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• W2012302334 abstract "We report the characterization of the effects of the A249S mutation located within the binding pocket of the primary quinone electron acceptor, QA, in the D2 subunit of photosystem II in Thermosynechococcus elongatus. This mutation shifts the redox potential of QA by ∼–60 mV. This mutant provides an opportunity to test the hypothesis, proposed earlier from herbicide-induced redox effects, that photoinhibition (light-induced damage of the photosynthetic apparatus) is modulated by the potential of QA. Thus the influence of the redox potential of QA on photoinhibition was investigated in vivo and in vitro. Compared with the wild-type, the A249S mutant showed an accelerated photoinhibition and an increase in singlet oxygen production. Measurements of thermoluminescence and of the fluorescence yield decay kinetics indicated that the charge-separated state involving QA was destabilized in the A249S mutant. These findings support the hypothesis that a decrease in the redox potential of QA causes an increase in singlet oxygen-mediated photoinhibition by favoring the back-reaction route that involves formation of the reaction center chlorophyll triplet. The kinetics of charge recombination are interpreted in terms of a dynamic structural heterogeneity in photosystem II that results in high and low potential forms of QA. The effect of the A249S mutation seems to reflect a shift in the structural equilibrium favoring the low potential form. We report the characterization of the effects of the A249S mutation located within the binding pocket of the primary quinone electron acceptor, QA, in the D2 subunit of photosystem II in Thermosynechococcus elongatus. This mutation shifts the redox potential of QA by ∼–60 mV. This mutant provides an opportunity to test the hypothesis, proposed earlier from herbicide-induced redox effects, that photoinhibition (light-induced damage of the photosynthetic apparatus) is modulated by the potential of QA. Thus the influence of the redox potential of QA on photoinhibition was investigated in vivo and in vitro. Compared with the wild-type, the A249S mutant showed an accelerated photoinhibition and an increase in singlet oxygen production. Measurements of thermoluminescence and of the fluorescence yield decay kinetics indicated that the charge-separated state involving QA was destabilized in the A249S mutant. These findings support the hypothesis that a decrease in the redox potential of QA causes an increase in singlet oxygen-mediated photoinhibition by favoring the back-reaction route that involves formation of the reaction center chlorophyll triplet. The kinetics of charge recombination are interpreted in terms of a dynamic structural heterogeneity in photosystem II that results in high and low potential forms of QA. The effect of the A249S mutation seems to reflect a shift in the structural equilibrium favoring the low potential form. Photosystem II (PSII), 2The abbreviations used are: PSII, photosystem II; P, primary electron donor; 3P, triplet state of the primary electron donor; QA, primary quinone acceptor; QB, secondary quinone acceptor; S0–4, redox states of the charge accumulating part of the water-oxidizing enzyme; Pheo, pheophytin (primary electron acceptor in PSII); BPheo, bacterial pheophytin (primary electron acceptor in bacterial reaction centers); TEMP, 2,2,6,6-tetramethylpiperidine; TEMPO, 2,2,6,6-tetramethylpiperidine-1-oxyl; WT′, T. elongatus strain that lacks the second gene coding for the D2 protein (ΔpsbD2 strain); OEC, oxygen-evolving complex; Chl, chlorophyll; MES, 4-morpholineethanesulfonic acid; DCMU, 13-(3,4-dichlorophenyl)-1,1-dimethylurea. 2The abbreviations used are: PSII, photosystem II; P, primary electron donor; 3P, triplet state of the primary electron donor; QA, primary quinone acceptor; QB, secondary quinone acceptor; S0–4, redox states of the charge accumulating part of the water-oxidizing enzyme; Pheo, pheophytin (primary electron acceptor in PSII); BPheo, bacterial pheophytin (primary electron acceptor in bacterial reaction centers); TEMP, 2,2,6,6-tetramethylpiperidine; TEMPO, 2,2,6,6-tetramethylpiperidine-1-oxyl; WT′, T. elongatus strain that lacks the second gene coding for the D2 protein (ΔpsbD2 strain); OEC, oxygen-evolving complex; Chl, chlorophyll; MES, 4-morpholineethanesulfonic acid; DCMU, 13-(3,4-dichlorophenyl)-1,1-dimethylurea. the water/plastoquinone oxidoreductase, uses light energy to extract four electrons from water, producing oxygen (1Joliot P. Barbieri G. Chabaud R. Photochem. Photobiol. 1969; 10: 309-329Crossref Scopus (515) Google Scholar, 2Kok B. Forbush B. Mc Gloin M. Photochem. Photobiol. 1970; 11: 457-475Crossref PubMed Scopus (1792) Google Scholar, 3Goussias C. Boussac A. Rutherford A.W. Philos. Trans. R. Soc. Lond. B Biol. Sci. 2002; 357: 1369-1381Crossref PubMed Scopus (144) Google Scholar, 4Britt R.D. Ort D.R. Yocum C.F. Oxygenic Photosynthesis: The Light Reactions. Kluwer, Dordrecht1996: 137-164Google Scholar). Each electron is transferred over a chain of redox cofactors to the terminal plastoquinone QB, which accepts two electrons and two protons (5Diner B.A. Babcock G.T. Ort D.R. Yocum C.F. Oxygenic Photosynthesis: The Light Reactions. Kluwer, Dordrecht1996: 213-247Google Scholar). The efficiency of PSII in converting light energy into a charge-separated state is remarkably high (5Diner B.A. Babcock G.T. Ort D.R. Yocum C.F. Oxygenic Photosynthesis: The Light Reactions. Kluwer, Dordrecht1996: 213-247Google Scholar). The univalent photochemistry must interface with the four-electron chemistry occurring at the electron donor side with the two-electron chemistry at the acceptor side. This is achieved by different mechanisms. On the donor side, the so-called oxygen-evolving complex (OEC) accumulates four redox equivalents before it extracts four electrons from two water molecules. Accumulation is necessary, because the energy needed to extract the electrons one by one from water requires more driving force than visible light provides (4Britt R.D. Ort D.R. Yocum C.F. Oxygenic Photosynthesis: The Light Reactions. Kluwer, Dordrecht1996: 137-164Google Scholar). During the catalytic cycle, the OEC exists in different oxidation states. These are designated S0, S1, S2, S3, and S4, where the subscript indicates the number of accumulated oxidation equivalents. The cycle is completed when the S4 state performs a 4-electron oxidation of water, with the OEC being returned to the most reduced of the so-called S states, S0. The OEC consists of four manganese ions, one calcium ion (3Goussias C. Boussac A. Rutherford A.W. Philos. Trans. R. Soc. Lond. B Biol. Sci. 2002; 357: 1369-1381Crossref PubMed Scopus (144) Google Scholar, 4Britt R.D. Ort D.R. Yocum C.F. Oxygenic Photosynthesis: The Light Reactions. Kluwer, Dordrecht1996: 137-164Google Scholar, 6Debus R.J. Biochim. Biophys. Acta. 1992; 1102: 269-352Crossref PubMed Scopus (1083) Google Scholar, 7Ferreira K.N. Iverson T.M. Maghlaoui K. Barber J. Iwata S. Science. 2004; 303: 1831-1838Crossref PubMed Scopus (2796) Google Scholar), and probably one chloride ion (Ref.8Wincencjusz H. Yocum C.F. van Gorkom H.J. Biochemistry. 1999; 38: 3719-3725Crossref PubMed Scopus (77) Google Scholar, but see also Ref. 9Olesen K. Andréasson L.-E. Biochemistry. 2003; 42: 2025-2035Crossref PubMed Scopus (89) Google Scholar). On the electron acceptor side, electron transfer involves two plastoquinones, the primary and secondary quinone acceptors (QA and QB). Although both are plastoquinones, their physical and chemical properties differ. QA is tightly bound and acts as a one-electron acceptor, with a short-lived semiquinone state that undergoes no observable protonation events during its lifetime (5Diner B.A. Babcock G.T. Ort D.R. Yocum C.F. Oxygenic Photosynthesis: The Light Reactions. Kluwer, Dordrecht1996: 213-247Google Scholar). QB, on the other hand, acts as a two-electron and two-proton acceptor with a stable semiquinone intermediate, QB– (5Diner B.A. Babcock G.T. Ort D.R. Yocum C.F. Oxygenic Photosynthesis: The Light Reactions. Kluwer, Dordrecht1996: 213-247Google Scholar, 10Bouges-Bocquet B. Biochim. Biophys. Acta. 1973; 314: 250-256Crossref PubMed Scopus (188) Google Scholar, 11Velthuys B.R. Amesz J. Biochim. Biophys. Acta. 1974; 333: 85Crossref PubMed Scopus (268) Google Scholar). Although the semiquinone state, QB–, is tightly bound, its quinone and quinol forms are exchangeable with the quinone pool in the membrane (12Wraight C.A. Israel J. Chem. 1981; 21: 348-354Crossref Scopus (189) Google Scholar, 13Velthuys B.R. FEBS Lett. 1981; 126: 277-281Crossref Scopus (237) Google Scholar). PSII is known to be susceptible to damage under high light intensities, a process called photoinhibition. Under physiological conditions the D1 protein of PSII has the highest turnover rate of all the proteins in the cell (14Mattoo A.K. Pick U. Hoffman-Falk H. Edelman M. Proc. Natl. Acad. Sci. U. S. A. 1981; 78: 1572-1576Crossref PubMed Scopus (220) Google Scholar). This is usually assumed to be the result of photodamage mediated by PSII photochemistry (Ref. 15Rutherford A.W. Krieger-Liszkay A. Trends Biochem. Sci. 2001; 26: 648-653Abstract Full Text Full Text PDF PubMed Scopus (234) Google Scholar; however, see also Ref. 16Hakala M. Tuominen I. Keranen M. Tyystjarvi T. Tyystjarvi E. Biochim. Biophys. Acta. 2005; 1706: 68-80Crossref PubMed Scopus (321) Google Scholar). It has been proposed that during charge recombination reactions, chlorophyll triplet-mediated, singlet oxygen (1O2) is produced and that this is the species responsible for the PSII damage (17Vass I. Styring S. Hundal T. Koivuniemi A. Aro E. Andersson B. Proc. Natl. Acad. Sci. U. S. A. 1992; 89: 1408-1412Crossref PubMed Scopus (445) Google Scholar, 18Macpherson A.N. Telfer A. Barber J. Truscott T.G. Biochim. Biophys. Acta. 1993; 1143: 301-309Crossref Scopus (179) Google Scholar, 19Hideg E. Spetea C. Vass I. Biochim. Biophys. Acta. 1994; 1186: 143-152Crossref Scopus (179) Google Scholar, 20Keren N. Berg A. van Kan P.J. Levanon H. Ohad I.I. Proc. Natl. Acad. Sci. U. S. A. 1997; 94: 1579-1584Crossref PubMed Scopus (242) Google Scholar, 21Trebst A. Depka B. Hollander-Czytko H. FEBS Lett. 2002; 516: 156-160Crossref PubMed Scopus (197) Google Scholar, 22Fufezan C. Rutherford A.W. Krieger-Liszkay A. FEBS Lett. 2002; 532: 407-410Crossref PubMed Scopus (149) Google Scholar). Conditions that favor charge recombination reactions (e.g. either high light intensities or excitation by single turnover flashes (23Keren N. Gong H. Ohad I. J. Biol. Chem. 1995; 270: 806-814Abstract Full Text Full Text PDF PubMed Scopus (92) Google Scholar)) induce a decrease in PSII activity. This is due to the fact that under photoinhibitory conditions the D1 protein is damaged and degraded faster than it is replaced (for review see Refs. 24Prasil O. Adir N. Ohad I. Barber J. The Photosystems: Structure, Function and Molecular Biology. Elsevier, Amsterdam, The Netherlands1992: 295-348Crossref Google Scholar and 25Aro E.M. Virgin I. Andersson B. Biochim. Biophys. Acta. 1993; 1143: 113-134Crossref PubMed Scopus (1882) Google Scholar). Different classes of herbicides induce different rates of photoinhibition. Phenolic herbicides stimulate the process of photoinhibition relative to urea-type herbicides, and under some in vitro conditions urea herbicides appear to afford some protection against photoinhibition relative to the herbicide-free material (26Nakajima Y. Yoshida S. Ono T. Plant Cell Physiol. 1996; 37: 673-680Crossref Scopus (31) Google Scholar, 27Komenda J. Koblizek M. Prasil O. Photosynth. Res. 2000; 63: 135-144Crossref PubMed Scopus (10) Google Scholar, 28Kyle D.J. Ohad I. Arntzen C.J. Proc. Natl. Acad. Sci. U. S. A. 1984; 81: 4070-4074Crossref PubMed Google Scholar, 29Pallett K.E. Dodge A.D. J. Exp. Bot. 1980; 31: 1051-1066Crossref Scopus (45) Google Scholar, 30Jansen M.A. Depka B. Trebst A. Edelman M. J. Biol. Chem. 1993; 268: 21246-21252Abstract Full Text PDF PubMed Google Scholar). This was explained by the fact that the binding of herbicides influences the midpoint potential of the redox couple QA/QA– (31Krieger-Liszkay A. Rutherford A.W. Biochemistry. 1998; 37: 17339-17344Crossref PubMed Scopus (149) Google Scholar). The urea type herbicide DCMU raises the midpoint potential of QA by 50 mV, whereas the phenolic herbicide bromoxynil lowers it by 45 mV (31Krieger-Liszkay A. Rutherford A.W. Biochemistry. 1998; 37: 17339-17344Crossref PubMed Scopus (149) Google Scholar). The relationship between the redox potential of QA and photoinhibition was explained in terms of the following model (15Rutherford A.W. Krieger-Liszkay A. Trends Biochem. Sci. 2001; 26: 648-653Abstract Full Text Full Text PDF PubMed Scopus (234) Google Scholar, 31Krieger-Liszkay A. Rutherford A.W. Biochemistry. 1998; 37: 17339-17344Crossref PubMed Scopus (149) Google Scholar). The back-reaction between the radical pair [P+⋅QA−⋅] and the ground state P occurs via two main pathways: an indirect pathway involving the repopulation of the [P+⋅Pheo−⋅] radical pair and a direct pathway involving direct tunneling to the ground state (Refs. 15Rutherford A.W. Krieger-Liszkay A. Trends Biochem. Sci. 2001; 26: 648-653Abstract Full Text Full Text PDF PubMed Scopus (234) Google Scholar, 31Krieger-Liszkay A. Rutherford A.W. Biochemistry. 1998; 37: 17339-17344Crossref PubMed Scopus (149) Google Scholar, 32Johnson G.N. Rutherford A.W. Krieger A. Biochim. Biophys. Acta. 1995; 1229: 202-207Crossref Scopus (161) Google Scholar; see also Ref. 33Van Gorkom H.J. Photosynth. Res. 1985; 6: 97-112Crossref PubMed Scopus (120) Google Scholar). The yields of both pathways are modulated by the difference in the free energy between the [P+⋅QA−⋅] and [P+⋅Pheo−⋅] radical pairs (Refs. 15Rutherford A.W. Krieger-Liszkay A. Trends Biochem. Sci. 2001; 26: 648-653Abstract Full Text Full Text PDF PubMed Scopus (234) Google Scholar, 31Krieger-Liszkay A. Rutherford A.W. Biochemistry. 1998; 37: 17339-17344Crossref PubMed Scopus (149) Google Scholar, 32Johnson G.N. Rutherford A.W. Krieger A. Biochim. Biophys. Acta. 1995; 1229: 202-207Crossref Scopus (161) Google Scholar; see also Ref. 33Van Gorkom H.J. Photosynth. Res. 1985; 6: 97-112Crossref PubMed Scopus (120) Google Scholar). If this gap is small, i.e. in the presence of a phenolic herbicide, the back-reaction occurs with a higher yield via the indirect pathway forming a significant yield of 3P. This 3P can then react with 3O2 forming the toxic singlet oxygen species, 1O2. When the energy gap is increased, i.e. in the presence of DCMU, the probability of forming the [P+⋅Pheo−⋅] state is lowered, and the chances are higher that the [P+⋅QA−⋅] radical pair decays via the direct and safe route. EPR-spin trapping experiments supported this model, because the yield of 1O2 formation was significantly lower in the presence of DCMU than in the presence of bromoxynil (22Fufezan C. Rutherford A.W. Krieger-Liszkay A. FEBS Lett. 2002; 532: 407-410Crossref PubMed Scopus (149) Google Scholar). To study further the influence of the redox potential of QA on the rate of photoinhibition we attempted to change its midpoint potential without using herbicides, thereby avoiding any potential difficulties from possible secondary effects of these chemicals. Thus we created mutants with a single point mutation in the QA binding pocket in Thermosynechococcus elongatus, the thermophilic cyanobacterium from which the structure of PSII was obtained by crystallographic methods (7Ferreira K.N. Iverson T.M. Maghlaoui K. Barber J. Iwata S. Science. 2004; 303: 1831-1838Crossref PubMed Scopus (2796) Google Scholar, 34Loll B. Kern J. Saenger W. Zouni A. Biesiadka J. Nature. 2005; 438: 1040-1044Crossref PubMed Scopus (1583) Google Scholar). One such mutant was used to study the influence of the midpoint potential of QA on charge recombination reactions and on photoinhibition. The results provide independent support (in the absence of herbicides) for the relationship between the redox potential and photoinhibition, and hence for the charge recombination model. In addition this study gives new mechanistic insights concerning the presence of a dynamic structural heterogeneity around the QA site and how this influences the redox potential. Strain and Standard Culture Conditions—T. elongatus cells were grown in a rotary shaker (120 rpm) at 45 °C under continuous illumination from fluorescent white lamps giving an intensity of ∼80 microeinstein m–2 s–1. Cells were grown in a DTN micro Einstein medium (35Roncel M. Boussac A. Zurita J.L. Bottin H. Sugiura M. Kirilovsky D. Ortega J.M. J. Biol. Inorg. Chem. 2003; 8: 206-216Crossref PubMed Scopus (66) Google Scholar) in an enriched CO2 atmosphere and in a 3-liter Erlenmeyer flask (1.5-liter culture). Cells were grown in the presence of spectinomycin (25 μgml–1) and streptomycin (10 μg ml–1). Mutant strains were grown additionally in the presence of the antibiotic kanamycin (40 μgml–1). Plasmids, in Vitro Mutagenesis, and Transformation of T. elongatus Cells—Site-directed mutations were performed using the Qiagen site-directed mutagenesis kit on the plasmid pUC18–43H (provided by Dr. M. Sugiura, see Ref. 36Sugiura M. Inoue Y. Plant Cell Physiol. 1999; 40: 1219-1231Crossref PubMed Scopus (148) Google Scholar). The pUC18-CP43H plasmid consists of the coding region for psbD1 and psbC (gene product CP43), with a sequence coding for His6 tag and the kanamycin resistance cassette. The construct is surrounded by the non-coding region of the psbD1 and psbC to provide a higher target sequence for the homologous recombination. The primers used to insert the point mutation in the pUC18-CP43H plasmid that resulted in the A249S mutation were 5′-GAA GAG ACC TAC TCG ATG GTG ACG AGT AAC CGG TTT TGG AGC CAA-3′ and its complementary strand. The A249S mutation was introduced by changing GCG to AGT (bold). Additionally, a conservative point mutation was introduced into the third position of the codon for Arg-251, changing CGT to CGG (underlined). Together these modifications introduced a new AgeI restriction site, which was used for fast screening of potential successfully transformed clones. Amplification of the mutated pUC18-H43 plasmid was done in Escherichia coli. The plasmid was purified and used to transform a strain of T. elongatus lacking the psbD2 gene by electroporation (provided by Dr. M. Sugiura, see Ref. 36Sugiura M. Inoue Y. Plant Cell Physiol. 1999; 40: 1219-1231Crossref PubMed Scopus (148) Google Scholar). The psbD2 gene codes the second copy of D2. We will further refer to the ΔpsbD2 strain as WT′. The transformation was done as described in Kirilovsky et al. (37Kirilovsky D. Roncel M. Boussac A. Wilson A. Zurita J.L. Ducruet J.M. Bottin H. Sugiura M. Ortega J.M. Rutherford A.W. J. Biol. Chem. 2004; 279: 52869-52880Abstract Full Text Full Text PDF PubMed Scopus (30) Google Scholar). Genomic DNA was isolated from T. elongatus cells essentially as described in Cai and Wolk (38Cai Y.P. Wolk C.P. J. Bacteriol. 1990; 172: 3138-3145Crossref PubMed Scopus (396) Google Scholar). PSII Core Complexes Preparation—PSII core complexes were prepared as described by Roncel et al. (35Roncel M. Boussac A. Zurita J.L. Bottin H. Sugiura M. Kirilovsky D. Ortega J.M. J. Biol. Inorg. Chem. 2003; 8: 206-216Crossref PubMed Scopus (66) Google Scholar) with the modifications described in Kirilovsky et al. (37Kirilovsky D. Roncel M. Boussac A. Wilson A. Zurita J.L. Ducruet J.M. Bottin H. Sugiura M. Ortega J.M. Rutherford A.W. J. Biol. Chem. 2004; 279: 52869-52880Abstract Full Text Full Text PDF PubMed Scopus (30) Google Scholar) except that the detergent concentration for the washing steps of the column was 0.03%. The preparations used in this work had an oxygen evolution activity of 2.2–3 mmol of O2 (mg of Chl)–1 h–1. Oxygen Evolution Measurements—Oxygen evolution was measured at 25 °C by polarography using a Clark-type oxygen electrode with saturating white light. Oxygen evolution in cells (10 μg of Chl.ml–1), thylakoid membranes (10 μg of Chl.ml–1), and PSII core complexes (5 μg of Chl.ml–1) was measured in 40 mm MES, pH 6.5, 15 mm MgCl2, 15 mm CaCl2, 10% (v/v) glycerol, 1 m glycinebetaine, and in the presence of 0.5 mm 2,6-dichloro-p-benzoquinone (dissolved in ethanol) as electron acceptor. Thermoluminescence Measurements—Thermoluminescence was measured with a customized apparatus as described in a previous study (39Ducruet J.-M. J. Exp. Bot. 2003; 54: 2419Crossref PubMed Scopus (113) Google Scholar). Cells were centrifuged and resuspended in a 40 mm MES (pH 6.5) buffer containing 15 mm MgCl2, 15 mm CaCl2, 10% glycerol, at a Chl concentration of 100 μgml–1. After dark adaptation (15 min) the cells were frozen to –80 °C in darkness and were maintained at that temperature for 30 min. After slow thawing, the cell suspension was incubated on ice in darkness. PSII complexes were measured at a concentration of 35 μg of Chl.ml–1. The particles were kept in complete darkness (for at least 30 min). To measure the thermoluminescence originating from the S2/3QB– charge recombination (B-Band) (40Rutherford A.W. Crofts A.R. Inoue Y. Biochim. Biophys. Acta. 1982; 682: 457-465Crossref Scopus (262) Google Scholar) the samples were incubated for 5 min in the dark at 40 °C and then flashed once or twice at 1 °C. To measure the thermoluminescence originating from the S2QA– recombination (Q-Band), the dark-adapted samples were flashed once and incubated with either 20 μm DCMU or 100 μm bromoxynil or without any additions at 40 °C for 3 min. After this pre-treatment T. elongatus cells were cooled down to –5 °C and flashed. Although one flash should be sufficient to introduce the charge-separated state in most of the reaction centers two flashes were needed in T. elongatus to obtain the largest Q band. This may reflect a large concentration of QB– in darkness (41Fufezan C. Zhang C. Krieger-Liszkay A. Rutherford A.W. Biochemistry. 2005; 44: 12780-12789Crossref PubMed Scopus (42) Google Scholar) that results in incomplete DCMU binding under the conditions of the experiment. For luminescence detection the samples were warmed at a constant rate (0.5 °C s–1) from 1 °C or –5 °C to 80 °C. When TL was measured in PSII cores the sample concentration was adjusted based on the amplitude of stable EPR signal TyrD·. Singlet Oxygen Measurements—Spin-trapping assays were performed in PSII particles at a concentration of 10 μg of Chl.ml–1. Samples were illuminated for up to 60 min with 1.5 millieinstein m–2 s–1 red light in the presence of 10 mm 2,2,6,6-tetramethyliperidine (TEMP), 4% (v/v) methanol, and buffer (40 mm MES, 15 mm MgCl2, 15 mm CaCl2, pH 6.5). If 1O2 is produced, it can react with TEMP and forms thereby the stable nitroxyl radical TEMPO, the amount of which is linearly correlated to the unsaturated EPR signal. To compare the rate of singlet oxygen production in different preparations from different batches, the signal sizes were normalized. This was done by calculating a factor between the size of the dark signal and the signal after 30- and 60-min illumination, respectively. In control experiments, the EPR signal present immediately after mixing the spin trap with PSII did not change its amplitude upon incubation in the dark for 60 min. EPR signals in dark controls result from impurities in the spin trap. X-band spectra were recorded at room temperature with a Bruker ESP 200 spectrometer at 9.7-GHz microwave frequency, 63-milliwatt microwave power, 100-kHz modulation frequency, and a modulation amplitude of 2 Gauss. Photoinhibition Assays in Vitro and in Vivo—Photoinhibition experiments in vivo were performed with a Chl concentration of 10 μg of Chl.ml–1. Cells were incubated at 30 °C in the presence of 34 μgml–1 chloramphenicol in a glass tube (3-cm diameter) while being stirred and with illumination from 3 Atralux spots of 150 watts (1 millieinstein m–2 s–1 each lamp). The effective light intensity encountered by each cell was diminished by approximately half due to the vessel used. For the photoinhibition experiments in vitro, PSII complexes were illuminated in a modulated fluorometer (PAM; Walz, Effelrich, Germany) adapted to a Hansatech oxygen electrode as previously described (42El Bissati K. Delphin E. Murata N. Etienne A. Kirilovsky D. Biochim. Biophys. Acta. 2000; 1457: 229-242Crossref PubMed Scopus (132) Google Scholar). All the experiments were carried out in a stirred cuvette of 1-cm diameter (32 °C) at a chlorophyll concentration of 3 μg of Chl.ml–1. The decrease of the yield of chlorophyll fluorescence was monitored in the modulated fluorometer during illumination with white light at 1.5 millieinstein m–2 s–1. Redox Titrations of QA Using Fluorescence Yield—Redox titrations were performed as described by Krieger et al. (43Krieger A. Rutherford A.W. Johnson G.N. Biochim. Biophys. Acta. 1995; 1229: 193-201Crossref Scopus (145) Google Scholar). Samples were diluted to a concentration of 15 μg of Chl.ml–1. The level of the variable fluorescence was used as a measurement for the reduction state of QA. During measurements the samples were kept under argon. No redox mediators were used (see Ref. 43Krieger A. Rutherford A.W. Johnson G.N. Biochim. Biophys. Acta. 1995; 1229: 193-201Crossref Scopus (145) Google Scholar). Potentials were measured with a combined Pt/AgCl/Ag electrode and normalized to the standard hydrogen electrode by calibrating the electrode with quinhydrone (a midpoint redox potential (Em) of 286 mV at pH 6.5, 25 °C). Reductive titrations were performed by gradual addition of sodium dithionite (dissolved in 40 mm MES, pH 6.5), oxidative titrations by addition of potassium ferricyanide. Addition of dithionite did not significantly change the pH of the medium. Fluorescence was measured with a PAM 101 fluorometer (Walz, Effeltrich, Germany) using a weak measuring light of 1.6 kHz. Measurement of the S2QA– Charge Recombination—The decrease of the maximal fluorescence yield in the seconds to minutes time range during photoinhibition was measured by using a modulated fluorometer (PAM, Walz). The decay of the fluorescence yield due to S2QA– charge recombination was measured using a double modulation fluorometer (PSI Instruments, Brno, Czech Republic) in the 1-ms to 30-s time range in whole cells (5 μg of Chl.ml–1), in the presence of 20 μm DCMU (dissolved in ethanol). To follow the reoxidation of QA–, a weak non-actinic probe flash was used. A small fraction of photons (∼12% in closed centers) of the probing flash is re-emitted as fluorescence. The variable fluorescence yield was measured with the same fluorometer. Measurements of the decay of S2QA– are not straightforward when measured as the decay of the fluorescence yield (44Joliot P. Joliot A. C. R. Hebd. Seances Acad. Sci. Paris. 1964; 258: 4622-4625PubMed Google Scholar, 45Lavergne J. Trissl H.W. Biophys. J. 1995; 68: 2474-2492Abstract Full Text PDF PubMed Scopus (224) Google Scholar, 46Cuni A. Xiong L. Sayre R. Rappaport F. Lavergne J. Phys. Chem. Chem. Phys. 2004; 6: 4825-4831Crossref Scopus (54) Google Scholar). It is known that the trapping efficiency of open reaction centers increases with the amount of closed reaction centers in the vicinity (44Joliot P. Joliot A. C. R. Hebd. Seances Acad. Sci. Paris. 1964; 258: 4622-4625PubMed Google Scholar, 45Lavergne J. Trissl H.W. Biophys. J. 1995; 68: 2474-2492Abstract Full Text PDF PubMed Scopus (224) Google Scholar). Consequently, a fluorescence probe pulse does not detect closed and opened reaction centers equally, but instead open centers are favored as a function of the fraction of closed centers according to Equation 1, F(c)=c1+J−J⋅c(Eq. 1) where c is the fraction of closed centers, F(c) is the measured fluorescence yield, J is the Joliot antenna connectivity factor (44Joliot P. Joliot A. C. R. Hebd. Seances Acad. Sci. Paris. 1964; 258: 4622-4625PubMed Google Scholar), and J + 1 is the average number of visited reaction centers per exciton. From Equation 1 it is easily deducible that 1) the observed decay kinetics of the fluorescence yield (i.e. F(c(t))) is non-linearly dependent on the re-oxidation kinetics of QA– (i.e. c(t)) if J > 0) and 2) the F(c(t)) is equal to the kinetics of the re-oxidation of QA– (i.e. c(t)) if J = 0). It was shown by Cuni et al. (46Cuni A. Xiong L. Sayre R. Rappaport F. Lavergne J. Phys. Chem. Chem. Phys. 2004; 6: 4825-4831Crossref Scopus (54) Google Scholar) that, if a weak enough excitation flash is applied, which induces charge separation in only a small proportion of the reaction centers (<15%), no correction is needed, just as if J = 0. This was always the case under our conditions. Under the conditions used, even with the lowest usable probe light intensity, the fluorescence yield probe flashes were only found to be non-actinic when a fluorescence probe pulse frequency of 5 Hz or less was used. However, with measuring pulses spaced at 200-ms intervals it is not possible to obtain a good measurement of the fast phase of decay. Therefore we had to use a different measuring protocol. In this protocol each sample was measured under four conditions resulting in four sets of experiments. Each set had a time base shifted by 50 ms up to 2 s and the same time base from that point forward. The kinetic traces (usually three) of each set were averaged, and the sets were merged together resulting in: (a) a higher temporal resolution for the fast kinetic phase and (b) a better signal-to-noise ratio for the slow kinetic phase. This protocol, however, introduces an uneven weighting of the data points, because each of the points from 0 to 2 s after the flash originate from only one set, whereas each point from 2 to 30 s represents the average of all four sets. To compensate for this effect the points from 2 to 30 s were weighted appropriately, i.e. th" @default.
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• W2012302334 title "Influence of the Redox Potential of the Primary Quinone Electron Acceptor on Photoinhibition in Photosystem II" @default.
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