Plant, Cell and Environment (2007) 30, 1240–1255
doi: 10.1111/j.1365-3040.2007.01704.x
Photosynthetic recovery following desiccation of desert
green algae (Chlorophyta) and their aquatic relatives
DENNIS W. GRAY1*, LOUISE A. LEWIS1 & ZOE G. CARDON1,2
1
Department of Ecology and Evolutionary Biology, University of Connecticut, Storrs, CT 06269-3043 and 2Center for
Integrative Geosciences, University of Connecticut, Storrs, CT 06269-2045, USA
ABSTRACT
Recent molecular data suggest that desert green algae have
evolved from freshwater ancestors at least 14 times in
three major classes (Chlorophyceae, Trebouxiophyceae
and Charophyceae), offering a unique opportunity to study
the adaptation of photosynthetic organisms to life on land
in a comparative phylogenetic framework. We examined
the photorecovery of phylogenetically matched desert and
aquatic algae after desiccation in darkness and under illumination. Desert algae survived desiccation for at least 4
weeks when dried in darkness, and recovered high levels of
photosynthetic quantum yield within 1 h of rehydration
in darkness. However, when 4 weeks of desiccation was
accompanied by illumination, three of six desert taxa lost
their ability to recover quantum yield during rehydration in
the dark. Aquatic algae, in contrast, recovered very little
during dark rehydration following even just 24 h of desiccation. Re-illuminating rehydrated algae produced a nearly
complete recovery of quantum yield in all desert and two of
five aquatic taxa. These contrasts provide physiological evidence that desert green algae possess mechanisms for photosynthetic recovery after desiccation distinct from those in
aquatic relatives, corroborating molecular evidence that
they are not happenstance, short-term visitors from aquatic
environments. Photosensitivity during desiccation among
desert algae further suggests that they may reside in protected microsites within crusts, and species specificity of
photosensitivity suggests that disturbances physically disrupting crusts could lead to shifts or losses of taxonomic
diversity within these habitats.
Key-words: chlorophyll fluorescence; desert crust; light
stress; photodamage; photoprotection.
INTRODUCTION
Living in a desert presents an enormous challenge for photosynthetic organisms. Deserts are dry, experience temperature extremes (hot and cold) and expose their inhabitants
Correspondence: D. W. Gray. Fax: +01 860 486 6364; e-mail:
[email protected]
*Present address: Crop Physiology Laboratory, Department of
Plants, Soils, and Climate, Utah State University, Logan, UT 843224820, USA.
1240
to intense solar radiation during periods of time when the
potential for dissipating absorbed energy through photosynthetic activity is limited by temperature or the lack of
water. Together these stresses make deserts some of the
least favourable places for unicellular photosynthetic
organisms to live. Yet, despite these stresses, many desert
regions contain a tremendous diversity of microscopic life,
much of which is organized into a biological crust at the soil
surface.
Desert biological soil crusts are typically dominated by
cyanobacteria, but may also contain bryophytes, lichens,
fungi and can even be dominated by free-living unicellular
green algae in moister, colder environments (Cameron
1960; Shields & Drouet 1962; Cameron 1964; Friedmann &
Ocampo-Paus 1967; Belnap, Budel & Lange 2001). During
periods of desiccation, many crust organisms lie dormant.
However, when water becomes available, they can quickly
become physiologically active (Rosentreter & Belnap 2001)
and play important ecological roles as primary producers
(Garcia-Pichel & Belnap 1996) and, in the case of the
cyanobacteria, by fixing nitrogen (Evans & Ehleringer 1993;
Belnap 2003).
Recent molecular studies have revealed surprising taxonomic diversity among unicellular green algae in biotic
crusts, spread across three major green algal classes (sensu
Mattox & Stewart 1984): Chlorophyceae, Trebouxiophyceae and Charophyceae (Lewis & Flechtner 2002).
Molecular data also indicate that these desert green algal
isolates are not simply incidental visitors blown in, for
example, on wind currents, but instead their DNA
sequences have evolved sufficiently to be considered taxa
clearly distinct from aquatic sister taxa (Lewis & Lewis
2005). Whether these desert algae possess distinct physiological features that distinguish them from their aquatic
counterparts is presently unknown.
The eukaryotic green desert algae offer a unique opportunity for studying the adaptation of photosynthetic organisms to terrestrial life. The traditional approach to
investigating the adaptation to land in photosynthetic
organisms has involved comparisons of photosynthetic
physiology in embryophytes (terrestrial green plants) and
aquatic algae. However, embryophytes are descendants of
a single evolutionary transition from an aquatic ancestor to
terrestrial descendants, so only one statistically independent comparison is encapsulated even in comparisons of
© 2007 The Authors
Journal compilation © 2007 Blackwell Publishing Ltd
Photorecovery after desiccation of green microalgae 1241
multiple land plants with multiple aquatic algae (Felsenstein 1985; Harvey & Pagel 1991). In contrast, the eukaryotic desert green algae represent multiple evolutionary
lineages that have arisen independently from aquatic ancestors at least 14 times (Lewis & Lewis 2005). By comparing
independently derived desert lineages with related aquatic
algae, it is possible to have multiple statistically independent comparisons that are not possible when terrestrial
plants are compared with algae. These evolutionarily independent transitions or replicated shifts to terrestrial life
seen in the desert green algae (Lewis & Flechtner 2002;
Lewis & Lewis 2005) facilitate the investigation of whether
shifts in habitat (aquatic to desert) are correlated with
changes in physiological traits in a statistically robust comparative phylogenetic framework.
In making the transition from an aquatic habitat to a
terrestrial one (desert crusts), unicellular algae have had to
cope with radically different ecological conditions. Compared to terrestrial environments, aquatic ones are relatively thermally stable (owing to the high heat capacity of
water). Terrestrial environments, on the other hand, lack
this thermal buffering and their inhabitants must be capable
of tolerating changes (often rapid) in temperature. In
aquatic environments, desiccation is rarely a problem, while
in terrestrial environments, desiccating conditions are the
norm. In terrestrial environments (especially deserts), these
periods of desiccation are interspersed with brief periods of
water availability following rain. Yet, it is important to recognize that some aquatic environments, such as intertidal
regions and ephemeral ponds, may experience desiccation
at regular intervals, blurring the distinction between aquatic
and terrestrial habitats. Aquatic and terrestrial habitats
also differ in the light environments their inhabitants are
exposed to. Because light is attenuated more rapidly
through a column of water than through a column of air,
aquatic environments typically have lower light intensities
than terrestrial ones. However, if an aquatic organism drifts
freely in the water column, the light environment it experiences may be more variable than the one it would experience in a terrestrial environment where mobility is more
limited, and organisms (algae in particular) are limited to
residing on surfaces. Given the differing ecological conditions likely experienced by aquatic and terrestrial organisms, it would be surprising if they did not possess
adaptations for their respective environments.
In this study, we attempt to better understand the physiological differences between desert and aquatic green
algae, in hopes of gaining insight into the evolutionary transition to land in photosynthetic organisms. Specifically, we
examined the response of multiple, independent lineages of
desert green algae and their aquatic relatives to two environmental stresses: desiccation and illumination during desiccation. By analysing data from desert and aquatic isolates
using a statistical comparative approach (Felsenstein 1985;
Harvey & Pagel 1991; Garland, Bennett & Rezende 2005),
any potentially confounding impacts of shared evolutionary
history among algae were removed. We examined desiccation tolerance in the dark and in light, and the rapidity with
which photosynthetic activity could recover after desiccation; these dynamics are a first step towards characterizing
potential damage-tolerance and damage-avoidance mechanisms in these desert green algae based on mechanisms
known in other photosynthetic organisms in the literature.
For example, the cyanobacterium Nostoc flageliforme
recovers photosynthetic ability slowly following rehydration and appears to rely on repair processes requiring
protein synthesis (Qiu et al. 2004), while recovery of photosynthetic activity in the cyanobacterium Microcoleus sp.
occurs rapidly following rehydration and does not require
de novo protein synthesis (Harel, Ohad & Kaplan 2004).
Within bryophytes, some species recover full photosynthetic ability within minutes of rehydration of the haploid
vegetative tissues (Marschall, Proctor & Smirnoff 1998;
Csintalan, Proctor & Tuba 1999; Proctor & Smirnoff 2000)
and tolerate very rapid drying rates (Oliver & Bewley 1997;
Oliver, Tuba & Mishler 2000), suggesting the existence of
strong, constitutively expressed protective mechanisms that
prevent damage to their photosynthetic systems during desiccation. In contrast, other species recover more slowly and
appear to require gradual drying to allow time for the synthesis of protective proteins such as ‘dehydrins’ (Oliver &
Bewley 1997; Oliver et al. 2000). This latter pattern is also
seen in the lycophyte ‘resurrection plant’ Selaginella
(Schwab, Schreiber & Heber 1989; Muslin & Homann 1992;
Oliver & Bewley 1997) and in certain angiosperms (Vicre,
Farrant & Driouich 2004).
Very little is known about the adaptations allowing
single-celled, eukaryotic green algae to survive in desert
crusts. However, because these desert algae are morphologically very simple (Lewis & Flechtner 2004) and lack
obvious structural features to distinguish them from their
aquatic relatives, their abilities to survive and grow in the
desert environment are likely the result of changes in physiology rather than persistent changes in gross morphology.
During desiccation, membranes may become physically disrupted by the loss of water (Bewley 1979; Crowe, Hoekstra
& Crowe 1992), and cells may suffer from both chemical
and oxidative damage when dry (Smirnoff 1993; Weissman,
Garty & Hochman 2005b). To cope with such potential
sources of damage, photosynthetic organisms more generally have evolved an array of physiological protective
mechanisms, including osmotic adjustments to the cytoplasm that reduce damage to membranes and organelles
(Crowe et al. 1992; Smirnoff 1992; Muller et al. 1997; Potts
1999), synthesis of proteins that protect cellular components during the desiccation process (Gwozdz, Bewley &
Tucker 1974; Oliver 1996), and the production of antioxidants and scavenging enzymes that neutralize reactive
oxygen species (ROS) generated during drying (Seel,
Hendry & Lee 1992; Kranner et al. 2003; Ledford & Niyogi
2005; Weissman, Garty & Hochman 2005a). Carotenoids,
especially those of the xanthophyll cycle, serve to harmlessly dissipate excess energy absorbed by chlorophyll in
terrestrial plants and algae (Bilger & Björkman 1990;
Demmig-Adams & Adams 1992; Gilmore 1997; Niyogi
1999; Bukhov et al. 2001a,b; Masojidek et al. 2004). In
© 2007 The Authors
Journal compilation © 2007 Blackwell Publishing Ltd, Plant, Cell and Environment, 30, 1240–1255
1242 D. W. Gray et al.
cyanobacteria, which lack a xanthophyll cycle, some species
synthesize screening pigments that shield them from UV
radiation (Bohm et al. 1995; Potts 1999). Finally, upon rehydration, desiccated organisms may repair damage by synthesizing new proteins to replace damaged ones (Gwozdz &
Bewley 1975; Oliver & Bewley 1984a,b; Oliver 1996; Oliver
& Bewley 1997; Oliver, Wood & O’Mahony 1997). Among
some aquatic green algae (such as Spirogyra and Volvox),
survival of desiccation is linked to switches in life stage; the
diploid zygote can tolerate adverse environmental conditions and then undergo meiosis to produce active, haploid
vegetative cells. However, such a pattern among desert
green algae as a strategy for tolerating desiccation has not
been generally documented. The degree to which photosynthetic organisms of all complexities rely on specific mechanisms to tolerate desiccation, and their relative reliance on
protection versus repair strategies, clearly varies considerably (Bewley & Oliver 1992; Oliver & Bewley 1997). The
potentially diverse mechanisms used by phylogenetically
diverse desert green algae are virtually unknown but are of
interest both for understanding the evolution of unicellular
life on land and for restoration efforts in arid lands where
crusts have been disrupted.
MATERIALS AND METHODS
Study organisms
In this study, we examine the physiology of 11 eukaryotic
green algae that live in contrasting habitats (desert and
aquatic) (Table 1). Desert algae were isolated from
samples of biological soil crusts taken from arid regions
of the southwestern USA, as part of the Biotic Crust
Project (http://hydrodictyon.eeb.uconn.edu/bcp/). Phyloge-
netic relationships among the desert algae were determined using previously published 18S rDNA or ITS
rDNA sequence data (Lewis & Flechtner 2004; Lewis &
Lewis 2005). This information was used to pair the desert
algae with their closest aquatic relatives available in
culture collections or in culture in our lab (Table 1).
DNA extraction, PCR, sequencing
18S rDNA sequence data from three of the aquatic green
algae were collected specifically for this study. Isolates of
Scenedesmus platydiscus (UTEX 2457), Chlamydopodium
sp. (MLO301CT) and Chlorella sp. (UTEX 318) were
grown on Bold’s Basal Medium (Bold 1949) agar slants
under a 12:12 L/D cycle at 20 °C. Cells were scraped off
the slants and concentrated by centrifugation at 3000 g.
Genomic DNA was extracted from the algal cells using a
modified CTAB extraction that included grinding the algal
cells in sterile sand (Shoup & Lewis 2003). The 18S rDNA
region was amplified using the primers SSU1 and SSU2, and
the resulting fragments were sequenced directly using
amplification and internal primers, including 284F and
1081R (Phillips & Fawley 2000), or others listed in Shoup &
Lewis (2003). Three new primers (660F 5′TATGGTGAG
TACTGCTATGGC3′, 817R 5′AGTCCTATCGTGTTAT
TCCATGC3′ and 918F 5′TGAAAGACGAACTACT
GCG3′) were designed in order to allow sequencing
through introns in the 18S rDNA gene of UTEX 2457.
Double-stranded PCR products were sequenced in 10 mL
volumes with the PRISM system (Applied Biosystems, Inc.,
Foster City, CA, USA) using the manufacturer’s directions.
Chromatograms from individual sequencing runs were first
trimmed, then assembled into consensus sequences in
Sequencher (GeneCodes Corp., Inc., Ann Arbor, MI,
Taxon
Habitata
Culture sourceb
GenBank accession number
Chlorophyceae
Bracteacoccus sp.
Bracteacoccus sp.
Scenedesmus platydiscus
Scenedesmus rotundus
Chlorogonium elongatum
Chlorosarcinopsis sp.
Chlamydopodium sp.
Monoraphidium braunii
D
D
A
D
A
D
A
A
BC2-1
CNP2-VF25
UTEX 2457
SEV3-VF49
UTEX 11
SEV2-VF1
MLO301CT
SAG 2006
AF516676
AF516677
EF159952
AF513373
U70589
AF516678
EF159950
AJ300527
Trebouxiophyceae
Chlorella sp.
Chlorella sp.
Myrmecia sp.
A
D
D
UTEX 318c
BC4-VF9
BC8-8
EF159951
AF516675
AF516674
Table 1. Sources of green algae (Chlorophyta), their origination habitats and
GenBank accession numbers corresponding
to the18S rDNA sequence data
The GenBank accession numbers listed in boldface indicate 18S rDNA data that are new to
this study.
a
Habitat from which the alga was isolated.
b
Isolates from the Biotic Crust Project, except for those labelled with the University of Texas
Culture Collection of Algae (UTEX) or the Sammlung von Algenkulturen der Universität
Göttingen (SAG) acronym.
c
This isolate was associated with the original UTEX 318, but is not Microthamnion.
D, desert; A, aquatic.
© 2007 The Authors
Journal compilation © 2007 Blackwell Publishing Ltd, Plant, Cell and Environment, 30, 1240–1255
Photorecovery after desiccation of green microalgae 1243
USA). Base call edits (when necessary) were carried out
manually. Most (>80%) of the reported base calls were
verified with sequencing reactions in both the forward and
reverse orientations; the remaining nucleotides were verified from independent sequencing reactions in the same
orientation.
Phylogenetic analyses
The 18S rDNA sequences from the 11 green algae isolates
listed in Table 1 were aligned by eye in the text editor of
PAUP* (Swofford 2002), excluding the intron sequences. In
addition to the ingroup sequences, in order to determine the
placement of the root for the comparative analysis, the 18S
rDNA sequences of two prasinophycean algae were used as
outgroup taxa (Nephroselmis olivacea, GenBank accession
number X74754, and Tetraselmis striata, X70802). The final
18S alignment consisted of 1770 characters, 97 of which
were excluded from the analysis because they could not be
aligned with certainty. Of the remaining 1673 characters,
133 were parsimony informative, and 1365 constant. The
alignment will be made available from TreeBASE (http://
www.treebase.org/).
Prior to the maximum likelihood (ML) analyses, the data
set was analysed using MODELTEST 3.06 (Posada & Crandall 1998). The favoured substitution model for this alignment of 18S data was determined to be TrN + I + G.
Parameter values were set as follows: relative base frequencies (pA = 0.2559, pC = 0.2162, pG = 0.2698, pT = 0.2581); relative rate matrix (rAC = 1.0000, rAG = 2.4720, rAT = 1.0000,
rCG = 1.0000, rCT = 5.1053, rGT = 1.0000); gamma shape,
Scenedesmus rotundus SEV3-VF49 (AF513373) D
100
1.00
Scenedesmus platydiscus UTEX 2457 (EF159952) A
Monoraphidium braunii SAG 2006 (AJ300527) A
92
1.00
Bracteacoccus sp. BC2-1 (AF516676) D
100
1.00
71
0.98
Bracteacoccus sp. CNP2-VF25 (AF516677) D
54
0.76
Chlorosarcinopsis sp. SEV2-VF1 (AF516678) D
Chlamydopodium sp. MLO301CT (EF159950) A
100
1.00
96
1.00
Chlorophyceae
58
0.67
0.6267, and proportion of invariant sites = 0.5048. ML
analyses were performed using PAUP* 4.b.10 (Swofford
2002) for UNIX. ML analyses used heuristic searches with
10 random additions of taxa, each followed by TBR branch
swapping. Bootstrap analysis (Felsenstein 1985) included
1000 replicates, with a single random addition of taxa for
each replicate, and under the same model as was used for
the heuristic searches.
Bayesian analyses utilized MrBayes 3.0b4 (Huelsenbeck
& Ronquist 2001). Two independent runs were performed
under the GTR + I + G model (determined to be the best
fitting model using MrModeltest v2, Nylander (2004).
Each run began with an independent random starting tree
and extended two million generations. Each run employed
three heated chains (temperature parameter of 0.2) in
addition to the cold chain. A flat Dirichlet prior for relative nucleotide frequencies and relative rate parameters, a
discrete uniform prior for topologies, and an exponential
distribution (mean of 1) for the gamma shape parameter
and all branch lengths were used. Trees were sampled
every 100 generations, yielding 2000 sampled trees per
run. History plots of the overall likelihood scores were
plotted to determine which trees were to be excluded as
burnin. Results were highly consistent between the two
runs, suggesting that each run was long enough to achieve
stationarity. Convergence was assessed by comparing splits
included in majority-rule consensus trees of each run
separately. The trees after burnin from both runs were
combined to generate a 50% majority-rule consensus tree.
The estimated posterior probability of a given split is
shown on the ML tree (Fig. 1).
67
0.98
Chlorella sp. BC4-VF9 (AF516675) D
67
0.98
Chlorella sp. UTEX 318 (EF159951) A
Myrmecia sp. BC8-8 (AF516674) D
Tetraselmis striata (X70802)
Nephroselmis olivacea (X74754)
0.01 substitutions/site
Trebouxiophyceae
Chlorogonium elongatum UTEX 11 (U70589) A
Figure 1. Maximum likelihood (ML)
phylogenetic tree of chlorophycean and
trebouxiophycean algae used in the
desiccation experiments, estimated from
18S rDNA sequences obtained from
algae listed in Table 1. Desert and aquatic
algae are indicated by the letters D and
A, respectively. Prasinophyte taxa
Nephroselmis and Tetraselmis were used
to orient the tree. Upper-case text
indicates the isolate numbers identifying
algal isolates. Text in parentheses
indicates the Genbank accession number
for the sequences associated with the
algal isolate. Boxes associated with nodes
of the tree indicate ML bootstrap support
(upper number) and Bayesian posterior
probabilities (lower number). Details of
the tree estimation are described in
Materials and Methods.
© 2007 The Authors
Journal compilation © 2007 Blackwell Publishing Ltd, Plant, Cell and Environment, 30, 1240–1255
1244 D. W. Gray et al.
Culture conditions
Algae were grown in Erlenmeyer flasks as aqueous unialgal cultures in a solution of Bold’s Basal medium supplemented with micronutrients (Trelease & Trelease 1935;
Bold 1949) and were allowed to colonize the surface of
porous glass beads (Siran carriers 1–2 mm diameter; Jaeger
Biotech Engineering, Inc., Costa Mesa, CA, USA) covering
the bottom of the flask. Cultures were maintained in an
Enconair model GR-27 walk in growth room (Enconair,
Winnipeg, Manitoba, Canada) under a 12:12 L/D cycle at a
constant temperature of 20 °C, a daytime photosynthetic
photon flux density (PPFD) of 130 mmol photons m-2 s-1
and were bubbled continuously with ambient air. Once
colonized, the algae-covered beads were transferred to
small plexiglass containers sized to be fit snugly into a
holder underneath the fiber optic from a PAM-2000 chlorophyll fluorometer (Heinz Walz GmbH, Effeltrich,
Germany) used for all fluorescence measurements. These
containers were made from square pieces of 0.635 cm thick
plexiglass measuring 1.905 cm on a side. A 1.27 cm diameter
hole was bored through the centre, and the bottom was
covered in fine mesh to create the floor of the container.
Nine containers fit easily into a Petri dish chamber.
Experimental design
The experiment utilized a two-way factorial design in which
algal samples were exposed to varying durations of desiccation and varying degrees of illumination during desiccation. Recovery of photosynthetic ability was assessed in
rehydrated algae by measuring the photosynthetic quantum
yield (efficiency of light utilization by photosystem II
[PSII]) using chlorophyll fluorescence (Genty, Briantais &
Baker 1989). Recovery of photosynthetic activity was initially measured in algae that were rehydrated in darkness,
and later measured in the same algal samples after they
were re-illuminated. This allowed us to compare differences
in the sensitivity of photosynthesis to desiccation and illumination in the desert and aquatic algae under study.
Each of the 11 algal taxa was assigned to three desiccation
treatments (24 h, 7 d and 4 weeks dry) and three illumination
treatments (dark, intermediate intensity and full intensity).
Each treatment combination was replicated three times for a
total of 27 samples for each of the 11 algae, resulting in 297
plexiglass sample containers housing algae on glass beads.
Within each illumination treatment, replicate samples
assigned to desiccation treatments were distributed across
three separate Petri dish chambers in which drying was
accomplished by pumping ambient air at 25% relative
humidity (RH) via tubing connected to aquarium pumps.
In the ‘dark’ treatment, drying chambers were covered in
aluminum foil to exclude light during desiccation and
storage. In the two ‘illuminated’ treatments (intermediate
intensity and full intensity), drying chambers were left
exposed to the illumination regime under which the algae
were grown in aqueous culture (130 mmol photons m-2 s-1
PPFD). Exposure to intermediate light intensity was
achieved by allowing the uppermost layer of beads to selfshade lower bead layers. Removing the upper layer immediately prior to rehydration allowed access to algae that had
been exposed to light levels intermediate between those of
the dark treatment and the upper bead layer of the full
intensity treatment. This removal of beads increased the
sample fiber optic distance by 2–3 mm and reduced the
intensity of the saturating flash for these samples. Examining the fluorescence induction kinetics obtained from these
samples, however, showed that fluorescence reached a peak
or plateau, indicating that the samples were receiving
flashes of sufficient intensity to fully saturate the algal photosystems. Algae were dried and stored under their respective illumination conditions for the duration of their
respective desiccation intervals.
Spot checks of the algae following 12–24 h of exposure to
the drying air stream showed that photosynthetic quantum
yield of the upper bead layers in both the dark and full
illumination treatments had declined to <0.1, indicating a
loss of photosynthetic activity. Algae were then allowed to
remain dry for an additional 24 h, 7 d or 4 weeks in the
growth chamber.
Following desiccation intervals of 24 h, 7 d or 4 weeks, the
algae were rehydrated by adding water to the Petri dish
housing the algae-covered beads in their plexiglass sample
containers. Water was allowed to wick up through the mesh
container bottom and throughout the algae-covered beads
by capillary action. This procedure minimized the physical
disturbance of the algae-covered beads during rehydration.
Rehydration took place initially in darkness, and the recovery of photosynthetic quantum yield was measured for each
plexiglass container 1, 24 and 48 h following the onset of
rehydration in darkness. Each chamber was secured at
a constant distance underneath the fiber optic of a PAM2000 fluorimeter (Heinz Walz GmbH), and measurements
were gathered as described further. Following rehydration
for 48 h in the dark, the algae were re-illuminated at
130 mmol photons m-2 s-1 PPFD on a regular 12:12 day–
night cycle, and photosynthetic quantum yields were measured following illumination of rehydrated algae for 1 and
5 d. Prior to measuring quantum yields of re-illuminated
samples, the algae were dark adapted for 6–8 h.
Chlorophyll fluorescence measurements
Chlorophyll fluorescence measurements (Fo, Fm, and Fv/Fm)
following standard nomenclature from van Kooten & Snel
(1990) and Maxwell & Johnson (2000) were made using the
measurement system described in detail in Gray, Cardon &
Lewis (2006). This system uses a PAM-2000 portable fluorometer (Heinz Walz GmbH) and a CR10X data logger
(Campbell Scientific, Inc., Logan, UT, USA) to implement a
protocol for simultaneously recording rapid fluorescence
induction kinetics, changes in steady-state fluorescence
and an improved method for measuring the true maximum
fluorescence (Fm) in algae. The measuring light intensity
was set to the lowest setting available with the PAM2000, corresponding to ~0.06 mmol photons m-2 s-1 and
© 2007 The Authors
Journal compilation © 2007 Blackwell Publishing Ltd, Plant, Cell and Environment, 30, 1240–1255
Photorecovery after desiccation of green microalgae 1245
modulated at a frequency of 600 Hz. Saturation pulses
were delivered at the lowest intensity halogen setting
(~1450 mmol photons m-2 s-1) and lasted for either 0.8 or
1.0 s. All fluorescence measurements were made on the
algae following a dark adaptation period of at least 6–8 h.
During fluorescence measurements, plexiglass containers
containing algae-covered beads were transferred in darkness to a sealed plexiglass measuring cuvette and were
placed into a fixed bracket. Air was directed over the
surface of the beads while they were held in place. A hole in
the lid of the cuvette admitted the PAM-2000 fiber optic to
a glass cover slip window directly above the algae. This
arrangement allowed repeatable placement of the fiber
optic at a fixed distance from the algae between sampling
periods. Maintaining such a fixed and repeatable geometry
is important for interpreting changes in Fo and Fm because
the fluorescence signal obtained from a sample is proportional to the sample to fiber optic distance. In preliminary
experiments, by repeatedly removing and replacing the
same algal sample between fluorescence measurements, we
were able to show that repeated Fo measurements taken on
a single sample varied by less than 2% using this cuvette
system (data not shown).This allowed us to track changes in
absolute levels of fluorescence (Fo and Fm), as well as the
ratio measurement Fv/Fm (which is not affected by changes
in fiber optic to sample distance) during the experiment.
Desiccation dynamics
In order to determine the rate and degree of drying experienced by the algal samples during desiccation, a drying curve
was measured for several replicate algal samples identical to
those used in the desiccation experiment. After placing a
plexiglass container of fully hydrated algae-covered beads in
the fluorescence measuring cuvette described earlier, dry air
at 25% RH was passed into the cuvette and across the
surface of the beads. Exhaust gas from the cuvette was
routed to a LI-6262 infrared gas analyser (IRGA) (LI-6262,
Li-Cor Inc., Lincoln, NE, USA), which measured water
vapour content of the air leaving the cuvette. Both quantum
yield and IRGA readings were monitored at 30 min intervals
for 24 h. This allowed us to observe the dynamics of water
loss from the system and changes in the quantum yield of the
algae during the drying process.
Statistical analyses
Analyses were carried out in two major ways, one incorporating phylogenetic information and one that did not.
Overall patterns of fluorescence and photosynthetic
recovery (ignoring lineages) were analysed with analysis of
variance (anova) using the GLM procedure of the statistical package SAS (1990). The original algal habitat (desert/
aquatic), illumination level during desiccation (darkness,
intermediate intensity and full intensity), length of desiccation (24 h, 7 d, 4 weeks), and time since rehydration (1 h
dark, 24 h dark, 48 h dark, 1 d re-illuminated, 5 d
re-illuminated) were treated as fixed main effects. Because
the same algal samples were sampled repeatedly during
rehydration, the time since rehydration effect was also
treated as a repeated factor in the analysis. The algal
‘species’ was treated as a random blocking factor nested
within the habitat main effect.
Comparative analyses were used to investigate the associations between habitat of origin and desiccation tolerance, and habitat of origin and resistance to photodamage
while desiccated, using phylogenetically corrected generalized least squares (PGLS) regression as implemented in
version 4.6b of the software package COMPARE (Martins
2004). PGLS allows testing of correlations between the
characters in a group of species (e.g. traits and habitat)
taking into account the underlying phylogenetic relationships among taxa. In PGLS, the parameter alpha sets the
rate of adaptation of a trait to the optimum. When alpha is
large, traits move instantly to new optima set by natural
selection, and there is no correlation between ancestors and
descendants. Under conditions of large alpha, the analysis
approximates a non-phylogenetically corrected least
squares regression (TIPS). TIPS assumes (potentially incorrectly) that all species in the analysis are evolutionarily
independent units of observation. When alpha is small,
PGLS approximates Felsenstein’s independent contrasts
(FIC), and phylogeny is a strong determinant of correlations among characters. The PGLS ML estimate of alpha is
usually intermediate to these two extremes (‘PGLS’).
To examine the differential impact of desiccation and
illumination during desiccation on photosynthetic performance in aquatic and desert algae, we derived two indices of
damage. The desiccation damage index (DDI) measures the
fraction of initial aqueous quantum yield lost following
desiccation in darkness. This is described in Eqn 1 where
YieldAQ(avg) represents the average quantum yield of an algal
taxon obtained from a random subsample (n = 3) of darkadapted replicates prior to application of the desiccation
treatment; Yielddarkdry(i) is the quantum yield of an individual
rehydrated algal sample that was previously dried and
stored in darkness, where the subscript (i) indicates different replicate samples. The total number of samples measured is indicated by j.
DDI =
1 j YieldAQ(avg) − Yielddarkdry(i)
∑
j 1
YieldAQ(avg)
(1)
The photodamage index (PDI) measures the additional
fraction of initial quantum yield that is lost when desiccated
algae are also exposed to illumination while they are dry.
This is described by Eqn 2, where YieldAQ(avg) represents the
average quantum yield of all dark-adapted algae samples
within a taxon prior to application of the desiccation treatment (n = 27); Yielddark(avg) represents the average quantum
yield of rehydrated algal isolate previously dried and stored
in darkness (n = 3); Yieldillum(i) is the quantum yield of an
individual rehydrated algal sample that was previously
dried and stored under illumination, where the subscript (i)
indicates different replicate samples. The total number of
samples measured is indicated by j.
© 2007 The Authors
Journal compilation © 2007 Blackwell Publishing Ltd, Plant, Cell and Environment, 30, 1240–1255
1246 D. W. Gray et al.
(2)
Both DDI and PDI scale algal responses to their initial
quantum yield, thereby allowing a fair comparison among
algae that may have inherently different quantum efficiencies of light utilization even when growing in aqueous
culture.
IRGA mV
Quantum yield
1500
0.7
0.6
0.5
Equilibration at
25% RH
1000
Bulk water lost
0.4
0.3
0.2
500
0.1
0
0
240
480
720
960
Quantum yield (Fv/Fm)
2000
1 j Yielddark(avg) − Yieldillum(i)
∑
j 1
YieldAQ(avg)
Airstream water content
(IRGA mV)
PDI =
0.0
1200 1440
Time elapsed (min)
RESULTS
Figure 2. Representative drying curve for desert alga
Sequencing and phylogenetic reconstruction
The resulting 18S rDNA regions ranged from 1664 to 5293
nucleotides in length. The S. platydiscus sequence contains
eight putative group IA introns, in positions 125, 989, 1534,
1976, 2840, 3484, 4278 and 4909 of the 18S rDNA region.
The entire fragment was PCR amplified in segments rather
than as a single amplification product. A 43-nucleotide
region of the 18S rDNA exon, present in the other algae
sequences, was not determined because of sequencing difficulty. The 43 undetermined positions, corresponding to
positions 4094–4135 in the sequence, were designated as the
nucleotide ambiguity code ‘N’ in the alignment. As this
region was not determined by sequencing, it is possible that
additional inserted sequences are present in the rDNA gene
of S. platydiscus. The newly obtained 18S consensus
sequences were subjected to BLAST searches (Altschul
et al. 1990) in order to screen for contaminant sequences
prior to phylogenetic analyses. The 18S sequences from
these isolates were deposited in GenBank under accession
numbers EF159950–EF159952.
ML analysis produced a phylogenetic tree with a score of
-lnL = 5066.9385 that groups the ingroup taxa into three
major lineages, each containing desert and aquatic derived
taxa (Fig. 1). Two of these lineages correspond to the clockwise and directly opposed flagellar apparatus orientation
groups in Chlorophyceae, and the third lineage contains
members of Trebouxiophyceae (Friedl 1995).
Desiccation dynamics
Water content of the air leaving the fluorescence cuvette
reached equilibrium shortly following placement of the
algal sample in the fluorescence cuvette. Humidity in the
cuvette remained constant for several hours, after which
water content of the air in the cuvette began to decline
precipitously over a 3 h period until the water content of
the exhaust air equilibrated at a dewpoint of 2 °C (relative
humidity of 25%). This pattern suggests an initial period in
which evaporation occurred from bulk water surrounding
the beads and was sufficient to maintain constant high
humidity in the chamber (Fig. 2). During this time, quantum
yield from the algae was very high. This was then followed
by a period of declining cuvette humidity as the remaining
water evaporated from the beads and from the algal cells.
As the chamber humidity declined, quantum yield of the
CNP2-VF25 placed in a flow-through cuvette with inlet air at
25% relative humidity and 20 °C. Water content of exhaust air
(closed squares) and algal quantum yield (open circles) was
measured at 30 min intervals for 24 h. Airstream water content
was measured with a LI-6262 infrared gas analyser (IRGA)
(Li-Cor Inc., Lincoln, NE, USA) which outputs a voltage signal
proportional to airstream water content. Quantum yield was
measured as described in the text. Similar drying patterns were
observed in desert and aquatic algae dried in darkness or under
illumination.
algal sample also declined; however, the decline in quantum
yield was delayed and did not take place until chamber
humidity had declined significantly. This delay might indicate a threshold water content below which algal physiology was adversely impacted. Once chamber humidity had
equilibrated with that of the incoming air (25% RH), algal
quantum yields also equilibrated at yields below 0.1, indicating a loss of photosynthetic activity in the desiccated
algae. The pattern of changes in quantum yield observed in
this drying curve closely matched the pattern of quantum
yield measurements observed in algae desiccated in Petri
dishes during the desiccation experiment. In both cases,
quantum yields began to decline after 12 h of exposure to a
desiccating airstream, and photosynthetic activity was lost
within 24 h.
Visible impacts of desiccation and illumination:
photobleaching
Following desiccation in darkness, dry beads colonized by
aquatic and desert algae remained extensively pigmented
regardless of the length of time they were desiccated (24 h,
7 d and 4 weeks). Beads colonized with desert and aquatic
algae also remained heavily pigmented when dried under
illumination for either 24 h or 7 d. However, following 4
weeks of desiccation under illumination, all the aquatic
algae were noticeably bleached of color compared to their
counterparts dried in darkness. This photobleaching was
also present to varying but lesser degrees in the desert
algae, with desert SEV3-VF49 and SEV2-VF1 showing the
least loss of color compared with samples dried under darkness. Photobleaching was limited to the uppermost layers of
beads, and darkly pigmented surfaces were present on
beads beneath the bleached top layer and frequently on the
underside of a bleached bead.
© 2007 The Authors
Journal compilation © 2007 Blackwell Publishing Ltd, Plant, Cell and Environment, 30, 1240–1255
Photorecovery after desiccation of green microalgae 1247
Table 2. Results of analysis of variance (anova) from the desiccation experiment examining differences in photosynthetic quantum yield
as a function of habitat of origin (habitat), illumination treatment (light), length of desiccation (dry) and rehydration interval (RH time)
Source
d.f.
Main effects
Habitat
Light
Dry
RH time
Interactions
Habitat ¥ light
Habitat ¥ dry time
Habitat ¥ RH time
Dry ¥ light
Dry ¥ RH
RH ¥ light
Habitat ¥ dry ¥ light
Habitat ¥ RH ¥ light
Dry ¥ RH ¥ light
Habitat ¥ dry ¥ RH ¥ light
MS
Error term
MSE
F
P
1
2
2
4
15.8199
1.13361
1.12877
3.44088
Alga (habitat)
Alga ¥ light(habitat)
Dry ¥ alga(habitat)
RH ¥ alga(habitat)
0.43545
0.059104
0.049507
0.121457
36.33
19.18
22.80
28.33
0.0002
<0.0001
<0.0001
<0.0001
2
2
4
4
5
8
4
8
10
10
0.18319
0.39981
0.13173
0.04745
0.02178
0.08551
0.04958
0.08965
0.00733
0.01150
Alga ¥ light(habitat)
Dry ¥ alga(habitat)
RH ¥ alga(habitat)
Dry ¥ alga ¥ light(habitat)
Dry ¥ RH ¥ alga(habitat)
RH ¥ alga ¥ light(habitat)
Dry ¥ alga ¥ light(habitat)
RH ¥ alga ¥ light(habitat)
Dry ¥ RH ¥ alga ¥ light(habitat)
Dry ¥ RH ¥ alga ¥ light(habitat)
0.059094
0.049482
0.121975
0.018321
0.008543
0.007027
0.018297
0.007027
0.003381
0.003374
3.10
8.08
1.08
2.59
2.55
12.17
2.71
12.76
2.17
3.41
0.0697
0.0031
0.3787
0.0529
0.0409
<0.0001
0.0454
<0.0001
0.0265
0.0008
d.f., degrees of freedom.
declined as algae remained dry for progressively longer
times, as indicated by the significant effect of dry time
(Table 2). In aquatic algae, most of the ability to recover
quantum yield upon rehydration was lost after 24 h of desiccation, while desert algae still recovered some quantum
yield even following 4 weeks of desiccation in darkness
(Fig. 3). These differences in the behaviour of desert and
Recovery of photosynthetic quantum yield
during rehydration of algae dried in darkness
Photosynthetic quantum yields measured on dry algae were
all <0.1, regardless of pigmentation, indicating a loss of
photosynthetic activity. The ability of algae to recover photosynthetic quantum yield when rehydrated in darkness
Correlation coefficient
Rehydration period Desiccation period TIPS
DDI
Dark 24 h
Illuminated 24 h
PDI
Dark 24 h
Illuminated 24 h
PGLS (alpha) FIC
Slope
24 h
7d
4 weeks
24 h
7d
4 weeks
-0.95 -0.95 (15.50)
-0.90 -0.90 (15.50)
-0.92 -0.93 (6.06)
NA
NA
-0.65 -0.66 (5.44)
-0.65 -0.72 (0.957)
-0.97 -0.58 ⫾ 0.13*
-0.94 -0.55 ⫾ 0.16*
-0.96 -0.50 ⫾ 0.13*
NA
NA
-0.69 -0.35 ⫾ 0.26*
-0.76 -0.35 ⫾ 0.22*
24 h
7 days
4 weeks
24 h
7d
4 weeks
0.59
0.61
0.93
NA
-0.29
-0.07
0.45
0.74
0.96
NA
-0.55
-0.14
0.59 (15.50)
0.66 (1.88)
0.93 (15.50)
NA
-0.37 (2.95)
0.06 (15.50)
Table 3. Results of phylogenetic
comparative analyses examining the
correlation between habitat of algal origin
and recovery of photosynthetic quantum
yield
0.09 ⫾ 0.08*
0.21 ⫾ 0.15*
0.45 ⫾ 0.11*
NA
-0.09 ⫾ 0.15 NS
0.01 ⫾ 0.14 NS
Results are presented for rehydration in darkness for 24 h and after re-illumination for 1 d
in algae dried for 24 h, 7 d, and 4 weeks.
Notes: The desiccation damage index (DDI) indicates the fraction of the initial aqueous
quantum yield that was lost as a result of desiccation. The photodamage index (PDI)
indicates the additional fraction of aqueous quantum yield that was lost as a result of
illumination.
*Comparisons yielding significant correlations.
PGLS, phylogenetically generalized least squares using the maximum likelihood (ML) estimate of alpha. FIC, approximation to Felsenstein independent contrasts; TIPS, nonphylogenetically corrected generalized least squares regression; NS, non-significant results;
NA, no data available.
© 2007 The Authors
Journal compilation © 2007 Blackwell Publishing Ltd, Plant, Cell and Environment, 30, 1240–1255
1248 D. W. Gray et al.
Desert algae
Aquatic algae
1 h post rehydration in darkness
(c) 0.8
Quantum yield (Fv/Fm)
Quantum yield (Fv/Fm)
(a) 0.8
1 h post rehydration in darkness
0.6
0.4
0.2
0.0
0.6
0.4
0.2
0.0
MLO301CT
SAG
2006
UTEX
2457
UTEX 11
UTEX
318
0.4
0.2
0.0
(d) 0.8
Quantum yield (Fv/Fm)
Quantum yield (F v/Fm)
(b) 0.8
24 h post rehydration in darkness
0.6
24 h post rehydration in darkness
0.6
0.4
0.2
0.0
BC2-1
CNP2VF25
Algal isolate
Legend:
Pre-desiccation
SEV3VF49
SEV2VF1
BC8-8
Algal isolate
Dry 24 h
Dry 7 d
aquatic algae were statistically significant as indicated by a
significant habitat effect (Table 2).
Within 1 h of the onset of rehydration in the dark, aquatic
algae desiccated for 24 h in darkness had recovered very
little (Fig. 3a). Following 24 h of rehydration, quantum
yields of aquatic algae were only slightly greater than yields
1 h post rehydration (Fig. 3a,b). In contrast, desert algae
desiccated for 24 h in darkness exhibited rapid recovery of
photosynthetic quantum yield upon rehydration, reaching
56–91% of their initial aqueous culture yield within 1 h
(Fig. 3c). After 24 h of rehydration in darkness, little additional recovery of yield was observed (Fig. 3d).
Recovery of photosynthetic quantum yield in
algae illuminated during desiccation
Recovery of photosynthetic quantum yield following rehydration was consistently lower in algae dried under illumination than in algae dried in darkness for equal lengths of
time, as indicated by a significant treatment effect in
Table 2. To ease visual comparisons in Fig. 4, results from
all aquatic and desert species, respectively, have been
grouped. Grouped data from dark-dried algae are shown in
black bars in Fig. 4, alongside data from beads that
were fully exposed to light during desiccation (full
illumination = white bars) and algae on beads that were
below the top-most bead layer during desiccation in the
light (intermediate illumination = grey bars). Data are separated into panels based on the length of time the algae
remained desiccated. Rehydration treatments (e.g. 1 and
24 h rehydration in the dark, as in Fig. 3, along with further
hydration in the light) are indicated along the x-axes of
Fig. 4.
Dry 4 weeks
BC4VF9
Figure 3. Recovery of photosynthetic
quantum yield in aquatic (a,b) and desert
(c,d) algae following rehydration in
darkness for 1 h (a,c) and 24 h (b,d).
Columns represent average quantum
yield following dark adaptation for 6–8 h.
Key to symbols: black bars = predesiccation; dark grey bars = algae
desiccated 24 h; light grey bars = algae
desiccated 7 d; white bars = algae
desiccated 4 weeks. Error bars represent
⫾1 SD with n = 3.
Aquatic algae had lost nearly all ability to recover
quantum yield following desiccation for as little as 24 h
(Fig. 3a,b), and illumination during desiccation tended to
reduce their ability to recover quantum yield still further
(Fig. 4a–c). The negative effect of light exposure during
desiccation was particularly pronounced in the desert algae
(Fig. 4d–f, grey and white bars), all of which had shown
rapid recovery to high quantum yields upon rehydration for
even just 1 h following all lengths of desiccation in darkness
(Figs 3c,d & 4d–f, black bars). Over time, as desert and
aquatic algae were rehydrated first in the dark and then
with supplemental light, the effect of having been desiccated in light became less obvious (Fig. 4), although the
aquatic algae were much more variable in their recovery in
the light (Fig. 4b,c). Unfortunately, for the algae dry only
1 d, recovery of quantum yield was not tracked after 48 h in
dark rehydration (Fig. 4a,d). Following 4 weeks of desiccation under illumination, the upper layer of most of the
desert algae exhibited no recovery even after 24 h of rehydration in darkness (Fig. 4f), although these same algal
species recovered significant quantum yields when dried 4
weeks in darkness (Figs 3c,d & 4f, black bars). Examining
quantum yields of algae on beads located beneath the
uppermost bleached surface layer revealed levels of recovery that were intermediate between those of algae dried in
darkness and the upper layers of algae dried under illumination (Fig. 4, grey bars).
Recovery of photosynthetic quantum yield in
algae re-illuminated following rehydration
The highly variable recovery among diverse aquatic algae
exposed to light during rehydration (right-most groups of
© 2007 The Authors
Journal compilation © 2007 Blackwell Publishing Ltd, Plant, Cell and Environment, 30, 1240–1255
Photorecovery after desiccation of green microalgae 1249
Desert algae
Aquatic algae
Darkness
Intermediate
Full
Quantum yield (Fv/Fm)
Dry 1 d
0.6
0.4
0.2
No data
No data
(d) 0.8
Quantum yield (Fv/Fm)
(a) 0.8
0.0
0.4
0.2
0.0
0.2
No data
No data
Dry 7 d
Figure 4. Recovery of photosynthetic
0.6
0.4
0.2
0.0
(c) 0.8
(f) 0.8
Quantum yield (Fv/Fm)
Dry 4 weeks
Quantum yield (Fv/Fm)
0.4
(e) 0.8
Dry 7 d
0.6
0.6
0.4
0.2
0.0
0.6
0.0
Quantum yield (Fv/Fm)
Quantum yield (Fv/Fm)
(b) 0.8
Darkness
Intermediate
Full
Dry 1 d
1h
dark
24 h
dark
1 d re5 d reilluminated illuminated
Time after rehydration
Dry 4 weeks
0.6
0.4
0.2
0.0
1h
dark
24 h
dark
1 d re5 d reilluminated illuminated
Time after rehydration
bars, Fig. 4b,c) is further explored in Fig. 5a,b. The aquatic
algae UTEX 11 and UTEX 318, which showed very low
recovery of photosynthetic quantum yield when rehydrated
in darkness (Figs 3a,b & 5a,b), exhibited dramatic increases
in quantum yield when they were re-illuminated for just 1 d
during rehydration (Fig. 5a,b). UTEX 318 showed almost
complete recovery of photosynthetic quantum yield following 5 d of illumination during rehydration (Fig. 5a,b,
hatched bars). In contrast, MLO301CT showed no recovery
from desiccation, no matter the rehydration treatment, and
SAG 2006 and UTEX 2457 were intermediate (Fig. 5a,b).
Upon re-illumination (after rehydration), the dramatic
increases in quantum yield observed for UTEX 318, UTEX
11 and the intermediate increase observed for UTEX 2457
coincided with declines in Fo of ~50% or more relative to
pre-illumination levels, whether algae had been desiccated
previously in the dark or under light (Fig. 6a,b). These
declines in Fo were accompanied by losses of pigmentation
easily visible to the naked eye 24 h after re-illuminating the
rehydrated algae (Fo was measured after overnight dark
adaptation of algae.). A contrasting pattern of gradual Fo
decrease during rehydration for MLO301CT and SAG 2006
was observed, particularly after desiccation in the dark
(Fig. 6a).These two algae did not exhibit easily visible losses
of pigmentation upon rehydration, although SAG 2006 did
quantum yield during rehydration of
aquatic (a–c) and desert (d–f) algae
exposed to various light intensities during
desiccation. Recovery following
desiccation for 24 h is shown in (a) and
(d); following 7 d is shown in (b) and (e),
or following 4 weeks is shown in (c) and
(f). Bars represent quantum yields of
rehydrated algae averaged over all
aquatic (a–c) or desert (d–f) isolates.
Error bars represent ⫾1 SD with n = 15
(aquatic) or n = 18 (desert). Data were
not collected for algae re-illuminated
following desiccation for 24 h (a,d). All
measurements were made following a
6–8 h period of dark adaptation. Key to
symbols: black bars = algae dried in
darkness, grey bars = upper bead layer of
algae dried under illumination, white
bars = lower bead layer of algae dried
under illumination.
exhibit a drop in Fo of nearly 50% upon re-illumination
when it had been desiccated in the light (Fig. 6b).
Patterns of recovery for desert algae dried under illumination were quite different from those for aquatic algae
(Fig. 5c,d). All desert algae dried under illumination for 4
weeks, and then rehydrated first in the dark, then with illumination, recovered a large proportion of the original
quantum yield (Fig. 5d). In all but one case (SEV3-VF49),
the majority of recovery occurred after light was applied
during rehydration (Fig. 5d, hatched bars). This behaviour
parallels that of aquatic alga UTEX 318 (Fig. 5b). In contrast, desert algae dried in the dark (Fig. 5c) recovered a
much higher proportion of the original quantum yield even
after just 1 h rehydration in the dark. Application of light
during rehydration was associated with further recovery
(Fig. 5c, hatched bars); however, the relatively large recovery of photosynthetic quantum yield attained by the desert
algae upon rehydration in darkness (Figs 3c,d; 4d–f & 5c)
made these increases less pronounced than those observed
in the aquatic algae (Figs 4b,c & 5a,b). Unlike in the aquatic
algae (Fig. 6a), Fo in dark-desiccated desert algae overall
remained much more constant during rehydration, even
when illumination was applied (Fig. 6c).
As noted previously, desert algae dried under illumination exhibited only partial recovery of quantum yield when
© 2007 The Authors
Journal compilation © 2007 Blackwell Publishing Ltd, Plant, Cell and Environment, 30, 1240–1255
1250 D. W. Gray et al.
Desert algae
Desiccated 4 weeks in darkness
Desiccated 4 weeks in darkness
0.6
0.4
0.2
0.0
Desiccated 4 weeks in full light
0.6
0.4
0.2
0.0
0.6
0.4
0.2
0.0
MLO301CT
SAG
2006
UTEX
2457
UTEX 11
Desiccated 4 weeks in full light
(d) 0.8
Quantum yield (Fv/Fm)
(b) 0.8
Quantum yield (Fv/Fm)
(c) 0.8
Quantum yield (Fv/Fm)
Quantum yield (Fv/Fm)
(a) 0.8
Aquatic algae
0.6
0.4
0.2
0.0
UTEX
318
BC2-1
CNP2VF25
24 h darkness
48 h darkness
Re-illumination 1 d
Re-illumination 5 d
Desert algae
(c)
1000
Fo (mV)
Fo (mV)
800
400
300
200
400
200
0
0
Desiccated 4 weeks in full light
Desiccated 4 weeks in full light
1000
(d)
500
800
400
Fo (mV)
Fo (mV)
600
100
300
200
600
400
200
100
0
photosynthetic quantum yield in BC2-1, CNP2-VF25 and
BC4-VF9 increased significantly (Fig. 5d, hatched bars), following a pattern similar to that exhibited by the aquatic alga
UTEX 318 (Fig. 5b). The swift recovery of quantum yield
when light was applied to BC2-1, CNP2-VF25 and
Desiccated 4 weeks in darkness
Desiccated 4 weeks in darkness
500
600
BC4VF9
1 h darkness
Aquatic algae
(b)
BC8-8
Pre-desiccation
rehydrated in darkness (Fig. 4d–f). Following 4 weeks of
desiccation under illumination, three of the desert algae
(BC2-1, CNP2-VF25 and BC4-VF9) exhibited no measurable recovery of quantum yield when rehydrated in darkness for up to 48 h (Fig. 5d). Yet, upon re-illumination,
(a) 600
SEV2VF1
Algal isolate
Algal isolate
Legend:
SEV3VF49
Figure 5. Recovery of photosynthetic
quantum yield following rehydration of
aquatic (a,b) or desert (c,d) algae
previously desiccated for 4 weeks in
darkness (a,c) or under full intensity
illumination of 130 mmol m-2 s-1 (b,d).
Columns represent average quantum
yield following 6–8 h dark adaptation.
Error bars represent ⫾1 SD with n = 3.
Key to symbols: black bars = predesiccation, dark grey = algae rehydrated
for 1 h in darkness, medium grey = algae
rehydrated for 24 h in darkness, light
grey = algae rehydrated for 48 h in
darkness, right hatched = algae
re-illuminated for 1 d, and left
hatched = algae re-illuminated for 5 d.
MLO301CT
SAG
2006
UTEX
2457
UTEX 11
UTEX
318
0
BC2-1
Algal isolate
Legend:
CNP2VF25
SEV3VF49
SEV2VF1
BC8-8
BC4VF9
Algal isolate
Pre-desiccaion
1 h darkness
24 h darkness
48 h darkness
Re-illumination 1 d
Re-illumination 5 d
Figure 6. Recovery of dark-adapted
steady-state fluorescence (Fo) following
rehydration of aquatic (a,b) or desert
(c,d) algae previously desiccated for 4
weeks in darkness (a,c) or under full
intensity illumination of 130 mmol m-2 s-1
(b,d). Columns represent average Fo
following 6–8 h dark adaptation. Error
bars represent ⫾1 SD with n = 3. Key to
symbols: black bars = pre-desiccation,
dark grey = algae rehydrated for 1 h in
darkness, medium grey = algae
rehydrated for 24 h in darkness, light
grey = algae rehydrated for 48 h in
darkness, right hatched = algae
re-illuminated for 1 d, and left
hatched = algae re-illuminated for 5 d.
© 2007 The Authors
Journal compilation © 2007 Blackwell Publishing Ltd, Plant, Cell and Environment, 30, 1240–1255
Photorecovery after desiccation of green microalgae 1251
Ecological differences
Comparing the impact of desiccation on the recovery of
quantum yield in desert algae with that of aquatic algae
shows that there is a very strong phylogenetically corrected
correlation (r = 0.9–0.95) between habitat of origin and the
capacity to recover quantum yield when rehydrated in
darkness (Fig. 7 & Table 3). These correlations were significant for all periods of desiccation. However, the strength
of this correlation declined markedly (r = 0.65) upon
re-illumination (Table 3), indicating that illumination can
ameliorate some of the damage caused by desiccation.
These patterns are clearly evident in Fig. 7a, which shows
that DDI is consistently higher in aquatic algae than in
desert algae, but that DDI declines in the aquatic algae
following re-illumination.
In addition to the damage incurred by the desert algae
upon desiccation, desert algae also lost an additional
25–55% of their initial aqueous quantum yield as a result of
being illuminated while dry. These reductions in quantum
yield due to illuminating dry algae (Fig. 7b) were comparable in magnitude to the reductions in yield due solely to
desiccation (Fig. 7a). In response to re-illumination, PDI of
the desert algae declined, exhibiting a response that was
similar to that observed for DDI in the aquatic algae, and
suggesting that re-illumination of rehydrated algae could
facilitate repair of damage caused by illuminating dry
desert algae.
In contrast, aquatic algae lost most of their initial
aqueous quantum yield following desiccation in darkness,
as indicated by very high DDI values (Fig. 7a). Because
little residual capacity for photosynthetic recovery was
present in aquatic algae following desiccation, illuminating
dry aquatic algae could only cause small additional losses in
quantum yield, and resulted in very low calculated PDI
values (Fig. 7b).
DISCUSSION
Unicellular green microalgae are common in microbiotic
crusts worldwide. Because they are very small (10–50 mm)
and rarely dominant, other photosynthetic organisms from
Desiccation damage index
(DDI)
(a)
(b)
Photodamage index
(PDI)
BC4-VF9 during rehydration was accompanied by declines
in Fo (Fig. 6d), and these declines in Fo occurred in conjunction with visibly noticeable losses of pigmentation that took
place upon re-ilumination. However, both the declines in Fo
and the loss of color in the desert algae were less severe
upon re-illumination than those observed in the aquatic
algae. Interestingly, these three desert algae (BC2-1, CNP2VF25 and BC4-VF9) were also the three desert taxa most
visibly bleached by desiccation in the light. When dried
under illumination, the three other desert algae (SEV2VF1, SEV3VF49 and BC8-8) exhibited a delayed and/or
low level of quantum yield recovery during rehydration in
darkness (Fig. 5d); re-illumination enhanced their recovery,
but Fo did not exhibit abrupt declines following
re-illumination (Fig. 6d).
1.0
0.8
0.6
0.4
0.2
0.0
1.0
Aquatic desiccated 24 h
Aquatic desiccated 7 d
Aquatic desiccated 4 weeks
Desert desiccated 24 h
Desert desiccated 7 d
Desert desiccated 4 weeks
0.8
0.6
0.4
0.2
0.0
1h
dark
24 h
dark
48 h
1d
5d
dark reillumination reillumination
Time after rehydration
Figure 7. Desiccation damage index (DDI) (a) and
photodamage index (PDI) (b) in desert (open symbols) and
aquatic (closed symbols) algae following rehydration in darkness
and following re-illumination of rehydrated algae. The DDI
represents the fraction of initial aqueous quantum yield lost as a
result of desiccation. The PDI represents the additional fraction
of initial aqueous quantum yield lost as a result of illumination.
Error bars represent ⫾1 SE with sample size n = 15–18. Symbols:
circles = algae dried for 24 h, squares = algae dried for 7 d,
triangles = algae dried for 4 weeks. X-axis labels correspond to
rehydration periods described in the text. Data were not
collected for rehydration intervals longer than 24 h in algae dried
for 24 h.
crusts, such as mosses, lichens and the cyanobacteria Microcoleus and Nostoc, have more often been the focus of physiological studies. Further, because desert green algae are
morphologically very similar to one another, their impressive diversity has only been revealed recently using molecular methods (e.g. Lewis & Lewis 2005), and exploration of
green algal contributions to crusts has focused on them as a
group, not at the species level. This impressive diversity has
been shown by Lewis & Lewis (2005) to be associated with
at least 14 evolutionarily independent transitions from
aquatic to desert habitats. This fact, coupled with the algae’s
simple coccoid morphology and their persistence in what
would appear to be very harsh environments, suggests these
algae offer an excellent opportunity to explore physiological tolerance and survival strategies that have evolved in
nature among unicellular eukaryotes making the transition
from water to land. By taking into account underlying
© 2007 The Authors
Journal compilation © 2007 Blackwell Publishing Ltd, Plant, Cell and Environment, 30, 1240–1255
1252 D. W. Gray et al.
phylogenetic relationships among algae, modern phylogenetic comparative methods aid in teasing out whether a
congruent pattern of occurrence in two characters (e.g. a
physiological trait and habitat) actually represents correlated evolutionary change in the two characters, or a chance
pattern of association (Martins 2004). Stronger evidence for
correlated evolution between characters is provided by
multiple independent coevolutionary events. Less convincing is a single evolutionary origin of an association between
characters, the presence of which in many species may be
easily explained by historical relationship. In our case,
having multiple data points, each representing an independent origin of desert organisms, ultimately will support a
more conclusive demonstration of correlated evolution of
physiological traits and a shift to the desert habitat.
The initial step must be to show that isolates of green
algae from deserts are not just transient visitors from
aquatic systems – they did not just blow in on the wind or
persist following a flood or rainstorm. Molecular evidence
suggests they are not transient aquatic visitors (Lewis &
Lewis 2005); now the physiological data presented here
further strengthen that conclusion. Desert green algae
were able to recover substantial photosynthetic quantum
yield within 1 h of rehydration (Figs 3c,d & 4d–f), particularly when desiccated in darkness (Figs 4d–f & 5c,d).
Aquatic algae were much more sensitive to desiccation
(Figs 3a,b; 4a–c; 5a,b; Tables 2 & 3). Interestingly, much of
the recovery achieved by the desert algae took place
within 1 h of rehydration (Figs 3c & 5c), a length of time
that is likely too short for the recovery to be explained by
massive synthesis of new proteins and construction of new
photosystems (Mattoo, Marder & Edelman 1989; Barber
& Andersson 1992). It suggests that the desert algae
possess well-developed mechanisms that protect their
photosynthetic machinery from desiccation-induced
damage. The rapid recovery of photosynthetic processes
observed in the desert algae parallels that seen in some
species of cyanobacteria (Harel et al. 2004) and mosses
(Proctor & Smirnoff 2000). In these organisms, recovery of
photosynthetic activity occurred within minutes of rehydration and did not require protein synthesis, leading to
the conclusion that recovery was due to the reassembly of
PSII from existing photosystem components that were
present and intact in the desiccated state (Proctor &
Smirnoff 2000; Harel et al. 2004). Determining whether
this is the case in desert green algae requires further
experimentation.
Despite their ability to withstand desiccation, the desert
algae examined in this study were susceptible to photodamage when dry. This result is interesting in that these
algae were exposed to rather low intensities of light
(130 mmol m-2 s-1), but they were isolated from desert crusts
where incident light would have approached full sun
(2000 mmol m-2 s-1) at the surface. When exposed to light
while they were desiccated, the desert algae recovered photosynthetic quantum yield more slowly upon rehydration
than they did when dried in darkness, and exhibited a dose
dependence of their response to illumination both in terms
of the intensity of incident light and the duration of illumination (Fig. 4d–f).
Not all desert algal species behaved similarly, however.
For BC2-1, CNP2-VF25 and BC4-VF9, exposure to light
during desiccation for 4 weeks led to the complete failure
to recover measurable photosynthetic quantum yield
even after 48 h of rehydration in darkness (Fig. 5d).
Re-illuminating these algae during rehydration stimulated a
dramatic increase in quantum yield (Fig. 5d) accompanied
by abrupt declines in Fo (Fig. 6d), which may reflect the
re-association or repair of photosystem components and the
establishment of a functioning system for dissipating
absorbed energy through the electron transport chain.
Heightened Fo has frequently been observed following stress
in a variety of photosynthetic organisms (Angelopoulos,
Dichio & Xiloyannis 1996; Briantais et al. 1996; Bartoskova,
Komenda & Naus 1999; Kouril et al. 2001) and is consistent
with a mechanism in which disruption of the electron transport chain reduces the potential for energy dissipation via
photochemistry, resulting in increased chlorophyll fluorescence. However, it is also of note that upon re-illuminating
the rehydrated desert algae, there was a visible (even to the
eye) loss of pigmentation in all samples, and the loss of color
was most pronounced in the three algae that exhibited the
lowest recovery of quantum yield in darkness (BC2-1,CNP2VF25 and BC4-VF9). An alternative explanation for the
declines in Fo upon re-illumination of rehydrated algae
noted earlier (Fig. 6d), then, may be that Fo declined as a
result of final photodestruction of chlorophyll in a subpopulation of already damaged or dead cells. The remaining
fluorescence signal, indicating improved quantum yield
upon re-illumination (Fig. 5d), would in this scenario be
derived from the surviving cells. Among those desert algae
that did recover quantum yield in darkness following longterm desiccation under illumination (SEV3-VF49, SEV2VF1 and BC8-8), the slow or delayed recovery they
exhibited (Fig. 5d) in comparison with dark dried samples
(Fig. 5c) was also consistent with a mechanism involving the
repair or construction of new photosystems.
Although aquatic algae rehydrated in darkness exhibited
little or no recovery of photosynthetic quantum yield
(Fig. 3a,b), quantum yield in a subset of these aquatic algal
species (UTEX 2457, UTEX 11, UTEX 318) recovered dramatically upon re-illumination (Fig. 5a,b). The concomitant
decline in Fo in these taxa (Fig. 6b) and the visible loss of
color noted after re-illumination parallel the patterns in
desert BC2-1, CNP2-VF25 and BC4-VF9 desiccated long
term in the light (Fig. 5d). Following rehydration in darkness, most of the cells in the UTEX 2457, UTEX 11 and
UTEX 318 samples apparently had retained large amounts
of chlorophyll, but they lacked functional PSII units able
to undergo charge separation (either dead or extremely
stressed cells). The presence of large amounts of chlorophyll would have resulted in the high Fo values observed in
the dark, but the lack of charge separation within PSII
would have resulted in low Fm values, little variable fluorescence and hence, low quantum yields (Krause & Weis 1991).
Upon re-illumination, this chlorophyll in dead or very
© 2007 The Authors
Journal compilation © 2007 Blackwell Publishing Ltd, Plant, Cell and Environment, 30, 1240–1255
Photorecovery after desiccation of green microalgae 1253
stressed cells likely was damaged, resulting in the overall
decline in Fo. However, at least some of the algal cells in
those samples of aquatic isolates appear to have survived
desiccation, and fluorescence from those survivors may
have dominated the high yield measurements 1 and 5 d
after re-illumination (Fig. 5b).
Ecological patterns and significance
Within their native habitat, desert crust algae must withstand numerous environmental stresses including periods
of intense desiccation, temperature extremes and intense
irradiance at the soil surface, potentially for months at a
time (Rosentreter & Belnap 2001). This capacity for rapidly
recovering photosynthetic ability would be advantageous in
allowing these desert algae to take advantage of brief
periods of water availability afforded by desert thunderstorms. Examining the cell morphology of these algae in
aqueous culture, while desiccated, and after rehydration
revealed no visible differences in cell morphology, suggesting that these algae withstand desiccation as vegetative cells
rather than as special resting stages or zygote spores.
In contrast to the behaviour of the desert algae, we
observed no photosynthetic recovery in closely related
aquatic algae rehydrated in darkness, and re-illumination
appeared to lead to recovery of only a sub-population of
cells from some aquatic taxa (Figs 5a,b & 6a,b). This suggests that the high degree of desiccation tolerance observed
in the desert algae is an adaptation to the desert environment rather than an ancestral characteristic of the evolutionary lineages to which the desert algae belong. This view
is supported by our comparative statistical analysis through
which we show a strong significant correlation between
habitat of origin and the ability to recover from desiccation
(Table 3). PGLS, TIPS and FIC all produced similar correlation coefficients in Table 3 because, with the suite of
species we used in the experiment, underlying lineagespecific traits were not driving results. Had all desert algae
been derived from a single transition from water to land by
one aquatic ancestor that possessed measurable desiccation
tolerance, then contrasting results from PGLS and TIPS
analyses would have indicated that the physiological traits
inherited by the related desert descendants were driving the
correlations between trait and habitat detected in the data.
Although the desert algae proved to be quite resistant to
desiccation, the degree to which they were impacted by
illumination (photodamage) is somewhat unexpected. The
desert algae examined in this study originated from crust
samples and clearly must be able to survive months of desiccation and illumination in crusts in the field. One possible
explanation for this pattern is that the desert green algae
may be able to acclimate to the higher light intensities
present under field conditions by synthesizing more photoprotective pigments, such as xanthophyll cycle pigments
(Bilger & Björkman 1990; Demmig-Adams & Adams 1992;
Gilmore 1997; Bukhov et al. 2001b; Masojidek et al. 2004).
Alternatively, the light sensitivity shown in this study may
indicate that in the field, these algae occupy micro-
environments within the crust where they are protected
from damaging light levels. The differential susceptibility to
illumination expressed by the taxa examined in this study
suggests there may be a complex spatial arrangement of
green algal species in the crust structured in response to the
attenuation of light with depth through the crust profile.
Vertical profiles of organisms are well-known characteristics
of biological crusts (Hu et al. 2003; Hoppert et al. 2004);
however, because many crust organisms lack readily discernable morphological features, most studies have focused on
the stratification of taxa grouped at higher taxonomic levels
rather than at the level of species. As noted earlier, the
problem of species identification is particularly acute for the
eukaryotic green algae. Determining the location of individual species within a crust will require the development of
molecular probes for in situ tagging of specific species.
Physical disturbances are well known to have deleterious
effects on crusts (Belnap & Eldridge 2001) and would represent an important threat to their green algal assemblages,
by potentially exposing light-sensitive algae inhabiting protected lower crust layers to high levels of illumination. This
enhanced illumination could lead to eventual loss of photosensitive taxa and reductions in algal biodiversity in the
disturbed crust fragments. In addition to potentially rendering these green algal communities susceptible to disturbance, their apparent sensitivity to illumination may have
important implications for designing remediation efforts
aimed at regenerating disturbed crusts. One technique that
has been used involves inoculating disturbed soil surfaces
with cultured crust organisms or a slurry made from soil
crust fragments (St. Clair, Johansen & Webb 1986; Belnap
1993). These inoculants have consistently enhanced the
rates of soil crust recovery following disturbance, and
cyanobacteria/green algae, mosses and lichens show
enhanced coverage and biomass in inoculated areas
(Belnap 1993; Belnap & Eldridge 2001). Lichens and
mosses also show enhanced species diversity (Belnap 1993;
Belnap & Eldridge 2001) in inoculated areas. Studies,
however, have not distinguished between the cyanobacterial and green algal components within crusts. Our results
suggest that spraying green algae cultured from desert
crusts directly onto soil surfaces as inoculants may prove
problematic for re-establishing these green algae given
their susceptibility to photodamage during desiccation.
ACKNOWLEDGMENTS
We thank S. Olm and D. Tyser for their valuable technical
assistance and P.O. Lewis for helpful discussions about phylogenetic comparative methods. We also thank two anonymous reviewers for helpful comments that greatly improved
the manuscript. This work was supported by a grant from
the National Aeronautics and Space Administration, Exobiology Program (EXB02-0042-0054) to L.A.L. and Z.G.C.
and a grant from the University of Connecticut Research
Foundation to Z.G.C. This is publication #10 from the
Center for Integrative Geosciences at the University of
Connecticut.
© 2007 The Authors
Journal compilation © 2007 Blackwell Publishing Ltd, Plant, Cell and Environment, 30, 1240–1255
1254 D. W. Gray et al.
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Received 25 January 2007; received in revised form 15 May 2007;
accepted for publication 29 May 2007
© 2007 The Authors
Journal compilation © 2007 Blackwell Publishing Ltd, Plant, Cell and Environment, 30, 1240–1255