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ENVIRONMENTAL STRESS AND ADAPTATION

The LPB1 Gene Is Important for Acclimation of Chlamydomonas reinhardtii to Phosphorus and Sulfur Deprivation^1,[w]

Department of Plant Biology, Carnegie Institution of Washington, Stanford, California 94305 (C.-W.C., J.L.M., A.R.G.); and Department of Biochemistry and Biophysics, University of California, San Francisco, California 94143–0448 (D.W.)

	ABSTRACT

TOP ABSTRACT RESULTS DISCUSSION MATERIALS AND METHODS LITERATURE CITED

 
Organisms exhibit a diverse set of responses when exposed to low-phosphate conditions. Some of these responses are specific for phosphorus limitation, including responses that enable cells to efficiently scavenge phosphate from internal and external stores via the production of high-affinity phosphate transporters and the synthesis of intracellular and extracellular phosphatases. Other responses are general and occur under a number of different environmental stresses, helping coordinate cellular metabolism and cell division with the growth potential of the cell. In this article, we describe the isolation and characterization of a mutant of Chlamydomonas reinhardtii, low-phosphate bleaching (lpb1), which dies more rapidly than wild-type cells during phosphorus limitation. The responses of this mutant to nitrogen limitation appear normal, although the strain is also somewhat more sensitive than wild-type cells to sulfur deprivation. Interestingly, depriving the cells of both nutrients simultaneously allows for sustained survival that is similar to that observed with wild-type cells. Furthermore, upon phosphorus deprivation, the lpb1 mutant, like wild-type cells, exhibits increased levels of mRNA encoding the PHOX alkaline phosphatase, the PTB2 phosphate transporter, and the regulatory element PSR1. The mutant strain is also able to synthesize the extracellular alkaline phosphatase activity upon phosphorus deprivation and the arylsulfatase upon sulfur deprivation, suggesting that the specific responses to phosphorus and sulfur deprivation are normal. The LPB1 gene was tagged by insertion of the ARG7 gene, which facilitated its isolation and characterization. This gene encodes a protein with strong similarity to expressed proteins in Arabidopsis (Arabidopsis thaliana) and predicted proteins in Oryza sativa and Parachlamydia. A domain in the protein contains some similarity to the superfamily of nucleotide-diphospho-sugar transferases, and it is likely to be localized to the chloroplast or mitochondrion based on programs that predict subcellular localization. While the precise catalytic role and physiological function of the putative protein is not known, it may function in some aspect of polysaccharide metabolism and/or influence phosphorus metabolism (either structural or regulatory) in a way that is critical for allowing the cells to acclimate to nutrient limitation conditions.

Many soil-dwelling organisms have evolved a suite of responses that enable them to survive extended exposure to conditions of low P availability. These responses can be divided into the P-specific responses and the general (or global) stress responses (Quisel et al., 1996; Davies and Grossman, 1998; Wykoff et al., 1998). P-specific responses facilitate mobilization and acquisition of Pi from both extracellular and intracellular sources (Quisel et al., 1996; Shimogawara et al., 1999), while the global responses allow for long-term survival by coordinating cellular metabolism and growth with nutrient availability.

Starvation of most plants and microorganisms for P results in elevated phosphatase activities (Juma and Tabatabai, 1988; Lee, 1988; Goldstein et al., 1989; Duff et al., 1991a, 1991b; Sachay et al., 1991; Tadano et al., 1993; Nakazato et al., 1997; Trull et al., 1997), which helps to mobilize internal Pi stores (e.g. from phytic acid and inositol phosphates in vacuoles) and to generate free Pi from external organic phosphates. Increased RNase production during P deprivation (Jost et al., 1991; Taylor and Green, 1991; Yen and Green, 1991; Löffler et al., 1992, 1993; Taylor et al., 1993; Bariola et al., 1994) may also boost Pi availability by hydrolyzing Pi from various RNA reservoirs (Green, 1994). Furthermore, the roots of specific plants such as rape (Brassica napus) may exude organic acids during P deprivation that lower the rhizosphere pH (Hoffland et al., 1992), facilitating solubilization and increased availability of soil Pi for transport into roots (Lopez-Bucio et al., 2000). P limitation can also result in the elevation of the V_max (but not necessarily alter the K_m) for Pi uptake into roots by triggering accumulation of high-affinity Pi uptake systems (Bieleski, 1973; Drew et al., 1984; Lefebvre and Clarkson, 1984; Schjorring and Jensen, 1984; McPharlin and Bieleski, 1987; Jungk et al., 1990; Lefebvre et al., 1990; Bieleski and Laluchli, 1992; Shimogawara and Usuda, 1995). While genes encoding plant Pi transporters have been isolated (Muchhal and Raghothama, 1999; Raghothama, 2000a, 2000b; Smith et al., 2000; Baek et al., 2001), the acquisition of Pi by plants can be complex because of interactions of plant roots with soil organisms such as mycorrhizal fungi (Rhizoctoni solani; Yao et al., 2002). Proliferation of mycorrhizal fungi helps mobilize soil Pi for uptake into the roots (Pfleger and Linderman, 1994; Rosewarne et al., 1999; Yao et al., 2002).

Morphological and biochemical changes in plants and microbes may also accompany P limitation. In P-starved plants, the root-to-shoot ratio may increase in conjunction with decreased lateral-root proliferation and the development of longer, more densely packed root hairs. P-starved black mustard (Brassica nigra) exhibits increased fixed carbon storage and elevated accumulation of lipids and phenolics. Furthermore, enzymes that serve a bypass function (Duff et al., 1989; Theodorou et al., 1991), helping to eliminate the need for phosphorylated adenylates, which can be limiting during P starvation, may replace other enzymes in metabolic pathways (e.g. phosphoenolpyruvate phosphatase may replace pyruvate kinase).

Like vascular plants, Chlamydomonas reinhardtii synthesizes a number of phosphatases (Lien and Knutsen, 1973; Matagne and Loppes, 1975; Loppes, 1976; Matagne et al., 1976; Patni et al., 1977; Loppes and Deltour, 1981; Quisel et al., 1996) and exhibits a change in the kinetics of Pi transport (Shimogawara et al., 1999) during P limitation. The cells synthesize an abundant extracellular phosphatase activity, with 85% to 90% associated with a 5' nucleotidase having an alkaline pH optimum and a monomeric, apparent molecular mass of 190 kD (Dumont et al., 1990; Quisel et al., 1996). Starved cells also display increased Pi uptake potential. The V_max for Pi increases by well over 10-fold and the K_m for Pi decreases from approximately 10 µM to between 0.1 and 0.3 µM (Shimogawara et al., 1999). Other genes that appear to be activated during P starvation encode enolase, pyruvate-formate lyase, and two ribosomal proteins, although some of these genes may also be induced under other growth-limiting conditions (Dumont et al., 1993).

Currently, little is known about regulatory elements that control P limitation responses in plants and algae. A number of mutants of C. reinhardtii have been isolated that exhibit aberrant phosphatase activity during P deprivation (Loppes and Matagne, 1973; Matagne and Loppes, 1975; Loppes, 1976; Loppes et al., 1977; Loppes and Deltour, 1981; Wykoff et al., 1999), and we have identified the altered gene in one such mutant. This gene, designated PSR1 (Phosphate Starvation Response), encodes a protein of 762 amino acids that is synthesized at elevated levels during P deprivation (Wykoff et al., 1999) and is a member of the MYB-CC subfamily of regulatory elements (Pabo and Sauer, 1992). A psr1 null mutant is unable to synthesize extracellular phosphatase in response to Pi deprivation and is aberrant for the other specific responses to P deprivation that were tested. Recently, PSR1 was shown to be important for the development of some general responses and survival of the cells during P deprivation under elevated light conditions (J. Moseley, C.-W. Chang, and A.R. Grossman, unpublished data). A similar regulatory element designated PHR1 was identified in Arabidopsis (Arabidopsis thaliana; Rubio et al., 2001).

In this study, we used a genetic screen to identify C. reinhardtii mutants that die more rapidly than wild-type cells during P limitation conditions. Two allelic mutants, low-P bleaching (lpb1-1) and lpb1-2, were isolated and the LPB1 gene was identified and sequenced. The predicted gene product has limited similarity to a UDP-Glc pyrophosphorylase (UDPGPase), which is involved in the synthesis or degradation of polysaccharides.

	RESULTS

TOP ABSTRACT RESULTS DISCUSSION MATERIALS AND METHODS LITERATURE CITED

 
We previously isolated the PSR1 gene, which regulates the specific responses of C. reinhardtii to P starvation (Wykoff et al., 1999) but also affects at least some of the general responses to P deprivation (J. Moseley, C.-W. Chang, and A.R. Grossman, unpublished data). In this article, we report the identification of a C. reinhardtii mutant that was still able to perform specific P-deprivation responses, but was screened for rapid death on solid medium following the imposition of P deprivation; this mutant, designated lpb1, is defined by two alleles. As shown in the colony assay (Fig. 1A), the lpb1 mutant bleaches and dies rapidly on solid medium lacking P. The loss of pigmentation or bleaching of the cells parallels cell death (data not shown). Quantification of viability of the mutant and wild-type strain in liquid medium is presented in Figure 2A. Approximately 30% of lpb1 mutant cells died after 2 d of P starvation, and after 5 d, less then 20% viable mutant cells remained. In contrast, essentially 100% of the wild-type cells were viable after 5 d of P deprivation (Fig. 2A).

View larger version (48K): [in this window] [in a new window]   Figure 1. Cell viability and phosphatase activity. A, Viability of the lpb1 mutant and wild-type cells after 1 week of growth on solid medium devoid of phosphate. The arrow denotes the bleached lpb1 colony. B, Qualitative analysis of phosphatase activity secreted by wild-type cells (wt), the lpb1 mutant, and the psr1 mutant. The psr1 strain represents a negative control; the mutant is unable to synthesize alkaline phosphatase in response to P deprivation (Wykoff et al., 1999). Cells were streaked onto TA solid medium prior to spraying the plates with the colorimetric phosphatase substrate 5-bromo-4-chloro-3-indolyl-P, which turns blue after phosphate is cleaved from the molecule by alkaline phophatase. The plates were allowed to develop for 2 h before recording the results.

View larger version (15K): [in this window] [in a new window]   Figure 2. Viability of wild-type cells, mutant strains, and the complemented lpb1 mutant in medium lacking P, S, N, or P plus S. A, Viability of wild-type cells (CC-125), lpb1, and lpb1 complemented with cos7 (containing the wild-type LPB1 gene) following P deprivation in liquid medium. B, Viability of wild-type cells and lpb1 following S deprivation in liquid medium. C, Viability of wild-type cells and lpb1 following N deprivation in liquid medium. D, Viability of wild-type cells and lpb1 following both P and S deprivation in liquid medium. A viability dye was used to distinguish viable from nonviable cells. This was confirmed by monitoring cell viability by plating the cells onto solid TAP medium (see "Materials and Methods" for further details).

While survival of the lbp1 mutant is specifically compromised during P or S starvation, this strain can still perform P and S deprivation-specific acclimation responses. The lpb1 mutant still exhibits the synthesis of an extracellular alkaline phosphatase activity (blue halo around the plated cells) during P starvation (Fig. 1B), and the accumulation of alkaline phosphatase activity in liquid cultures of the mutant cells following elimination of P from the medium is about 60% of that of wild-type cells (Fig. 3A). The mutant strain also develops relatively high levels of arylsulfatase (ARS) activity during S deprivation (Fig. 3B). Furthermore, both the mutant and wild-type strains display increased accumulation of transcripts encoding the PHOX phosphatase (Fig. 4A), the PTB2 phosphate transporter (Fig. 4B), and the regulatory element PSR1 (Fig. 4C) after 12 h of P deprivation. These transcripts were previously shown to increase during P deprivation (Wykoff et al., 1999; J. Moseley, C.-W. Chang, and A.R. Grossman, unpublished data). The high level of PHOX transcript at 24 h after the initiation of P deprivation of wild-type cells relative to the lpb1 mutant may reflect a rapid turnover of the transcript when the mutant cells begin to die.

View larger version (24K): [in this window] [in a new window]   Figure 3. Measurements of alkaline phosphatase and ARS activities following exposure of wild-type cells and the lpb1 mutant to P and S deprivation. A, Quantification of alkaline phosphatase activity associated with wild-type cells (CC-125) and lpb1 (lpb1) following P or P and S deprivation. B, Quantification of ARS activity associated with wild-type cells (CC-125) and lpb1 (lpb1) following S or S and P deprivation. Alkaline phosphatase and ARS activities were determined by monitoring the hydrolysis of -nitrophenol phosphate and -nitrophenol sulfate, respectively, at OD410. The absorbance readings were converted to nmol PNPP/mg chl/min.

View larger version (17K): [in this window] [in a new window]   Figure 4. Change in mRNA levels of P deprivation-responsive genes following exposure of wild-type cells and the lpb1 mutant to medium lacking P. A to C, The PHOX (A), PTB2 (B), and PSR1 (C) mRNA levels over a period of 24 h of P deprivation. Levels of the mRNA were quantified by qPCR, and the threshold cycle number or CT value was normalized to the amount of constitutively expressed CBLP transcript. The fold change was calculated relative to the amount of transcript at time 0 for wild-type cells.

View larger version (18K): [in this window] [in a new window]   Figure 5. Viability of P-starved wild-type cells (CC-125) and the lpb1 mutant (lpb1) in the presence of DCMU. DCMU was added to 1 µm DCMU, which eliminated all photosynthetic O2 evolution (data not shown).

View larger version (85K): [in this window] [in a new window]   Figure 6. The LPB1 gene and predicted gene product. A, Structure of LPB1 cDNA and genomic clones. The black rectangles represent the exons, while the lines represent introns. The rectangle at the beginning of the gene sequence represents the part of the cDNA that was deduced by RNA ligase-mediated RACE (see "Materials and Methods"). The site of insertion of the ARG7 gene is marked above the genomic sequence. B, Alignment of predicted LPB1 polypeptide sequence with homologous polypeptides encoded by an expressed sequence of Arabidopsis and hypothetical proteins of O. sativa and Parachlamydia sp. Amino acids in a blackened background represent identical or conserved residues for all four sequences, while amino acids in a gray background represent identical or conserved residues for two or three of the four sequences.

LPB1 mRNA was not detected by RNA-blot hybridization using RNA isolated from wild-type cells grown under either nutrient-replete or P starvation conditions. However, we were able to detect low-level constitutive expression from the gene using quantitative PCR (qPCR; data not shown); the level of RNA did not appear to change during P deprivation.

	DISCUSSION

TOP ABSTRACT RESULTS DISCUSSION MATERIALS AND METHODS LITERATURE CITED

 
In this study, we identified a C. reinhardtii mutant, lpb1, that still performs the specific P-deprivation responses, as evaluated by the induction of alkaline phosphatase activity and the accumulation of mRNAs encoding the PTB2 phosphate transporter, the PHOX phosphatase, and the regulatory protein PSR1, but dies much more rapidly than wild-type cells during P starvation. The mutant also accumulates ARS activity following the initiation of S starvation, but was more sensitive than wild-type cells to conditions of S depletion. The accumulation of the ARS and alkaline phosphatase activities were somewhat lower in the mutant than in the wild-type strains, but the reduced levels may reflect the inability of the cells to survive extended periods of P or S stress relative to the wild-type strain. When lpb1 mutant cells were starved simultaneously for P and S, the development of ARS activity, but not alkaline phosphatase activity, was observed. It was previously demonstrated that acclimation of C. reinhardtii to S deprivation occurs more rapidly than acclimation to P deprivation. Generally, it takes longer for cells to respond to P than to S deprivation; this is exemplified by the less rapid decline in photosynthetic O₂ evolution during P deprivation, as was previously reported (Wykoff et al., 1998). Therefore, the more rapid response of the cells to S deprivation leads to domination of S-deprivation over P-deprivation responses. However, the dual-stress conditions did enable the mutant to survive for a longer time than either of the single-stress conditions, suggesting that there is some combination of S and P-deprivation responses occurring in the cells; the combination of these responses must have a synergistic role in maintaining cell viability.

One of the general responses to nutrient deprivation in C. reinhardtii is a marked decline in photosynthetic activity (Wykoff et al., 1998), and it was previously demonstrated that this decline was critical for sustaining the viability of the cells during nutrient deprivation (Davies et al., 1996). Therefore, the death of lpb1 during P deprivation may be a consequence of the inability of the mutant strain to properly modulate photosynthetic electron flow. However, the loss of photosynthetic activity during P deprivation is similar in the two strains (data not shown), and while DCMU, an inhibitor of photosynthetic electron flow, did delay of the death of lpb1 to some extent, it did not entirely rescue the viability phenotype. These results suggest that there is some other cause(s) for the death of the lpb1 mutant during P deprivation.

Genes encoding proteins similar to the predicted LPB1 polypeptide were identified in other organisms; one is an expressed protein of Arabidopsis and the other two are hypothetical proteins, one in O. sativa and the other in Parachlamydia sp. (UWE25). Furthermore, these sequences have domains with some similarity to sequences of UDP-acetylhexosamine pyrophosphorylase and UDPGPase, enzymes that catalyze the formation of UDPGlcNAc and UDPG, respectively. Indeed, LPB1 appears to be part of a larger family of proteins that includes UDPGPase, UDP-acetylhexosamine pyrophosphorylase (Peneff et al., 2001), and Suc phosphate synthase (Eimert et al., 1996). The UDPGlcNAc and UDPG sugar donors are key metabolites required for glycoprotein and glycolipid synthesis (UDPGlcNAc) as well as for the synthesis of Suc and various polysaccharides associated with carbon storage, cell wall structure, and protein modification (UDPG; Kleczkowski, 1994; Kleczkowski et al., 2004). Furthermore, in Arabidopsis, both expression from the UGP gene (gene encoding UDPGPase) and UDPGPase activity were shown to be up-regulated by conditions leading to P deprivation (Ciereszko et al., 2001; Repetto et al., 2003); the activity may be involved in optimizing the use of P resources (by generating pyrophosphate that could be hydrolyzed by pyrophosphatases to Pi) in times of need.

A number of observations suggest that it is unlikely that LPB1 has either UDP-acetylhexosamine pyrophosphorylase or UDPGPase activity. UDPGPase is 97% to 99.6% identical among mammalian proteins and 33% to 54% identical between mammalian and nonmammalian proteins (Flores-Diaz et al., 1997). Gene models with much greater conservation with the nucleotide-diphospho-sugar transferases are found in the draft C. reinhardtii genome (http://genome.jgi-psf.org/chlre2/chlre2.home.html; model C_40043 is the best match to UDPGP, and model C_100106 is the best match to UDP-acetylhexosamine pyrophosphorylase). The identity between LBP1 and potential UDPGPase homologs of C. reinhardtii and other organisms, including those of plants, is only 8% to 12% over the entire protein sequence and 25% to 29% over the regions that align among the proteins (Supplemental Fig. 1). Furthermore, there are three clearly defined, conserved motifs of UDPGPase (Bork and Koonin, 1996; Flores-Diaz et al., 1997). These motifs are defined as regions of 30 amino acids, in which at least 11 amino acids are strictly conserved in all homologs (Flores-Diaz et al., 1997). LPB1 has 8/24 identical amino acids for conserved motif 1 and 9/21 identical amino acids for conserved motif 2 (Fig. 7). Also, LPB1 and its Arabidopsis, O. sativa, and Parachlamydia sp. putative homologs lack the five conserved Lys residues that were previously shown to be important for substrate binding or catalysis (Katsube et al., 1991; Kazuta et al., 1991). Finally, UDPGPase in both plant and animal systems occurs almost exclusively in the cytosol, with activity sometimes associated with Golgi membranes (Simcox et al., 1977; Azzar et al., 1978; Nishimura and Beevers, 1979; Persat et al., 1983, 1984; Kleczkowski, 1994). Based on programs designed to predict subcellular locations of nuclear-encoded polypeptides, LPB1 appears to be localized in an organelle. Predictions from a variety of different localization programs are presented in Table I. For C. reinhardtii, LPB1 is predicted to localize to the chloroplast or mitochondrion, depending on the prediction tool used. In contrast, the prediction programs place the Arabidopsis LPB1-like protein in the chloroplast. Since different prediction tools used for these analyses are based on different algorithms, it is highly probable that LPB1 of C. reinhardtii and Arabidopsis are organelle localized. Frequently, there are difficulties in predicting whether a precursor polypeptide of C. reinhardtii is targeted to the mitrochondrion or chloroplast, probably because the algorithms used have not been trained extensively enough with polypeptides previously shown to localize to each of the organelles. It is also worth noting that the predicted cleavage sites for the LPB1 polypeptides of C. reinhardtii and Arabidopsis are in the same position with respect to the sequence alignment (the cleavage site of LPB1 predicted by MitoProtII is at amino acid 86, while the cleavage site of the Arabidopsis protein predicted by ChloroP is at amino acid 72). The lack of a targeting peptide in UWE25 of Parachlamydia (but present on the predicted sequences of Arabidopsis, O. sativa, and C. reinhardtii) is apparent from the alignment shown in Figure 6B. UWE25 is an endosymbiotic Chlamydial species that is thought to have diverged from the Chamydiaceae about 700 million years ago. Interestingly, a number of putative plant genes with significant similarity to UWE25 have been localized to the chloroplast, suggesting cyanobacterial anscestry (Horn et al., 2004). Similarly, there is only limited similarity between LPB1 and UDP-acetylhexosamine pyrophosphorylase.

View larger version (44K): [in this window] [in a new window]   Figure 7. Sequence similarity between two conserved domains of UDPGPases from human (Swissprot Q07131), Caenorhabditis elegans (NCBI accession no. U58727), barley (Hordeum vulgare; NCBI accession no. X91347), Arabidopsis (NCBI accession no. BAB10518), and gene model C_40034 from the draft C. reinhardtii genome, with the similar domains of C. reinhardtii LPB1 and the putative LPB1 homolog in Arabidopsis (locus id, At3g56040).

	MATERIALS AND METHODS

TOP ABSTRACT RESULTS DISCUSSION MATERIALS AND METHODS LITERATURE CITED

 

Strains and Growth Condition

The wild strain of Chlamydomonas reinhardtii used in these studies was CC-125. The lpb1 mutant was generated in the CC-425 background by transformation with pJD67. Selected transformants were backcrossed to CC-125 five times to ensure genetic homogeneity of the two strains. The standard growth medium was Tris-acetate phosphate (TAP; Harris, 1989), and liquid cultures were grown in Erlenmeyer flasks with gentle agitation on a rotary shaker (120 rpm) and illuminated at 80 µmol photon m^–2 s^–1 from cool-white fluorescent lamps at 27°C. To achieve P deprivation, cells in mid-logarithmic phase of growth (1–3x10⁶ cells/mL) were pelleted by centrifugation (3,000g, 5 min), washed twice with TAP medium lacking P (TA; Quisel et al., 1996), and then resuspended in TA to 1x10⁶ cells/mL. This protocol was modified when performing viability measurements; the cells were resuspended in either TA or TAP medium at 2.5x10⁴ cells/mL to allow for a longer periods of sustained growth. To prepare TA solid medium, we used 0.5% agarose instead of 1.2% agar (Shimogawara et al., 1999). To impose S and N starvation on the cells, TAP lacking sulfate (chloride salts were substituted for the sulfate salts) and TAP lacking ammonium (potassium chloride and molybdic acid were substituted for ammonium chloride and ammonium molybdate, respectively) liquid medium was used for washing the cells, in a manner similar to that described for achieving P deprivation.

Insertional Mutagenesis and Screening for Low-P-Bleaching Mutants

The plasmid pJD67, harboring the ARG7 gene encoding arginosuccinate lyase (Davies et al., 1994), was linearized by digestion with HindIII and transformed (Kindle, 1990) into the arg auxotrophic strain CC-425. Transformants that could grow on medium lacking Arg were spread at low density on solid TAP medium containing low Pi (10 µM) and grown for 7 d (Shimogawara et al., 1999). Those transformants that rapidly bleached were subjected to a secondary screen on solid TAP medium to eliminate pigment mutants and on high-salt medium lacking acetate (Harris, 1989) to eliminate those mutants that were defective for photosynthesis.

Viability and Activity Assays

Cells were counted in a hemocytometer, and cell viability was determined by staining with a solution of 0.0125% phenosafranin, 0.0125% methylene blue, 2.5 mM potassium phosphate, and 2.5% ethanol, similar to the assay previously described by Davies et al. (1996). Dead cells stained blue, whereas live cells excluded the dyes and remained green. The production of alkaline phosphatase activity by colonies growing on solid medium was evaluated by spraying colonies with an aqueous solution of 10 mM 5-bromo-4-chloro-3-indolyl-P (X-Pi; Sigma-Aldrich, St. Louis). Alkaline phosphatase activity in liquid cultures was measured as previously described (Shimogawara et al., 1999), except that the reaction was incubated at 37°C for 15 min. ARS activity was measured as described by de Hostos et al. (1988), except that the reaction was incubated at 37°C for 15 min.

In Vivo Measurements of Oxygen Evolution

Light-dependent O₂ evolution and dark respiration were measured with a Clark-type oxygen electrode (Hansatech, King's Lynn, UK) and reported by the Oxygraph system, as previously described (Wykoff et al., 1998).

Transformation and Complementation of lpb1

The lpb1 mutant was generated by introduction of the ARG7 gene (Wykoff et al., 1999) into CC-425 (cw15, nit1, arg). The ARG phenotype and the introduced ARG7 gene cosegregated with the lpb1 phenotype (death during P deprivation), suggesting that the inserted pJD67 DNA was responsible for sensitivity of the mutant to P deprivation. Plasmid rescue (Davies et al., 1996) was used to isolate DNA flanking the inserted vector; genomic DNA of the transformant was digested with SacI, and the fragments generated were self-ligated, transformed into Escherichia coli, and transformants selected for kanamycin resistance. A 300-bp region of the DNA flanking the vector sequence was amplified by PCR and used to screen a wild-type cosmid library (Purton and Rochaix, 1994; Zhang et al., 1994). Three cosmids were isolated, all of which shared identical restriction fragment profiles. The lpb1 mutant strain (nit1 cw15) was cotransformed with pMN24 (NIT1) and the identified cosmids containing the putative LPB1 gene, transformants were selected for growth on nitrate, and each of the transformants was screened for growth characteristics on TA plates; several of the cotransformed strains showed growth characteristics that were similar to those observed for the wild-type strain. The cosmid was able to rescue the lpb phenotype even after it was digested with EcoRI, KpnI, or XbaI. These enzymes were used to identify fragments of the insert DNA harboring the gene responsible for rescuing the mutant phenotype; a 9-kb XbaI-EcoRI complementing fragment was further characterized.

Isolation of LPB1 cDNA

Based on information generated by the C. reinhardtii genome project (http://genome.jgi-psf.org/chlre2/chlre2.home.html), we identified four 5' end-truncated LPB1 cDNA clones. Because the longest cDNA clone was 2.2 kb and did not contain the full-length sequence, RNA ligase-mediated RACE (GeneRacer kit, Invitrogene, Carlsbad, CA) was used to recover a full-length cDNA sequence. Total RNA was extracted from P-starved cells for 2, 4, 8, 12, and 24 h and treated with DNase I at 37°C for 1 h. The protocol described by the manufacture was followed except that we did not dephosphorylate the nucleic acid fragments by calf intestine phosphatase and, because the primers used had high-GC content, the reverse transcription was performed at 42°C for 30 min and then 50°C for 20 min. Furthermore, because the lpb1 transcript was not very abundant, nested PCR was necessary to obtain a 1.9-kb cDNA fragment containing the 5' region of the gene.

qPCR

qPCR was performed in the Lightcycler (Roche Applied Science, Indianapolis) using the SYBR green I RNA amplification kit. Total RNA was extracted from C. reinhardtii cells deprived of P for 4, 8, 12, 24, and 48 h (Shrager et al., 2003) as described previously (Wykoff et al., 1999), and polyA RNA was purified (polyA extraction kit, Ambion, Austin, TX), quantified using the ribogreen RNA quantification kit (Molecular Probes, Eugene, OR), and 10 ng of polyA RNA was used for each real-time qPCR reaction. The reaction mixture also included carrier RNA (RNA from bacteriophage MS2). The primers used for the CBLP transcript were 5'GTCATCCACTGCCTGTGCTT and 5'CCTTCTTGCTGGTGATGTTG. An intron is positioned between the latter two primers, and amplification with these primers would detect the presence of contaminating genomic DNA (based on amplicon sizes). The primer pairs used for qPCR were 5'CTAGAATGGAAACTGGTTGATG and 5'GTTACACGGAGAAAGCTATCA, 5'CAACAGCAGCAACAAGAGCA and 5'ATCACCGAAGTCAAAGTCCC, and 5'-TTCCGTTTCCGTTCTCTGAC-3' and 5'-CCCTGCATCTTGTTCTCCAG-3' for PTB2, PSR1, and PHOX, respectively. In total, three pairs of primers were used for amplification of the LPB1 transcript: (1) 5'GCAGTTGCAGTGAAGGAAAAGG and 5'TGTTCCTACAGCTCCCCTATGC, (2) 5'TGATGTACCCCAATTACTGCCA and 5'CAAATGTAFCACCTCTCAGCGA, and (3) 5'CCGGCTTCATCTTCCTGTTCCA and 5'CCCATCACATTCTCCGCGTACA. The sizes of amplicons associated with each of these transcripts were approximately 100 bp, which yields optimal PCR efficiency. The annealing temperature used for amplifications was 53°C. Quantification of relative transcript levels was accomplished by the 2e^–DDCt method (Livak and Schmittgen, 2001), using CBLP (Schloss, 1990) as an internal control gene (endogenous reference; the RNA level from this gene remained constant during P deprivation).

Sequence data from this article have been deposited with the EMBL/GenBank data libraries under accession numbers AY946347, AY946348, NP567031, AC112159, and YP007323.

	ACKNOWLEDGMENTS

 
This is a Carnegie Institution Publication (no. 1684).

Received January 13, 2005; returned for revision March 4, 2005; accepted March 7, 2005.

	FOOTNOTES

 
¹ This work was supported by the U.S. Department of Agriculture (grant no. 2002–35301–12178) and by the Carnegie Institution of Washington.

^[w] The online version of this article contains Web-only data.

Article, publication date, and citation information can be found at www.plantphysiol.org/cgi/doi/10.1104/pp.105.059550.

^* Corresponding author; e-mail arthurg{at}stanford.edu; fax 650–325–6857.

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