Plant, Cell and Environment (2008) 31, 1813–1824
doi: 10.1111/j.1365-3040.2008.01887.x
Chloroplast biogenesis during rehydration of the
resurrection plant Xerophyta humilis: parallels to the
etioplast–chloroplast transition
ROBERT A. INGLE1*, HELEN COLLETT1*, KEREN COOPER1, YUICHIRO TAKAHASHI2, JILL M. FARRANT1 &
NICOLA ILLING1
1
Department of Molecular and Cell Biology, University of Cape Town, Private Bag, Rondebosch 7701, South Africa and
The Graduate School of Natural Science and Technology, Okayama University, 3-1-1 Tsushima-naka, Okayama 700-8530,
Japan
2
ABSTRACT
De-etiolation of dark-grown seedlings is a commonly used
experimental system to study the mechanisms of chloroplast biogenesis, including the stacking of thylakoid membranes into grana, the response of the nuclear-chloroplast
transcriptome to light, and the ordered synthesis and
assembly of photosystem II (PSII). Here, we present the
xeroplast to chloroplast transition during rehydration of the
resurrection plant Xerophyta humilis as a novel system for
studying chloroplast biogenesis, and investigate the role of
light in this process. Xeroplasts are characterized by the
presence of numerous large and small membrane-bound
vesicles and the complete absence of thylakoid membranes.
While the initial assembly of stromal thylakoid membranes
occurs independently of light, the formation of grana is light
dependent. Recovery of photosynthetic activity is rapid in
plants rehydrated in the light and correlates with the lightdependent synthesis of the D1 protein, but does not require
de novo chlorophyll biosynthesis. Light-dependent synthesis of the chlorophyll-binding protein Lhcb2 and digalactosyldiacylglycerol synthase 1 correlated with the formation
of grana and with the increased PSII activity. Our results
suggest that the molecular mechanisms underlying photomorphogenic development may also function in desiccation
tolerance in poikilochlorophyllous resurrection plants.
Key-words: desiccation tolerance; photosynthesis; resurrection plant; xeroplast.
INTRODUCTION
The light reactions of photosynthesis couple the absorption
of light by chlorophyll to the generation of energy and
reducing power to drive the fixation of CO2 (Nelson &
Yocum 2006). Operation of the light reactions inevitably
leads to the formation of reactive oxygen species (ROS),
Correspondence: N. Illing. Fax: +27 21 689 7573; e-mail:
[email protected]
*These authors contributed equally to this work.
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd
initially the superoxide radical (O2•-) and singlet oxygen
(1O2) (Ivanov & Khorobrykh 2003; Møller, Jensen &
Hansson 2007), and chloroplasts contain several antioxidant systems to scavenge ROS (Apel & Hirt 2004; Foyer &
Noctor 2005). The equilibrium between ROS production
and scavenging can be perturbed by environmental stresses
leading to a rapid increase in ROS concentration (Apel &
Hirt 2004; Møller et al. 2007). Under water-deficit stress,
especially under high-light conditions, the excitation energy
harvested by chlorophyll can greatly exceed the demand of
the Calvin cycle for ATP and NADPH, leading to overreduction of the electron transport chain and enhanced generation of ROS (Smirnoff 1993; Apel & Hirt 2004; Møller
et al. 2007). While ROS play critical roles in cell signalling
(Kovtun et al. 2000; Foyer & Noctor 2005), they can also
cause extensive oxidative damage to macromolecules such
as lipids, proteins and nucleic acids (Møller et al. 2007).
Resurrection plants, which are able to tolerate the loss of
95% of protoplasmic water and recover full metabolic
activity in existing tissues upon rehydration, avoid a toxic
build-up of ROS by a controlled and reversible shutdown of
photosynthesis early on during the drying process (Sherwin
& Farrant 1998; Farrant 2000).
Angiosperm resurrection plants can be classified into two
groups based on the mechanisms they utilize to shut down
photosynthesis during desiccation. Homoiochlorophyllous
species, such as Craterostigma, retain their chlorophyll and
rely on pigment production and morphological changes,
such as leaf folding, to prevent light–chlorophyll interactions during desiccation (Sherwin & Farrant 1998; Farrant
2000). In contrast, poikilochlorophyllous resurrection
plants, such as Xerophyta, dismantle thylakoid membranes
and break down chlorophyll during drying (Tuba et al. 1996;
Sherwin & Farrant 1998; Farrant 2000).
Recent studies have indicated that down-regulation of
photosystem II (PSII) subunit expression also occurs in
poikilochlorophyllous species during desiccation (Collett
et al. 2004; Ingle et al. 2007). PSII is a large protein complex
located predominately in the granal thylakoid membranes
of the chloroplast, and contains approximately 25 protein
subunits encoded by the psb genes (Mullineaux 2005;
1813
1814 R. A. Ingle et al.
Nelson & Yocum 2006). Six psb genes were previously
identified as desiccation down-regulated in a small-scale
microarray analysis of Xerophyta humilis gene expression.
These included psbA, which encodes the D1 subunit of the
PSII core complex, and psbO and psbP, which encode components of the oxygen-evolving complex (OEC). A reduction in protein levels of several PSII subunits in Xerophyta
viscosa at 55% relative water content (RWC) correlated
with the cessation of photosynthetic activity in this species
(Ingle et al. 2007).
Upon rehydration, photochemical activity recovers
rapidly in Xerophyta species (Sherwin & Farrant 1996),
suggesting that they have evolved mechanisms to allow the
rapid biogenesis and assembly of both thylakoid membranes and the photosynthetic apparatus. The molecular
basis of this process and the signalling events involved
are unclear, although the role of water availability is obvious. Strikingly, similar processes occur in the etioplast–
chloroplast transition during photomorphogenesis when
light acts as the signal for chlorophyll biosynthesis, formation and stacking of thylakoid membranes, and translation
of several PSII mRNAs including psbA, psbB and psbC
(Klein & Mullet 1987; von Wettstein, Gough & Kannangara
1995; Baena-Gonzalez & Aro 2002). Here, we present the
reassembly of chloroplasts during rehydration of X. humilis
as a novel system to study chloroplast biogenesis, and demonstrate the role of light in several key events in this
process.
Determination of RWC
Absolute water content (AWC) of leaf samples was calculated using the formula (fresh biomass–dry biomass)/dry
biomass. RWC was calculated using the formula (AWC ¥
100)/AWC at full turgor (determined after bagging the
control plants overnight after watering). Ten leaf samples
were taken at each time point from each treatment group
for determination of RWC.
Determination of chlorophyll content
The leaf samples were cut into small pieces, and chlorophyll
was extracted in 100% acetone for 4 d at 4 °C. Total
chlorophyll (a + b) content (mg g DW-1) was determined
spectrophotometrically using the equation (7.05 ¥ A661.6) +
(18.09 ¥ A644.8) as described in Lichtenthaler (1987).
Measurement of PSII operating efficiency
The quantum yield of photosystem II (FPSII), the proportion of light absorbed by the PSII antennae used in photochemistry (Genty, Briantais & Baker 1989), was determined
by measurement of chlorophyll fluorescence using a PAM2100 portable chlorophyll fluorometer (Heinz-Walz GmbH,
Effeltrich, Germany). The leaf samples were light adapted
at a photosynthetic flux of ~50 mmol m-2 s-1 for 15 min prior
to measurement of FPSII.
CO2 measurements
MATERIALS AND METHODS
Plant material and culture
Xerophyta humilis plants were collected from Borakalalo
National Park (Limpopo Province, South Africa), potted
and grown under glasshouse conditions as described in
Sherwin & Farrant (1996). Prior to this study, the plants
were transferred to a controlled environment room with
a photosynthetic flux of ~200 mmol m-2 s-1 under a 16 h
light/8 h dark cycle at 25 °C. The plants were dried down
by withholding water for 2 weeks, and then kept in a
desiccated state for a further 2 weeks prior to rehydration. Hydrated (control) plants were regularly watered
throughout. Light-excluding boxes were placed over the
plants the previous evening for the rehydration in the
dark experiments.
Rehydration time course
Desiccated plants were rehydrated under a normal 16 h
light/8 h dark cycle or in continuous darkness, beginning 1 h
prior to ‘dawn’. Each rehydration experiment spanned a
36 h time course as previous studies had suggested a substantial recovery of PSII activity in X. humilis within this
time frame (Sherwin & Farrant 1996). Tissue samples were
collected immediately prior to rewatering and at 3, 6, 9, 12,
15, 18, 24 and 36 h post-watering and from the hydrated
(control) plants at the same time points.
The rate of net CO2 assimilation or release was determined
using an LI-6400 portable photosynthesis system (Li-Cor
Biosciences, Lincoln, NE, USA), operated at an ambient
CO2 concentration of 350 ppm. The parameters A and
Rd were calculated using the equations described by von
Caemmerer & Farquhar (1981).
Chloroplast ultrastructural studies
Chloroplast ultrastucture was examined using transmission
electron microscopy as previously described in Cooper &
Farrant (2002). Briefly, small pieces of leaf tissue (approximately 2 mm2) were excised from the middle of four different leaves, and RWC was determined for each leaf. Fixation
was carried out in 2.5% glutaraldehyde in 0.1 m phosphate
buffer (pH 7.4) containing 0.5% caffeine, and samples were
postfixed in 1% osmium in phosphate buffer.After dehydration in a graded ethanol series, the tissue was infiltrated with
epoxy resin over 4 d. The 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. Sections
were stained with 2% uranyl aceate and 1% lead citrate, and
were viewed with a transmission electron microscope. The
dimensions of chloroplasts, vesicles, plastoglobuli, starch
bodies, thylakoid membranes and grana were measured
with Image-Pro 6.2 (Mediacybernetics, Bethesda, MD,
USA). Measurements were made on three images per RWC,
from at least two different leaves, for both light regimes.
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Plant, Cell and Environment, 31, 1813–1824
Chloroplast biogenesis during rehydration of X. humilis 1815
Isolation of partial psbD, psbS and dgd1
cDNAs from X. humilis
First-strand cDNA was synthesized from 2 mg of total RNA
using a SuperScript™ II Reverse Transcriptase system
(Invitrogen, Carlsbad, CA, USA). One microlitre of the
resulting cDNA was used as the template in a standard
30 mL PCR reaction. PCR primers were designed to conserved regions with low amino-acid codon degeneracy
based on known homologs in the National Center for Biotechnology Information (NCBI) databases. The X. humilis
psbD (~1 kb) and psbS (~0.5 kb) partial cDNAs were isolated using the following primers psbD: 5′-GACTGG
TTACGRAGGGACCG-3′ and 5′-GGTAGAACCTCCTC
CTCATCAGGGA-3′ (annealing temperature, 58 °C), and
psbS: 5′-GTNGGYCGYGTTGCYATG-3′ and 5′-ATNG
CRGCRANGAAGAAGAA-3′ (annealing temperature,
62 °C). A 103 bp sequence from the 3′ end of dgd1 was
previously isolated in a differential display PCR screen
(Collett, unpublished data). An additional 1.4 kb of
5′sequence was isolated by RT-PCR using the degenerate
primer 5′-ACAACAGCNAGTCTTCCNTGGATG-3′ in
combination with the 3′ gene-specific primer 5′-GAAA
TTGACATTTGTACCTGGC-3′. The resulting PCR
fragments were cloned into the pGEM-T-Easy vector
(Promega, Madison, WI, USA) and were sequenced. Nucleotide sequences were deposited in GenBank (see further
discussion for accession numbers).
Northern blot analysis
Total RNA was isolated from the leaf tissue using
TriReagent (Molecular Research Centre, Inc., Cincinnati,
OH, USA). For northern blot analysis, 20 mg of total RNA
was transferred onto nitrocellulose membrane following
formaldehyde gel electrophoresis. Blots were prehybridized
in buffer containing 50 mm sodium phosphate buffer (pH
6.8), 5 ¥ SSC, 5 ¥ Denhardt’s solution, 50% formamide,
0.1% (w/v) sodium dodecyl sulphate (SDS), 0.1% (w/v)
sodium pyrophosphate and 50 mg mL-1 salmon sperm
DNA, and were then probed overnight at 42 °C with 32Plabelled cDNA probes prepared using the Megaprime kit
(Amersham Pharmacia Biotech, Piscataway, NJ, USA). The
blots were washed for 2 ¥ 10 min in 1 ¥ SSC, 0.1% (w/v)
SDS at RT, followed by 2 ¥ 10 min in 0.5 ¥ SSC, 0.1% (w/v)
SDS at 55 °C, and were exposed to autoradiography film.
The following X. humilis cDNA clones were used in northern blot analysis: psbA (AF545583), psbD (DQ067928),
psbO (DV767869), psbP (AF545584), psbR (AY146990),
psbS (DQ067929), psbT (DV850415), psbY (DV768147)
and DGD1 (AY186241).
Western blot analysis
Total protein was isolated from the leaf tissue as previously
described (Ingle, Smith & Sweetlove 2005). Thirty micrograms of total protein was separated on 12% sodium dodecyl
sulphate–polyacrylamide gel electrophoresis (SDS–PAGE)
gels and was transferred to nitrocellulose membrane. Membranes were blocked for 2 h in 1 ¥ TBST containing 10% w/v
non-fat milk powder. Primary antibodies were diluted in
1 ¥ TBST [digalactosyldiacylglycerol synthase 1 (DGD1)
1:1000, Lhc2b 1:2000 and D1 1:4000] containing 10% w/v
non-fat milk powder. Blots were incubated with primary
antibody for 2 h, followed by 3 ¥ 5 min washes in 1 ¥ TBST
and incubation with secondary antibody (rabbit IgG HRP,
1:5000 dilution) for 1.5 h. Bands were detected using chemiluminescence as described by Durrant & Fowler (1994).
RESULTS
Chloroplast biogenesis during rehydration of
X. humilis
Chloroplast ultrastructure undergoes major modifications
during rehydration in Xerophyta species. We defined six
stages of chloroplast biogenesis during rehydration on the
basis of detailed measurements made of chloroplasts,
membrane-bound vesicles, plastoglobules, starch granules,
assembling thylakoid membranes, single thylakoids and
grana (summarized in Table 1 and Fig. 1). The membranebound vesicles could be divided into two populations. The
larger vesicles (diameter range, 60–200 nm) were consistently less stained by osmium than the smaller vesicles
(diameter, 20–60 nm). These membrane-bound vesicles differed from the oval osmophilic plastoglobules (diameter,
25–85 nm), which were present at all stages of chloroplast
biogenesis (Fig. 1). We have coined the term ‘xeroplast’ to
describe the stage 1 chloroplasts that were present in dry
leaves. Xeroplasts are characterized by the presence of
numerous large and small membrane-bound vesicles, with a
length-to-width ratio (L/W) between 1.0 and 2.0, and a
chloroplast L/W ratio between 1.0 and 1.6. The smaller
membrane-bound vesicles were often clustered together in
string-like arrays in xeroplasts (Fig. 1).
Stage 2 of chloroplast biogenesis was characterized by a
change in the size distribution of the membrane-bound
vesicles. The more osmophobic larger membrane-bound
vesicles maintained an L/W ratio between 1.0 and 2.0, while
the smaller membrane-bound vesicles started to elongate to
an L/W ratio between 2.0 and 7.0.These smaller membranebound vesicles were frequently found in a head-to-toe
arrangement (Fig. 1). Chloroplasts started to lengthen at
stage 3 (L/W ratio of >2), and were characterized by the first
appearance of a single starch body, the presence of scattered larger membrane-bound vesicles (L/W ratio between
1.0 and 2.0) and smaller membrane-bound vesicles (L/W
ratio between 2.0 and 7.0), which were frequently linked
together (Fig. 1). The larger membrane-bound vesicles and
the plastoglobules were usually found in association with
these strings, which we suggest are the emerging prothylakoid membranes. The larger membrane-bound vesicles
had disappeared by stage 4, and the small, elongated
membrane-bound vesicles were assembled into single thylakoid membranes. Starch bodies were now more numerous
and larger (Fig. 1). Grana were first visible in stage 5 and
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Plant, Cell and Environment, 31, 1813–1824
Present
2.0–7.0
2.0–7.0
Absent
Absent
Present
2.0–7.0
We used the quantum yield of PSII (FPSII) as a measure
of PSII photochemical activity (Genty et al. 1989; Baker &
Rosenquist 2004), and net CO2 assimilation to monitor the
recovery of photosynthesis during the rehydration process.
The recovery of FPSII was rapid under the 16 h L/8 h D
cycle, beginning within 12 h of rewatering, and reaching
90% of the control (fully hydrated) plant values at 15 h
(Fig. 2a). In parallel with the recovery in FPSII, net CO2
assimilation was first detected at 12 h post-watering
(Fig. 2b). As expected, no CO2 assimilation was observed in
the plants rehydrated in constant darkness; however, a
modest recovery in FPSII (~25%) was observed in these
plants (Fig. 2a).
Few or none
Absent
Absent
1.0–1.6
>2.0
>2.0
>2.0
>2.0
2
3
4
5
6
L/W, length-to-width ratios.
1.0–2.0
1.0–1.6
1
2.0–7.0
1.0–2.0
Chloroplast
L/W
2.0–7.0
Absent
1.0–2.0
Many
1.0–2.0
Few
1.0–2.0
Few or none
Few or none
Photosynthetic activity resumes rapidly in the
presence of light
De novo chlorophyll biosynthesis is not
required for resumption of photosynthesis
Stage
1.0–2.0
were assembled from three to four thylakoid membranes
(Fig. 1). Further stacking of thylakoids occurred and by
stage 6, and grana were more frequent and far thicker, with
more than seven thylakoids stacked together (Fig. 1).
The populations of chloroplasts for each RWC point
during rehydration were categorized according to this
staging system to see whether there was a difference in
chloroplast biogenesis in leaves rehydrated in the dark
compared with the light. The rate of change in RWC values
during rehydration was not affected by the light regime
(Supporting Information Fig. S1). Chloroplast biogenesis
following rehydration in X. humilis was rapid, and there was
no significant difference in the rate of biogenesis from stage
1 through stage 4 for plants rehydrated in the dark compared with the light (Table 2). The majority of chloroplasts
had changed from stage 1 to stage 2 within 6 h of rehydration, even though the RWC on average was still less than
40%. The leaves of the plants were fully hydrated by 12 h,
and contained a mixed population of chloroplasts at different stages of biogenesis. After 12 h, the remaining chloroplasts in the leaves rehydrated in the dark progressed to
stage 4, but no further (Supporting Information Fig. S2). In
contrast, under the 16 h L/8 h D cycle, a few chloroplasts
with clear grana (stage 5) were first detectable at 12 h following rehydration.The proportions of chloroplasts in stage
5 and stage 6 steadily increased with time following rehydration in the light.
1.2–1.4
1.2–1.4
Present (layer of <4
thylakoids)
Mature (layer of >4
thylakoids)
Present
1.2–1.4
Absent
Present, but as strings of
small vesicles
Present
Present
Present, few and small
Absent
Absent
1.2–1.4
Absent
Absent
Absent
1.2–1.4
Absent
Absent
Absent
1.2–1.4
Grana
Thylakoids
(width < 10 nm)
Membrane-bound
vesicle (width,
10–40 nm)
L/W
Membrane-bound
vesicles (width,
20–60 nm)
L/W
Unstained membranebound vesicles (width,
60–200 nm)
L/W
Table 1. Ultrastructural features that characterize the stages 1 to 6 of chloroplast biogenesis during rehydration in Xerophyta humilis
Dark plastoglobules
(diameter range from
25 to 85 nm)
L/W
Starch grains
1816 R. A. Ingle et al.
Xerophyta species break down the majority of their chlorophyll during desiccation and resynthesize it upon rewatering (Sherwin & Farrant 1996; Farrant et al. 2003; Supporting
Information Fig. S3a). In etioplasts, synthesis of chlorophyllide from the precursor protochlorophyllide occurs only in
response to light (von Wettstein et al. 1995). Similarly, no
increase in chlorophyll content was detected in X. humilis
plants rehydrated in the dark (Fig. 2c, Supporting Information Fig. S3b). In contrast, under the 16 h L/8 h D cycle,
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Plant, Cell and Environment, 31, 1813–1824
Chloroplast biogenesis during rehydration of X. humilis 1817
(a)
(b)
(c)
(d)
(e)
(f)
(g)
(h)
(i)
(j)
Figure 1. Transmission electron micrographs illustrating
ultrastructural features characterizing different stages of
chloroplast biogenesis during rehydration in Xerophyta
humilis. Stages 1 to 4 of chloroplasts rehydrated in the dark
(a,c,e,g) are compared with stages 1 to 4 of chloroplasts
rehydrated in the light (b,d,f,h). Stages 5 and 6 were only
observed in leaves rehydrated in the light (i,j). Insets are
magnified to illustrate assembly of thylakoid membranes.
Scale bars = 1 mm. l, large vesicle; o, oval vesicle; e, small
elongated vesicle; p, plastoglobule; s, starch grain; t, thylakoid;
g, grana.
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Plant, Cell and Environment, 31, 1813–1824
1818 R. A. Ingle et al.
Dark
Stage 1
Stage 2
Stage 3
Stage 4
Stage 5
Stage 6
16 h light/8 h dark
Stage 1
Stage 2
Stage 3
Stage 4
Stage 5
Stage 6
0h
3h
100
67
33
100
92
8
6h
9h
12 h
26 h
30 h
34 h
51 h
75
25
100
25
67
8
25
75
100
100
100
67
33
8
92
100
25
75
33
17
33
17
Table 2. Percentage of chloroplasts in
different stages of biogenesis following
rehydration in the dark or under a 16 h
light/8 h dark cycle
8
58
33
58
33
Chloroplasts were staged in 12 independent images, from two different leaves, per time
point.
See Table 1 for classification of stage.
chlorophyll content increased to approximately 40% of
that in the control plants by 36 h post-watering (Fig. 2c).
However, net CO2 assimilation (and an almost total recovery of FPSII) was detected just after 12 h and prior to any
increase in chlorophyll content (Fig. 1a–c). This suggests
that the residual chlorophyll present in desiccated tissue
(<1 mg g DW-1) is sufficient for the resumption of photosynthetic activity during rehydration.
Water availability is the primary signal for psb
gene expression during rehydration
Down-regulation of psb gene expression occurs at both the
mRNA and protein level during desiccation in Xerophyta
species (Collett et al. 2004; Ingle et al. 2007). We thus analysed psb transcript levels during rehydration to determine
whether light is required for transcriptional activation of
these genes. In addition to the six psb genes we previously
identified as desiccation down-regulated (Collett et al.
2004), we isolated partial cDNA clones for psbD and psbS
from X. humilis. psbD encodes the D2 subunit required for
the assembly of the PSII core complex (Baena-Gonzalez &
Aro 2002), while PsbS is involved in non-photochemical
quenching (Li et al. 2000) and thus might be important
during rehydration to limit damage from excess excitation
energy. The mRNA abundance of these eight psb genes was
followed during the rehydration process by northern blot
analysis (Fig. 3). Under a 16 h L/8 h D cycle, mRNA levels
of seven of these eight genes had increased by 6 h postwatering and peaked after 9 h, that is, prior to net CO2
assimilation, while psbA mRNA levels increased only at
12 h post-watering and peaked at 24 h. A similar pattern of
gene expression was observed for psbO, psbP, psbR, psbTn
and psbY in the plants rehydrated under constant darkness.
Transcript levels increased to similar maximum levels albeit
more slowly, peaking at 24 h after rewatering. In contrast,
the increase in psbS expression was reduced, and no
increase in psbA or psbD mRNA levels was detected in the
plants rehydrated in the dark (Fig. 3) during the course of
the experiment. However, both psbA and psbD mRNA
were detectable in desiccated tissue, suggesting that they
can be stably stored (Fig. 3).
Light is required for translation of the D1
reaction centre protein
While psbA mRNA is present in the etioplasts of darkgrown plants, synthesis of the D1 protein is light dependent
(Müller & Eichacker 1999; Zhang & Aro 2002). Given the
apparent parallels between the etioplast–chloroplast transition and chloroplast biogenesis during rehydration, we
examined D1 protein levels by Western blotting. While
psbA mRNA was present at all time points including in
desiccated tissue (Fig. 3), the D1 protein was only detectable from 12 h post-watering in plants rehydrated in the
presence of light (Fig. 4) and correlated with the resumption of photosynthetic activity. The modest recovery in
FPSII (~25%) observed for plants rehydrated under constant darkness (Fig. 2a) is presumably because of basal
levels of the D1 protein, which are not detectable by
Western blot analysis.
Light is required for the synthesis of two other
chloroplast proteins implicated in PSII activity
We also investigated whether the synthesis of two proteins,
Lhcb2 and digalactosyldiacylglycerol synthase 1 (DGD1),
believed to play important roles in the stability and activity
of PSII and in the stacking of thylakoid membranes was
light dependent. In addition to their role in light harvesting,
the LHCII proteins are thought to play a role in the formation of granal stacks through protein–protein interactions
between LHCII and PSII complexes in adjacent thylakoid
membranes (Mullineaux 2005). Western analysis of one of
the LHCII proteins (Lhcb2) revealed that its synthesis was
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Plant, Cell and Environment, 31, 1813–1824
Chloroplast biogenesis during rehydration of X. humilis 1819
(a)
16 h L/8 h D
0.9
0.8
0
F PSII
0.7
6
9 12 24 36
Dark
6
9
12 24 36 h
0.6
psbA
0.5
0.4
psbD
0.3
0.2
psbO
0.1
01
0
psbP
0
6
12
18
24
30
36
psbR
h post-watering
mmol CO2 m2 s–1
(b)
6
p
psbS
4
psbTn
2
psbY
0
–2
18S rRNA
–4
Figure 3. Northern analysis of psb gene expression in
–6
0
6
12
18
24
30
36
h post-watering
(c)
mg chlorophyll g DW–1
9
Xerophyta humilis plants rehydrated under a 16 h light/8 h dark
cycle or in constant darkness. Twenty micrograms of total RNA
was probed with 32P-labelled partial cDNA probes for eight psb
genes from X. humilis. 18S rRNA signal intensity indicates equal
loading of the RNA samples.
8
7
6
5
4
3
2
1
0
0
6
12
18
24
30
36
h post-watering
Figure 2. Recovery of photosystem II (PSII) quantum yield (a),
CO2 assimilation (b) and chlorophyll content (c) in Xerophyta
humilis plants rehydrated under a 16 h light/8 h dark cycle (䊐) or
in constant darkness (䉱). Relative water content, PSII quantum
yield and chlorophyll content are also shown for control plants
not subjected to dehydration. The horizontal bar indicates the
16 h light (white)/8 h dark (black) cycle operating in the growth
chamber. 䉬, Represents the control hydrated X. humilis grown
under the same light/dark cycle conditions. Values indicated are
means ⫾ SD (n = 10), and the results shown are for one
experiment representative of three.
respectively (Dörmann & Benning 2003). In addition to
its role in contributing to the formation of the protonimpermeable bilayer, a small fraction of the DGDG pool
is thought to play a critical role in stabilizing PSII and in
stabilizing the formation of the LHCII trimers in the lightharvesting antenna (Dörmann et al. 1995; Steffen et al.
2005). DGD1 is the major enzyme catalyzing the conversion of MGDG into DGDG, and a 103 bp fragment of a
DGD1 homolog was previously identified as dehydration
up-regulated in X. humilis in a differential display PCR
screen (Collett, unpublished data). In the present study,
a 1.4 kb partial cDNA of this gene was obtained using
RT-PCR, and the predicted amino acid sequence shows 80
Dark
16 h L/8hD
0
6
9
12 24 36
6
9
12 24 36 h
D1 (PsbA)
Ponceau S
dependent on the presence of light (Fig. 5) and correlated
with the formation of granal stacks in plants rehydrated
under the 16 h L/8 h D cycle.
Two
galactolipids,
monogalactosyldiacylglycerol
(MGDG) and digalactosyldiacylglycerol (DGDG), constitute up to 50 and 20% of lipids in the thylakoid membranes,
Figure 4. Western analysis of D1 protein levels during
rehydration of Xerophyta humilis. Thirty micrograms of total
protein was probed with a D1 antibody. Equal loading of the gel
was verified by Ponceau S staining of the membrane after protein
transfer. The results shown are from one experiment
representative of three.
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Plant, Cell and Environment, 31, 1813–1824
1820 R. A. Ingle et al.
16 h L/8 h D
0 12
24
Dark
36
12 24
36 h
Lhcb2
Figure 5. Western analysis of Lhcb2 protein levels during
rehydration of Xerophyta humilis. Thirty micrograms of total
protein was probed with a polyclonal antibody to Lhcb2. Equal
loading of the gel was verified by Ponceau S staining of the
membrane after protein transfer. The results shown are from one
experiment representative of three.
and 78% identity to the rice and Arabidopsis homologs,
respectively (data not shown). Northern analysis confirmed
that DGD1 is up-regulated in X. humilis during desiccation,
with transcript levels detectable only at RWC below 30%
(Fig. 6a). DGD1 mRNA was stably stored in desiccated
tissue, and transcript levels decreased within the first 36 h of
rewatering irrespective of the presence of light (Fig. 6b).
However, Western blot analysis revealed a marked difference in DGD1 protein levels between the two treatment
groups. Despite the presence of the DGD1 transcript, no
DGD1 protein was detected in plants rehydrated in the dark.
In contrast, DGD1 protein was detected at 6 h post-watering
in plants rehydrated under the 16 h light/8 h dark cycle, with
levels peaking at 9 h. Levels of this protein declined rapidly,
and by 12 h post-watering, no DGD1 protein was detectable
by immunoblotting.
DISCUSSION
Two experimental systems have been previously used to
describe chloroplast development in higher plants, namely
the development of proplastids during seed germination
and the maturation of etioplasts in plants initially grown in
the dark. Here, we present the xeroplast to chloroplast transition in X. humilis during rehydration as a novel system
(a)
100
63
48
27
to study chloroplast biogenesis. This species carries out a
controlled and reversible shutdown of photosynthesis early
on during dehydration. However, upon rehydration, rapid
reassembly of the thylakoid membranes and photosynthetic
apparatus occurs. As is the case in both proplastid development and etioplast maturation, light plays an important
signalling role in this process.
The plastids in desiccated X. humilis leaves, which we
have termed ‘xeroplasts’, differ substantially from both
proplastids and etioplasts. Proplastids are small spherical
organelles (0.2–1 mm in diameter), which originate maternally, and are maintained in an undifferentiated state in the
developing embryo (Mullet 1998; Vothknecht & Westhoff
2001). Etioplasts differentiate from proplastids in seedlings
grown in the dark, and are characterized by the presence
of well-developed paracrystalline prolamellar bodies with
single thylakoids extending into the stroma (Robertson &
Laetsch 1974). Xeroplasts in X. humilis are larger (0.7–2 mm
in diameter) than proplastids, but lack the paracrystalline
prolamellar bodies of etioplasts, which form a reserve of
membrane material that is rapidly rearranged into thylakoid membranes upon exposure to light (Robertson &
Laetsch 1974).
The ultrastructural changes that accompany the development of proplastids into chloroplasts include changes in
chloroplast shape, starch formation, lamellar extension and
granal development, and have been used to define clear
stages of development (Whatley 1974). Stages of the basic
pathway include (1) a proplastid stage; (2) an amyloplast
stage in which starch granules appear; (3) an amoeboid stage
in which plastids become elongated and folded; (4) a stage of
plastid elongation where there is development of perforated
stroma lamellae and later incipient grana; and (5) a maturation stage when the aligned lamellae become continuous,
and grana increase in number and depth of stacking
(Whatley 1977). A final stage, gerontoplast, can be defined
which occurs when leaves senesce (Vothknecht & Westhoff
2001).These senescent chloroplasts are characterized by the
presence of large, unstained membrane-bound vesicles, and
osmophilic globules occupy much of the interior (Whatley
1974). Xeroplasts resemble these senescent chloroplasts in
6 % RWC
dgd1 mRNA
18S rRNA
16 h L/8 h D
(b)
0
6
9
12
24
Dark
36
6
9
12
24
36 h
dgd1 mRNA
DGD1 protein
Figure 6. Digalactosyldiacylglycerol
synthase 1 (DGD1) expression in
Xerophyta humilis during dehydration
(a) and rehydration (b). For Northern
analysis, 20 mg of total RNA was probed
with a 32P-labelled partial dgd1 cDNA
probe. Western analysis was carried out
on 30 mg of total protein with a DGD1
polyclonal antibody.
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Plant, Cell and Environment, 31, 1813–1824
Chloroplast biogenesis during rehydration of X. humilis 1821
that they too are filled with unstained membrane-bound
vesicles and osmophilic globules (Fig. 1). However, unlike
gerontoplasts, xeroplasts are not senescent, and are able to
differentiate back into chloroplasts upon rehydration.
Although similar ultrastructural changes are seen during
xeroplast development, they are more rapid and do not
occur in the same order as the proplastid-chloroplast developmental sequence. We identified six distinct stages in the
xeroplast–chloroplast transition (Fig. 1, Table 1). The first
change in ultrastructure organization of xeroplasts was
observed within 3 h of rehydration when many of the smaller
unstained membrane-bound vesicles start to elongate to
form the precursors to thylakoid membranes (stage 2). Stage
3 is characterized by the elongation of chloroplasts and by
the first appearance of starch bodies while thylakoid precursor membranes are still being formed (Fig. 1).There is a clear
correlation between the sequential disappearance of the
unstained membrane-bound vesicles and the appearance of
the prothylakoid membranes, suggesting that in contrast to
proplastids (Muhlethaler & Frey-Wyssling 1959;Vothknecht
& Westhoff 2001), the thylakoid membranes are not derived
from the inner chloroplast membrane. No membrane-bound
vesicles remain by stage 4, and immature, perforated stromal
thylakoids are clearly visible. Progression from stage 1 to
stage 4 occurs at the same rate in X. humilis rehydrated
under a 16 h L/8 h D cycle or in the dark (Fig. 1, Table 2).
However, while thylakoid reassembly in X. humilis is
independent of light, the formation of grana is light dependent with only single appressed thylakoids observed in
plants rehydrated in the dark, that is, chloroplast development arrests at stage 4 (Fig. 1, Supporting Information
Fig. S2). During photomorphogenesis, light acts as a signal
for the formation of thylakoids and granal stacks in etioplasts (Lopez-Juez & Pyke 2005).This process is mediated at
least in part by phytochrome, as Arabidopsis mutants lacking
the chromophore phytochromibilin display reduced granal
formation during de-etiolation (Chory et al. 1989). Interestingly, chloroplasts from X. humilis plants rehydrated in the
dark (Fig. 1) resembled those observed in several darkgrown Arabidopsis constitutive photomorphogenic mutants;
the development of chloroplasts in dark-grown cop mutants
is similarly stalled at stage 4 with no more than two layers of
thylakoid structures being observed (Deng, Caspar & Quail
1991; Kwok et al. 1996). Thus, the light requirement for
granal formation is conserved between etioplast-chloroplast
and xeroplast-chloroplast development.
Grana are not present in bacteria or algae, and are therefore not essential for oxygenic photosynthesis. It has been
suggested that their evolution allowed the formation of
larger LHCII complexes in higher plants without restricting
quinone diffusion (Mullineaux 2005). Accordingly, PSII
complexes in unstacked thylakoids have been shown to
have smaller LHC than those in granal stacks (Armond &
Arntzen 1977). The LHCII proteins themselves have been
suggested to play a role in the formation of grana via interactions with PSII complexes and other LHCII proteins in
adjacent thylakoid membranes (Mullineaux 2005; Standfuss
et al. 2005). In X. humilis, light-dependent synthesis of
Lhcb2 was found to correlate with the formation of grana in
plants rehydrated under the l6 h L/8 h D cycle (Figs 1 & 5).
Photosynthetic activity resumed rapidly in plants rehydrated under the 16 h L/8 h D cycle, with net CO2 assimilation recorded by 12 h post-watering, correlating with a
~65% recovery in FPSII (Fig. 2). Interestingly, while net
CO2 assimilation did not occur in plants rehydrated in the
dark, a modest recovery in FPSII was observed. The relationship between FPSII and the rate of linear electron
flow through PSII can be complicated under environmental
stress, and the proportion of active PSII centres cannot be
determined by measurement of FPSII (Maxwell & Johnson
2000). Nonetheless, the partial recovery of FPSII suggests
that some degree of assembly of the PSII core complex may
occur even in the absence of light. Further study is required
to determine whether this is in fact the case.
Light is required for chlorophyll biosynthesis during
proplastid development, etioplast–chloroplast transition
(Baena-Gonzalez & Aro 2002) and also xeroplast–
chloroplast transition (Fig. 2c). However, we found that
the resumption of photosynthesis occurred prior to any
increase in chlorophyll content (Fig. 2). While the chlorophyll content of X. humilis leaves declines dramatically
during dehydration, a residual amount (approximately 10%
of that present in fully hydrated plants) is present in desiccated leaf tissue, and is apparently sufficient for the initial
recovery of photosynthetic activity. Free chlorophyll is a
potent generator of ROS (Hutin et al. 2003), and the
residual chlorophyll present in desiccated tissue may be
bound in a protein–chlorophyll complex to prevent photooxidative damage. The primary chlorophyll-binding proteins in plants are the LHC proteins, but these are not
detectable in desiccated tissue of Xerophyta species (Fig. 5,
Ingle et al. 2007). An alternative candidate might be the
early light-inducible proteins (ELIPS). These proteins can
bind chlorophyll and are transiently expressed during
de-etiolation or under high-light conditions (Grimm, Kruse
& Kloppstech 1989; Adamska et al. 1999). Interestingly, an
ELIP-like protein (dsp 22) has been identified as desiccation up-regulated in Craterostigma plantagineum and has
been shown to associate with PSII protein–pigment complexes (Alamillo & Bartels 2001).
We also investigated whether light was required for the
transcription of eight psb genes encoding subunits of PSII,
because the down-regulation of psb gene expression occurs
at both the mRNA and protein level during dehydration
(Collett et al. 2004; Ingle et al. 2007). With the exception of
psbA and psbD, water was the primary signal for mRNA
accumulation, although the rate of synthesis was delayed in
the absence of light (Fig. 3). In contrast, while both psbA
and psbD mRNA are apparently stably stored in desiccated
tissue, no increase in mRNA levels was observed in the
absence of light (Fig. 3). Interestingly, there was little difference between the transcriptional activation of nuclear
(psbO, P, R, S, Tn and Y) or plastid (psbA and D) encoded
psb genes, suggesting that despite the apparently disorganized state of the xeroplast in desiccated tissue (Fig. 1), it
remains transcriptionally competent.
© 2008 The Authors
Journal compilation © 2008 Blackwell Publishing Ltd, Plant, Cell and Environment, 31, 1813–1824
1822 R. A. Ingle et al.
While psbA mRNA was constitutively present,
synthesis of the D1 protein was light dependent and was
first detectable 12 h after rehydration under a 16 h L/8 h D
cycle (Fig. 4), correlating with the resumption of net CO2
assimilation. PSII complex assembly during the etioplast–
chloroplast transition has been well characterized (BaenaGonzalez & Aro 2002; Blomqvist, Ryberg & Sundqvist
2006). Many of the PSII protein subunits accumulate in the
dark including D2, cytochrome b559 and components of the
OEC (Müller & Eichacker 1999; Baena-Gonzalez & Aro
2002). However, accumulation of the D1 polypeptide occurs
only after illumination and involves both light-dependent
translation initiation and stabilization of the D1 protein by
binding of chlorophyll a (Müller & Eichacker 1999; Zhang
& Aro 2002). In addition, there is evidence to suggest that a
precomplex containing D2 and cytochrome b559 is required
to act as an acceptor for the elongating D1 polypeptide
ensuring co-translational incorporation of the protein into
PSII (Müller & Eichacker 1999). While it is unclear whether
the incorporation of D1 into PSII during rehydration in X.
humilis requires the presence of a D2/cytochrome b559
acceptor, the light dependence of D1 protein synthesis suggests that the PSII reassembly may occur by the same route
as that of the etioplast–chloroplast transition.
Light was also found to play a role in the posttranscriptional regulation of DGD1, which catalyzes the
synthesis of the galactolipid DGDG. DGD1 protein was
transiently present at 6 and 9 h post-watering in plants rehydrated under the 16 h L/8 h D cycle, but no DGD1 protein
was detected in plants rehydrated in the dark (Fig. 6b).
While light-responsive DGD1 expression has not been previously reported, a transient peak in mRNA and protein
levels of MGDG synthase has been previously reported
in cucumber during de-etiolation (Yamaryo et al. 2003).
MGDG is the major galactolipid of the thylakoid membrane and also a substrate for the DGD1 enzyme.
During thylakoid biosynthesis in proplastids, MGDG
and DGDG are synthesized on the inner and outer chloroplast membranes, and are transported to the developing
thylakoids by the vesicle-inducing plastid protein VIPP1
(Dörmann & Benning 2003). Specific fractions of the
DGDG pool are also thought to play a role in the stabilizing
PSII (Dörmann et al. 1995; Steffen et al. 2005; Sakurai et al.
2007) and the trimerization of LHCII proteins that form the
light-harvesting antenna of PSII (Nussberger et al. 1993; Liu
et al. 2004; Holzl et al. 2006).As the stockpile of MGDG and
DGDG in the membrane-bound vesicles in xeroplasts is
apparently sufficient for thylakoid assembly (irrespective of
the presence of the DGD1 protein), the DGD1 produced in
the light may serve to synthesize DGDG that plays a role in
the stabilization of the PSII–LHCII supercomplexes.
In summary, we present the xeroplast–chloroplast transition as a novel system for studying chloroplast biogenesis in
higher plants. While the ultrastructural changes that occur
in the chloroplast differ from those in proplastids or etioplasts, there are also striking similarities, with light being
required for grana formation, chlorophyll biosynthesis, and
the synthesis of D1 and Lhcb2. It has been previously
suggested that resurrection plants have co-opted aspects of
seed desiccation tolerance into their vegetative desiccation
tolerance as evidenced by the expression of ‘seed-specific’
genes in their vegetative tissues (Illing et al. 2005). We
propose that in X. humilis, the molecular mechanisms
involved in photomorphogenesis may also be utilized in the
desiccation tolerance programme, supporting the hypothesis that vegetative desiccation tolerance is based primarily
on altered patterns of gene regulation rather than on the
presence of novel genes.
ACKNOWLEDGMENTS
We thank Borakalalo National Parks for allowing plant
collection, and are grateful to the following people for
the donation of antibodies: D1, Eva Aro (University of
Turku, Finland), and DGD1, Peter Dörmann (Max Planck
Institute for Molecular Plant Physiology, Germany). This
research was funded from grants provided by the University of Cape Town and the National Research Foundation of
South Africa.
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Figure S1. Recovery of relative water content (RWC) of
leaves used in electron micrographs characterizing changes
in ultrastructure associated with chloroplast biogenesis
during Xerophyta humilis rehydration. X. humilis plants
were rehydrated under a 16 h light/8 h dark cycle (䊐) or in
constant darkness (䉱).
Figure S2. Xeroplasts halted at stage 4 of chloroplast development when rehydrated in the dark for different lengths of
time. Scale bars = 1 mm.
Figure S3. Chlorophyll biosynthesis during rehydration.
Xerophyta humilis plants during rehydration under a 16 h
light/8 h dark cycle (a). Comparison of plants rehydrated
under 16 h light/8 h dark or under constant darkness for
48 h (b).
Received 7 August 2008; received in revised form 22 August 2008;
accepted for publication 24 August 2008
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