Logo of eukcellPermissionsJournals.ASM.orgJournalEC ArticleJournal InfoAuthorsReviewers
Eukaryot Cell. 2007 Jun; 6(6): 919–930.
Published online 2007 Apr 13. doi:  10.1128/EC.00207-06
PMCID: PMC1951523

Acclimation to Singlet Oxygen Stress in Chlamydomonas reinhardtii[down-pointing small open triangle]


In an aerobic environment, responding to oxidative cues is critical for physiological adaptation (acclimation) to changing environmental conditions. The unicellular alga Chlamydomonas reinhardtii was tested for the ability to acclimate to specific forms of oxidative stress. Acclimation was defined as the ability of a sublethal pretreatment with a reactive oxygen species to activate defense responses that subsequently enhance survival of that stress. C. reinhardtii exhibited a strong acclimation response to rose bengal, a photosensitizing dye that produces singlet oxygen. This acclimation was dependent upon photosensitization and occurred only when pretreatment was administered in the light. Shifting cells from low light to high light also enhanced resistance to singlet oxygen, suggesting an overlap in high-light and singlet oxygen response pathways. Microarray analysis of RNA levels indicated that a relatively small number of genes respond to sublethal levels of singlet oxygen. Constitutive overexpression of either of two such genes, a glutathione peroxidase gene and a glutathione S-transferase gene, was sufficient to enhance singlet oxygen resistance. Escherichia coli and Saccharomyces cerevisiae exhibit well-defined responses to reactive oxygen but did not acclimate to singlet oxygen, possibly reflecting the relative importance of singlet oxygen stress for photosynthetic organisms.

Reactive oxygen species (ROS) production is an unavoidable consequence of life in an aerobic environment, and reliance upon oxygenic photosynthesis presents plants and algae with sources of ROS not generally shared by their nonphotosynthetic counterparts. Nearly any form of biotic or abiotic stress affects the chloroplast, where the photosynthetic electron transport chain brings together photosensitizing pigments, redox-active electron carriers, and oxygen generation in a polyunsaturated lipid environment. Disruptions in the balance between incoming excitation energy and terminal electron acceptors can result in ROS production and eventual cell death. High-light (HL) stress, for example, leads to increased production of singlet oxygen (1O2*), hydrogen peroxide, and superoxide in the chloroplast (31, 44), while hypersensitive responses to tobacco mosaic virus in tobacco result in down-regulation of the proteins necessary to repair ROS-mediated damage to photosystem II (71). Understanding how plants and algae respond to ROS and limit ROS-induced damage is therefore necessary to piece together responses to biotic and abiotic stress.

As a result of the capacity of ROS for damaging cellular constituents, including proteins, nucleic acids, and membranes (41), ROS are often cast in a purely destructive role. Evidence is emerging, however, that sublethal levels of ROS can be important signaling intermediates (4, 30), activating pathways that bolster defense responses and enhance survival of subsequent stress (11, 13, 47, 78). For example, in the yeast Saccharomyces cerevisiae, sublethal levels of hydrogen peroxide activate the YAP1 (yeast activator protein 1) transcription factor (15, 85), which then promotes expression of a number of antioxidant-related genes (35, 54), including thioredoxin (51), glutathione peroxidase (48), and gamma-glutamylcysteine synthetase (GSH1), the rate-limiting enzyme in glutathione biosynthesis (86). S. cerevisiae also acclimates to superoxide (25, 49) and lipid hydroperoxides (17), and these responses are often characterized by specificity to the form of the original stress (2, 80). GSH1, for example, is induced by both superoxide and peroxide, but loss of YAP1 abolishes peroxide induction of GSH1, while leaving superoxide induction intact (76). Acclimation responses to hydrogen peroxide and superoxide are also regulated by different response regulons in Escherichia coli (40, 78, 81). The lessons learned from E. coli and S. cerevisiae indicate that the nature of ROS signaling depends on the chemical identity of the ROS. Therefore, to understand the mechanisms by which cells sense and respond to oxidative stress, it is necessary to investigate responses to individual ROS.

Although ROS sensors in E. coli and S. cerevisiae have been characterized, the absence of obvious homologs of these sensors in algae and plants suggests that mechanisms for responding to ROS may differ in photosynthetic organisms (reviewed in reference 5). Furthermore, the abundance of photosensitizing pigments required for photosynthesis means that plants and algae may be subject to oxidative stresses, such as 1O2*, that are not as important for nonphotosynthetic organisms. Despite the possible importance of 1O2* stress responses in photosynthetic organisms, little is known about what systems may exist to counteract 1O2* damage. 1O2* is a highly reactive, excited state of oxygen that can be formed when excited triplet chlorophyll (3Chl*) in photosystem II interacts with ground-state oxygen. Environmental stress that upsets the balance between light harvesting and energy utilization lengthens the lifetime of chlorophyll (1Chl*) (reaction 1), increasing the likelihood that 1Chl* will undergo intersystem crossing to form 3Chl* (reaction 2). 3Chl* is longer-lived than 1Chl* and reacts more readily with ground-state 3O2 (reaction 3). The physical interaction between 3Chl* and oxygen produces 1O2* (reaction 3), liberating oxygen from the spin restriction that normally limits its reactivity with singlet-state biological molecules (39).

The three reactions are as follows: reaction 1, 1Chl + light → 1Chl*; reaction 2, 1Chl* → 3Chl*; reaction 3, 3Chl* + 3O21Chl + 1O2*.

While pigments, such as chlorophyll and protochlorophyllide, can generate 1O2* endogenously, exogenous photosensitizing dyes, such as rose bengal (RB), generate 1O2* as well (77). 1O2* is highly reactive and can modify lipids (36), nucleic acids (58), and proteins (14). Experiments using lipophilic photosensitizers in E. coli established that a 1O2* molecule could not travel more than 0.07 μm within a cell before either being quenched or reacting with another molecule (60), but recent work using a microscope capable of detecting near-infrared phosphorescence from 1O2* has indicated that 1O2* generated in the cytoplasm is capable of moving across cell membranes (75).

Despite the transience of 1O2*, several lines of evidence indicate that 1O2* can impact gene expression in photosynthetic organisms. Previous work in the single-celled alga Chlamydomonas reinhardtii established 1O2*-mediated regulation of a putative glutathione peroxidase gene (GPXH) (21, 23, 55). The photosynthetic proteobacterium Rhodobacter sphaeroides also induces a glutathione peroxidase in response to singlet oxygen (37), and multiple R. sphaeroides operons have been identified that appear to be under 1O2* control (3). Recently, work with protochlorophyllide-accumulating flu mutants in Arabidopsis thaliana has shown that 1O2* generated by protochlorophyllide accumulation in the chloroplast can trigger gene expression changes in the nucleus, many of which are specific to singlet oxygen and are not mimicked by treatment with hydrogen peroxide or superoxide (34, 63). 1O2* responses in flu mutants include growth arrest and programmed cell death, both of which are controlled by the nucleus-encoded, chloroplast-localized protein EX1 (EXECUTER 1) (83). Despite this array of physiological responses to singlet oxygen, acclimation to singlet oxygen has not yet been demonstrated in any of these systems.

To learn more about how C. reinhardtii responds to photooxidative stress, we assayed for the ability to acclimate to specific forms of ROS. We found that sublethal levels of 1O2* triggered a clear enhancement of defenses against 1O2*. Characterization of this response revealed that the abundance of transcripts of a small subset of genes was enhanced in response to 1O2* pretreatment. Constitutive overexpression of either of two of these genes—a glutathione peroxidase gene and a glutathione S-transferase gene—was sufficient to promote 1O2* resistance. The inability of S. cerevisiae and E. coli to acclimate to 1O2* suggests the importance of 1O2* responses for photosynthetic organisms.


Strains and growth conditions.

The wild-type C. reinhardtii strain used in this work was 4A+ (16). The pc1 y7 double mutant strain was produced by crossing CC-2471 (pc1 mt+) to CC-1174 (y7 mt−) and selecting for strains that failed to green in both the light and dark (28). The pc1 and y7 parental strains were obtained from the C. reinhardtii Stock Center.

Cells were grown photoautotrophically in minimal (HS) medium or photoheterotrophically in acetate-containing (TAP) medium under low-light (LL) conditions (50 μmol photons m−2 s−1) as described previously (7). Cultures were grown to mid-exponential phase (1 × 106 to 2 × 106 cells/ml). For LL to HL transitions, cells were shifted to 500 μmol photons m−2 s−1. RB (Sigma), hydrogen peroxide (EM Science), methyl viologen (Sigma), neutral red (Sigma), tert-butyl hydroperoxide (Sigma), and metronidazole (Sigma) were each dissolved in water and added directly to the growth medium immediately prior to use. Deuterium oxide, neutral red, and tert-butyl hydroperoxide assays were carried out in 100-μl volumes in 96-well trays at 50 to 60 μmol photons m−2 s−1. Deuterium oxide experiments were performed in TAP medium containing 95% (vol/vol) deuterium oxide (Sigma). Cells were incubated in this mix for 2 h prior to RB treatment.

Experiments with E. coli were performed using strain DH5α in Luria broth at 37°C. Experiments with S. cerevisiae were performed at 30°C in yeast extract-peptone-dextrose (YPD) medium using strain YPH500 (73). Maximum pretreatment concentrations used were the highest concentrations of RB that did not result in cell death.

Tocopherol and pigment analysis.

Wild-type cells were grown photoautotrophically in 100-ml cultures under LL. RB treatments were administered as described above, and 2-ml samples were taken for acetone extraction and high-performance liquid chromatography determination of tocopherol and pigment content as described previously (6).

RNA isolation.

Samples were harvested by centrifugation (3,200 × g, 4°C, 3 min) and then resuspended in 0.1 volume of H2O at 4°C. An equal volume of 2× lysis buffer (0.6 M NaCl, 10 mM EDTA, 100 mM Tris HCl [pH 8.0], 4% [wt/vol] sodium dodecyl sulfate [SDS]) was added to the cell suspension, which was subsequently incubated at 65°C for 5 min. Then, 0.132 volume of 2 M KCl was added, and the samples were incubated on ice for 15 min. Samples were then centrifuged at 12,000 × g for 10 min at 4°C. Supernatants were extracted twice with phenol-chloroform and once with chloroform before precipitating RNA overnight using 0.33 volume of 8 M LiCl at 4°C. This was followed by a final ethanol precipitation. RNA quality was assessed using the ratio of absorbance at 260 nm and 280 nm and by ethidium bromide staining following gel electrophoresis. RNA for both microarrays and RNA gel blot analysis was isolated in this manner.

Microarray experimental design.

Cultures (100 ml) of C. reinhardtii 4A+ were grown photoautotrophically at 50 μmol photons m−2 s−1 until they reached a density of 1.5 × 106 cells/ml. “Pretreated” samples were treated with 2 μM RB, whereas “unpretreated” cultures received a mock inoculation with an equal volume of water. Cells were incubated for 2 h before RNA was harvested.

Microarray slides were printed as part of the C. reinhardtii Genome Project (46, 72). Fragments corresponding to the last 400 base pairs at the 3′ ends of 2,761 C. reinhardtii cDNAs were amplified and spotted onto polyamine-coated slides (Corning, Acton, MA). Each slide contained four replicate spots arrayed in four distinct grids. A total of four slides were used, encompassing two biological replicates, each with a dye-switch control.

Microarray probe labeling.

Reverse transcription of RNA samples was carried out at 42°C for 2 h in the presence of deoxynucleoside triphosphates containing a 1:1 ratio of amino-allyl dUTP to dTTP. Samples were treated with EDTA to stop the reaction and with NaOH to destroy RNA. After neutralizing the sample with HCl, cDNA was purified using a Microcon 30 column, dried, and resuspended in 0.1 M Na2HCO3 buffer (pH 9.0). RNA samples were labeled using Post-Labeling Reactive Dye Packs from Amersham Biosciences (Amersham, Little Chalfont, Buckinghamshire, United Kingdom). Probes were purified individually using QIAquick columns (QIAGEN, Valencia, CA) and then combined.

Microarray hybridization conditions.

Slides were blocked in a succinic anhydride-sodium borate solution for 20 min and then rehydrated in a boiling water bath for 1 min. Slides were then rinsed in ethanol and dried by brief centrifugation. Prehybridization was carried out for 20 min at 50°C in a solution containing 3.5× SSC (1× SSC is 0.15 M NaCl plus 0.015 M sodium citrate), 0.1% (wt/vol) SDS, and 10 mg/ml bovine serum albumin. After prehybridization, slides were rinsed with water and then with isopropanol and dried. For hybridization, a 2× hybridization buffer containing 6× SSC, 0.2% (wt/vol) SDS, 0.1 mg/ml poly(dA), and 0.1 mg/ml yeast tRNA was prepared and mixed with an equal volume of labeled cDNA. Slides were hybridized overnight in a 50°C water bath. The next day, slides were washed once in 2× SSC, 0.03% (wt/vol) SDS, once in 1× SSC, and once in 0.05× SSC, each for 5 min. Slides were then dried and stored until scanning.

Microarray image analysis.

Slides were scanned on an ArrayWoRx Biochip Reader and quantitated using the SoftWoRx Tracker Microarray program, both from Applied Precision, LLC (Issaquah, WA). Each represented gene had to have a valid data point from each biological replicate to be considered further. Abnormal spots were flagged manually and excluded from further analysis, as was any spot in which the mean spot intensity did not exceed two times the median background intensity, or in which the signal-to-noise ratio value was less than 1. Spots that were >20% saturated were also excluded.

Data analysis.

Each slide contained four replicates of each spot arrayed in separate grids. For data normalization, each grid (containing a single set of 2,761 C. reinhardtii cDNA PCR products) was normalized individually. Preliminary analysis indicated that considerably fewer than 5% of the data points showed a greater than twofold change, suggesting that scaling could be an appropriate method for normalization of this data set. For comparison, data were also normalized manually in Excel (Microsoft). The median background intensity was subtracted from the mean spot intensity in each channel. A regression line was fit to a Cy3 versus Cy5 plot, and Cy3 values were divided by the slope of the line. Tailing was often observed at high and low intensity values. To account for this tailing, linear scaling and intensity-dependent normalization were performed using the SNOMAD program with a span of 0.7 and a trim of 0.1 (10). Data generated by both normalization methods were nearly identical. The data presented in Table Table11 are ratios calculated from SNOMAD-normalized data.

Differentially expressed genes in microarray analyses of cells pretreated with RB

To account for the many levels of replication in our experimental design, statistical analysis (analysis of variance) was performed using mixed-model analysis of variance performed by the J/maanova program (9), with treatment (pretreated or not pretreated) and dye as fixed effects and array, biological sample, and spot (referring to the four replicate spots present on each array) as random effects. Data deleted using the criteria described in the preceding paragraph were imputed using the nearest-neighbor algorithm provided in the Statistical Analysis of Microarrays program (82) before performing analysis of variance. The data presented in Table Table11 represent those genes for which the difference between pretreated and unpretreated expression levels exceeded 1.5-fold and the false discovery rate-adjusted tabulated P value supporting differential expression was less than 1 × 10−4. P values were calculated using the Fs test statistic described by Cui et al. (12).

RNA gel blot analysis.

Primers were designed based upon the expressed sequence tag (EST) contig assembly sequences (72). The primers used to amplify GSTS1 were 5′-TTACGACTTCCTCCGCACTC-3′ and 5′-CGGGACCAGACCTGTTTCTTG-3′. The primers used to amplify PHC8 were 5′-CAGTCGCCTACCACAATTCAC-3′ and 5′-TGGCCTCATCTTCTCACCTTC-3′. Primers for amplification of a portion of contig number 20021010.5327 (referred to as 5327 hereafter) were 5′-GCCAGACTTGTTGTCTTATTACCAT-3′ and 5′-CTGTATTTGCTGTGTAAGGGTTTG-3′. The primers used to amplify GSTS2 were designed based on the contig 20021010.3547. GSTS2 primer sequences were 5′-AAGGCCTACTACCAGGACAAGAC-3′ and 5′-CTGTAAACCAAACGACTTCAAGG-3′. PCR products were cloned into the pGEM-T Easy vector (Promega, Madison, WI). GPXH probes were generated from vectors described by Leisinger et al. (55, 56). The APX1 probe was described by Ledford et al. (53). To generate RNA probes, vector inserts were transcribed in the antisense direction in the presence of digoxigenin-labeled dUTP (Roche Molecular Biochemicals, Germany).

RNA from C. reinhardtii cells was prepared as described above, and 5 μg of RNA per sample were fractionated using denaturing gel electrophoresis (67). Blots were hybridized in DIG-EasyHyb solution (Roche Molecular Biochemicals, Germany) at 67°C, and high-stringency washes were carried out at 68°C using 0.2× SSC, 0.1% (wt/vol) SDS.

Overexpression of GPXH and GSTS1.

PSAD flanking sequences were used to drive constitutive overexpression of GPXH and GSTS1 (24). GPXH cDNA was amplified from the vector described by Leisinger et al. (56), using primers designed to engineer an NdeI site at the beginning and an EcoRI site at the end. The GPXH_NdeI forward primer sequence was 5′-TCACAACAAGCCCATATGGCGAACCCCGAGTTTTACG-3′, and the GPXH_EcoRI reverse primer sequence was 5′-CAGCTGCTGCCAGAATTCTTAGTTACGCGTTC-3′ (restriction sites are underlined). Similarly, GSTS1 was amplified from genomic DNA using the following primers: GSTS1_NdeI 5′-TCACAACAAGCCCATATGGCCCCCAAGCTGTA-3′ and GSTS1_EcoRI 5′-CAGCTGCTGCCAGAATTCTTACGCGTCTGGCC-3′. PCR products were digested with EcoRI and NdeI and cloned into the pSL18 vector containing PSAD 5′ and 3′ untranslated regions as well as a paromomycin-selectable marker (24, 64).

ProPSAD:GPXH and ProPSAD:GSTS1 were transformed into C. reinhardtii 4A+ by the method of Dent et al. (16). Transformants were selected on TAP plates containing 10 μg/ml paromomycin (Sigma).

Microarray data accession numbers.

Raw microarray data have been deposited into the National Center for Biotechnology Information Gene Expression Omnibus database at http://www.ncbi.nlm.nih.gov/geo under series accession number GSE4681.


C. reinhardtii exhibits a robust acclimation response to 1O2*.

To learn more about how photosynthetic organisms respond to photooxidative stress, the model alga, C. reinhardtii, was tested for the ability to acclimate to specific ROS. Acclimation was scored as the ability of pretreatment with a sublethal level of a ROS-generating compound to induce defense responses that enhanced survival of a subsequent challenge with higher concentrations of ROS (Fig. (Fig.1A).1A). We tested hydrogen peroxide, tert-butyl hydroperoxide (an organic peroxide capable of triggering lipid peroxidation chain reactions), metronidazole (a compound which generates superoxide only in the chloroplast) (70), and RB (a xanthene dye that produces 1O2* upon excitation by light) for the ability to induce an acclimation response. The response to RB was the most dramatic, with a significant proportion of pretreated cells surviving concentrations that killed all unpretreated cells (Fig. (Fig.1B1B and Fig. Fig.2A,2A, leftmost photo).

FIG. 1.
C. reinhardtii acclimates to singlet oxygen. (A) Schematic of acclimation assays. C. reinhardtii was assayed for an ability to acclimate to oxidative stress following pretreatment with sublethal levels of that stress. Cells were pretreated and challenged ...
FIG. 2.
Light and solvent dependency of responses to RB. (A) Light dependency of acclimation to RB. Cells were pretreated and challenged in the light (50 μmol photons m−2 s−1; indicated by a sun) or the dark (indicated by a moon). Pretreatment ...

Acclimation to RB is a response to 1O2*.

RB produces 1O2* only when excited by visible light, with peak absorption occurring between 450 and 580 nm (77). To ensure that RB toxicity during the challenge treatment was the result of 1O2* production, the entire experiment was repeated in the dark. RB was not toxic in the dark at these concentrations (Fig. (Fig.2A,2A, middle photo), indicating that the toxicity of the challenge was dependent upon the 1O2* produced during photosensitization. To determine whether the acclimation response was a response to 1O2* or to the presence of a xenobiotic compound, the pretreatment was conducted in the dark followed by a challenge in the light. Cells that were pretreated in the dark failed to acclimate (Fig. (Fig.2A,2A, rightmost photo), indicating that acclimation does not occur in the absence of 1O2* production.

The lifetime of 1O2* is approximately 10 times longer in deuterium oxide than it is in H2O (59). Therefore, if RB toxicity is dependent upon 1O2*, then toxicity should be enhanced in deuterium oxide. As shown in Fig. Fig.2B,2B, ,11 μM RB was lethal to cells treated with RB in deuterium oxide, whereas the viability of cells treated with RB in water was unaffected at that concentration.

Acclimation to 1O2* is rapidly induced and transient.

Pretreatment times and concentrations were varied to determine how quickly acclimation is induced (Fig. (Fig.3A).3A). Acclimation could be triggered by as little as 0.25 μM RB, and the effect increased with longer incubation times and higher concentrations of RB. A small degree of acclimation was evident with only 30 min of pretreatment, and the degree of 1O2* resistance increased with longer pretreatment time.

FIG. 3.
Acclimation to 1O2* is rapid and transient. (A) Cells were grown photoautotrophically to mid-exponential phase, then pretreated with various RB concentrations and durations (duration of pretreatment is indicated below each photo), and challenged ...

In yeast, acclimation to hydrogen peroxide is a transient physiological adaptation that is lost less than 2 h following the pretreatment (13). To find out whether the 1O2* resistance induced by pretreatment was caused by transient changes in cell physiology or reflected a permanent, adaptive response, cells that were pretreated (2 μM RB, 2 h) and challenged (8 μM RB, 1 h) were washed to rid the medium of RB and incubated for 24 h before repeating the pretreatment and challenge (Fig. (Fig.3B).3B). Twenty-four hours later, these cells behaved as if they were no longer acclimated to 1O2* (Fig. (Fig.3B,3B, right photo), and another pretreatment was necessary to induce resistance to 8 μM RB. Based on these data, acclimation to 1O2* is likely the product of pretreatment-induced, transient changes in cell physiology that enhance 1O2* resistance.

Acclimation is specific to 1O2*.

To explore possible overlap between responses to different ROS, we then tested the ability of 1O2* pretreatment to induce acclimation to other ROS-generating compounds, including hydrogen peroxide, methyl viologen, metronidazole, tert-butyl hydroperoxide, and neutral red (another photosensitizing dye). The only evidence of cross-tolerance as the result of 1O2* pretreatment was observed in the case of neutral red (Fig. (Fig.4A),4A), which has been shown to produce 1O2* in treated C. reinhardtii thylakoids (23). Interestingly, pretreatment with RB increased methyl viologen sensitivity but had no impact on sensitivity to metronidazole. This could be related to differences in chemical structure or sites of action between these two compounds (70). The experiment was also reversed, this time pretreating with neutral red, hydrogen peroxide, methyl viologen, metronidazole, and tert-butyl hydroperoxide, and challenging with RB. Only the neutral red pretreatment was able to enhance resistance to RB (data not shown). These results suggest that the acclimation occurs specifically in response to 1O2* and activates defenses that are specific to protection against 1O2*-mediated damage.

FIG. 4.
Cross-acclimation between 1O2* and other sources of oxidative stress. (A) Cross-acclimation between 1O2* and other ROS. Cells were grown photoautotrophically, pretreated (+) with 2 μM RB for 2 h at 50 μmol photons ...

HL treatment induces acclimation to 1O2* stress.

1O2* is produced endogenously during exposure to HL when triplet chlorophyll interacts with ground-state oxygen. To find out whether there is an overlap between the response to HL and acclimation to 1O2* stress, unpretreated (LL-grown) cells were shifted to HL and then assayed for resistance to 1O2*. Cells that had been pretreated with HL for 1 h were more resistant to 1O2* (Fig. (Fig.4B4B).

Acclimation to 1O2* stress does not alter the composition or content of carotenoids or vitamin E.

Several small-molecule antioxidants are capable of quenching 1O2* or scavenging 1O2*-produced damage products (69). Included in this group of antioxidants are carotenoids and tocopherols (vitamin E). In C. reinhardtii, it has previously been shown that alterations in carotenoid composition can affect RB sensitivity (6, 7), but there were no changes in β-carotene, lutein, or the xanthophyll cycle pigments zeaxanthin, antheraxanthin, and violaxanthin during the 2-hour pretreatment or the 1-hour challenge (Fig. (Fig.5).5). Similarly, no changes in vitamin E content or composition were observed (Fig. (Fig.5).5). Based on these data, changes in the abundance or composition of carotenoids and vitamin E do not account for the increased resistance to 1O2* seen following pretreatment.

FIG. 5.
Acclimation does not involve changes in cellular carotenoid or α-tocopherol content or composition. Photoautotrophically grown cells were pretreated with 2 μM RB for 2 h at 50 μmol photons m−2 s−1 and then challenged ...

Microarray analysis of gene expression during acclimation to 1O2* stress.

In addition to small-molecule antioxidants, expression of nucleus-encoded antioxidant enzymes could enhance survival of 1O2*stress. To identify the genes that changed expression in response to the 1O2* pretreatment, we used the cDNA microarrays generated as part of the C. reinhardtii Genome Project (46, 72). These partial-genome arrays (v1.0) had 2,761 spots, representing approximately 20% of the genome. RNA isolated from pretreated cells was compared directly to RNA from mock-pretreated cells. Four slides were used to assay two biological replicates, each with a dye-switch control. In addition, each slide contained four replicate spots.

Although many genes encoding proteins with possible roles in antioxidant metabolism were included on these partial-genome arrays (53), only a small number of genes changed expression in response to the sublethal pretreatment with RB (Fig. (Fig.6A).6A). Each of these genes is listed in Table Table1.1. Among those genes that increased expression were a glutathione peroxidase (GPXH) and a glutathione S-transferase (GSTS1). A cytosolic thioredoxin (TRXH) that has been shown to play a role in resistance to DNA alkylating agents (68) also increased expression, as did a gene encoding a predicted protein with 40% sequence identity to pherophorins from Volvox carteri (38) (PHC8). In addition, a gene with no sequence similarity to genes of known function (5327) increased expression. Among those genes that decreased expression during pretreatment were two genes related to the carbon-concentrating mechanism, the periplasmic carbonic anhydrase (CAH1) and a chloroplast envelope carrier protein (CCP1), both of which are induced in response to low CO2 concentrations (33, 45, 65).

FIG. 6.
Pretreatment with 1O2* causes changes in gene expression. (A) Volcano plot of microarray results comparing pretreated cells with unpretreated cells. Log2-transformed gene expression ratios are plotted on the x axis against log10-transformed P ...

Gene expression changes following RB pretreatment are in response to 1O2*.

To determine whether the gene expression changes observed in the microarray experiment were in response to 1O2* or simply to the presence of a xenobiotic compound (RB), RNA gel blot analysis of GPXH, GSTS1, PHC8, and 5327 was performed using RNA isolated from heterotrophically grown cells treated with RB in the light and in the dark (Fig. (Fig.6B).6B). In all cases, gene expression changed only when cells were treated with RB in the light, indicating that these changes are in response to photosensitized 1O2* production. GSTS2, another putative sigma class glutathione S-transferase with high sequence similarity to GSTS1 was also assayed, as was expression of an ascorbate peroxidase (APX1) that was not present on the array. Neither of these genes showed altered RNA levels in response to 1O2* when cells were grown heterotrophically (Fig. (Fig.6B).6B). CAH1 transcript was not detected (data not shown), likely because of the presence of acetate in the medium (19).

Endogenous photosensitizers also induce GPXH and GSTS1 expression.

Although RB appeared to be affecting cell physiology through the production of 1O2*, we wanted to know whether these same changes would occur in response to an endogenous source of 1O2*. To that end, we evaluated gene expression changes in a chlorophyll biosynthesis mutant. In C. reinhardtii, there is both a light-dependent and a light-independent pathway for the conversion of protochlorophyllide to chlorophyllide (28). The pc1 y7 double mutant is blocked in both pathways, leading to the accumulation of protochlorophyllide and inability to grow under even LL conditions (57). In the presence of light, accumulated protochlorophyllide can act as an endogenous photosensitizer and generate 1O2* within the chloroplast (63). RNA was isolated from pc1 y7 and wild-type cells following a shift from the dark to LL, and RNA gel blot analysis was used to evaluate gene expression changes immediately following this shift. Transcript abundance of both GPXH and GSTS1 increased to higher levels in the pc1 y7 mutant relative to the wild type in response to the transfer from dark to LL (Fig. (Fig.7A).7A). GPXH exhibited a more rapid response, with transcript abundance increasing after only 30 min in the light, whereas GSTS1 expression exhibited a slower response, increasing after 3 h.

FIG. 7.
Impact of endogenous photosensitizers on GPXH and GSTS1 expression. (A) RNA gel blot analysis of pc1 y7 cells transferred from dark to LL. The protochlorophyllide-accumulating mutant pc1 y7 and the wild type (WT) were grown in TAP medium in the dark and ...

Given the cross-tolerance observed between HL and 1O2* produced exogenously by RB (Fig. (Fig.4B),4B), it was possible that the genes that increased expression in response to 1O2* could also be induced by HL treatment of wild-type cells. Previous work has shown that GPXH responds rapidly to changes in light intensity (21, 53). GSTS1 expression was also tested. Expression of both genes was enhanced following exposure to HL, with GPXH again showing a more rapid induction. Induction of GPXH occurred within 30 min, followed by increased GSTS1 expression within 3 h (Fig. (Fig.7B7B).

Specificity of 1O2*-induced gene expression changes.

To determine whether other ROS affect expression of the 1O2*-responsive genes, gene expression changes in response to hydrogen peroxide, metronidazole, and tert-butyl hydroperoxide were monitored. As previously reported (55, 56), GPXH expression did not respond as strongly or as quickly to hydrogen peroxide or the superoxide generators metronidazole and methyl viologen but did respond slowly to tert-butyl hydroperoxide, an organic peroxide (Fig. (Fig.8).8). All other 1O2*-induced genes were also induced by these additional ROS. Interestingly, GSTS2 and APX1, which did not increase expression in response to 1O2* in heterotrophically grown cells (Fig. (Fig.6),6), did respond to 1O2* in photoautotrophically grown cells (Fig. (Fig.8).8). CAH1, which decreased expression in response to 1O2* in the microarray analysis (Table (Table1),1), also decreased expression in response to all other ROS tested (Fig. (Fig.88).

FIG. 8.
Oxidative stress-induced changes in gene expression. Cells were grown photoautotrophically at 50 μmol photons m−2 s−1 and treated with the indicated compounds (tBOOH, tert-butyl hydroperoxide; MZ, metronidazole). Concentrations ...

Constitutive overexpression of either GPXH or GSTS1 is sufficient to enhance resistance to 1O2*.

To investigate the possible connection between enhanced 1O2* resistance and increased expression of GPXH or GSTS1, each gene was constitutively overexpressed using the PSAD promoter (24). Overexpression constructs were transformed into wild-type cells, and transformants were screened based on GPXH or GSTS1 expression (Fig. 9A and B). Because the PSAD 3′ and 5′ untranslated regions are smaller than the untranslated regions flanking the endogenous GPXH gene (362 bp versus 721 bp), ProPSAD:GPXH transformants have two GPXH transcripts, the smaller band produced from the transgene and the larger band corresponding to transcript from the endogenous gene. Of the 16 transformants screened for each construct, 12 overexpressed GPXH, and 8 overexpressed GSTS1. A subset of these is shown in Fig. Fig.99.

FIG. 9.
Constitutive overexpression of GPXH and GSTS1 confers resistance to 1O2*. (A) RNA gel blot analysis of ProPSAD:GPXH transformants. RNA was extracted from heterotrophically grown paromomycin-resistant transformants containing the ProPSAD:GPXH overexpression ...

ProPSAD:GPXH and ProPSAD:GSTS1 transformants were then assayed for sensitivity to 1O2* by plating cells onto agar medium containing either RB or neutral red (Fig. (Fig.9C).9C). Constitutive overexpression of either GPXH or GSTS1 was sufficient to confer enhanced resistance to 1O2*, as evidenced by the ability of ProPSAD:GPXH and ProPSAD:GSTS1 strains to withstand higher concentrations of RB and neutral red (Fig. (Fig.9C).9C). ProPSAD:GPXH and ProPSAD:GSTS1 strains were also more resistant than the wild type to RB in liquid culture, but the level of resistance was not as high as that induced by pretreatment with RB (data not shown).

E. coli and S. cerevisiae do not acclimate to 1O2* stress.

There are large mutant collections for E. coli and S. cerevisiae, two model organisms in which oxidative stress responses have been extensively investigated. In hopes of taking advantage of the tools available in these systems, E. coli and S. cerevisiae were tested for the ability to acclimate to RB-induced 1O2* stress. Concentrations of RB were established for both E. coli and S. cerevisiae that would be sublethal (for pretreatments) over different periods of exposure ranging from 15 min to 5 h. No condition was found that enhanced survival after challenge, and instead the pretreatment often had an additive, deleterious effect when followed by a challenge (Fig. (Fig.1010).

FIG. 10.
Screens for acclimation to 1O2* in E. coli and S. cerevisiae. E. coli and S. cerevisiae cultures were pretreated with the indicated RB concentrations for 2 h, then challenged for 1 h, and plated.


C. reinhardtii exhibits an acclimation response to 1O2*.

Responses to 1O2* have recently been characterized in several photosynthetic organisms, including C. reinhardtii (21, 55), A. thaliana (63, 83), and R. sphaeroides (3, 37), but how these organisms sense 1O2* remains a mystery. We have found that C. reinhardtii exhibits a physiological acclimation in response to 1O2* which is summarized in Fig. Fig.11.11. This response is characterized by the ability of a sublethal 1O2* pretreatment to transiently enhance 1O2* resistance (Fig. (Fig.11 to to3)3) and can be triggered by both exogenous photosensitizing dyes, including RB, or endogenous pigments, such as chlorophyll (Fig. (Fig.4).4). While acclimation to hydrogen peroxide, superoxide, and lipid peroxides has been demonstrated in E. coli and S. cerevisiae, C. reinhardtii is, to our knowledge, the first example of an organism that exhibits an acclimation response to 1O2*. This has been independently verified in a very recent study by Fischer et al. (20). Furthermore, E. coli and S. cerevisiae did not acclimate to 1O2* under our conditions (Fig. (Fig.10),10), although for E. coli we cannot rule out the possibility that an acclimation response might have been obscured by the high DNA damage sensitivity of the recA mutant strain that was used. 1O2* is a particularly important ROS for photosynthetic organisms because of their reliance upon photosensitizing pigments for light harvesting (50), and acclimation to 1O2* in C. reinhardtii may reflect the relative importance of 1O2* stress for this photosynthetic alga. It will be interesting to examine acclimation in other photosynthetic organisms, such as A. thaliana and Synechocystis sp. strain PCC 6803.

FIG. 11.
Model of acclimation to 1O2* in C. reinhardtii. Pchlide, protochlorophyllide; LOOH, lipid hydroperoxide.

The response to 1O2* was remarkably specific, and no evidence of cross acclimation with other forms of ROS was observed (Fig. (Fig.4A).4A). This is particularly significant because approximately 1% of photosensitization reactions involve electron transfer to oxygen, generating superoxide radicals (26, 52). This is the case regardless of whether the source is an exogenous dye, such as RB, or an endogenous pigment, such as chlorophyll or protochlorophyllide (27), but the absence of cross-acclimation between superoxide and 1O2* argues that acclimation is a response to 1O2* and not superoxide. Furthermore, deuterium oxide, which lengthens the half-life of 1O2*, enhanced RB toxicity (Fig. (Fig.2B),2B), also indicating that 1O2* is responsible for the cell death observed in unacclimated cells. Deuterium oxide enhancement of toxicity does not definitively implicate 1O2*; although, because deuterium oxide assays are widely used to test for the involvement of 1O2* (66), it is possible that deuterium oxide slows the decay of other ROS as well (8, 62). Nevertheless, this result taken together with the absence of cross-acclimation between superoxide and 1O2* suggests that acclimation to RB is a specific response to 1O2* production.

The ability of HL to induce acclimation to 1O2* stress demonstrates an intriguing overlap between 1O2* responses and HL exposure (Fig. (Fig.4B)4B) and suggests that endogenous 1O2* (produced by excited chlorophyll) can induce acclimation to exogenous 1O2* (produced by RB) (Fig. (Fig.11).11). In its natural environment, C. reinhardtii would regularly experience photon flux densities equivalent to our HL treatment, which represents ~25% of full sunlight, and rapid fluctuations in light intensity similar to the 10-fold increase used in our experiments occur frequently in nature. A number of ROS are produced in response to HL, including hydrogen peroxide, superoxide, and 1O2* (29, 31, 44). The fact that methyl viologen, metronidazole, hydrogen peroxide, and tert-butyl hydroperoxide did not induce acclimation to 1O2* (Fig. (Fig.4A)4A) implies that the HL-generated signal is likely mediated by 1O2* rather than hydrogen peroxide, lipid peroxides, or superoxide. However, because tert-butyl hydroperoxide is a shorter molecule than naturally occurring lipids, a role for lipid peroxides cannot be completely ruled out.

Changes in gene expression, but not carotenoid or vitamin E composition, occur during acclimation.

Carotenoids are efficient 1O2* quenchers, and they increase 1O2* resistance when overexpressed in E. coli (79). A C. reinhardtii double mutant, npq1 lor1 mutant that is unable to synthesize lutein and zeaxanthin is also more sensitive than the wild type to 1O2* (7). Furthermore, creating a triple mutant containing npq1, lor1, and npq2, which causes cells to accumulate zeaxanthin, restores RB tolerance to wild-type levels (6). Tocopherols (vitamin E) also play a role in 1O2* defense, and the tocopherol-deficient vte1 mutant in A. thaliana accumulates more lipid peroxides in response to 1O2* than the wild type (43), but despite the potential for carotenoids and tocopherols to protect against 1O2*, changes in carotenoid and tocopherol composition or content did not accompany acclimation to 1O2* in C. reinhardtii (Fig. (Fig.5).5). Glutathione can also be an important component of antioxidant defenses against 1O2* by providing reducing power for lipid peroxide scavenging enzymes, but previous work has established that neither glutathione content nor glutathione redox state is altered by RB treatment under conditions similar to those used in this work (1 μM RB for 20 to 120 min at 120 μmol photons m−2 s−1) (22).

Instead, acclimation to 1O2* was associated with changes in nuclear gene expression. Microarray experiments using the v1.0 C. reinhardtii cDNA arrays (46, 72) detected only 14 genes that changed expression in response to pretreatment with 1O2* (Fig. (Fig.6A6A and Table Table1).1). Six of the 14 genes (GPXH, GSTS1, PHC8, 5327, CAH1, and THI4a) were also tested by RNA gel blot analysis (Fig. (Fig.6B6B and and8;8; also data not shown), and the changes in gene expression were confirmed in each case. The arrays used in our analysis cover approximately 20% of the genome. Extrapolating from these results to the full genome yields an estimated 70 1O2*-regulated genes in the C. reinhardtii genome.

Gene expression changes in response to the pretreatment were light dependent, indicating that transcript abundance changed in response to 1O2*, and not merely in reaction to the presence of a xenobiotic compound, such as RB (Fig. (Fig.6B).6B). This was particularly interesting in the case of GSTS1. No longer relegated only to the role of xenobiotic detoxification, glutathione S-transferases are now known to be a diverse group of enzymes responsible for detoxifying endogenous compounds, including lipid peroxides (1, 74). Some play a direct role in signaling (1). In mammalian systems, prostaglandin H synthase-2, also a member of the sigma class of glutathione S-transferases, is induced by ROS (18). Activation of GSTS1 by RB and endogenous photosensitizers only in the light suggests that transcript abundance of this gene is regulated by oxidative stress rather than the mere presence of foreign chemicals (Fig. (Fig.6B6B and and7).7). The signal that triggers enhanced GSTS1 expression was not specific to 1O2*, however, and induction of both GSTS1 and GSTS2 occurred in response to each ROS tested (Fig. (Fig.8).8). 1O2* induction of APX1 and GSTS2 was more complicated, occurring only in photoautotrophically grown cultures (Fig. (Fig.6B6B and and88).

Decreased expression of the periplasmic carbonic anhydrase gene, CAH1, was also observed in response to each of the ROS tested (Fig. (Fig.8).8). CAH1 transcript levels respond rapidly to changes in CO2, and mRNA abundance decreases within an hour after the start of CO2 supplementation (32). The signal that triggers these changes in C. reinhardtii is as yet unknown, and the effect of 1O2* on CAH1 and CCP1 expression might indicate some cross talk between oxidative stress and regulation of the carbon-concentrating mechanism. In the marine diatom Phaeodactylum tricornutum, increases in cyclic AMP (cAMP) have been suggested to repress transcription of a chloroplastic carbonic anhydrase gene (42). Interestingly, a sequence motif similar to the mammalian cAMP response element has been identified within a region of the GPXH promoter that is responsible for increased transcription of this gene in response to 1O2* in C. reinhardtii (55).

Of all the genes that changed expression in response to 1O2*, only GPXH showed a stronger, more rapid response to 1O2* than to the other ROS tested (Fig. (Fig.8).8). This result confirms previously published work showing that GPXH is induced by photosensitizing dyes and organic hydroperoxides, but not by the superoxide-generating herbicides metronidazole and methyl viologen (55). Because GPXH and GSTS1 encode proteins with potential antioxidant function, it was possible that changes in expression of these genes could affect 1O2* resistance in C. reinhardtii. Overexpression of either GPXH or GSTS1 was sufficient to enhance 1O2* resistance (Fig. (Fig.9).9). However, pretreatment with other ROS also enhance GSTS1 expression (Fig. (Fig.8)8) without increasing 1O2* resistance (data not shown), suggesting that transient increases in GSTS1 transcript cannot be sufficient to induce 1O2* resistance and implying an additional level of regulation. This added layer of regulation could be at the level of translation, and singlet oxygen has been previously shown to affect translation elongation of the D1 protein in Synechocystis sp. strain PCC 6803 (61).

How do GPXH and GSTS1 enhance resistance to 1O2*? Sequence alignments with glutathione peroxidases from other organisms showed that GPXH exhibits features of phospholipid hydroperoxide glutathione peroxidases (data not shown) (5). Given that some glutathione S-transferases also function as lipid peroxidases, the simplest explanation is that overexpressing GPXH and GSTS1 protects cells from 1O2* by enhancing lipid peroxidase activity, but other possible functions for these two genes certainly have not been ruled out. For example, both glutathione peroxidases and glutathione S-transferases have been shown to play direct signaling roles as well (15, 84). Biochemical characterization of GPXH and GSTS1, as well as loss-of-function mutants or RNA interference lines, would be useful to determine the functions of these two genes during 1O2* acclimation.

Model of acclimation to 1O2*.

Overall, this study has allowed us to derive a model of 1O2* acclimation (Fig. (Fig.11)11) in which 1O2* from exogenous dyes, such as RB (Fig. (Fig.11 and and2),2), or endogenous pigments, such as chlorophyll or protochlorophyllide (Fig. (Fig.7),7), activates a signal transduction pathway that increases expression of GPXH, GSTS1, as well as other genes (Table (Table1).1). How the 1O2* signal is perceived and converted to enhanced gene expression remains a mystery. There are three obvious possibilities: 1O2* could itself directly modify a protein sensor; a by-product of 1O2* damage, such as a lipid peroxide or protein peroxide, could interact with the sensor protein; or 1O2* could activate a sensor by perturbing the redox state of the cell (Fig. (Fig.11).11). At present, there is no definitive evidence for or against any of these possibilities. Regulation of GPXH expression by 1O2* has been mapped to two regions of the promoter (55), one of which contains a putative cAMP-response element that is also found in predicted introns of both GSTS1 and GSTS2 (data not shown). Future work evaluating the role of this element in 1O2* signaling could be valuable for piecing together how this short-lived signal is sensed.

HL-induced acclimation to 1O2* stress demonstrates the presence of a chloroplast-to-nucleus retrograde signaling pathway capable of activating the acclimation response. Studies of 1O2* responses in the flu mutant of A. thaliana have also demonstrated a 1O2*-activated retrograde signaling pathway (63, 83). In A. thaliana, this pathway is mediated by EX1. flu ex1 double mutants produce as much 1O2* as flu single mutants but do not experience either growth arrest or cell death in response to light/dark cycles (83). This suggests a signaling rather than antioxidant role for the chloroplast-localized EX1 and also indicates that, in the flu mutant, cell death in response to 1O2* is genetically programmed rather than the direct product of oxidative damage. Whether this response to 1O2* is conserved in C. reinhardtii is currently unknown, but there is a gene with sequence similarity to EX1 in the current release of the C. reinhardtii genome. Future work will address the role of this gene in C. reinhardtii.

This work expands our knowledge of biological responses to 1O2* and raises questions about the nature of the sensing and signaling pathways involved in acclimation to 1O2*. The physiological and molecular characterization described here opens the door for genetic approaches to dissect these pathways. The strong acclimation to 1O2* exhibited by C. reinhardtii makes it an ideal model photosynthetic eukaryote in which to pursue studies of 1O2*.


We gratefully acknowledge Chung-Soon Im and Arthur Grossman for assistance with microarray analyses, Trevor Starr for assistance with S. cerevisiae experiments, and Setsuko Wakao for critical reading of the manuscript.

This work was supported by grants from the National Institutes of Health (GM071908) and the University of California Toxic Substances Research and Teaching Program (03T-1) to K.K.N. H.K.L. was supported in part by a National Institutes of Health Predoctoral Genetics training grant (T32-GM07127).


[down-pointing small open triangle]Published ahead of print on 13 April 2007.


1. Agianian, B., P. A. Tucker, A. Schouten, K. Leonard, B. Bullard, and P. Gros. 2003. Structure of a Drosophila sigma class glutathione S-transferase reveals a novel active site topography suited for lipid peroxidation products. J. Mol. Biol. 326:151-165. [PubMed]
2. Alic, N., T. Felder, M. D. Temple, C. Gloeckner, V. J. Higgins, P. Briza, and I. W. Dawes. 2004. Genome-wide transcriptional responses to a lipid hydroperoxide: adaptation occurs without induction of oxidant defenses. Free Radic. Biol. Med. 37:23-35. [PubMed]
3. Anthony, J. R., K. L. Warczak, and T. J. Donohue. 2005. A transcriptional response to singlet oxygen, a toxic byproduct of photosynthesis. Proc. Natl. Acad. Sci. USA 102:6502-6507. [PMC free article] [PubMed]
4. Apel, K., and H. Hirt. 2004. Reactive oxygen species: metabolism, oxidative stress, and signal transduction. Annu. Rev. Plant Biol. 55:373-399. [PubMed]
5. Avery, A. M., and S. V. Avery. 2001. Saccharomyces cerevisiae expresses three phospholipid hydroperoxide glutathione peroxidases. J. Biol. Chem. 276:33730-33735. [PubMed]
6. Baroli, I., A. D. Do, T. Yamane, and K. K. Niyogi. 2003. Zeaxanthin accumulation in the absence of a functional xanthophyll cycle protects Chlamydomonas reinhardtii from photooxidative stress. Plant Cell 15:992-1008. [PMC free article] [PubMed]
7. Baroli, I., B. L. Gutman, H. K. Ledford, J. W. Shin, B. L. Chin, M. Havaux, and K. K. Niyogi. 2004. Photo-oxidative stress in a xanthophyll-deficient mutant of Chlamydomonas. J. Biol. Chem. 279:6337-6344. [PubMed]
8. Bielski, B. H. J., and E. Saito. 1971. Deuterium isotope effect on the decay kinetics of perhydroxyl radical. J. Phys. Chem. 75:2263-2266.
9. Churchill, G. A. 2004. Using ANOVA to analyze microarray data. BioTechniques 37:173-177. [PubMed]
10. Colantuoni, C., G. Henry, S. Zeger, and J. Pevsner. 2002. SNOMAD (Standardization and NOrmalization of MicroArray Data): web-accessible gene expression data analysis. Bioinformatics 18:1540-1541. [PubMed]
11. Collinson, L. P., and I. W. Dawes. 1992. Inducibility of the response of yeast cells to peroxide stress. J. Gen. Microbiol. 138:329-335. [PubMed]
12. Cui, X., J. T. G. Hwang, J. Qui, N. J. Blades, and G. A. Churchill. 2005. Improved statistical tests for differential gene expression by shrinking variance components estimates. Biostatistics 6:59-75. [PubMed]
13. Davies, J. M., C. V. Lowry, and K. J. Davies. 1995. Transient adaptation to oxidative stress in yeast. Arch. Biochem. Biophys. 317:1-6. [PubMed]
14. Davies, M. J. 2004. Reactive species formed on proteins exposed to singlet oxygen. Photochem. Photobiol. Sci. 3:17-25. [PubMed]
15. Delaunay, A., D. Pflieger, M. B. Barrault, J. Vinh, and M. B. Toledano. 2002. A thiol peroxidase is an H2O2 receptor and redox-transducer in gene activation. Cell 111:471-481. [PubMed]
16. Dent, R. M., C. M. Haglund, B. L. Chin, M. C. Kobayashi, and K. K. Niyogi. 2005. Functional genomics of eukaryotic photosynthesis using insertional mutagenesis of Chlamydomonas reinhardtii. Plant Physiol. 137:545-556. [PMC free article] [PubMed]
17. Evans, M. V., H. E. Turton, C. M. Grant, and I. W. Dawes. 1998. Toxicity of linoleic acid hydroperoxide to Saccharomyces cerevisiae: involvement of a respiration-related process for maximal sensitivity and adaptive response. J. Bacteriol. 180:483-490. [PMC free article] [PubMed]
18. Feng, L., Y. Xia, G. E. Garcia, D. Hwang, and C. B. Wilson. 1995. Involvement of reactive oxygen intermediates in cyclooxygenase-2 expression induced by interleukin-1, tumor necrosis factor-alpha, and lipopolysaccharide. J. Clin. Investig. 95:1669-1675. [PMC free article] [PubMed]
19. Fett, J. P., and J. R. Coleman. 1994. Regulation of periplasmic carbonic anhydrase expression in Chlamydomonas reinhardtii by acetate and pH. Plant Physiol. 106:103-108. [PMC free article] [PubMed]
20. Fischer, B. B., R. Dayer, M. Wiesendanger, and R. I. L. Eggen. 10 April 2007. Independent regulation of the GPXH gene expression by primary and secondary effects of high light stress in Chlamydomonas reinhardtii. Physiol. Plant. doi:.10.1111/j.1399-3054.2007.00901.x [Cross Ref]
21. Fischer, B. B., R. I. L. Eggen, A. Trebst, and A. Krieger-Liszkay. 2006. The glutathione peroxidase homologous gene Gpxh in Chlamydomonas reinhardtii is upregulated by singlet oxygen produced in photosystem II. Planta 223:583-590. [PubMed]
22. Fischer, B. B., A. Krieger-Liszkay, and R. I. L. Eggen. 2005. Oxidative stress induced by the photosensitizers neutral red (type I) or rose bengal (type II) in the light causes different molecular responses in Chlamydomonas reinhardtii. Plant Sci. 168:747-759.
23. Fischer, B. B., A. Krieger-Liszkay, and R. I. L. Eggen. 2004. Photosensitizers neutral red (type I) and rose bengal (type II) cause light-dependent toxicity in Chlamydomonas reinhardtii and induce the Gpxh gene via increased singlet oxygen formation. Environ. Sci. Technol. 38:6307-6313. [PubMed]
24. Fischer, N., and J. D. Rochaix. 2001. The flanking regions of PsaD drive efficient gene expression in the nucleus of the green alga Chlamydomonas reinhardtii. Mol. Genet. Genomics 265:888-894. [PubMed]
25. Flattery-O'Brien, J., L. P. Collinson, and I. W. Dawes. 1993. Saccharomyces cerevisiae has an inducible response to menadione which differs from that to hydrogen peroxide. J. Gen. Microbiol. 139:501-507. [PubMed]
26. Foote, C. S. 1991. Definition of type I and type II photosensitized oxidation. Photochem. Photobiol. 54:659. [PubMed]
27. Foote, C. S. 1968. Mechanisms of photosensitized oxidation. Science 162:963-970. [PubMed]
28. Ford, C., S. Mitchell, and W.-Y. Wang. 1981. Protochlorophyllide photoconversion mutants of Chlamydomonas reinhardtii. Mol. Gen. Genet. 184:460-464.
29. Förster, B., C. B. Osmond, and B. J. Pogson. 2005. Improved survival of very high light and oxidative stress is conferred by spontaneous gain-of-function mutations in Chlamydomonas. Biochim. Biophys. Acta 1709:45-57. [PubMed]
30. Foyer, C. H., and G. Noctor. 2005. Redox homeostasis and antioxidant signaling: a metabolic interface between stress perception and physiological responses. Plant Cell 17:1866-1875. [PMC free article] [PubMed]
31. Fryer, M. J., K. Oxborough, P. M. Mullineaux, and N. R. Baker. 2002. Imaging of photo-oxidative stress responses in leaves. J. Exp. Bot. 53:1249-1254. [PubMed]
32. Fujiwara, S., H. Fukuzawa, A. Tachiki, and S. Miyachi. 1990. Structure and differential expression of two genes encoding carbonic anhydrase in Chlamydomonas reinhardtii. Proc. Natl. Acad. Sci. USA 87:9779-9783. [PMC free article] [PubMed]
33. Fukuzawa, H., S. Fujiwara, Y. Yamamoto, M. L. Dionisio-Sese, and S. Miyachi. 1990. cDNA cloning, sequence, and expression of carbonic anhydrase in Chlamydomonas reinhardtii: regulation by environmental CO2 concentration. Proc. Natl. Acad. Sci. USA 87:4383-4387. [PMC free article] [PubMed]
34. Gadjev, I., S. Vanderauwera, T. S. Gechev, C. Laloi, I. N. Minkov, V. Shulaev, K. Apel, D. Inzé, R. Mittler, and F. Van Breusegem. 2006. Transcriptomic footprints disclose specificity of reactive oxygen species signaling in Arabidopsis. Plant Physiol. 141:436-445. [PMC free article] [PubMed]
35. Gasch, A. P., P. T. Spellman, C. M. Kao, O. Carmel-Harel, M. B. Eisen, G. Storz, D. Botstein, and P. O. Brown. 2000. Genomic expression programs in the response of yeast cells to environmental changes. Mol. Biol. Cell 11:4241-4257. [PMC free article] [PubMed]
36. Girotti, A. W., and T. Kriska. 2004. Role of lipid hydroperoxides in photo-oxidative stress signaling. Antioxid. Redox Signal. 6:301-310. [PubMed]
37. Glaeser, J., and G. Klug. 2005. Photo-oxidative stress in Rhodobacter sphaeroides: protective role of carotenoids and expression of selected genes. Microbiology 151:1927-1938. [PubMed]
38. Godl, K., A. Hallmann, A. Rappel, and M. Sumper. 1995. Pherophorins: a family of extracellular matrix glycoproteins from Volvox structurally related to the sex-inducing pheromone. Planta 196:781-787. [PubMed]
39. Green, M. J., and H. A. Hill. 1984. Chemistry of dioxygen. Methods Enzymol. 105:3-22. [PubMed]
40. Greenberg, J. T., P. Monach, J. H. Chou, P. D. Josephy, and B. Demple. 1990. Positive control of a global antioxidant defense regulon activated by superoxide-generating agents in Escherichia coli. Proc. Natl. Acad. Sci. USA 87:6181-6185. [PMC free article] [PubMed]
41. Halliwell, B., and J. M. C. Gutteridge. 1984. Oxygen toxicity, oxygen radicals, transition metals and disease. Biochem. J. 219:1-14. [PMC free article] [PubMed]
42. Harada, H., K. Nakajima, K. Sakaue, and Y. Matsuda. 2006. CO2 sensing at ocean surface mediated by cAMP in a marine diatom. Plant Physiol. 142:1318-1328. [PMC free article] [PubMed]
43. Havaux, M., F. Eymery, S. Porfirova, P. Rey, and P. Dörmann. 2005. Vitamin E protects against photoinhibition and photo-oxidative stress in Arabidopsis thaliana. Plant Cell 17:3451-3469. [PMC free article] [PubMed]
44. Hideg, E., C. Barta, T. Kalai, I. Vass, K. Hideg, and K. Asada. 2002. Detection of singlet oxygen and superoxide with fluorescent sensors in leaves under stress by photoinhibition or UV radiation. Plant Cell Physiol. 43:1154-1164. [PubMed]
45. Im, C. S., and A. R. Grossman. 2002. Identification and regulation of high light-induced genes in Chlamydomonas reinhardtii. Plant J. 30:301-313. [PubMed]
46. Im, C. S., Z. Zhang, J. Shrager, C. W. Chang, and A. R. Grossman. 2003. Analysis of light and CO2 regulation in Chlamydomonas reinhardtii using genome-wide approaches. Photosynth. Res. 75:111-125. [PubMed]
47. Imlay, J. A., and S. Linn. 1987. Mutagenesis and stress responses induced in Escherichia coli by hydrogen peroxide. J. Bacteriol. 169:2967-2976. [PMC free article] [PubMed]
48. Inoue, Y., T. Matsuda, K. Sugiyama, S. Izawa, and A. Kimura. 1999. Genetic analysis of glutathione peroxidase in oxidative stress response of Saccharomyces cerevisiae. J. Biol. Chem. 274:27002-27009. [PubMed]
49. Jamieson, D. J. 1992. Saccharomyces cerevisiae has distinct adaptive responses to both hydrogen peroxide and menadione. J. Bacteriol. 174:6678-6681. [PMC free article] [PubMed]
50. Krieger-Liszkay, A. 2004. Singlet oxygen production in photosynthesis. J. Exp. Bot. 56:337-346. [PubMed]
51. Kuge, S., and N. Jones. 1994. YAP1 dependent activation of TRX2 is essential for the response of Saccharomyces cerevisiae to oxidative stress by hydroperoxides. EMBO J. 13:655-664. [PMC free article] [PubMed]
52. Lambert, C., T. Serna, and T. G. Truscott. 1990. Rose Bengal radicals and their reactivity. J. Chem. Soc. Faraday Trans. 86:3879-3882.
53. Ledford, H. K., I. Baroli, J. W. Shin, B. B. Fischer, R. I. L. Eggen, and K. K. Niyogi. 2004. Comparative profiling of lipid-soluble antioxidants and transcripts reveals two phases of photo-oxidative stress in a xanthophyll-deficient mutant of Chlamydomonas reinhardtii. Mol. Genet. Genomics 272:470-479. [PubMed]
54. Lee, J., C. Godon, G. Lagniel, D. Spector, J. Garin, J. Labarre, and M. B. Toledano. 1999. Yap1 and Skn7 control two specialized oxidative stress response regulons in yeast. J. Biol. Chem. 274:16040-16046. [PubMed]
55. Leisinger, U., K. Rüfenacht, B. Fischer, M. Pesaro, A. Spengler, A. J. B. Zehnder, and R. I. L. Eggen. 2001. The glutathione peroxidase homologous gene from Chlamydomonas reinhardtii is transcriptionally up-regulated by singlet oxygen. Plant Mol. Biol. 46:395-408. [PubMed]
56. Leisinger, U., K. Rufenacht, A. J. B. Zehnder, and R. I. L. Eggen. 1999. Structure of a glutathione peroxidase homologous gene involved in the oxidative stress response in Chlamydomonas reinhardtii. Plant Sci. 149:139-149.
57. Li, J., and M. P. Timko. 1996. The pc-1 phenotype of Chlamydomonas reinhardtii results from a deletion mutation in the nuclear gene for NADPH:protochlorophyllide oxidoreductase. Plant Mol. Biol. 30:15-37. [PubMed]
58. Martinez, G. R., A. P. Loureiro, S. A. Marques, S. Miyamoto, L. F. Yamaguchi, J. Onuki, E. A. Almeida, C. C. Garcia, L. F. Barbosa, M. H. Medeiros, and P. Di Mascio. 2003. Oxidative and alkylating damage in DNA. Mutat. Res. 544:115-127. [PubMed]
59. Merkel, P. B., R. Nilsson, and D. R. Kearns. 1971. Deuterium effects on singlet oxygen lifetimes in solutions. A new test of singlet oxygen reactions. J. Am. Chem. Soc. 94:1030-1031.
60. Moan, J. 1990. On the diffusion length of singlet oxygen in cells and tissues. J. Photochem. Photobiol. 6:343-347.
61. Nishiyama, Y., S. I. Allakhverdiev, H. Yamamoto, H. Hayashi, and N. Murata. 2004. Singlet oxygen inhibits the repair of photosystem II by suppressing the translation elongation of the D1 protein in Synechocystis sp. PCC 6803. Biochemistry 43:11321-11330. [PubMed]
62. Okajima, T., and T. Ohsaka. 2003. Chemiluminescence of lucigenin by electrogenerated superoxide ions in aqueous solutions. Luminescence 18:49-57. [PubMed]
63. op den Camp, R. G., D. Przybyla, C. Ochsenbein, C. Laloi, C. Kim, A. Danon, D. Wagner, E. Hideg, C. Göbel, I. Feussner, M. Nater, and K. Apel. 2003. Rapid induction of distinct stress responses after the release of singlet oxygen in Arabidopsis. Plant Cell 15:2320-2332. [PMC free article] [PubMed]
64. Pollock, S. V., S. L. Colombo, D. L. Prout, Jr., A. C. Godfrey, and J. V. Moroney. 2003. Rubisco activase is required for optimal photosynthesis in the green alga Chlamydomonas reinhardtii in a low-CO2 atmosphere. Plant Physiol. 133:1854-1861. [PMC free article] [PubMed]
65. Pollock, S. V., D. L. Prout, A. C. Godfrey, S. D. Lemaire, and J. V. Moroney. 2004. The Chlamydomonas reinhardtii proteins Ccp1 and Ccp2 are required for long-term growth, but are not necessary for efficient photosynthesis in a low-CO2 environment. Plant Mol. Biol. 56:125-132. [PubMed]
66. Ryter, S. W., and R. M. Tyrrell. 1998. Singlet molecular oxygen (1O2): a possible effector of eukaryotic gene expression. Free Radic. Biol. Med. 24:1520-1534. [PubMed]
67. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
68. Sarkar, N., S. Lemaire, D. Wu-Scharf, E. Issakidis-Bourguet, and H. Cerutti. 2005. Functional specialization of Chlamydomonas reinhardtii cytosolic thioredoxin h1 in the response to alkylation-induced DNA damage. Eukaryot. Cell 4:262-273. [PMC free article] [PubMed]
69. Schäfer, F. Q., H. P. Wang, E. E. Kelley, K. L. Cueno, S. M. Martin, and G. R. Buettner. 2002. Comparing beta-carotene, vitamin E and nitric oxide as membrane antioxidants. Biol. Chem. 383:671-681. [PubMed]
70. Schmidt, G. W., K. S. Matlin, and N.-H. Chua. 1977. A rapid procedure for selective enrichment of photosynthetic electron transport mutants. Proc. Natl. Acad. Sci. USA 74:610-614. [PMC free article] [PubMed]
71. Seo, S., M. Okamoto, T. Iwai, M. Iwano, K. Fukui, A. Isogai, N. Nakajima, and Y. Ohashi. 2000. Reduced levels of chloroplast FtsH protein in tobacco mosaic virus-infected tobacco leaves accelerate the hypersensitive reaction. Plant Cell 12:917-932. [PMC free article] [PubMed]
72. Shrager, J., C. Hauser, C. W. Chang, E. H. Harris, J. Davies, J. McDermott, R. Tamse, Z. Zhang, and A. R. Grossman. 2003. Chlamydomonas reinhardtii genome project. A guide to the generation and use of the cDNA information. Plant Physiol. 131:401-408. [PMC free article] [PubMed]
73. Sikorski, R. S., and P. Hieter. 1989. A system of shuttle vectors and yeast host strains designed for efficient manipulation of DNA in Saccharomyces cerevisiae. Genetics 122:19-27. [PMC free article] [PubMed]
74. Singh, S. P., J. A. Coronella, H. Benes, B. J. Cochrane, and P. Zimniak. 2001. Catalytic function of Drosophila melanogaster glutathione S-transferase DmGSTS1-1 (GST-2) in conjugation of lipid peroxidation end products. Eur. J. Biochem. 268:2912-2923. [PubMed]
75. Snyder, J. W., E. Skovsen, J. D. C. Lambert, and P. R. Ogilby. 2005. Subcellular, time-resolved studies of singlet oxygen in single cells. J. Am. Chem. Soc. 127:14558-14559. [PubMed]
76. Stephen, D. W., S. L. Rivers, and D. J. Jamieson. 1995. The role of the YAP1 and YAP2 genes in the regulation of the adaptive oxidative stress responses of Saccharomyces cerevisiae. Mol. Microbiol. 16:415-423. [PubMed]
77. Stiel, H., K. Teuchner, A. Paul, D. Leupold, and I. E. Kochevar. 1996. Quantitative comparison of excited state properties and intensity-dependent photosensitization by rose bengal. J. Photochem. Photobiol. B 33:245-254. [PubMed]
78. Storz, G., L. A. Tartaglia, and B. N. Ames. 1990. Transcriptional regulator of oxidative stress-inducible genes: direct activation by oxidation. Science 248:189-194. [PubMed]
79. Tatsuzawa, H., T. Maruyama, N. Misawa, K. Fujimori, and M. Nakano. 2000. Quenching of singlet oxygen by carotenoids produced in Escherichia coli—attenuation of singlet oxygen-mediated bacterial killing by carotenoids. FEBS Lett. 484:280-284. [PubMed]
80. Thorpe, G. W., C. S. Fong, N. Alic, V. J. Higgins, and I. W. Dawes. 2004. Cells have distinct mechanisms to maintain protection against different reactive oxygen species. Proc. Natl. Acad. Sci. USA 101:6564-6569. [PMC free article] [PubMed]
81. Tsaneva, I. R., and B. Weiss. 1990. soxR, a locus governing a superoxide response regulon in Escherichia coli K-12. J. Bacteriol. 172:4197-4205. [PMC free article] [PubMed]
82. Tusher, V. G., R. Tibshirani, and G. Chu. 2001. Significance analysis of microarrays applied to the ionizing radiation response. Proc. Natl. Acad. Sci. USA 98:5116-5121. [PMC free article] [PubMed]
83. Wagner, D., D. Przybyla, R. op den Camp, C. Kim, F. Landgraf, K. P. Lee, M. Würsch, C. Laloi, M. Nater, E. Hideg, and K. Apel. 2004. The genetic basis of singlet oxygen-induced stress responses of Arabidopsis thaliana. Science 306:1183-1185. [PubMed]
84. Wang, T., P. Arifoglu, Z. Ronai, and K. D. Tew. 2001. Glutathione S-transferase P1-1 (GSTP1-1) inhibits c-Jun N-terminal kinase (JNK1) signaling through interaction with the C terminus. J. Biol. Chem. 276:20999-21003. [PubMed]
85. Wood, M. J., G. Storz, and N. Tjandra. 2004. Structural basis for redox regulation of Yap1 transcription factor localization. Nature 430:917-921. [PubMed]
86. Wu, A. L., and W. S. Moye-Rowley. 1994. GSH1, which encodes gamma-glutamylcysteine synthetase, is a target gene for yAP-1 transcriptional regulation. Mol. Cell. Biol. 14:5832-5839. [PMC free article] [PubMed]

Articles from Eukaryotic Cell are provided here courtesy of American Society for Microbiology (ASM)
PubReader format: click here to try


Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...


Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...