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Carnegie Institution, Department of Plant Biology, 260 Panama Street, Stanford, California 94305,1 Department of Statistics, Sequoia Hall, 390 Serra Mall, Stanford University, Stanford, California 94305-40652
Received 4 August 2005/ Accepted 31 October 2005
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ABSTRACT |
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INTRODUCTION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Eukaryotic signal transduction pathways for sensing and responding to P starvation have been best studied in the yeast Saccharomyces cerevisiae. P-deficient S. cerevisiae cells increase their capacity for Pi uptake by expressing a high-affinity Pi transport system, mainly the Pho84p H+/Pi symporter, which operates at acidic pHs (3, 6). The Pho89p protein is a Na+/Pi cotransporter and may be utilized at more alkaline pHs (29). Pi that is covalently bound to organic molecules may be mobilized by cleavage by an acid phosphatase encoded by the PHO5 gene (reviewed by Vogel and Hinnen [50] and by Oshima [36]). There are many genes that are coregulated under P starvation conditions in S. cerevisiae, forming the PHO regulon (35). Changes in gene expression that accompany P starvation of S. cerevisiae are regulated by a system comprised of a cyclin (Pho80) and a cyclin-dependent kinase (Pho85). Under P-replete conditions, the kinase activity of Pho80/Pho85 catalyzes hyperphosphorylation of the Pho4 transcription factor, which is then excluded from the nucleus. When P becomes limiting, the cyclin-dependent kinase inhibitor Pho81 is activated, preventing Pho80/Pho85 kinase activity and leading to hypophosphorylation of Pho4 and its subsequent import into the nucleus. Once in the nucleus, Pho4 and Pho2 interact, populating the promoters of target genes and activating gene expression (reviewed by Lenburg and O'Shea [26]).
In photosynthetic eukaryotes, Pi is also critical for chloroplast function. Pi must be transported from the cytoplasm of the cell, across the double membrane of the plastid envelope, and into the chloroplast stroma, where it is used to satisfy the biosynthetic and energetic requirements of the organelle. Pi is needed for both DNA and RNA synthesis within the chloroplast, as the plastids contain their own genomes and ribosomes, and each chloroplast usually has multiple chromosome copies. Photosynthetic generation of ATP and the synthesis of phospholipids represent other critical chloroplast activities requiring Pi, and the phosphorylation of polypeptides of the photosynthetic apparatus modulates the properties of light-harvesting functions in response to changing environmental conditions (55). Furthermore, P limitation leads to depletion of the pool of phosphorylated intermediates in the pentose-phosphate cycle, which results in a marked reduction in photosynthetic carbon fixation (5, 22). Studies with Chlamydomonas reinhardtii have demonstrated that P and sulfur deprivation causes a similar loss of photosynthetic electron transport activity, the consequence of a combination of reduced photosystem II (PS II) abundance, accumulation of PS II QB nonreducing centers, an increase in nonphotochemical quenching, and an increase in the tendency of the cells to be in state II (53). These processes all redirect absorbed excitation energy away from PS II and linear electron transport. The observations described above support the idea that when nutrient availability limits the growth of the cell, a suite of mechanisms are mobilized to manage/dissipate excess absorbed light energy, reducing the potential of the system to generate reactive oxygen species which could cause severe cellular damage.
Little is known about the regulatory pathways by which plants and algae sense and respond to P deficiency. Genetic screens for C. reinhardtii mutants with a diminished Pi uptake capacity or reduced alkaline phosphatase activity during P-limited growth (46, 54) led to the discovery of the phosphorus starvation response 1 (PSR1) gene, which encodes a nucleus-localized factor with a MYB-like DNA binding domain and a coiled-coil protein-protein interaction domain (54). PSR1 is required for normal growth and the synthesis of alkaline phosphatase following starvation of the cells for P. Furthermore, the abundance of the PSR1 transcript and its protein product also increases during P starvation (54). Subsequent studies with Arabidopsis thaliana revealed that an analogous gene, phosphate starvation response 1 (PHR1), functions to elicit changes in root growth and morphology, anthocyanin production, and the activities of several P deficiency-inducible genes. These results suggest that the key regulators of the P deprivation responses in plants and C. reinhardtii (and perhaps other algae) are very similar, suggesting the possibility of common downstream signaling factors (42). However, it is not known whether PSR1 and PHR1 have dual functions, acting as both the sensor of intracellular Pi levels and the transcription factor that elicits altered gene expression. Recent findings have implicated an Arabidopsis SUMO E3 ligase, SIZ1, in the regulation of PHR1 activity (31), suggesting a multilevel control of plant responses to Pi deficiency.
In this study, we combine genome-based, molecular, and physiological approaches to examine P starvation responses in a wild-type C. reinhardtii strain and its isogenic psr1 mutant. Potential P deprivation-responsive genes were identified by an examination of genomic information and by cDNA-based microarray analyses. A number of new P deficiency-responsive target genes were identified, many encoding proteins that would enable the cells to better cope with excess absorbed excitation energy, limiting the extent of potential photodamage during P deprivation; these stress-related genes may be directly under PSR1 control. The importance of these genes was suggested by experiments in which elevated light levels caused a dramatic loss of viability of psr1 mutant, but not wild-type, cells during P deprivation.
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MATERIALS AND METHODS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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150 rpm) at 25°C, with continuous
illumination (40 µmol photons m–2
s–1). P-free Tris-acetate (TA) medium was prepared
by substituting 1.5 mM potassium chloride for potassium phosphate, as
described by Quisel et al.
(39). To starve cells for
P, cells were initially grown in TAP medium, harvested by
centrifugation (4,000 x g, 5 min), washed twice with
TA medium, and resuspended to a final cell density of 1 x
106 cell/ml in TA medium. P-starved cultures were maintained
under the same growth conditions as the unstarved cells, except that
all cultures were sampled 4, 12, 24, and 48 h after the
initiation of P deprivation. For RNA preparation, cells were harvested
by centrifugation in a model HN-SII clinical centrifuge (International
Equipment Co.) at three-fourths the maximum speed (5,800 rpm)
for 10 min, and cell pellets were immediately frozen in liquid
N2 and stored at –80°C. Strains for
high-light-intensity experiments were grown in liquid and solid TA and
TAP media and in TA medium supplemented with 10 µM
glucose-1-phosphate (Sigma Chemical Co., St. Louis, MO) at
27°C, with continuous illumination at 600 to 800 µmol
photons m–2
s–1. RNA preparation. Cell pellets were maintained in liquid N2. Each pellet was simultaneously thawed and homogenized in 8 ml of homemade Trizol reagent (Invitrogen Co., Carlsbad, CA) by continuously vortexing the suspension; the homogeneous suspension was then incubated at room temperature for at least 10 min. Chloroform (1.6 ml) was added to the suspension, and the tubes were shaken vigorously for 1 min and incubated at room temperature for an additional 2 to 3 min. Phase separation was achieved by centrifugation at 12,000 x g for 15 min, and 0.5 volume (3 to 4 ml) of 0.8 M sodium citrate-1.2 M NaCl was added to the aqueous phase, followed by the addition of 0.5 volume (compared to the initial volume) of isopropanol to precipitate nucleic acids. Precipitations were done at room temperature for 10 min, and the nucleic acid precipitate was collected by centrifugation at 12,000 x g for 10 min. Nucleic acids were washed with 8 ml of 75% ethanol, dried, and dissolved in diethyl pyrocarbonate-treated, double-distilled water. Poly(A) RNA was prepared using a MicroPoly(A) Pure small-scale mRNA purification kit (Ambion, Austin, TX) and was quantified fluorimetrically using a RiboGreen RNA quantification kit (Molecular Probes, Eugene OR) and a Tecan SPECTRAFluor fluorimeter (Zurich, Switzerland).
Preparation of fluorescent microarray probes. The incorporation of Cy3-dUTP and Cy5-dUTP (Amersham Pharmacia Biotech, Piscataway, NJ) was performed as described by Zhang et al. (56), with the following modifications: (i) 200 to 500 ng of poly(A) RNA was used as the template for labeling reactions; (ii) the deoxynucleoside triphosphate mixture contained 1 mM dTTP and 2.5 mM (each) of dATP, dCTP, and dGTP; (iii) labeling reactions were incubated at 42°C for 30 min, 45°C for 20 min, and 50°C for 30 min; (iv) incorporation of the fluorescent dyes into cDNA was not examined; (v) the reference RNA was isolated from log-phase cells of wild-type strain CC-125 prior to P deprivation; and (vi) samples prepared from P-starved (4, 12, 24, and 48 h) CC-125 and P-replete (0 h) and P-starved (4, 12, 24, and 48 h) psr1-1 cells were compared to the reference sample.
Preparation, hybridization, washing, and scanning of microarrays.
The printing and storage of array
slides (v1.1) were performed as described by Zhang et al.
(56) for the "2.7
k array," except that 3,079 different cDNA probes were printed
on each slide. Additional details about the array are given at
http://nostoc.stanford.edu/jeff/lab/chlamyarray/index.html
and in Table
1 and Table S1 in the supplemental material.
Slides were hydrated by holding them above a 90°C water bath
for 3 s and were then immediately dried on a 100°C hot
plate for 5 s. DNAs were cross-linked by UV radiation at 300
mJ in a UV Stratalinker 1800 (Stratagene, La Jolla, CA).
Prehybridization was performed by baking the slides at 65°C for
10 min, followed by a 30- to 60-min incubation at 50°C in
prehybridization buffer containing 3x SSC (1x SSC is
0.15 M NaCl plus 0.015 M sodium citrate), 0.1% sodium dodecyl sulfate,
and 0.1 mg/ml bovine serum albumin. Slides were washed with water and
then isopropanol, each for 2 min at room temperature, and then dried by
centrifugation for 5 min in a SpeedVac Plus, model SC210A (Savant,
Holbrook, NY). Hybridization, washing, and scanning of the slides were
performed as described by Zhang et al.
(56), except that the
hybridization solution was prepared by mixing 20 µl of
hybridization buffer with 20 µl of the labeled probes, and the
fluorescence images from 12 complete array sets (three slides per time
point, with four copies of each cDNA per slide) were analyzed. Two
independent sets of RNA samples were used for array analyses; a dye
swap was performed using one of the RNA
samples.
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0.05 and a multiple testing correction adjustment using the
method of Benjamini and Hochberg
(2). Gene identification. To identify specific genes in the C. reinhardtii genome, either sequence information from GenBank for previously isolated and characterized genes or sequence information from cDNAs or genomic DNAs from other organisms (mainly A. thaliana and cyanobacteria) was used to perform BLAST alignments against C. reinhardtii genomic (http://genome.jgi-psf.org/chlre2) and expressed sequence tag (EST) database (http://www.biology.duke.edu/chlamy_genome/search.html) sequences. Candidate orthologous and paralogous predicted proteins were aligned with each other and evaluated using CLUSTALW (48); identity shading was performed using GeneDoc software (33).
Quantitative real-time PCR.
Isolated
total RNA was treated with RNase-free DNase I (Ambion Inc., Austin, TX)
and then extracted with phenol-chloroform to isolate the RNA. Real-time
quantitative PCRs (qPCR) for all genes except PTOX1,
PTOX2, GAP1, and DHAR were performed by
using 0.1 µg of DNase-treated total RNA and an iScript One-Step
RT-PCR kit with SYBR green (Bio-Rad Laboratories, Hercules, CA). The
amplifications were performed using either an iCycler IQ or Chromo4
real-time PCR thermocycler (Bio-Rad Laboratories, Hercules, CA), with
the following cycling conditions: (i) 50°C for 30 min for cDNA
synthesis, (ii) 95°C for 5 min to denature reverse
transcriptase, and (iii) 40 to 42 cycles of 95°C for 15 or
30 s and 60°C for 30 s, with fluorescence
detection after the 60°C annealing/extension step. Melting
curve analysis was performed on all PCRs to ensure that single DNA
species were amplified, and product sizes were verified by agarose gel
electrophoresis. Discrete products were not obtained with the one-step
system for the PTOX1, PTOX2, GAP1, and
DHAR transcripts; consequently, two-step qPCR analysis was
performed for these transcripts. For cDNA synthesis, 1 µg of
DNase I-treated total RNA was reverse transcribed by using a
Superscript II kit (Invitrogen, La Jolla, CA) as described by the
manufacturer. qPCR was performed using either a DyNAmo Hot Start SYBR
green qPCR kit (MJ Research Inc., Waltham, MA) or IQ SYBR green
supermix (Bio-Rad Laboratories, Hercules, CA) and analyzed with the
Mx3005P qPCR system (Stratagene, La Jolla, CA) or the Chromo4 system
(Bio-Rad Laboratories, Hercules, CA). Cycling conditions included an
initial incubation at 95°C for 10 min followed by 40 to 45
cycles of 94°C for 10 s, 55°C to 60°C
for 15 s, and 72°C for 10 to 15 s. The
relative expression ratio of a target gene was calculated based on the
2–
CT
method (28), using the
average cycle threshold (CT) calculated from
duplicate measurements. Relative expression ratios from two independent
experiments are reported. The CBLP gene was used as a control
gene, and each primer was designed by Primer3 software to distinguish
the different isozymes.
Primers for real-time PCR were as follows: CBLP, forward (5'-CTTCTCGC CCATGACCAC-3'), reverse (5'-CCCACCAGGTTGTTCTTCAG-3'); PHOX, forward (5'-TTCCGTTTCCGTTCTCTGAC-3'), reverse (5'-CCCTGCATCTT GTTCTCCAG-3'); PTB1, forward (5'-GCTTCTGCATCGAGCTGTC-3'), reverse (5'-TTCCCTCCATCAGACCAATG-3'); PBT2, forward (5'-TGTGCTCGTGCATTCTCTTC-3'), reverse (5'-CCCTTGGTGAACACGAAGTAA-3'); PTB3, forward (5'-GCGAGAACTCCTACGTCCTG-3'), reverse (5'-AGTCCAGTCGCTGTTGGAAG-3'); PTB4, forward (5'-CCAACCTGGCAATCTACATG-3'), reverse (5'-GCCTTGTTCGAGTCCCAGT-3'); PTB5, forward (5'-CTCAACCCAGTTGGCAATTTACTTT-3'), reverse (5'-GCCTTGTTCGAGTCCCAGT-3'); PTB6, forward (5'-TTCTTCCTGAACCGCATCTT-3'), reverse (5'-GCTCTTCGCTCCCTTGTAGA-3'); AOX1, forward (5'-TGGGCTCCATAGCTTCGGTTC-3'), reverse (5'-GTGGCAGCGGGCTTGTGG-3'); AOX2, forward (5'-AGCATGTCAGGGCTTGTGT-3'), reverse (5'-GGCATCCTTGACATGACTCAG-3'); PTOX1, forward (5'-GACAGCCTGGACGCAGTG-3'), reverse (5'-GCAGTTGTTGATGAGCTTGC-3'); PTOX2, forward (5'-GGACAAGGCGTTGGAGAAC-3'), reverse (5'-CCGACACAGACAGCTCCAG-3'); GAP1, forward (5'-CTGGCGTGTTCACTGACATC-3'), reverse (5'-TTGACGCCCATCACATACAT-3'); GAP2, forward (5'-TTGAGTCTGTGGCGTACCTG-3'), reverse (5'-CCCTCAATGGACTTGCAGTT-3'); GAP3, forward (5'-CCGTGTCCTCCAAGTCTGG-3'), reverse (5'-CAGCAGCGCAGGAAGTTG-3'); HYD1, forward (5'-GCGACTGGTTCTGTGTGGAC-3'), reverse (5'-CATTGGATTGTCCCACTCG-3'); HYD2, forward (5'-GCGAGTGGTTCTGTGTGAGC-3'), reverse (5'-CTGGTCCCAGTCACTGTCG-3'); GPLV1, forward (5'-GTGGACAGCGTCAACAACAT-3'), reverse (5'-ACTCCGCCTGGTCCTTGTAG-3'); GPLV2, forward (5'-ACTTCGGCTGGGAGGATTAC-3'), reverse (5'-CCACTCCGTCTGGTTCTTGT-3'); DHAR, forward (5'-GCCCAAGCTATACCATGCTG-3'), reverse (5'-TAGTCCACATGCTGCCACTC-3').
Immunological detection of PHOX. C. reinhardtii cell cultures were concentrated by centrifugation and adjusted to a chlorophyll concentration of 1 mg/ml. The chlorophyll concentration was estimated according to the absorbance at 652 nm of 5-µl aliquots of cell suspension that were extracted in an 80% acetone-20% methanol mixture. Whole cell proteins equivalent to 10 µg of chlorophyll were solubilized in 1x Laemmli sample loading buffer, separated by discontinuous sodium dodecyl sulfate-polyacrylamide gel electrophoresis using a 7.5% polyacrylamide resolving gel, and transferred to polyvinylidene difluoride membranes (Pierce Biotechnology, Rockford, IL). Membranes were blocked for 1 h in a solution containing 5% dry milk and then incubated for at least 2 h with a 1/500 dilution of anti-PHOX antiserum (18) in 1% milk. A 1/3,000 dilution of alkaline phosphatase-conjugated goat anti-rabbit antiserum in 1% milk was used as the secondary antibody, and the alkaline phosphatase colorimetric reaction was performed as described by Sambrook et al. (44).
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RESULTS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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2-fold (Fig.
1A), 2.5-fold (Fig.
1B), and 3-fold
(Fig. 1C), specifically in
CC-125, in the psr1-1 mutant, or in both strains. We
chose to concentrate on those transcripts that exhibited a change of
2.5-fold or more following exposure of cells to P deprivation
conditions. Table 1 shows
the genes encoding the transcripts that changed by 2.5-fold in
either wild-type cells or psr1-1 cells; to be
included in the table, this change had to be observed for at least one
point during the time course of P deprivation, and it had to pass
Student's t test for significance. The transcripts of 235
genes exhibited changes of 2.5-fold in wild-type cells during
P deprivation, corresponding to somewhat less than 10% of the genes on
the array. Of these, transcripts of 152 genes differentially
accumulated in wild-type cells, but not in the psr1-1
mutant; such genes are candidates for being directly regulated by PSR1.
Furthermore, there were 29 transcripts in psr1-1
cells, but not in wild-type cells, that changed in abundance during P
deprivation. The genes encoding these transcripts most likely respond
to secondary stress conditions that result from the inability of
psr1-1 cells to acclimate properly to P deficiency.
An additional 83 transcripts differentially accumulated in both CC-125
and the psr1-1 mutant during P deprivation, with the
majority (53) being
regulated in the same direction (either up or down) in the two strains,
suggesting that PSR1 is not involved (or has minimal involvement) in
the control of these genes. There was one transcript that increased by
>2.5-fold in CC-125 and decreased in the
psr1-1 mutant; conversely, there were 30 transcripts
that increased by 2.5-fold in the psr1-1 mutant and
decreased in the wild-type strain. These responses may also be primary
or secondary responses to PSR1 production in the cell.
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A second set of six genes that may be under PSR1 control encode potential "electron valves," which are enzymes that may serve to protect the cell against oxidative damage resulting from overexcitation of the photosynthetic and respiratory electron transfer chains. Genes in this category encode AOX1, an isoform of a mitochondrial alternative oxidase, PTOX1, a terminal oxidase present in the plastid, and HYD2, an iron hydrogenase that can serve as an alternative electron acceptor for PS I. Interestingly, the transcripts for a plastid-targeted isoform of the glyceraldehyde-3-phosphate dehydrogenase (GAP; Calvin cycle enzyme) gene, GAP1, and for a starch phosphorylase gene, GPLV, also increase in a PSR1-dependent manner. Increasing carbon fixation coupled with the synthesis and storage of starch is another way that the cell can eliminate excess reductant generated by photosynthetic electron transport and diminish the probability of producing reactive oxygen species.
There are also 26 transcripts that increase in a PSR1-independent manner during P deprivation; the levels of these transcripts change to a similar extent in wild-type cells and the psr1-1 mutant following the imposition of P deprivation. A number of genes encoding proteases and chaperones, listed under the heading "proteolysis/stress-related proteins" in Table 1, behave in this fashion. While we cannot formally rule out the possibility that genes which are regulated similarly in both the wild-type and psr1 mutant strains may respond to the centrifugation and washing steps that precede the induction of P deprivation, in many cases elevated levels of the transcripts from these genes were observed at earlier time points following P starvation in the psr1-1 mutant than in wild-type cells. Also, many of the responses were sustained over the 48-h time course of P deprivation. This suggests that these responses are neither induced by mechanical stress nor under PSR1 control but that the management of P resources (e.g., scavenging P from external and internal stores) in the mutant may be less effective than that in wild-type cells, which in turn can result in more rapid manifestation of PSR1-independent P deprivation responses. The levels of many transcripts encoding proteins required for photosynthesis also declined to similar extents in both strains. Overall, while a distinct subset of genes that respond to P deprivation conditions appears to be controlled by PSR1, a similar number of genes that are differentially expressed during P starvation are regulated by mechanisms that do not involve this transcription factor.
Genes potentially regulated by P availability based on genomic sequences. To analyze the expression of C. reinhardtii genes involved in the acquisition of Pi from the environment (specific responses), we surveyed the draft genome sequence for genes encoding potential secreted phosphatase enzymes and Pi transporters. A single gene encoding a potential phosphatase and multiple genes encoding potential Pi transporters were identified. Changes in the levels of the transcripts from these genes were examined using qPCR following exposure of cells to P starvation.
(i) PHOX, a calcium-dependent alkaline phosphatase.
Using TBLASTN, we identified a gene on
scaffold 86 of the C. reinhardtii version 3 genome sequence
with strong similarity (E value = 0.0) to the PHOX alkaline
phosphatase gene of Volvox carteri
(18), as shown in the
alignment in Fig.
2. PHOX is likely to be the abundant, inducible, extracellular alkaline
phosphatase of C. reinhardtii that was characterized by Quisel
et al. (39). This enzyme
has a molecular mass of 190 kDa, and its activity was
demonstrated to be Ca2+ dependent and to peak in the
basic pH range. The gene model on scaffold 86
(fgenesh1_pg.C_scaffold_86000013) has 57%
identity with the PHOX gene of V. carteri (Fig.
2). A sequence gap on the
genome scaffold coincides with a region close to the C-terminal end of
the predicted C. reinhardtii protein that is missing in the
alignment with V. carteri PHOX. Furthermore, no other
significant matches to PHOX were identified in the genome. These
results suggest that C. reinhardtii may have a single
PHOX gene homolog and that this single gene is interrupted by
a sequence gap in the draft genome
sequence.
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As shown in Table 2, for wild-type cells (CC-125) transcripts encoding many of the predicted transporters increased after 24 h of P deprivation. Significant increases in PTB2, PTB3, PTB4, and PTB5 mRNAs were observed for wild-type cells after 24 h of P deprivation, with little increase or a declining abundance in the psr1-1 mutant subjected to the same conditions. These results demonstrate that PSR1 is involved in controlling the abundance of these transcripts and that this control is probably exerted at the level of transcription (since PSR1 is a MYB domain-containing transcription factor). Interestingly, PTB2-PTB3 and PTB5-PTB4 are contiguous, with the transcript from the proximal gene of each pair (PTB2 and PTB5) being most strongly elevated after 24 h of P starvation. No significant increase in the level of PTB1 mRNA was observed during P stress, which is in agreement with the report of Kobayashi et al. (24), and the PTB6 transcript could not be detected (data not shown).
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The qPCR analysis demonstrated
that PHOX transcript levels markedly increase by 24
h of P depletion (Table
2). This increase is under
the control of PSR1, and the extent of the increase may reflect the
very low basal level of the transcript present in P-replete cells. To
confirm that the C. reinhardtii PHOX gene encodes the major
inducible alkaline phosphatase activity, immunoblot analysis was
performed on whole-cell extracts from Pi-replete versus
P-starved wild-type and psr1-1 mutant strains with
antiserum raised to a V. carteri PHOX peptide (Fig.
5) (18). The immunoblot
revealed a polypeptide of 190 kDa that cross-reacts with
anti-PHOX antibodies; this polypeptide was observed in extracts of
wild-type cells starved for P (but not in extracts of cells grown on
complete medium), and the band was not detected for the
psr1-1 strain under any conditions tested. The
cross-reacting band was not detected on a control blot without the
anti-PHOX antibody (data not shown), excluding the possibility
that residual alkaline phosphatase activity remained in the C.
reinhardtii protein extract after the proteins were transferred to
the membrane (since an alkaline phosphatase color reaction was used to
detect the secondary antibody). Together, these data strongly suggest
that C. reinhardtii PHOX encodes the major inducible
alkaline phosphatase activity of C.
reinhardtii.
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5- to 10-fold) in the psr1-1 strain,
possibly reflecting some contribution of PSR1 to the maintenance of
basal levels of HYD expression. PSR1-dependent regulation was
also observed for two genes encoding the starch phosphorylase homologs
GPLV1 and GPLV2. Based on both microarray and qPCR data, the
GPLV1 transcript was elevated four- to ninefold in wild-type
cells after 24 h of P deprivation but remained approximately
constant in the psr1-1 mutant during P deprivation.
Similarly, the GPLV2 transcript increased threefold in
wild-type cells and remained constant in the psr1-1
strain during P deprivation. Therefore, in most of the cases that we
have examined, where transcript abundance for one member of a gene
family is potentially regulated by PSR1, the regulation of the other is
independent of PSR1.
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Sensitivity of psr1-1 mutant to high light intensity. The discovery that the mRNAs of a variety of genes with a putative electron valve function increased during P deprivation in a PSR1-dependent manner raised questions concerning the role that PSR1 might play in controlling general responses to P deficiency. Therefore, we compared the sensitivities of the psr1-1 mutant, wild-type cells, and a PSR1-complemented strain to elevated light intensities during P deprivation (Fig. 6). In moderate light (40 µmol photons m–2 s–1) on TA medium supplemented with a limiting amount of a hydrolyzable Pi source (10 µM glucose-1-phosphate), the psr1-1 mutant survived but grew more slowly than wild-type cells or the complemented strain and also arrested at a lower cell density. In contrast, when the three strains were exposed to high light intensities (700 to 800 µmol photons m–2 s–1) for 3 days during P deprivation, the psr1-1 mutant became bleached and died, while little loss of viability occurred for wild-type cells and the complemented mutant (Fig. 6A). Viability staining of cultures grown in liquid TA medium under high light intensities confirmed that 80 to 90% of psr1-1 cells died by day 6 of P depletion, whereas 80% of the wild-type cells and the complemented cells survived these conditions (Fig. 6B).
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DISCUSSION |
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In initial studies of psr1 mutants, no increased sensitivity to moderate light intensities was observed, and accordingly, PSR1 was not considered to play a role in protecting cells against photodamage. However, the microarray data demonstrated that a class of genes encoding potential "electron valves" appeared to be activated during P starvation in a PSR1-dependent manner. These genes encode enzymes that assist in the dissipation of potentially harmful species that may accumulate as a consequence of photosynthetic and respiratory electron transfer reactions under conditions that might restrict the use of products generated by these reactions. Microarray analyses identified transcripts from 210 genes that exhibited differential accumulation during P deprivation of threefold or greater in the wild-type and psr1-1 strains (Fig. 1). This represents approximately two-thirds the number of transcripts that exhibited a threefold change during sulfur starvation (56), although some transcripts are common between the two sets of data. The commonality may reflect responses to general intracellular stress conditions that may be generated by both types of nutrient deprivation.
Pi-scavenging genes. A total of 10 potential Pi transporters were identified in the C. reinhardtii genome (Fig. 3 and 4); at least six are differentially expressed during P deprivation. The transcripts for four of the PTB and one of the PTA genes increased in abundance in P-starved cells. Increased levels of these transcripts appeared to require PSR1. It is not clear which of these transporters (or groups of transporters) contributes most significantly to the high-affinity Pi uptake observed in P-deprived cells. In S. cerevisiae, which grows best in low-pH environments, the major secreted phosphatase, PHO5, has an acidic pH optimum, and the dominant high-affinity transporter is the Pho84 H+/Pi symporter (PTA type) (3, 6). The S. cerevisiae Pho89 Na+/Pi cotransporter (PTB type) may play a less significant role in Pi uptake, as it operates best in alkaline conditions (29). However, the situation may be very different for C. reinhardtii, which thrives in neutral or basic environments and secretes a predominant alkaline phosphatase during P deprivation (39). This may, in part, explain the bias towards activation of the PTB genes in C. reinhardtii; like Pho89, the PTB enzymes would be expected to display superior Pi uptake at an alkaline pH. Currently, there is no definitive evidence regarding the cellular location of the PTB and PTA proteins, but TargetP v1.01 predicts with high confidence that all of the PTB proteins are routed along the secretory pathway (Table 4), making these strong candidates for components of a plasma membrane (or vacuolar membrane)-associated Pi starvation-inducible Pi uptake system. Based on both microarray and qPCR analyses, PTB2 and PTB5 transcripts increase in abundance >20- and several hundredfold, respectively, following exposure of cells to P deprivation conditions (Tables 1 and 2). No change in Pi uptake characteristics was observed with a ptb1 mutant of C. reinhardtii (15), implying that the gene product is not essential for the elevated Pi uptake observed when cells begin to experience P limitations. Congruent with this result, the PTB1 transcript level does not respond significantly to P deprivation conditions. Kobayashi et al. (24) have suggested that PTB1 may be involved in intracellular Pi transport. The paired genomic clusters of the PTB2/PTB3 and PTB4/PTB5 genes, the high sequence identity between the members of the pairs, and the presence of numerous, scattered PTB gene fragments in close proximity suggest that this gene family has undergone a recent expansion. The different patterns of transcript abundance in response to P limitation defined for the individual members of this gene family suggest that the encoded proteins may play divergent roles in Pi uptake and distribution within the cell.
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The C. reinhardtii PHOX gene encodes a homolog of the PHOX protein of V. carteri, and the level of mRNA encoding this protein increases dramatically during P deprivation (Table 2). Immunological analyses using V. carteri PHOX antibodies suggest that this protein represents the previously characterized secreted, Ca2+-dependent alkaline phosphatase of C. reinhardtii (39).
C. reinhardtii also maintains internal stores of polyphosphate in cytosolic organelles, akin to the polyphosphate bodies of fungi and the acidocalciosomes found in trypanosomatids and apicomplexan parasites (43). Although nothing is known about the synthesis and regulation of polyphosphate in C. reinhardtii, it is interesting that the mRNA encoding VCX1, a potential vacuolar H+/Ca2+ antiporter (Table 1), also increased during P deprivation. In yeast, this protein is responsible for controlling cytosolic Ca2+ levels in the presence of high extracellular concentrations of Ca2+ (30). This finding raises the possibility that the VCX1 protein serves to maintain vacuolar sequestration of the Ca2+ that is released from polyphosphate bodies in the vacuole as Pi is enzymatically generated during P deprivation-triggered polyphosphate degradation.
Stress-related gene expression. A number of transcripts encoding stress-related proteins increased during P starvation in both the wild-type strain and the psr1-1 mutant; the levels of these transcripts do not appear to be under PSR1 control. These transcripts include those encoding two putative 22-kDa chloroplast-localized heat shock proteins, a glutathione S-transferase, an E3 ubiquitin ligase, and two protease homologs (Table 1, proteolysis/stress-related proteins). Chaperones and proteases may be required to sequester and eliminate misfolded or damaged proteins (9, 13). While most of these transcripts only significantly increased at the final time point following elimination of P from the growth medium (48 h) in the wild-type strain, they became elevated by 12 h of P deprivation in the psr1-1 mutant. These results suggest that while both strains show a generalized stress response, the mutant senses more extreme stress conditions at earlier times after the elimination of P from the medium; this more rapid response may reflect the inability of the mutant to scavenge Pi from both intracellular and extracellular resources.
A number of physiological responses have been shown to be elicited by P starvation. With respect to photosynthetic activity, there is a depletion of PS II reaction centers, accumulation of QB nonreducing centers, increased nonphotochemical quenching, and transition of the energy transfer characteristics of the antenna chlorophyll from state I to state II. These characteristics are indicators of excess excitation of the photosynthetic apparatus and suggest that the photosynthetic electron transport chain is hyperreduced, even under conditions of moderate light intensity (53). Gene expression analysis supports this idea, since, in addition to the P starvation-responsive chaperones and proteases, we observed increases in some transcripts that were also elevated at high light intensities (21). For example, there was an increase in transcripts encoding the granule-bound starch synthase I (STA2), which may help to relieve excess excitation by stimulating starch synthesis, and LHCSR2, a protein related to light-harvesting complex (LHC) antenna proteins that may have a photoprotective function (Table 1) (21, 41). The transcripts for these proteins are also elevated during sulfur deprivation (56). Although the kinetics of increase in STA2 mRNA were similar for wild-type cells and the psr1-1 mutant, the increase in transcript abundance was observed earlier in psr1-1 mutant cells than in wild-type cells (12 h versus 24 h). Furthermore, the increased transcript levels were sustained in the mutant cells, even up to 48 h; at 48 h, the mRNA level in wild-type cells dropped to below the 2.5-fold threshold (Table 1). These results again suggest that the signal transduction pathway that regulates some of the deprivation-induced genes becomes active more rapidly and that the response is sustained longer following the initiation of P starvation of psr1-1 than that of wild-type cells.
Electron valves and radical-scavenging enzymes. In addition to potential stress-related transcripts that increase in both wild-type cells and psr1-1 mutant cells, another set of transcripts encoding proteins implicated in photoprotection is elevated exclusively in wild-type cells during P deprivation. These photoprotective proteins may act as "electron valves" that serve to drain electrons from the photosynthetic and mitochondrial electron transfer chains, decreasing the potential dangers associated with hyperreduction of electron carriers and the generation of triplet chlorophyll molecules. An enhanced capacity for alternative respiration in iron-starved C. reinhardtii has been reported previously (51), and mutants of tobacco plants with a defective alternative oxidase were unable to sustain normal respiratory rates upon P deprivation (37). We observed increased transcript levels from the alternative oxidase gene AOX1 in wild-type cells starved for P for 24 h. In contrast, the psr1-1 mutant exhibited a transient increase in AOX1 mRNA after 4 h of P deprivation, but this increase was not maintained. Increased alternative oxidase activity during P starvation may perform a number of functions, including preventing the accumulation of reactive oxygen species in the mitochondria, allowing continued flux of metabolites through the tricarboxylic acid cycle for generating organic carbon "backbones," and controlling carotenoid biosynthesis and the rate of ATP formation under conditions in which the availability of Pi may be limiting (7, 23, 25). Increased levels of the AOX1 transcript were previously reported during S starvation (56), and upon transfer of cells from growth on ammonium to growth on nitrate (1). Surprisingly, AOX1 is the only transcript encoding a known mitochondrial protein that was significantly upregulated in these experiments, suggesting that the control of mitochondrial processes during P deprivation may be regulated largely by posttranscriptional mechanisms.
Increased levels of transcripts encoding PTOX1, a putative plastid terminal oxidase, were also observed in wild-type P-starved cells, but not in the psr1-1 mutant. The PTOX1 protein may represent the terminal oxidase activity responsible for chlororespiration (8, 38). This terminal oxidase may relieve redox pressure associated with photosynthetic electron transfer by withdrawing electrons from plastoquinol and combining them with O2. PTOX1 is a homolog of a plastid terminal oxidase in plants, encoded by the IMMUTANS gene, which is also implicated in carotenoid synthesis (23), raising the possibility that PTOX1 induction during P starvation in C. reinhardtii might play a role in the production of photoprotective carotenoids.
A third potential electron valve that may increase during P deprivation in cells with active PSR1 is the Fe hydrogenase, encoded by HYD2 (HYDB). The C. reinhardtii hydrogenases catalyze the transfer of electrons from PS I to water to produce H2 gas under anaerobic/microaerobic conditions, and they are believed to help reoxidize the plastoquinol pool when the cytoplasm of the cell is highly reducing, especially during anaerobic or hypoxic growth (14, 16, 20). The hydrogenase activity might increase plastoquinol oxidation during P starvation, relieving the block on linear electron transfer and helping to prevent the production of reactive oxygen species. The activation of HYD2 gene expression might be a by-product of the production of a microaerobic environment within cells as oxygen evolution declines during P deprivation and respiration consumes oxygen faster than it is being produced. However, were this the case, it would be expected that HYD1 expression would be induced as well, but this was not observed in either microarray or qPCR experiments. HYD2 mRNA abundance is much greater in wild-type cells than in the psr1-1 mutant (compare transcript levels in Tables 1 and 3), suggesting that PSR1 plays a role in maintaining expression of this gene (or the conditions required for its expression).
As discussed above, C. reinhardtii shows increased STA2 mRNA expression during P deprivation; the STA2 protein is involved in the synthesis of starch. Elevating starch synthesis can help control the accumulation of excess reductant in the cell and limit the production of reactive oxygen species. However, P starvation of wild-type cells also leads to elevated mRNA for both GPLV, encoding a starch phosphorylase, and an alpha amylase gene. Both of these gene products are involved in the conversion of starch to sugar. Therefore, the starved cells may also have an increased capacity to actively degrade storage starch, which could be important for maintaining the flux through the starch synthesis pathway. Finally, the transcript for dehydroascorbate reductase also increases in P-starved wild-type cells but not in the psr1-1 mutant. This enzyme is involved in recycling of ascorbate, an antioxidant that plays an important role in scavenging reactive oxygen species in the chloroplast. Together, the results presented above demonstrate that increases in mRNAs for proteins that have an electron valve function and that help the cells manage an elevated redox state appear to be normal aspects of the P starvation responses of C. reinhardtii. Although a causative relationship between the activation of these pathways and resistance to high light intensities remains to be established, it is intriguing that conditions of high light intensity are lethal to the psr1 mutant and that the transcripts from many of these genes do not significantly increase in the psr1-1 mutant.
Photosynthesis. A number of genes represented on the array encoding proteins with photosynthetic functions appear to be differentially expressed during P starvation. While down-regulation of PS II function and abundance has been demonstrated previously (53), we did not observe changes in mRNA levels for nuclear genes encoding structural subunits of PS II during P starvation. Instead, transcripts from several LHCA genes, encoding components of the LHC of PS I (LHC I), and PSAE and PSAK genes, encoding subunits of PS I, declined in both P-starved wild-type cells and psr1-1 mutant cells. A reduced level of the CHL27 (CRD1) transcript was also observed. CHL27 encodes a component of the chlorophyll biosynthetic enzyme Mg-protoporphyrin IX monomethylester cyclase (49) and has been reported to play a role in controlling the interaction between the PS I reaction center and LHC I. Therefore, reduced levels of CHL27 protein would be expected to result in a weaker energetic connection between the LHC I antennae and core PS I (32). The transcripts for proteins that are minor subunits of the cytochrome b6/f complex, i.e., PETO and PETN, also appear to decline in wild-type cells, with the PETM and PETC transcripts showing similar trends (although the level of the PETM and PETC mRNAs only declined by 50% over the time course [see Table S1 in the supplemental material]). Indeed, transcripts encoding all PSB, PSA, and PET proteins represented by elements on the array declined in both wild-type cells and the psr1-1 strain.
Aberrant gene expression in the psr1-1 mutant. Twenty-nine transcripts exhibited increased accumulation in psr1-1 but not wild-type cells during P starvation (Fig. 1). The increased levels of these transcripts may be a consequence of the more extreme stress conditions experienced by the psr1-1 mutant than by wild-type cells since the mutant would not be able to properly acclimate as P levels in the medium declined. The aberrant metabolic status of the mutant strain may also trigger specific signaling pathways in addition to more general stress response pathways. Increased levels of transcripts encoding phytoene synthase, a key regulatory enzyme in carotenoid biosynthesis, and homogentisic acid geranylgeranyl transferase, an enzyme required for tocopherol biosynthesis, may reflect the cell's response to elevated reducing conditions and a greater tendency to form reactive oxygen molecules, both of which are associated with nutrient deprivation conditions (34). Interestingly, the psr1-1 mutant also exhibited increased mRNAs for a subset of genes associated with sulfur acquisition and assimilation (Table 1, sulfur metabolism). This finding may reflect a change in the energy status of the cell and the inability of the mutant to efficiently take up and activate SO42–. However, since we observed an increase in transcript levels for only a subset of the genes associated with sulfur-deficient conditions, it is possible that there is a more specific link between the P and S deprivation pathways. Understanding these links may help us to decipher the networks of interactions among stress responses and the hierarchy of responses observed.
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ACKNOWLEDGMENTS |
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We thank Chung-Soon Im, Wirulda Pootakham, Steve Pollock, Nakako Shibagaki, and Jeff Shrager for technical assistance and helpful comments on the manuscript and Armin Hallmann and Sabeeha Merchant for the kind gift of antibodies. We also thank the Joint Genome Institute for providing access to a prerelease of version 3 of the Chlamydomonas draft genome sequence.
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FOOTNOTES |
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Supplemental material for this article may be found at http://ec.asm.org/.
This is Carnegie Institution publication no. 1725.
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