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J. Biol. Chem., Vol. 279, Issue 8, 6337-6344, February 20, 2004
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¶¶||**
From the
Department of Plant and Microbial Biology, University of California, Berkeley, California 94720-3102 and Commissariat à l'Energie Atomique Cadarache, Département d'Ecophysiologie Végétale et de Microbiologie, Laboratoire d'Ecophysiologie de la Photosynthèse, Unité Mixte de Recherche CNRS Commissariat à l'Energie Atomique 163, Université Méditérranée Commissariat à l'Energie Atomique 1000, F-13108 Saint-Paul-lez-Durance, France
Received for publication, November 26, 2003
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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-carotene and some array of oxygenated carotene derivatives, the xanthophylls. In green tissue, the xanthophylls are located in the light-harvesting antennae of the membrane-bound photosystem I (PSI)1 and photosystem II (PSII) complexes. They aid in light harvesting and are essential for maintaining the health of the photosynthetic apparatus by participating in energy dissipation under excessive light and by preventing photo-oxidative damage to the thylakoid membrane through quenching of excited intermediates, such as triplet chlorophylls and singlet oxygen (1-3).
The contents of -carotene and xanthophylls in the chloroplasts of plants and algae are tightly regulated in response to the environment and to the developmental stage of the organism, and their accumulation is coordinated with the biogenesis and assembly of the photosynthetic apparatus. Carotenoid deficiency, caused either by mutation or by the application of herbicides, has devastating effects on chloroplast integrity, interferes with thylakoid membrane biogenesis, and influences the expression of nuclear genes that encode the protein components of the chloroplast (4).
In plants and green algae, the presence of either zeaxanthin or lutein in the chloroplast is necessary for protection against photo-oxidative stress. Zeaxanthin is the only xanthophyll that accumulates exclusively under light stress. Work with mutants impaired in the accumulation of lutein and zeaxanthin has shown that when plants and algae are under high light stress, these pigments are required for the efficient transition of the light-harvesting antenna from a conformation that favors light harvesting to one that allows for thermal dissipation of part of the excess excitation energy (5-7). This thermal dissipation of excess light energy is known as nonphotochemical quenching (NPQ) (8). The process of NPQ is accompanied by the deepoxidation of existing violaxanthin to zeaxanthin in the so-called xanthophyll cycle (9). However, zeaxanthin may also prevent photo-oxidative stress by a mechanism separate from NPQ, such as through direct quenching of excited intermediates and/or scavenging of free radicals (10, 11). It has been suggested as well that zeaxanthin may exert its protective effect by making the thylakoid membrane less permeable to oxygen (12).
Two xanthophyll-deficient mutants of Chlamydomonas reinhardtii are defective in the development of NPQ when exposed to high light (HL), but neither mutation has a lethal effect on growth in HL conditions (5). The npq1 mutant is impaired in the HL-induced de-epoxidation of violaxanthin to antheraxanthin and zeaxanthin, such that zeaxanthin accumulation in HL is prevented, but the mutant can survive the HL treatment (13). The lor1 mutation prevents the constitutive accumulation of lutein and its derivative loroxanthin in Chlamydomonas (14). Even though the lor1 strain has a normal xanthophyll cycle, it is partially defective in NPQ, and it can survive in HL (5). The double mutant npq1 lor1, which accumulates violaxanthin and neoxanthin as the only xanthophylls irrespective of light conditions, shows a severe lack of NPQ and undergoes irreversible bleaching at a photon flux density (PFD) of 500 µmol of photons m-2 s-1, a condition in which the wild type and the single mutants grow normally (5). Suppressors that restore growth of npq1 lor1 in HL include npq2 mutants that cause constitutive accumulation of zeaxanthin, demonstrating that zeaxanthin is sufficient for survival and growth in HL (11).
In this work, we analyzed the behavior of the low light-grown npq1 lor1 double mutant under conditions of oxidative stress and during exposure to HL leading to photobleaching. We monitored the changes in growth, photosynthetic activity, and oxidative stress status in the double mutant in comparison with the wild-type strain. Our results suggest that photobleaching of npq1 lor1 in HL is because of excessive formation of reactive oxygen species (ROS) in the thylakoid membrane due to the absence of the antioxidants, zeaxanthin and lutein. Our results also rule out the possibility that photobleaching of npq1 lor1 is initiated by an enhanced susceptibility to photodamage of the PSII reaction center.
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EXPERIMENTAL PROCEDURES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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To test the effect of pro-oxidants and anaerobiosis on growth in the light, cells were grown in HS medium until they reached a density of 1-2 x 106 cells/ml. At this point, the cells were concentrated by centrifugation to 1 x 107 cells/ml, serial dilutions were prepared, and the cells were immediately spotted onto HS agar medium with or without pro-oxidants. The concentrations of pro-oxidants that were used supported at least partial growth of the wild-type strain. All additives were water-soluble. Rose bengal (Sigma), hydrogen peroxide (Fisher), methyl viologen (Sigma), and metronidazole (Sigma) plates were prepared 1 day prior to use. The plates were incubated in LL for 7 days with the exception of metronidazole plates, which were incubated for 10 days. Anaerobic experiments were carried out in anaerobic BBL GasPak Pouches (BD Biosciences). In this case, the cells were spotted onto plates that were then incubated for 1 day at LL before being transferred to an intermediate PFD of 250 µmol of photons m-2 s-1 (moderate light (ML)). The incubation in LL allowed the pouches to become anaerobic before the cells were exposed to a higher light intensity.
Measurements of Fluorescence and Oxygen Evolution—Chlorophyll (Chl) fluorescence was measured using an FMS2 pulse-amplitude-modulation fluorometer (Hansatech, King's Lynn, UK). The cells, contained in 10-ml culture samples, were deposited onto 2.5-mm diameter 12-µm pore size nitrocellulose filters (Millipore, Bedford, MA) by filtration and were dark-adapted in a moist Petri dish for 15 min prior to the measurements. To avoid variation in fluorescence parameters due to state transitions, the cells were exposed to 8 min of illumination with weak far-red light (light-emitting diode source of 735 nm peak wavelength) prior to determination of Fv/Fm. The light-saturated rate of oxygen evolution was measured with a polarographic Clark-type oxygen electrode (Hansatech) at 25 °C. Intact cells were used in their original growth medium with the addition of 4 mM NaHCO3 as a terminal electron acceptor. Illumination with white light was provided by an LS2 lamp (Hansatech), and the intensity of the incident light was varied with neutral-density filters (Melles-Griot, Irvine, CA). Dark respiration was measured first, followed by measurements of the rate of photosynthesis at three sequentially increasing PFDs (550, 750, and 1000 µmol of photons m-2 s-1) to account for changes in the light intensity necessary for saturation of photosynthesis after the transfer from LL to HL. Only the light-saturated rates of oxygen evolution are shown. The total cellular Chl content and Chl a:Chl b ratios were determined spectrophotometrically in 90% acetone (16).
Measurement of Oxidative Stress Parameters—ROS were measured using the cell-permeant fluorescent dye, 5-(-6)-chloromethyl-2',7'-dichlorodihydrofluorescein diacetate, acetyl ester (CM-H2DCFDA; Molecular Probes, Eugene, OR). The 5-ml culture samples were harvested by centrifugation at 3200 x g for 5 min and resuspended in 750 µl of HS medium (freshly adjusted to pH 5.7 with acetic acid) to bring the total volume to 1 ml. 1.5 µl of CM-H2DCFDA (freshly dissolved in Me2SO; final concentration 10 µM) or a Me2SO-only control were added to the cell samples, which were incubated at room temperature for 15 min in the dark. The cells were then centrifuged and resuspended as before, and fluorescence of 105 cells was measured on an EPICS XL-MCL flow cytometer (Beckman-Coulter, Miami, FL).
The extent of lipid peroxidation in cells was estimated by measuring the formation of thiobarbituric acid-reactive substances (TBARS) and by thermoluminescence. For TBARS determination, 10-ml culture aliquots were taken before and at time intervals after exposure to HL. Butylated hydroxytoluene was added to the samples at a final concentration of 0.01% (w/v) to terminate lipid peroxidation chain reactions, and the content of TBARS was measured as described by Baroli et al. (11). Thermoluminescence measurements were performed with a custom-built apparatus (17). The cells were deposited on 2.5-mm diameter 12-µm pore size nitrocellulose filters (Millipore), which were placed in the copper chamber of the apparatus. The samples were heated from 25 to 150 °C at a rate of 6 °C min-1. Temperature was controlled using a thermocouple mounted between the sample and the heating element. The luminescence signal was recorded during sample heating with a compact red-extended photomultiplier (H5701-50, Hamamatsu, Bridgewater, NJ) shielded with an RG665 glass filter (Schott, Yonkers, NY). Both the sample temperature and thermoluminescence were recorded by a computer using a DaqPad-1200 data acquisition system (National Instruments, Austin, TX). The amplitude of the 135 °C thermoluminescence band was used as an index of lipid peroxidation (10, 18).
Preparation of Protein Extracts, SDS-PAGE, and Immunoblotting—To prepare whole-cell protein extracts for SDS-PAGE, cells were harvested from 15-ml culture aliquots by centrifugation at 3200 x g for 4 min at 4 °C. Protein sample preparation, SDS-PAGE, and immunoblotting were performed as described in Ref. 11. The anti-D1 and anti-LHCII antibodies were kindly provided by A. Melis (University of California, Berkeley, CA) and F.-A. Wollman (Institut de Biologie Physico-Chimique, Paris, France), respectively.
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RESULTS |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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14 h. Fig. 2A shows that, when exposed to HL, the rate of increase in cell density slowed down but persisted in both strains. The wild-type culture had an initial slight decrease in cell viability, as assayed by colony forming units, after transfer to HL (Fig. 2B). The apparent viability of wild-type cells then increased rapidly between 3 h and 6 h, after which viability was maintained at a level that was consistent with the total cell density. In contrast to the wild type, the npq1 lor1 cells showed a substantial decline in viability that was already evident after 6 h in HL. By 24 h of HL treatment, only 20% of the initial cells were able to form colonies (Fig. 2B), but there was a slight increase in cell viability between 24 and 48 h.
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3.5 compared with a value of 2.6 for the wild type (see also Ref. 5). The Chl a:Chl b ratio did not change significantly in either strain during the HL treatment (data not shown). The cellular total Chl content of LL-grown cells was 25% lower in the double mutant compared with the wild type (Fig. 3A). During the first 6 h of HL treatment, the Chl content in the wild type initially decreased 20% and then stabilized at the lower value. In the double mutant, the Chl content decreased progressively upon exposure to HL until it reached only 12% of the initial value after 48 h of HL treatment. At this point, the npq1 lor1 culture contained a mixed population composed of bleached cells, which were devoid of thylakoid membranes when observed with transmission electron microscopy (data not shown), and cells containing at least some Chl. These two kinds of cells could be distinguished by flow cytometry (see below).
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0.65 for the rest of the treatment. The decline in PSII efficiency was not accompanied by a decrease in photosynthetic oxygen evolution, which remained the same during the first 6 h of treatment and then showed a progressive increase followed by stabilization after 24 h in HL (Fig. 3C).
Loss of Photosynthetic Proteins during Exposure to HL—The loss of Chl and photosynthetic activity in the double mutant was indicative of damage to the thylakoid membrane. To characterize the kinetics of this photodamage, we monitored the content of specific photosynthetic proteins. Fig. 4 shows the contents of the PSII reaction center D1 protein and the peripheral LHCII as a function of time in HL as determined by immunoblotting. The wild type showed no significant change in the cellular level of either protein during exposure to HL. In contrast, after a lag of 6 h, the cellular content of the D1 protein decreased dramatically (half-time 6 h) in the double mutant such that only 30% of the initial D1 content remained after 48 h in HL (Fig. 4A). LHCII proteins in npq1 lor1 cells were also lost in HL, but more gradually, with a half-time of 24 h.
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9-fold during exposure of the double mutant to HL, whereas the wild type showed only a slight increase in lipid peroxidation (Fig. 6A). The oxidation of lipids in the proximity of Chl can also be monitored by the luminescence signal emanating from cells heated to temperatures higher than 70 °C (18, 21). Fig. 6B shows that, compared with that of the wild type, in the double mutant, the thermoluminescence signal peaking at 135-145 °C increased considerably during HL illumination, consistent with the TBARS results (Fig. 6A).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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Response of Wild-type Chlamydomonas Cells to a Transition from LL to HL—A light intensity of 500 µmol of photons m-2 s-1 did not appear to be excessive for growth of wild-type Chlamydomonas, but it caused some initial symptoms of light stress. When grown in LL and transferred to HL, the wild type responded with a slowing down of the cell duplication rate (doubling time of 14 h in LL versus >24 h after transfer to HL, Fig. 2A) and an initial slight decrease in cell viability. The number of cells able to form colonies then rose substantially between 6 and 12 h in HL (Fig. 2B), despite the more gradual increase in cell density (Fig. 2A). The discrepancy between cell density and apparent cell viability might be explained by an increase in the plating efficiency of wild-type cells occurring within the first 6 h after transfer from LL to HL.
After the initial stress period in which the Chl/cell and Fv/Fm decreased slightly but significantly (Fig. 3, A and B, respectively), the wild-type cells stabilized at lower values of both parameters. In contrast, the light shift did not have any negative effect on the rate of photosynthetic electron transport, measured as oxygen evolution in intact cells (Fig. 3, C and D), or on the cellular content of the PSII D1 protein (Fig. 4A) consistent with the notion that the HL treatment is not photoinhibitory to the wild type.
The LHCII protein level showed a transient decrease of 10% during the first 12 h in HL, but it stabilized at a level almost identical to the initial one (Fig. 4B). A similar response of the major LHC proteins has been observed in the green alga Dunaliella salina (22). The fact that the Chl a:Chl b ratio remained constant after the shift from LL to HL suggests that there was no preferential down-regulation of the LHC proteins relative to the content of reaction centers within the first 48 h in HL. A similar lack of LHC down-regulation has been observed in wild-type Arabidopsis immediately following transfer from LL to HL (23).
Photo-oxidative Stress in npq1 lor1 Cells—Why does the npq1 lor1 double mutant bleach in HL? Upon transfer from LL to HL, symptoms of light stress in npq1 lor1 cells were first manifested within 3 h as declines in PSII efficiency, measured as Fv/Fm, and photosynthetic oxygen evolution (Fig. 3). The addition of acetate as an organic carbon source did not rescue growth of npq1 lor1 in HL (Fig. 1), however, which would indicate that essential processes other than photosynthesis are being affected by the HL treatment. Cell viability began to decrease between 3 and 6 h (Fig. 2B) followed by loss of D1 and LHCII proteins after6hinHL (Fig. 4) and accumulation of lipid peroxidation products after 12 h in HL (Fig. 6). The fact that lowered oxygen tensions reduced the extent of bleaching in npq1 lor1 at least partially (Fig. 1) suggests the involvement of ROS, yet significantly higher levels of ROS were detected only after at least 12 h in HL (Fig. 7A). Because the intracellular CM-H2DCF probe for ROS is a soluble molecule, it is conceivable that ROS are generated initially in chloroplast membranes where photosynthesis and other processes would be subject to photo-oxidative damage and are not readily detected by the probe until later time points when antioxidant defenses are overwhelmed.
The lack of NPQ in the mutant might be expected to increase ROS levels in thylakoids, primarily resulting in photodamage to PSII that leads to a more general photo-oxidative bleaching. However, npq1 lor1 was not more susceptible than the wild type to PSII photodamage, as indicated by turnover of the D1 protein (Fig. 5). In addition, the recently described npq5 mutant of Chlamydomonas, which has normal xanthophyll composition but lacks NPQ, survives excess light (24). Taken together, these results show that lack of NPQ and enhanced rates of photodamage to PSII are not the primary underlying cause of the HL sensitivity of npq1 lor1, and they point instead to a more general deficiency in antioxidant capacity as the causative factor.
Formation of ROS in the chloroplast is consistent with the widespread damage to the thylakoid membrane that we observed when npq1 lor1 was exposed to HL (see particularly Figs. 4 and 6). The lack of NPQ could increase the yield of singlet oxygen in the Chl antenna of PSII. In addition, higher levels of superoxide and hydrogen peroxide could be produced at PSI due to lack of NPQ. We hoped to provide insights into these possibilities by examining the effect on npq1 lor1 of pro-oxidants that produce different ROS. The double mutant was considerably more sensitive than the wild type to the singlet oxygen generator rose bengal and to the superoxide generators metronidazole and methyl viologen but not to H2O2 (Fig. 8). Thus, we can exclude formation of H2O2 as a cause of enhanced oxidative damage in the mutant. Toxicity of metronidazole and methyl viologen is the result of the ability of these molecules to accept electrons from ferredoxin and subsequently donate electrons to oxygen, generating superoxide. Metronidazole is toxic to Chlamydomonas only in the light, and PSI-deficient mutants are more resistant to the inhibitor, supporting the idea that toxicity occurs because of superoxide generation at the acceptor side of PSI (25). Methyl viologen is capable also of killing Chlamydomonas cells in the dark (25), indicating that its toxicity does not derive entirely from photochemical superoxide production in the chloroplast. Nevertheless, in LL-grown cells, the presence of lutein in the chloroplast of wild-type cells is apparently sufficient to protect against toxicity of methyl viologen (Fig. 8). The ROS indicator dye that we used, CM-H2DCFDA, does not allow a distinction between singlet oxygen and superoxide in vivo and attempts to measure singlet oxygen with a specific probe (DanePy) (26) in intact cells failed because of interfering background fluorescence of Chlamydomonas cells.2 Both singlet oxygen and superoxide could be produced by the mutant in HL.
However, other results reported here indicate that singlet oxygen could be the main ROS produced in HL-exposed npq1 lor1. For example, HL treatment of the mutant led to extensive formation of lipid peroxides (Fig. 6A), a known product of singlet oxygen attack on membrane lipids (27). In addition, it is likely that some of the high-temperature thermoluminescence signal that was detected in npq1 lor1 at wavelengths of >665 nm (Fig. 6B) is attributable to singlet oxygen. This red luminescence can be because of both singlet oxygen and excitation energy transfer from triplet carbonyls to Chl (28). Given that the Chl content of the mutant was strongly reduced in HL (Fig. 3A), the high thermoluminescence signal might be due, at least in part, to singlet oxygen. These results, along with the fact that zeaxanthin and lutein are well known as efficient quenchers of singlet oxygen (reviewed in Ref. 3), are consistent with the hypothesis that higher levels of singlet oxygen are produced in npq1 lor1 in HL. To test this hypothesis, experiments will be performed to examine singlet oxygen-regulated gene expression (29) in npq1 lor1 in response to HL.
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FOOTNOTES |
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Present address: Molecular Plant Physiology Group, Research School of Biological Sciences, Australian National University, P. O. Box 475, Canberra ACT 2601, Australia.
¶ These authors contributed equally to this paper.
|| Supported by National Institutes of Health Predoctoral Genetics Training Grant T32-GM07127.
** Present address: The Johns Hopkins University School of Medicine, 720 Rutland Ave., Baltimore, MD 21205.
To whom correspondence should be addressed. Tel.: 510-643-6602; Fax: 510-642-4995; E-mail: niyogi{at}nature.berkeley.edu.
1 The abbreviations used are: PSI, photosystem I; PSII, photosystem II; Chl, chlorophyll; Fv/Fm, maximum photochemical efficiency of PSII in the dark-adapted state; HL, high light; LHCII, major light-harvesting complex of PSII; LL, low light; ML, moderate light; NPQ, nonphotochemical quenching of Chl fluorescence; PFD, photon flux density; ROS, reactive oxygen species; TBARS, thiobarbituric acid-reactive substances; TAP, Tris-acetate-phosphate; HS, high salt; CM-H2DCFDA, 5-(-6)-chloromethyl-2',7'-dichlorodihydrofluorescein diacetate, acetyl ester.
2 E. Hideg, I. Baroli, and K. K. Niyogi, unpublished results.
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ACKNOWLEDGMENTS |
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REFERENCES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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