, B) and the fully
hydrated state (C, D). No ¯uorescence ascribable to the alexa¯uor 568
probe can be detected in the immunological controls (A, C) in which
BSA was substituted for the TIP 3;1 antibody. When TIP 3;1 labelled
with alexa¯uor 568 was added (B, D), small vacuoles (arrowed)
present in dehydrated tissue (B) become labelled in red. No small
vacuoles nor TIP 3;1 labelling was evident in the hydrated leaves (D).
The magni®cation bars represent 10 mm.
and volume (Table 3) to yield concentrations (Table 4)
invokes the assumption that the density of the dry leaf
material is homogeneous. This assumption is not true
(otherwise separation on a density gradient would not
occur), but the range of densities on the gradient (1.3±1.6 g
cm±3) was not large and would not appreciably affect the
patterns of metabolite distributions calculated.
A large proportion of the small vacuoles were packed in
the BSCs (Table 3), and these structures contained
approximately equal amounts of protein and free proline,
and somewhat more sucrose. These constituents total
nearly 13% of the dry mass of the dehydrated leaves;
however, since these small vacuoles occupy over 30% of
the leaf tissue (Table 3) there must be many more, as yet
unidenti®ed, metabolites in these small vacuoles. From the
insert to Fig. 1, it was clear that many of the small vacuoles
are of different electron density and hence it is suggested
that the distribution of the metabolites is not uniform in
each vacuole. It is interesting to note that most of the
proline was found in the small vacuoles and not the
cytoplasm (Table 4) and hence may not act an osmolyte in
this environment. Sucrose concentration in the dry leaf was
high (Table 2) and most of this was located in the
cytoplasm of the mesophyll/epidermal cells (Table 4).
Unfortunately, differences in the composition of the
cytoplasm of the different cell types or the composition
of individual organelles could not be determined using
these techniques.
Downloaded from http://jxb.oxfordjournals.org at Main Library L610 Lawrence Livermore Nat Lab on 4 November 2009
Mechanical stabilization in desiccated tissues 657
Fig. 4. Concentrations of proline (A1, A2), sugars (B1, B2), sucrose (®lled squares), glucose (®lled triangles), and fructose (®lled circles), and
protein (C1, C2) in inner desiccation-tolerant leaves of E. nindensis during dehydration (left panel) and rehydration (right panel). Error bars
represent standard deviations, n >3.
Discussion
Resurrection plants are thought to survive the mechanical
stresses associated with desiccation using one of two
strategies. Extensive cell wall folding, which should
reduce tensions on structures enclosed by the wall, has
been reported in some resurrection plants (Hallam and
Luff, 1980; Vicre et al., 1999). Alternatively, cells
maintain their original volume and become packed with
vacuoles ®lled with non-aqueous matter (Dalla Vecchia
et al., 1998; Farrant, 2000). In E. nindensis, both strategies
are employed; the thicker-walled BSCs maintain volume
and abundant small vacuoles replace the water of the
hydrated cells (Fig. 1A), and the thinner-walled mesophyll
and epidermal cells become reduced in volume and
extensive cell wall folding is observed (Fig. 1B).
The importance of cell wall folding in the mechanical
stabilization of desiccated tissues, as observed in the
mesophyll and epidermal cells (Fig. 1B) have been
investigated in both this and other resurrection plants
Downloaded from http://jxb.oxfordjournals.org at Main Library L610 Lawrence Livermore Nat Lab on 4 November 2009
658 Vander Willigen et al.
(Vicre et al., 1999; Vander Willigen et al., 2001a).
However, the signi®cance of abundant small non-aqueous
vacuoles in desiccated tissues such as those observed in the
BSCs of E. nindensis (Fig. 1A; Table 1) has not been
studied. Due to dif®culties in investigating the ultrastructure of partially hydrated or dry tissues (Platt et al., 1997),
it was not possible to determine how and from where these
vacuoles originated. However, it was evident that at least
some de novo membrane synthesis must have occurred
since the total tonoplast length per cell increased on
Fig. 5. Distribution among fractions 1±8 of the marker enzymes
(solid bars, cytosolic marker; grey bars, vacuolar marker) and TIP 3;1
antibodies (open bars) after non-aqueous fractionation of dry
desiccation-tolerant inner E. nindensis leaves (5% RWC).
dehydration (Table 1). A relatively large surface area-tovolume ratio has been suggested to be important to the
mechanical stabilization of dehydrated cells (Iljin, 1957).
Since this author reported that small cells were reported to
be more tolerant to drying than larger cells, it is probable
that in desiccated vegetative tissues numerous small
vacuoles are mechanically more stable than one large
one in desiccated tissues.
There are numerous reports of an increase in metabolites
in resurrection plants on drying (Fig. 4; Tymms and Gaff,
1979; Bianchi et al., 1991; Ghasempour et al., 1998;
Whittaker et al., 2001), but the subcellular localization of
these has not been established previously. Non-aqueous
fractionation has become a useful technique for analysing
subcellular biochemistry (Heineke et al., 1997; Farre et al.,
2001) and it is, in fact, particularly well suited to
desiccated material in which the addition of aqueous
solvents is undesirable (to avoid rehydration). This technique has not yet been validated in heterogeneous tissues
and a number of assumptions were necessary before any
®nal metabolite distributions could be calculated. Thus
there is a greater degree of uncertainty in the results
obtained and caution in their interpretation is necessary.
Using non-aqueous fractionation, it was found that there
were almost equal amounts of proline and protein and
slightly more sucrose in the small vacuoles in E. nindensis
(Table 4), but there is a signi®cant proportion of other, as
yet unidenti®ed, constituents as well. Proline accumulation
Table 2. Subcellular distribution of metabolites in dry (5% RWC) desiccation-tolerant inner leaves of E. nindensis separated
using non-aqueous fractionation
Metabolites in each fraction were measured spectrophotmetrically and the subcellular distribution calculated by comparing the metabolite and
marker enzyme distribution. The results represent the means 6standard deviation of the measurements on three different fractionations with
different tissue samples, each pooled from at least four individual plants.
Proportional distribution in each compartment
Protein
Free proline
Glucose
Fructose
Sucrose
Cytoplasm
Small
vacuole
Central
vacuole
5164
1862
5962
6062
7264
3063
5363
1762
1661
1563
1963
2962
2562
2463
1464
Concentration in leaf
(g/100 g)
Mass percentage in each compartment
Cytoplasm
Small
vacuole
Central
vacuole
12.760.7
7.660.6
0.0360.01
0.1660.09
13.661.1
6.4
1.4
0.02
0.1
5.6
3.9
4.0
0
0.02
5.6
2.4
2.2
0
0.04
5.2
Table 3. The relative volumes of the various tissue types in the dry desiccation-tolerant E. nindensis leaves calculated from
transmission electron micrographs
The ratio of BSC to mesophyll and epidermal relative volume was 32.763.8:21.662.6 calculated from light micrographs. The results represent
the means 6standard deviation of measurements from more than 30 micrographs from four different sections from at least two different
specimens per sample. It was assumed that none of the vacuoles in the BSC were central vacuoles.
Central vacuole
Small vacuole
Cytoplasm
Contribution of each compartment to tissue type
Contribution of each compartment in each tissue to total leaf volume
Bundle sheath cells
Mesophyll and epidermis
Bundle sheath cells
Mesophyll and epidermis
0
9368
263
3965
762
4166
0
31
1
8
1
9
Downloaded from http://jxb.oxfordjournals.org at Main Library L610 Lawrence Livermore Nat Lab on 4 November 2009
Mechanical stabilization in desiccated tissues 659
rehydration before photosynthesis and autotrophism can
resume (Hambler, 1961). Although the accumulation of
metabolites has been reported for both poikilochlorophyllous and homoiochlorophyllous (retain chlorophyll and
chloroplast structure on drying) resurrection plants, the
formation of non-aqueous vacuoles in the poikilochlorophyllous species may act not only in membrane stabilization and water replacement, but as a store of energy for the
initial stages of rehydration. In E. nindensis, it is in the
BSC in which abundant small vacuoles form (Fig. 1;
Table 1). Since it is in the vascular bundles where the
majority of photosynthetic carbon reduction will occur, it
is possible that the accumulation of the metabolites
illustrated in Fig. 4, are necessary to reconstitute the
subcellular structure of these cells. It is likely that this may
be necessary for metabolism in the BSCs before those of
the mesophyll and epidermal tissues.
Not only are the BSCs the site of photosynthetic carbon
reduction, but they are also in closest proximity to the
vascular tissues. As a consequence, these cells will be the
®rst to experience the inrush of water on rehydration.
Membrane tearing on rehydration is thought to be a cause
of irreparable damage in many desiccation-sensitive
tissues (Pammenter and Berjak, 1999). The presence of a
TIP 3;1 on the membranes of the small vacuoles in the
dehydrated BSC cell of E. nindensis (Fig. 3) may be
important to ensure high membrane water permeability for
rehydration. The TIP 3;1 from P. vulgaris (the antibody
which cross-reacts with the protein from desiccated inner
leaves of E. nindensis) is an aquaporin since it is permeable
to water, the activity of which is regulated at the protein
level by turgor-dependent phosphorylation (Maurel et al.,
1995). Other TIP 3;1s expressed during the dehydration
stages of seed maturation have been proposed to be
involved in maintaining membrane permeability during
this phase (Maurel et al., 1995). Since the expression of
TIP 3;1 in seeds is developmentally regulated, appearing
coincident with the onset of desiccation tolerance, it has
been presumed to play a protective/stabilizing role in
those tissues (Johnson et al., 1990). However, since the
E. nindensis TIP 3;1 is found only in very dry tissues, its
is thought to act as an osmolyte during water de®cits and
has also recently been shown to be associated with the
formation of small vacuoles in Saccharomyces cerevisiae
(Maggio et al., 2002). Since the accumulation of proline
(Fig. 4A) was at a similar stage in drying to the appearance
of the small vacuoles (Table 1), it is possible that this
solute was incorporated into the vacuoles as they were
forming, and hence may not be acting as an osmolyte in
this circumstance. Although proline is generally associated
with the cytoplasm, it has been observed in the lytic
vacuoles of potato tubers (Farre et al., 2001). Of the
metabolites measured, sucrose was found to contribute the
highest mass in the small vacuoles of the BSC. However, it
was in the cytoplasm of the mesophyll and epidermal cells
where most of the sucrose was located, making up more
than all the total mass of all the other measured metabolites
in these tissues. Sugars are known to be important in the
formation of glasses and the stabilization of membranes.
While the importance of vitri®cation in conferring
desiccation tolerance has been questioned (Buitink et al.,
2002), it is still believed to contribute towards the
stabilization of tissue once they have become dry, and
hence to longevity in the desiccated state (Leopold et al.,
1994).
The protein storage vacuoles found in orthodox,
desiccation-tolerant seeds (Pernollet, 1978), are very
similar in appearance to the small vacuoles found in
desiccation-tolerant vegetative tissues (Fig. 1). Unlike the
seed protein vacuoles, it seems that the non-aqueous
vacuoles of E. nindensis are ®lled with a number of
different constituents. Although their composition does
differ from that of the cytoplasm and lytic vacuoles
(Table 4), no particular metabolites investigated in this
study were exclusively associated with them.
Although the formation of small vacuoles and the
maintenance of cell volume on dehydration is not exclusive to poikilochlorophyllous resurrection plants such as
E. nindensis, it is more commonly recorded in this group
of desiccation-tolerant plants (Farrant, 2000; Vander
Willigen et al., 2001a). These plants lose chlorophyll on
dehydration and hence this must be resynthesized on
Table 4. The relative distributions of the measured metabolites among the subcellular compartments of the bundle sheath cells
and mesophyll and epidermal tissues of desiccated leaves of E. nindensis
These data are calculated as g/100 g tissue and derived from Tables 2 and 3, as explained in the text.
Bundle sheath cells
Protein
Free proline
Glucose
Fructose
Sucrose
Mesophyll and epidermis
Cytoplasm
Small
vacuole
Central
vacuole
Cytoplasm
Small
vacuole
Central
vacuole
0.46
0.10
0.00
0.01
1.97
3.68
3.83
0.00
0.02
5.35
0
0
0
0
0
5.96
1.31
0.02
0.09
25.58
0.18
0.18
0.00
0.00
0.26
2.44
2.23
0.01
0.04
5.23
Downloaded from http://jxb.oxfordjournals.org at Main Library L610 Lawrence Livermore Nat Lab on 4 November 2009
660 Vander Willigen et al.
role in desiccation tolerance cannot yet be conclusively
deduced.
There have also been suggestions that this TIP isoform
may be permeable to glycerol and small proteins (Maurel
and Chrispeels, 2001). During germination, TIP 3;1s from
seeds are thought to be associated with the mobilization of
the contents of the protein storage vacuoles with which
they are associated (Johnson et al., 1990; Maurel et al.,
1997). The TIP 3;1 found in E. nindensis is associated with
the abundant small vacuoles (Fig. 3) and may thus be
important in mobilizing the proteins, sugars, proline, and
other solutes which accumulate therein (Table 4). Jauh
et al. (1999) proposed that some TIPs play a role in
determining the function of a vacuole; the TIP 3;1 seen on
protein storage vacuoles are gradually replaced by TIP 1;3s
during seed germination in P. vulgaris, rede®ning the
function and structure of those vacuoles. The proposed TIP
in E. nindensis (Figs 2, 3) might have a similar function.
The appearance of this possible TIP 3;1 in desiccated
inner leaves of E. nindensis is the only account of this TIP
isoform in vegetative tissues. To date, no expression of all
other TIP 3;1s known in the seeds of numerous plants have
been reported to be expressed subsequent to seed maturation (Maurel et al., 1997), although desiccated plant
tissues were not examined in those studies. However, this
antibody has been shown to cross-react with high speci®city to TIP 3;1s in seeds, but not vegetative tissue, from a
number of species. This TIP isoform may well be absent
from hydrated vegetative tissues, but it may be important
in desiccation-tolerant vegetative tissue.
Acknowledgements
The authors thank Dr JC Rogers for the antibodies and K McGrath
and K Cooper for technical assistance. Anglo American is acknowledged for allowing plant collections on their property. University
Research Council (UCT) and National Research Foundation (NRF)
research grants to JMF ®nanced the running costs and CVW was
kindly supported by scholarships from UCT, NRF, and the Claude
Harris Leon Foundation.
References
Barrieu F, Marty-Mazarazs D, Thomas D, Chaumont F,
Charbonnier M. 1999. Desiccation and osmotic stress increase
the abundance of mRNA of the tonoplast aquaporin BobTIP26-1
in cauli¯ower cells. Planta 209, 77±86.
Bianchi G, Murelli C, Bochicchio A, Vazzana C. 1991. Changes
in low-molecular weight substances in Boea hydroscopica in
response to desiccation and rehydration. Phytochemistry 30, 461±
466.
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.
Buitink J, Folkert A, Hoekstra FA, Leprince O. 2002.
Biochemistry and biophysics of tolerance systems. In: Black M,
Pritchard HW, eds. Desiccation and survival in plants. Drying
without dying. New York: CABI Publishing, 293±318.
Chomzynski P, Sacchi N. 1987. Single-step method of RNA
isolation by acid guanidinum thiocynate±phenol-chloroform
extraction. Analytical Biochemistry 162, 156.
Dalla Vecchia FD, Asmar TE, Calamassi R, Rascio N, Vazzana
C. 1998. Morphological and ultrastructural aspects of dehydration
and rehydration in leaves of Sporobolus sta®anus. Plant Growth
Regulation 24, 219±228.
Farrant JM. 2000. A comparison of mechanisms of desiccationtolerance among three angiosperm resurrection plant species.
Plant Ecology 151, 29±39.
Farre E, Tiessen A, Roessner U, Geigenberger P, Trethewey
RN, Willmitzer L. 2001. Analysis of the compartmentation of
glycolytic intermediates, nucleotides, sugars, organic acids,
amino acids, and sugar alcohols in potato tubers using a nonaqueous fractionation method. Plant Physiology 127, 685±700.
Gaff DF. 1971. Desiccation-tolerant ¯owering plants of Southern
Africa. Science 174, 1033±1034.
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.
Hallam ND, Luff SE. 1980. Fine structural changes in the
mesophyll tissue of the leaves of Xerophyta villosa during
desiccation. Botanical Gazette 141, 173±179.
Hambler DJ. 1961. A poikilohydrous, poikilochlorophyllous
angiosperm from Africa. Nature 191, 1415±1416.
Hartung W, Schiller P, Dietz K-J. 1998. Physiology of
poikilohydric plants. Progress in Botany 59, 299±327.
Heineke D, Lohous G, Winter H. 1997. Compartmentation of C/N
metabolism. In: Foyer CH, Quick WP, eds. A molecular approach
to primary metabolism in higher plants. London: Tailor & Francis
Ltd, 205±217.
Henzler T, Waterhouse RN, Smyth AJ, Carajal M, Cooke DT,
Schaffner R, Steudle E, Clarkson DT. 1999. Diurnal variations
in hydraulic conductivity and root pressure can be correlated with
the expression of putative aquaporins in the roots of Lotus
japonicus. Planta 210, 50±60.
Iljin WS. 1957. Drought resistance in plants and physiological
processes. Annual Review of Plant Physiology 8, 257±274.
Jauh G-Y, Fischer AM, Grimes HD, Ryan CA, Rogers JC. 1998.
d-Tonoplast intrinsic protein de®nes unique plant vacuole
functions. Proceedings of the National Academy of Sciences,
USA 95, 12995±12999.
Jauh G-Y, Phillips TE, Rogers JC. 1999. Tonoplast intrinsic
protein isoforms as markers for vacuolar functions. The Plant
Cell 11, 1867±1882.
Johannson I, Larsson C, Ek B, Kjellbom P. 1996. The major
intergral proteins of spinach leaf plasma membranes are putative
aquaporins and are phosphorylated in response to Ca2+ and
apoplastic water potential. The Plant Cell 8, 1181±1191.
Johannson I, Karlsson M, Shukla VK, Chrispeels MJ, Larsson
C, Kjellbom P. 1998. Water transport activity of the plasma
membrane aquaporin PM28A is regulated by phosphyorylation.
The Plant Cell 10, 451±459.
Johnson KD, Herman EM, Chrispeels MJ. 1989. An abundant,
highly conserved tonoplast protein in seeds. Plant Physiology 91,
1006±1013.
Johnson KD, Hofte H, Chrispeels MJ. 1990. An intrinsic
tonoplast protein of protein storage vacuoles in seeds is
structurally related to a bacterial solute transport (GlpF). The
Plant Cell 2, 525±532.
Kirsh H-H, Vera-Estrella R, Golldack D, Quigley F,
Michalowski CB, Bronwyn J, Barkla BJ, Bonhert HJ. 2000.
Downloaded from http://jxb.oxfordjournals.org at Main Library L610 Lawrence Livermore Nat Lab on 4 November 2009
Mechanical stabilization in desiccated tissues 661
Expression of water channel proteins in Mesembryanthemum
crystallinum. Plant Physiology 123, 111±124.
Kingston-Smith AH, Galtier N, Pollock CJ, Foyer CH. 1998.
Soluble acid invertase activity is dependent of species differences
in leaf carbohydrates, diurnal sugar pro®les and paths of phloem
loading in leaves. New Phytologist 139, 283±292.
Leopold AC, Sun WQ, Bernal-Lugo I. 1994. The glassy state in
seeds: analysis and function. Seed Science Research 4, 267±274.
Liu Q, Umeda M, Uchimiya H. 1994. Isolation and expression
analysis of two rice genes encoding the major intrinsic protein.
Plant Molecular Biology 26, 2003±2004.
Maggio A, Miyazaki S, Veronese P, Fujita T, Ibeas JI, Damsz B,
Narasimhan ML, Hasegawa PM, Joly RJ, Bressan RA. 2002.
Does proline accumulation play an active role in stress-induced
growth reduction? The Plant Journal 31, 699±712.
Magne C, Larher F. 1992. High sugar content of extracts interferes
with colimetric determination of amino acids and free proline.
Analytical Biochemistry 200, 115±118.
Mariaux J-B, Bockel C, Salamini F, Bartels D. 1998.
Desiccation- and abscisic acid-responsive genes encoding major
intrinsic proteins (MIPs) from the resurrection plant
Craterostigma plantagineum. Plant Molecular Biology 38,
1089±1099.
Maurel C, Chrispeels MJ. 2001. Aquaporins. A molecular entry
into plant water relations. Plant Physiology 125, 135±138.
Maurel C, Kado RT, Guern J, Chrispeels MJ. 1995.
Phosphorylation regulates the water channel activity of the
seed-speci®c aquaporin a-TIP. EMBO Journal 14, 3028±3035.
Maurel C, Tacnet F, Gucku J, Guern J, Ripoche P. 1997.
Puri®ed vesicles to tobacco cell vacuolar and plasma membranes
exhibit dramatically different water permeability and water
channel activity. Proceedings of the National Academy of
Sciences, USA 94, 7103±7108.
Meryman HT. 1974. Freezing injury and its prevention in living
cells. Annual Review of Biophysics and Bioengineering 3, 341±
363.
Mitra BN, Yoshino R, Morio T, Yokoyama M, Maeda M,
Urushihara H, Tanaka Y. 2001. Loss of a member of the
aquaporin gene family, aqp A affects spore dormancy in
Dictyostelium. Gene 251, 131±139.
Neale AD, Blomstedt CK, Bronson P, Le TN, Guthridge K,
Evans D, Gaff DF, Hamill JD. 2000. The isolation of genes from
the resurrection grass Sporobolus stap®anus which are induced
during severe drought stress. Plant, Cell and Environment 23,
265±277.
Norwood M, Truesdale MR, Richter A, Scott P. 2000.
Photosynthetic carbohydrate accumulation in the resurrection
plant Craterostigma plantagineum. Journal of Experimental
Botany 51, 159±165.
Pammenter NW, Berjak P. 1999. A review of recalcitrant seed
physiology in relation to desiccation-tolerance mechanisms. Seed
Science and Technology 9, 13±37.
Pernollet JC. 1978. Protein bodies of seeds: ultrastructure,
biochemistry, biosynthesis and degradation. Planta 17, 1473±
1480.
Platt KA, Oliver MJ, Thompson WW. 1997. Importance of the
®xative for reliable ultrastructural preservation of poikilohydric
plant tissues. Observation on dry, partially dry, and fully hydrated
tissues of Selaginella lepidophylla. Annals of Botany 80, 599±
610.
Riens B, Lohaus, G, Heineke D, Heldt HW. 1991. Amino acid
and sucrose content determined in the cytosolic, chloroplastic,
and vacuolar compartments and in the phloem sap of spinach
leaves. Plant Physiology 97, 227±233.
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.
Sarda X, Tousch D, Ferrare K, Legrand E, Dupuis JM,
Cassedelbart F, Lamaze T. 1997. Two TIP-like genes
encoding aquaporins are expressed in sun¯ower guard cells.
The Plant Journal 12, 1103±1111.
Sherwin HW, Farrant JM. 1996. Difference in rehydration of
three desiccation-tolerant angiosperm species. Annals of Botany
78, 703±710.
Spurr AR. 1969. A low viscosity epoxy resin embedding medium
for electron microscopy. Journal of Ultrastructural Research 26,
31±43.
Stitt M, Lilley RM, Gerhardt R, Heldt HW. 1989. Metabolite
levels in speci®c cells and subcellular compartments of plant
leaves. Methods in Enzymology 174, 518±550.
Tymms MJ, Gaff DF. 1979. Proline accumulation during water
stress in resurrection plants. Journal of Experimental Botany 30,
165±168.
Vander Willigen C, Farrant JM, Pammenter NW. 2001a.
Anomalous pressure volume curves of resurrection plants do
not suggest negative turgor. Annals of Botany 88, 537±543.
Vander Willigen C, Pammenter NW, Mundree SG, Farrant.
2001b. Some physiological comparisons between the resurrection
grass, Eragrostis nindensis, and the related desiccation-sensitive
species, E. curvula. Plant Growth Regulation 35, 121±129.
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.
Whittaker A, Bochicchio A, Vazzana C, Lindsey G, Farrant J.
2001. Changes in leaf hexokinase activity and metabolite levels in
response to drying in the desiccation-tolerant species Sporobolus
sta®anus and Xerophyta viscosa. Journal of Experimental Botany
52, 961±969.
Winter H, Robinson DG, Heldt HW. 1993. Subcellular volumes
and metabolite concentrations in barley leaves. Planta 191, 180±
190.
Yamada S, Katsuhara M, Kelly WB, Michalowski CB, Bonhert
HJ. 1995. A family of transcripts encoding water channel
proteins: tissue-speci®c expression in the common ice plant. The
Plant Cell 7, 1129±1142.
Yamada S, Komori T, Myers T, Kuwata S, Kubo T, Imaseki H.
1997. Expression of plasma membrane water channel genes under
water stress in Nicotiana excelsior. Plant Cell Physiology 38,
1226±1231.
Downloaded from http://jxb.oxfordjournals.org at Main Library L610 Lawrence Livermore Nat Lab on 4 November 2009