Planta (2012) 235:729–745 DOI 10.1007/s00425-011-1537-2

ORIGINAL ARTICLE

Nitrogen deprivation results in photosynthetic hydrogen production in Chlamydomonas reinhardtii Gabriele Philipps • Thomas Happe Anja Hemschemeier

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Received: 27 June 2011 / Accepted: 6 October 2011 / Published online: 22 October 2011 Ó Springer-Verlag 2011

Abstract The unicellular green alga Chlamydomonas reinhardtii is able to use photosynthetically provided electrons for the production of molecular hydrogen by an [FeFe]hydrogenase HYD1 accepting electrons from ferredoxin PetF. Despite the severe sensitivity of HYD1 towards oxygen, a sustained and relatively high photosynthetic hydrogen evolution capacity is established in C. reinhardtii cultures when deprived of sulfur. One of the major electron sources for proton reduction under this condition is the oxidation of starch and subsequent non-photochemical transfer of electrons to the plastoquinone pool. Here we report on the induction of photosynthetic hydrogen production by Chlamydomonas upon nitrogen starvation, a nutritional condition known to trigger the accumulation of large deposits of starch and lipids in the green alga. Photochemistry of photosystem II initially remained on a higher level in nitrogen-starved cells, resulting in a 2-day delay of the onset of hydrogen production compared with sulfur-deprived cells. Furthermore, though nitrogen-depleted cells accumulated large amounts of starch, both hydrogen yields and the extent of starch degradation were significantly lower than upon sulfur

G. Philipps AG Photobiotechnologie, Fakulta¨t fu¨r Biologie und Biotechnologie, Lehrstuhl fu¨r Biochemie der Pflanzen, Ruhr-Universita¨t Bochum, ND2/132, 44780 Bochum, Germany T. Happe AG Photobiotechnologie, Fakulta¨t fu¨r Biologie und Biotechnologie, Lehrstuhl fu¨r Biochemie der Pflanzen, Ruhr-Universita¨t Bochum, ND2/169, 44780 Bochum, Germany A. Hemschemeier (&) AG Photobiotechnologie, Fakulta¨t fu¨r Biologie und Biotechnologie, Lehrstuhl fu¨r Biochemie der Pflanzen, Ruhr-Universita¨t Bochum, ND2/134, 44780 Bochum, Germany e-mail: [email protected]

deficiency. Starch breakdown rates in nitrogen or sulfurstarved cultures transferred to darkness were comparable in both nutritional conditions. Methyl viologen treatment of illuminated cells significantly enhanced the efficiency of photosystem II photochemistry in sulfur-depleted cells, but had a minor effect on nitrogen-starved algae. Both the degradation of the cytochrome b6f complex which occurs in C. reinhardtii upon nitrogen starvation and lower ferredoxin amounts might create a bottleneck impeding the conversion of carbohydrate reserves into hydrogen evolution. Keywords Chlamydomonas reinhardtii  Hydrogen production  Nitrogen deprivation  Photosynthesis  Starch Abbreviations ATP Adenosine triphosphate Chl Chlorophyll DCMU 3-(3,4-dichlorophenyl)-1,1-dimethylurea LHCII Light-harvesting complex of PSII NAD(P)H Nicotinamide adenine dinucleotide (phosphate) PQ Plastoquinone PSI, PSII Photosystem I, II Rubisco Ribulose-1,5-bisphosphate carboxylase/ oxygenase

Introduction The unicellular green alga Chlamydomonas reinhardtii is a photosynthetic flagellate that has been used as a model for studying principles of oxygenic photosynthesis, flagella function, cell cycle and circadian rhythms as well as responses to nutrient deprivation for decades (Grossman 2000; Harris 2001; Merchant et al. 2006, 2007; Grossman

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et al. 2007; Stern et al. 2008). For several years, the microalga is also being investigated in terms of biotechnological applications with special emphasis on photosynthetic lipid accumulation and production of molecular hydrogen (H2) (Melis 2007; Ghirardi et al. 2009; Wang et al. 2009; Siaut et al. 2011). It is known since the late 1930s that some species of unicellular green algae are able to metabolize H2 both in the dark and in connection with photosynthetic electron transport (reviewed by Melis and Happe 2004). The biochemical and genetic foundations of H2 evolution were revealed more than 50 years later, when the hydrogenase enzyme HYD1 of C. reinhardtii, responsible for proton reduction and H2 oxidation, was purified and analyzed (Roessler and Lien 1984; Happe and Naber 1993; Happe et al. 1994) and the HYD1 gene was isolated (Happe and Kaminski 2002). HYD1 has been shown to be located in the chloroplast (Happe et al. 1994) and to accept electrons from photosynthetic ferredoxin PetF (Happe and Naber 1993; Winkler et al. 2009). C. reinhardtii produces H2 in the dark, probably by means of fermentative ferredoxin reduction (Gfeller and Gibbs 1984; Philipps et al. 2011), but the exact routes of electron transfer to the hydrogenase in the dark are still unknown. In the light, HYD1 is coupled to photosynthetic electron transport via reduction of PetF by photosystem I (PSI) (Stuart and Gaffron 1972; Gfeller and Gibbs 1985; Winkler et al. 2009). A H2 metabolism in Chlamydomonas can only be observed under anaerobic conditions, since [FeFe]-hydrogenases are very sensitive towards molecular oxygen (O2) (Ghirardi et al. 1997; Goldet et al. 2009; Stripp et al. 2009). Algal cells which were adapted to anoxic conditions by flushing the suspension with argon or nitrogen gas in the dark accumulate HYD1 transcripts and synthesize the hydrogenase proteins, resulting in a rapidly increasing in vitro hydrogenase activity (Happe and Naber 1993; Happe and Kaminski 2002; Forestier et al. 2003). Cells thus pretreated and transferred to sudden illumination will evolve significant amounts of H2 gas for a short period of time until photosynthetic O2 evolution by photosystem II (PSII) inhibits the hydrogenase enzymes and the Calvin cycle has taken over its function as the major electron sink of photosynthetic electron transport (Stuart and Gaffron 1972; Cinco et al. 1993). Since this artificially induced H2 metabolism of C. reinhardtii is of a very transient nature, it was not regarded to have biotechnological significance until the year 2000, when a study showed that sulfur (S-) deprived Chlamydomonas cultures establish the capacity of evolving considerable amounts of H2 for several days (Melis et al. 2000). Upon sulfur deficiency, illuminated cultures of the microalga undergo metabolic changes which provide the three most important preconditions for sustained production of significant amounts of H2 gas, namely anaerobic conditions, a substantial resource of low-potential electrons and

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the diminishment of competing electron sinks (reviewed for example by Melis 2007; Ghysels and Franck 2010; Hemschemeier and Happe 2011). Anaerobic (O2-free) or rather hypoxic (O2-limited) conditions are established in sealed algal cultures because the lack of sulfur results in a gradual inactivation and degradation of PSII complexes (Wykoff et al. 1998). While PSII-catalyzed O2 evolution decreases, respiratory O2 uptake remains on a higher level (Melis et al. 2000; Hemschemeier et al. 2008a), resulting in consumption of O2 in closed flasks. As soon as the O2 concentration in the algal cells is sufficiently low, HYD1 transcript and HYD1 protein accumulate and H2 evolution can be detected (Melis et al. 2000; Zhang et al. 2002; Hemschemeier et al. 2008b; Jacobs et al. 2009). Electron sources for H2 generation by S-depleted C. reinhardtii cells are both a residual water oxidizing activity of PSII and the oxidation of starch (Melis et al. 2000; Fouchard et al. 2005; Kruse et al. 2005; Hemschemeier et al. 2008a; Chochois et al. 2009). The latter is accumulated in S-deficient algae as a general response to nutrient starvation (Grossman 2000). When an S-deprived Chlamydomonas culture has passed to the hypoxic, H2-producing stage, electrons probably resulting from oxidative starch degradation are transferred to the PQ pool by the Nda2 enzyme, a plastidic NAD(P)H dehydrogenase of the class II type (Mus et al. 2005; Jans et al. 2008; Desplats et al. 2009). Electrons are then transported via the cytochrome b6f complex and PSI to ferredoxin PetF, which transfers them to the [FeFe]-hydrogenase. In the absence or upon severe deficiency of macronutrients, C. reinhardtii cells stop growth and carbon dioxide (CO2) fixation (Grossman 2000; Melis et al. 2000; Zhang et al. 2002; Hemschemeier et al. 2008a). Ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) protein is degraded rapidly in S-depleted Chlamydomonas cultures (Zhang et al. 2002), and H2 production has been proposed to be an alternative electron sink for the cells, which allows them to maintain photosynthetic electron transport in order to prevent lightinduced oxidative stress and to produce chemical energy in form of ATP (Melis et al. 2000; Happe et al. 2002; Melis 2007; Hemschemeier et al. 2008a). To date, S deficiency is the only condition which has been reported to induce a prolonged and significant H2 evolution in illuminated wild-type C. reinhardtii, though alternative protocols using genetically altered strains have been developed in order to circumvent this severe stress condition. One of these approaches allowed the diminishment of PSII amounts by controllable production of Nac2, a factor required for PsbD (D2) accumulation (Surzycki et al. 2007). Another approach made use of antisense constructs to down-regulate the expression of the SULP gene encoding a chloroplast-envelope sulfate transporter

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and achieved H2 photoproduction in the presence of 100 lM sulfate (Chen et al. 2005). In this study, we show that Chlamydomonas also evolved considerable amounts of H2 when incubated in sealed flasks in the absence of a nitrogen source. However, hydrogenase activity, H2 evolution and accompanying fermentative pathways were established about 48 h later than in S-deprived algal suspensions, and neither H2 nor formate and ethanol were produced in amounts as observed in S-deficient cultures. Though the starch content of Chlamydomonas cells was about twofold higher under N- than under S deprivation, indicating a higher potential for H2 generation, starch was degraded much later in N-deficient cultures and stayed on a high level. Analysis of the photosynthetic apparatus of N-starved C. reinhardtii cultures indicated that the delay in the establishment of a H2 metabolism was due to prolonged PSII activity and that the N starvation induced degradation of the cytochrome b6f complex might be a reason for the inability of the cells to efficiently channel electrons from carbohydrate oxidation to the hydrogenase.

Materials and methods Strains and growth conditions Chlamydomonas reinhardtii wild-type strain CC-124 (137c mt- nit1- nit2-) was obtained from the Chlamydomonas culture collection at Duke University, Durham, NC (http:// www.chlamy.org/strains.html) and grown photoheterotrophically in Tris acetate phosphate (TAP) medium (Harris 1989) on a horizontal shaker with bottom-up illumination of an intensity of 100 lmol photons m-2 s-1 at 20°C. To induce nitrogen and sulfur deprivation, cells were grown until they had reached a chlorophyll (Chl) content of about 15 lg Chl ml-1, harvested by centrifugation (1,250g for 3 min at 20°C) and washed once with S- or N-free medium. Sulfur- and nitrogen-free media were prepared using the standard TAP-medium recipe and replacing sulfate by the respective chloride salts (TAP-S) and NH4Cl by KCl (TAP-N). After a second centrifugation step, cells were resuspended in 280 ml of TAP-S or TAP-N medium to obtain a cell density of about 5 9 106 cells ml-1 and transferred to square glass bottles with a total volume of 325 ml. Flasks were sealed with gas-tight rubber stoppers (Suba seals 37, Sigma-Aldrich, http://www.sigmaaldrich. com) and then incubated under constant one-side illumination of 60 lmol photons m-2 s-1 at 23°C on a magnetic stirrer. When indicated, 28 ll of DCMU dissolved in ethanol was added to the cultures to reach a final concentration of 10 lM. The same volume of ethanol was added to control flasks. Analysis of dark fermentation capacity was done by incubating Chlamydomonas cells in TAP-S and

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TAP-N in the light as described above until they had reached the beginning of the H2 photoproduction phase as judged from small amounts (0.02 lmol H2 ml-1 gas) of H2 in the gas phase. Then, the flasks were flushed with argon for 5 min to reset both O2 and H2 concentrations in the cultures, before the bottles were wrapped in aluminum foil and incubated for further 24 h in the dark. In all cases, samples of the cell suspensions were taken by piercing the seals with gas-tight syringes. Determination of cell number, pigment concentration and starch content Cell numbers were determined by adding 10 ll of 0.25% (w/v) iodine in ethanol to 1 ml of cell suspension and then counting the cells using a hemocytometer. Chlorophyll and carotenoid concentrations were analyzed using 1 ml cell suspension as described before (Wellburn 1994). Cell pellets obtained after acetone extraction were further processed for quantification of starch by staining with iodine solution (Southgate 1976). Cell pellets were resuspended in 300 ll double-distilled water and boiled for 20 min to solubilize starch. After centrifugation at high speed for 1 min, 100 ll of the supernatant were mixed with 800 ll water and 100 ll of potassium iodine solution containing 0.75% (w/v) KI and 0.15% (w/v) I2. Absorbance was measured spectrophotometrically at k = 580 nm. Analyses of hydrogenase activity, H2, CO2 and O2 content of the gas phase as well as fermentation products and acetate in the medium In vitro hydrogenase activity of cell suspension aliquots and the hydrogen and oxygen concentrations in the headspace above S- or N-deprived C. reinhardtii cultures were determined as reported previously (Hemschemeier et al. 2009; Philipps et al. 2011). Briefly, to determine total in vitro hydrogenase activity of the cells, 500 ll of S- or N-deprived C. reinhardtii cells were withdrawn from the incubation flasks with a syringe and injected into 1.5 ml of an Argon-saturated reaction mixture containing 1% Triton X-100, 100 mM sodium dithionite and 10 mM methyl viologen in 50 mM potassium phosphate buffer, pH 6.8. The mixture was incubated in a shaking water bath at 37°C for 20 min before injecting 200 ll of the headspace into a gas-chromatograph (Shimadzu GC-2010, http://www. shimadzu.eu), equipped with a PLOT fused silica-coated ˚ molecular sieve column (10 m 9 0.32 mm, pore size 5 A from Varian, http://www.varian.de) to quantify H2 content. H2 and O2 in the headspace of the culture flasks were quantified by injecting 200 ll of the gas phase above the cell suspensions into the same gas-chromatograph. Concentrations of formate, ethanol and acetate in the medium

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were determined using assays from r-Biopharm (http:// www.r-biopharm.com, formate: Cat.-No. 10 979 732 035, ethanol: Cat.-No. 10 176 290 035, acetate: Cat.-No.: 10 148 261 035), and CO2 concentration in the gaseous phase was analyzed by mass-spectrometry as described in Philipps et al. (2011). Characterization of photosynthetic parameters Photosynthetic O2 evolution and respiratory O2 consumption rates were determined using a Clark-type oxygen electrode (model respire 1 from Hansatech, http://www. hansatech-instruments.com). One-ml aliquots of N- or S-depleted Chlamydomonas cultures were withdrawn from the incubation flasks and transferred to the measuring chamber. Cells were illuminated using cool-white fluorescence bulbs at a light intensity of 60 lmol photons m-2 s-1 at 23°C for 5 min before recording O2 consumption rates in shaded cultures and O2 production rates upon illumination for 5 min each. Gross rates of photosynthetic O2 evolution were calculated by adding the absolute value of respiratory O2 consumption measured in the dark to O2 evolution rates detected in illuminated cells. Pulse amplitude modulated (PAM) chlorophyll fluorescence measurements were conducted using a DUAL-PAM100 measuring device (Walz, Germany, http://www.walz. com). The terminology used throughout the article is according to Maxwell and Johnson (2000). F0 was determined in aliquots of N- or S-starved C. reinhardtii cultures incubated in the measuring cuvette in the dark for 5 min. The intensity of the measuring beam was 0.2 lmol of photons m-2 s-1. Fm was determined by application of a 300 ms saturating pulse of 10,000 lmol of photons m-2 s-1. Ft was determined after the cells had been illuminated with an actinic light source of 95 lmol of photons m-2 s-1 for 230 s, during which saturating pulses were applied every 20 s to analyze Fm0 values. Fv/Fm as an indicator of the maximum quantum efficiency of PSII was calculated using F0 and Fm values according to the formula (Fm-F0)/Fm. For calculating the efficiency of PSII photochemistry in illuminated cells (UPSII = (Fm0 -Ft)/Fm0 ) (Genty et al. 1989), Fm0 and Ft values observed after 210 s of applying actinic light were used (Maxwell and Johnson 2000). When indicated, methyl viologen as artificial electron acceptor of PSI (Dodge 1971; Hiyama and Ke 1971) was added to the same aliquots of S- or N-deprived C. reinhardtii cultures after the recording of fluorescence parameters as described above. After the actinic light had been switched off, a methyl viologen solution was added to the cells to reach a final concentration of 1 mM, and the cells were incubated for 5 min in the dark until the same measurements as outlined above were repeated to obtain F0, Fm, Ft and Fm0 in methyl viologen-treated cells.

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The recording of fluorescence emission spectra at 77 K was conducted using a single-beam luminescence spectrometer (Aminco Bowman Series 2, Spectronic Instruments, Rochester, NY, USA) connected to a computer running the software for AB2 luminescence spectrometer (v4.2). 0.2 ml of N- or S-deprived C. reinhardtii cells were removed from the sealed flasks with a syringe, diluted 1:4 with the respective medium, transferred to the measuring cuvette and directly frozen in liquid nitrogen. Frozen samples were excited using light of k = 435 nm, and a fluorescence emission spectrum from k = 650 to 750 nm was recorded. Values were corrected using the correction option provided by the instruments software and the ratio of fluorescence emission at k = 685 nm (PSII) to fluorescence emission at k = 715 nm (PSI) was calculated. Immunoblot analyses Preparation of crude protein extracts was conducted by removing 1 ml of cell suspension from the incubation flasks, harvesting the cells by centrifugation (1 min at 14,100g at room temperature) and resuspending the cell pellet in 59 protein lysis buffer (0.25 M TrisHCl, pH 8.0; 25% glycerol; 7.5% SDS; 0.25 mg ml-1 bromophenol blue; 12.5% (v/v) 2-mercaptoethanol). After heating the samples at 90°C for 5 min they were stored at -20°C. Prior to loading them on gels, samples were again incubated for 5 min at 90°C and cell debris was pelleted by centrifugation (1 min at 14,100g at room temperature). The supernatant was separated by SDS-PAGE (sodium dodecyl sulfate polyacrylamide gel electrophoresis) (Laemmli and Favre 1973) or Tris-Tricine gels (Schagger 2006) using 5% collecting and 10 or 16% separating gels. Blotting, washing and antibody hybridization steps were conducted as described before (Hemschemeier et al. 2008b; Philipps et al. 2011). Polyclonal antibodies used were anti-C. reinhardtii-HYD1 (Happe et al. 1994) (1:5,000), anti-C. reinhardtii-PsaD (Fischer et al. 1997) (1:5,000), anti-Arabidopsis-PsbA (Park and Rodermel 2004) (1:5,000), anti-C. reinhardtii-cytochrome f (Cyt f) (Bulte and Wollman 1992) (1:20,000) and anti-spinach-ferredoxin (Bohme 1977) (1:2,000). Statistical analysis of results All physiological data shown are means of at least three independent experiments. Error bars indicate the standard deviation. Tables below Figs. 6 and 7 show the percent changes of the mean values. Immunoblot analyses were conducted using protein samples from three independent experiments. In each case, one representative immunoblot is shown in Figs. 1, 4 and 7.

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SDS PAGE Fig. 1 Hydrogen accumulation (a), in vitro hydrogenase activity (b) and HYD1 immunoblot analyses (c) of nitrogen and sulfurdepleted, sealed C. reinhardtii cultures. a H2 concentration in the headspace of TAP-S (black circles) and TAP-N (gray circles) cultures was quantified by gas-chromatography. b In vitro hydrogenase activity was determined using the artificial [FeFe]-hydrogenase electron donor methyl viologen in lysed cell samples withdrawn from the incubation flasks at the depicted time-points. a and b Mean values of a biological triplicate are shown, error bars indicate standard deviations. c Accumulation of hydrogenase protein in raw extracts of N- and S-starved algal cells was analyzed by immunoblotting using a polyclonal HYD1 antibody. An amount corresponding to 0.5 9 106 cells was loaded on each lane. A representative immunoblot of one experiment is shown. Coomassie stained SDS gels are shown as loading control

When photoheterotrophically grown C. reinhardtii cells were subjected to N- or S-deprivation and incubated in sealed flasks under moderate light intensities (60 lmol photons m-2 s-1), cultures from both nutritional conditions evolved H2 gas (Fig. 1a). Under the conditions applied, H2 was detectable in the gas phase of S-depleted cells after 32 h (Fig. 1a). The onset of H2 evolution was paralleled by the development of in vitro hydrogenase activity assayed in cell samples withdrawn from the incubation flasks at the indicated time points (Fig. 1b). A very low hydrogenase enzyme activity (0.004 lmol H2 9 106 cells-1 h-1) was measured in S-deprived cells after 1 day, and it increased rapidly in the following 24 h (Fig. 1b). In contrast, detectable accumulation of H2 gas (Fig. 1a) and the establishment of in vitro hydrogenase activity (Fig. 1b) did not occur earlier than 72 h after the transfer of C. reinhardtii cells to nitrogen-free medium. N-deficient algal cultures did furthermore not accumulate H2 in the same amounts as did S-deprived cell suspensions (Fig. 1a). While the H2 concentration in the headspace of the latter reached 35 lmol H2 ml gas-1 on average after 96 h of medium exchange, the former produced only 20.4 lmol H2 ml gas-1 and reached this level not until 192 h after the transfer to TAP-N medium (Fig. 1a). In vitro hydrogenase activity of N-starved Chlamydomonas cells reached about 3.4 lmol H2 9 106 cells-1 h-1 and therewith almost the same level as detected in S-deficient cultures (3.95 lmol H2 9 106 cells-1 h-1) (Fig. 1b). In nitrogen-deficient cells, however, hydrogenase enzyme activity reached its maximum only 192 h after nitrogen had been excluded from the medium, while the maximum in vitro hydrogenase activity of S-starved cultures was measured after 120 h (Fig. 1b). Notably, while in vitro hydrogenase activity paralleled the accumulation and stagnation of H2 gas in the headspace of S-deprived algal cultures, hydrogenase enzyme activity increased further in N-starved cells after 144 h when the amount of H2 in the gas phase was already close to the maximum (Fig. 1a, b). In both S- and N-deprived C. reinhardtii cultures, in vitro hydrogenase activity was paralleled by the accumulation of HYD1 protein as detected by immunoblotting (Fig. 1c). Sulfur-deprived H2-evolving C. reinhardtii cultures have been reported to excrete formate and ethanol and to accumulate CO2 (Tsygankov et al. 2002; Kosourov et al. 2003; Hemschemeier et al. 2008a, b). In the experimental setup applied in this study, Chlamydomonas cultures

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Nitrogen-deprived cells exhibit prolonged PSII activity and a delay in O2 consumption Both N- and S-depleted C. reinhardtii cultures gradually consumed O2 from the gas phase. However, while a continuous decrease of O2 concentrations in the headspace of S-deficient cell suspensions was observable from the beginning of the experiment, N-starved cells showed a delay in O2 consumption, and even a transient accumulation of O2 could be detected. The O2 concentration in the headspace of S-depleted Chlamydomonas cultures decreased by 50% within 24 h, and after 72 h no detectable amounts of O2 were left. In the gas phase of N-depleted cells, the O2 concentration reached about 5 lmol O2 ml gas-1 after 48–72 h. In S-deprived cells, the same O2 content of the gas phase was sufficiently low to allow the onset of H2-production (compare Figs. 1a and 3a). In contrast to cultures incubated in TAP-S medium, N-starved algae did not consume the O2 present in the headspace completely. After 144 h, 2 lmol O2 ml gas phase-1 were left, and this concentration stayed constant until the end of the experiment. The delay in the net consumption of O2 observed in N-depleted relative to S-deficient C. reinhardtii cell suspensions was paralleled by a moderately slower decrease in photosynthetic O2 evolution rates and a higher fraction of

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started to accumulate detectable amounts of formate and ethanol in the medium as well as CO2 in the gas phase after 1 day of being incubated in S-free medium (Fig. 2a–c) and thus simultaneously to H2 evolution (Fig. 1a). Nitrogendeprived C. reinhardtii cells did also produce formate, ethanol and CO2. Formate and ethanol were detectable in supernatants of N-starved algae after 72 h (Fig. 2a, b). Formate and ethanol concentrations reached 0.25 lmol formate ml-1 and 0.06 lmol ethanol ml-1 after 192 h of nutrient deprivation, and thus only 17 and 4.4%, respectively, of the levels detected in S-deficient cultures (Fig. 2a, b). CO2 accumulated in the gas phase of N-deprived algal suspensions up to 34 nmol ml gas-1, which was roughly one-fourth of the concentration detected in the headspace of sulfur-depleted cultures (Fig. 2c). Amounts of acetate in the medium showed the same trend as has been reported before in case of S- (Melis et al. 2000) and N deficiency (Work et al. 2010), though in the latter study, cells were not sealed. In both S- and N-starved cultures analyzed here, the acetate content of the medium decreased to roughly half of the original amount. 0.59 ± 0.06 g acetate l-1 remained in the supernatant of S-deficient cells after 48 h of incubation, and 0.45 ± 0.03 g acetate l-1 was present in the medium of N-starved C. reinhardtii cultures after 72 h. In both cases, the acetate concentration stayed constant afterwards (data not shown).

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time [h] Fig. 2 Concentrations of formate (a) and ethanol (b) in the medium and of carbon dioxide (c) in the headspace of N- and S-starved algal suspensions. a and b Formate and ethanol were determined in the supernatant of TAP-S (black circles) and TAP-N (gray circles) algal cultures at the depicted time-points. Mean values of three independent experiments are shown. c CO2 was quantified in the headspace of the nutrient-deficient cell cultures by mass spectrometry from biological triplicates. Error bars indicate standard deviations

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intact and open PSII centers as observed in PAM chlorophyll fluorescence measurements (Figs. 3b, c, 4a). Photosynthetic O2 evolution and respiratory O2 uptake rates were examined using a Clark-type O2-electrode and showed the well-reported changes in S-deprived Chlamydomonas cultures (Wykoff et al. 1998; Melis et al. 2000). Gross O2 production rates in illuminated cell samples decreased by 52% within 24 h and declined further to 19% of the original rate during the next day (Fig. 3b). Meanwhile, O2 consumption rates in the dark decreased by 39% during the first day of sulfur depletion and stayed roughly on this level until the end of the experiment (Fig. 3b). In N-deprived cells, photosynthetic O2 evolution rates did decrease, too, but this decline occurred slower than in S-depleted cells (compare Fig. 3b with c). After 24 h of incubation in TAPN medium, gross O2 production rates of the algae had decreased by 44% from 262 to 146 nmol O2 9 106 cells-1 h-1, and after 48 h, 30% of the original O2 evolution rates were still measurable (Fig. 3c). Respiratory O2 uptake rates of N-starved cells as examined in cell samples in the dark declined from 120 to 82 nmol O2 9 106 cells-1 h-1 during the first 24 h and remained on this level (Fig. 3c). The capacity for higher photosynthetic O2 evolution in nitrogen- versus sulfur-starved C. reinhardtii cells was reflected by a higher proportion of intact and photochemically active PSII complexes as determined by PAM chlorophyll fluorescence analyses (Fig. 4a). The maximum quantum efficiency of PSII (Fv/Fm) (Maxwell and Johnson 2000) measured in dark-adapted cells decreased moderately from 0.59 to 0.52 within 24 h in S-deprived algal cultures, and then dropped rapidly to 0.25 during the following day (Fig. 4a). The efficiency of PSII photochemistry in illuminated cells, which can be referred from the value UPSII determined in light-adapted culture aliquots (Maxwell and Johnson 2000), decreased more rapidly and more strongly than Fv/Fm in Chlamydomonas upon S deficiency. UPSII was at a value of 0.16 after 24 h and at 0.11 after 48 h of treatment and further decreased thereafter (Fig. 4a). After 120 h, a value of 0.05 was determined, and UPSII stayed on this level until the end of the experiment (192 h) (Fig. 4a). The gradual loss of PSII activity was paralleled by decreasing amounts of PsbA (D1) protein (Fig. 4b). The decline of both values, Fv/Fm and UPSII, as well as the loss of PsbA protein has been observed in C. reinhardtii upon S deprivation before (Wykoff et al. 1998; Melis et al. 2000; Zhang et al. 2002; Antal et al. 2003). In N-deprived cells, both Fv/Fm and UPSII decreased slower (Fig. 4a). The maximum quantum efficiency of PSII centers (Fv/Fm) declined gradually from 0.59 to 0.46 within the first 2 days after transfer to N-free medium and reached values below 0.2 only after 120 h of incubation (Fig. 4a). The slower loss of active PSII centers observed upon

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time [h] Fig. 3 Oxygen concentration in the headspace of N- or S-deprived C. reinhardtii cultures (a) and gross photosynthetic O2 evolution as well as respiratory O2 uptake rates in S- (b) or N-deficient (c) cell suspensions. a O2 in the gas phase of TAP-S (black circles) and TAP-N (gray circles) cultures were quantified by injecting gas samples into a gas-chromatograph (n = 3). b and c O2 evolution (black circles, gray circles) and uptake rates (black triangles, gray triangles) were determined in the light and in darkness, respectively. Cell samples were transferred from the incubation flasks into the chamber of a Clark-type O2 electrode. Gross photosynthetic O2 production rates were calculated by adding the absolute values of O2 uptake in the dark to O2 evolution in the light. Values shown are means of four independent analyses. Error bars indicate standard deviations

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24 48 72 96 120 144 168 192

24 48 72 96 120 144 168 192

time [h]

time [h]

b

0

24 48 72 96 120 144 168 192 h

PsbA

0

24 48 72 96 120 144 168 192 h -N

-S

SDS PAGE

Fig. 4 PSII quantum yields (a) and PsbA (D1) amounts as detected by immunoblotting (b) in S- and N-depleted Chlamydomonas cultures. a Maximum quantum yield of PSII (Fv/Fm = (Fm-F0)/ Fm) determined in dark-adapted TAP-S (black squares) and TAP-N (gray squares) culture aliquots and efficiency of PSII photochemistry in illuminated cells (UPSII = (Fm0 -Ft)/Fm0 ) (TAP-S black circles, TAP-N gray circles). F0 was determined in aliquots of C. reinhardtii cultures incubated in the measuring cuvette in the dark for 5 min; Fm

was achieved by applying a saturating pulse in the dark. Ft was read after 230 s of illumination with actinic light; Fm0 values were determined using the saturating pulse at 210 s after the actinic light had been switched on. Depicted values and standard deviations are calculated from biological triplicates. b Immunoblot analyses were performed on crude protein extracts using PsbA antibody. Coomassie stained SDS gels are shown as loading control

nitrogen deprivation was not paralleled by an equally stepwise loss of PsbA (D1) protein detected by immunoblot analyses (Fig. 4b). Rather, the decrease in D1-protein levels was similar in N- versus S-deprived C. reinhardtii cells (Fig. 4b). PSII photochemical efficiency as indicated by UPSII did also decrease more slowly in N-starved C. reinhardtii cultures and reached a value of 0.11 only after 72 h of incubation under N-limited conditions, while this value was determined in S-deprived cells after 48 h already (Fig. 4a). While PSII activity as referred from gross photosynthetic O2 evolution rates as well as from Fv/Fm and UPSII decreased slower in N- than in S-depleted cells, it dropped to hardly detectable values after 120 h of N starvation (Figs. 3c, 4a). In S-deficient cells, in contrast, PSII activity remained low but quite stable until the end of the experiment (192 h after transfer to TAP-S medium) as referred from all applied analyses (Figs. 3b, 4a). It has been reported before that sulfur-depleted C. reinhardtii cells exhibit transfer of their light-harvesting complexes II (LHCII) to PSI (Wykoff et al. 1998) during a process called state transitions (reviewed by Lemeille and Rochaix 2010). The extent to which the mobile LHCII antennae are associated with either PSII or PSI can be estimated by determining the fluorescence yield at 685 nm (PSII) and 715 nm (PSI) at low temperatures (77 K), and a decrease of the 685 nm:715 nm ratio reflects transition to state II and vice versa. Upon the experimental conditions applied in this study, the 685 nm:715 nm ratio of

fluorescence measured at 77 K in Chlamydomonas culture aliquots withdrawn from S- or N-deprived cell suspensions showed an initial increase from 1.1 to 1.5 in the first 24 h of treatment. In the following 2 days, it decreased steadily

123

2.0

ratio E 685 : E 715

1.5

1.0

0.5

0.0 0

6

24

48

72

96

120

168

216

time [h] Fig. 5 Analysis of state transitions by 77 K fluorescence spectroscopy. S- (black bars) or N-depleted (gray bars) Chlamydomonas culture aliquots were withdrawn from the incubation flasks at the depicted time-points, quickly diluted 1:4 in TAP-S and TAP-N medium, respectively, and immediately frozen in liquid nitrogen. Fluorescence emission spectra were recorded from k = 650 to 750 nm at 77 K and the ratio of fluorescence emission at k = 686 nm (PSII) to fluorescence emission at k = 715 nm (PSI) was calculated from biological triplicates

Planta (2012) 235:729–745

737

Table 1 Cell numbers and pigment concentrations of sulfur- and nitrogen-deprived C. reinhardtii cultures during the time-course of nutrient starvation Time (h)

Cell count (106 cells ml-1)

Chlorophyll content (lg 9 106 cells-1)

Chlorophyll a ? b (lg ml-1)

Carotenoids (lg ml-1)

-S

-S

-S

-S

-N

-N

-N

-N

0

5.1 ± 0.4

4.6 ± 0.4

1.4 ± 0.2

1.6 ± 0.1

7.1 ± 0.4

7.3 ± 0.3

1.1 ± 0.2

1.3 ± 0.3

24

5.8 ± 0.5

5.8 ± 0.5

2.4 ± 0.2

1.7 ± 0.2

13.6 ± 1.5

10.0 ± 0.3

2.2 ± 0.7

2.1 ± 0.4

48

6.3 ± 0.4

6.0 ± 0.5

2.3 ± 0.2

1.5 ± 0.2

14.2 ± 1.6

9.1 ± 0.9

2.5 ± 0.9

2.2 ± 0.8

72

6.0 ± 0.6

5.8 ± 0.4

2.3 ± 0.5

1.3 ± 0.2

13.5 ± 1.7

7.3 ± 0.8

2.5 ± 0.7

2.2 ± 0.5

96

6.1 ± 0.6

5.5 ± 0.4

2.3 ± 0.6

1.4 ± 0.4

13.8 ± 3.0

7.5 ± 1.7

2.6 ± 1.1

2.4 ± 1.0

120

5.9 ± 0.3

5.3 ± 0.6

2.2 ± 0.4

1.3 ± 0.2

12.8 ± 2.1

7.1 ± 1.4

2.6 ± 0.8

2.2 ± 0.8

144 168

6.2 ± 0.6 5.7 ± 0.5

5.5 ± 0.6 5.4 ± 0.5

2.3 ± 0.5 2.4 ± 0.4

1.4 ± 0.2 1.4 ± 0.2

14.2 ± 1.8 13.7 ± 1.5

7.4 ± 0.8 7.3 ± 0.8

2.9 ± 0.7 2.8 ± 0.6

2.4 ± 0.4 2.6 ± 0.4

192

5.6 ± 0.7

5.3 ± 0.7

2.4 ± 0.5

1.3 ± 0.2

13.2 ± 1.4

7.0 ± 0.5

2.8 ± 0.4

2.5 ± 0.2

Time (h)

-1

-1

Chlorophyll a (lg ml )

Chlorophyll b (lg ml )

Chlorophyll a:b ratio

Carotenoids:chlorophyll ratio

-S

-S

-S

-S

-N

-N

-N

-N

0

4.9 ± 0.4

5.0 ± 0.5

2.2 ± 0.5

2.3 ± 0.3

2.3 ± 0.7

2.3 ± 0.5

0.16 ± 0.03

0.18 ± 0.03

24

9.0 ± 1.8

6.7 ± 0.6

4.7 ± 0.4

3.3 ± 0.3

1.9 ± 0.5

2.1 ± 0.4

0.16 ± 0.03

0.21 ± 0.04

48

9.3 ± 1.9

5.8 ± 0.9

4.9 ± 0.4

3.3 ± 0.2

1.9 ± 0.5

1.7 ± 0.3

0.17 ± 0.04

0.24 ± 0.06 0.31 ± 0.05

72

9.3 ± 2.0

5.0 ± 0.8

4.1 ± 0.5

2.3 ± 0.2

2.3 ± 0.7

2.3 ± 0.4

0.19 ± 0.03

96

9.7 ± 2.8

5.6 ± 1.6

4.1 ± 0.3

2.0 ± 0.2

2.4 ± 0.6

2.8 ± 0.7

0.18 ± 0.03

0.32 ± 0.06

120

9.0 ± 2.0

5.4 ± 1.4

3.8 ± 0.1

1.7 ± 0.1

2.3 ± 0.5

3.2 ± 0.9

0.20 ± 0.03

0.31 ± 0.06

144

10.2 ± 1.9

5.8 ± 0.9

3.9 ± 0.3

1.6 ± 0.3

2.6 ± 0.6

3.9 ± 1.2

0.20 ± 0.03

0.32 ± 0.04

168

9.7 ± 1.3

6.0 ± 0.9

4.0 ± 0.3

1.4 ± 0.2

2.5 ± 0.2

4.4 ± 1.2

0.21 ± 0.02

0.36 ± 0.03

192

9.7 ± 1.6

5.7 ± 0.6

3.5 ± 0.3

1.2 ± 0.2

2.9 ± 0.7

4.8 ± 1.2

0.21 ± 0.01

0.37 ± 0.05

Determination of Chl a, Chl b and total carotenoids was performed in pellets from 1 ml of algal suspension extracted by 1 ml of 80% acetone according to Wellburn (1994). Cells were counted using a hemocytometer. Mean values ± standard deviation were obtained from six independent experiments

to values around 0.6 and stayed rather constant until the end of the experiment (Fig. 5). While both S- and N-deficient cells showed the same trend of the development of the 685 nm:715 nm ratio, it remained slightly higher in nitrogen-starved cells after the maximum at 24 h (Fig. 5). Different changes of pigment composition of S- and N-starved algal cultures were observed during the timecourse of nutrient starvation. Cellular chlorophyll content increased from 1.4 to 2.4 lg Chl 9 106 cells-1 within the first 24 h of sulfur starvation and stayed on this level (Table 1). In N-deficient Chlamydomonas cultures, after an initial increase from 1.6 to 1.7 lg Chl 9 106 cells-1, the cellular chlorophyll content decreased to 1.3 lg Chl 9 106 cells-1 and stayed roughly on this level until the end of the experiment. N-deprived algae showed an increase of the Chl a:Chl b ratio from 2.3 (0 h) to 4.8 (192 h), which was mostly due to decreased Chl b amounts (Table 1). The carotenoid to chlorophyll ratio of N-deficient cells increased from 0.18 to 0.37 (Table 1). Both ratios increased only slightly in S-deficient algal cultures (Table 1).

Nitrogen-starved algae accumulate more starch, but do hardly degrade it Both sulfur- and nitrogen-deficient algal cultures accumulated substantial amounts of starch (Fig. 6a). In S-deprived cells, starch content increased from 4 lg ml cell suspension-1 to 101 lg ml-1 within 48 h (Fig. 6a). Afterwards, a constant decrease of starch levels down to 25% of the maximum was observed (Fig. 6a, b), a phenomenon described before to appear in S-deprived, H2 evolving C. reinhardtii cells (Melis et al. 2000; Tsygankov et al. 2002; Zhang et al. 2002; Fouchard et al. 2005). In N-starved algal cells, starch accumulated constantly during the first 96 h and reached 188 lg ml cell suspension-1, which was roughly twice as much as determined in sulfur-starved cells (Fig. 6a). Afterwards, however, starch levels decreased only moderately in N-deprived Chlamydomonas cultures (Fig. 6a, b). While in S-deficient cells, the concentration of starch decreased by 65% during 4 days after the maximum starch level had been detected, starch content decreased by only 13% during the first 4 days after the highest starch

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(Table 2). In sulfur-starved Chlamydomonas cells, the starch content decreased from 76.2 to 4.6 lg starch ml cell suspension-1 (-71.6 lg), and in N-deficient cells, it decreased by 55.3 lg from 119.9 to 64.4 lg starch ml cell suspension-1 (Table 2). In supernatants withdrawn 24 h after the transfer to darkness from S- and N-deprived cultures, similar concentrations of ethanol were detected (1.4 lmol ethanol ml-1 in case of S- and 1.0 lmol ethanol ml-1 in case of N-starved cells). The supernatant of cells from both nutritional conditions showed an increase of acetate levels by 1.1 lmol acetate ml-1 (Table 2). In contrast, about four times more formate accumulated in the medium of S-deficient cells compared with N-free cultures (1.1 lmol formate ml-1 versus 0.3 lmol formate ml-1) (Table 2). S-deprived C. reinhardtii cells transferred to darkness accumulated 0.05 lmol H2 ml-1 headspace (Table 2), while nitrogen-starved and shaded cells accumulated roughly the 30-fold amount of H2 (Table 2).

a 250

µg starch · ml -1

200 150 100 50 0 0

24

48

72

96

120

144

168

192

time [h]

b h

0

24

48

72

96

120

144

168

-S

4

89

100

81

60

45

35

32

192 25

-N

2

45

82

96

100

97

91

90

87

Fig. 6 Starch content (a) and percent values of starch levels relative to the highest value measured (set to 100%) (b). a Starch was quantified in pellets of 1 ml C. reinhardtii cell suspension aliquots by staining with iodine solution. Results from S-deficient cultures are shown in black, and gray bars indicate results from N-deprived cells (n = 3). b The percentage of the amount of starch relative to the highest value measured (set to 100%, shaded in gray) is shown for each time-point

content was measured in nitrogen-depleted algal cells (Fig. 6a, b). After 192 h of incubation in either medium, 25 lg starch ml-1 was left in sulfur-, and 164 lg starch ml-1 in nitrogen-deprived cells (Fig. 6). In order to determine whether N starvation resulted in a general impairment of starch breakdown capacity, the latter was analyzed in the dark after the cells had built up their starch reserves in the light. The culture flasks were transferred to darkness and the gaseous composition of the headspace as well as secreted fermentative products and starch content of the cells was analyzed after 24 h

A positive effect of methyl viologen on PSII photochemistry is more pronounced in S- than in N-deprived cells It has been concluded from various studies that the oxidative degradation of starch is a major electron source for H2 production besides residual PSII water oxidation activity in sulfur-depleted C. reinhardtii wild-type cells (Melis et al. 2000; Fouchard et al. 2005). This seemed not to be the case in N-starved algal cultures, as the high starch levels were hardly degraded under the conditions applied in this study (see above, Fig. 6a, b). Notably, no significant change of H2 yields was observed when 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU), a specific PSII inhibitor, was added to N-deficient cells after about 72 h of incubation in N-free medium, when H2 was just detectable in the headspace (data not shown). In order to assess the capacity of the photosynthetic electron transport chain

Table 2 Starch content as well as fermentative products in the supernatant of C. reinhardtii cultures which had been incubated in S- or N-free medium in the light for 32 h (-S) and 68 h (-N) and then flushed with argon and transferred to darkness Starch content (lg ml-1)

Ethanol (nmol ml-1)

Formate (nmol ml-1)

Acetate (nmol ml-1)

H2 (lmol ml-1)

-S 0h

76.2 ± 0.7

24 h

4.6 ± 4.0

BD

BD

1.44 ± 0.17

1.07 ± 0.07

9.19 ± 1.39 10.34 ± 0.84

0.05 ± 0.02

-N 0h

119.9 ± 2.2

24 h

64.5 ± 2.0

BD

BD

7.71 ± 0.76

1.00 ± 0.06

0.27 ± 0.20

8.79 ± 0.27

1.50 ± 0.50

Cell samples were removed from the flasks immediately after Argon flushing (0 h) and after 24 h of dark incubation. Cells were harvested by centrifugation. Starch was quantified in the cell pellets, fermentative products in the supernatants. H2 was determined in the headspace of the cultures at the same time points. Mean values ± standard deviation were obtained from three independent experiments BD Below detection

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a

739

0.7

0.7 -S

-N

0.6

quantum yield

quantum yield

0.6 0.5 0.4 0.3 0.2 0.1 0.0

0.5 0.4 0.3 0.2 0.1 0.0

0

24

48

0

72 96 120 144 168 192

24

48

b

72 96 120 144 168 192

time [h]

time [h] 0

24

48

72

96

120

144

168

192

h

0

24

48

72

96

120

144

168

192

-S 93

172

226

234

221

269

264

268

288

-N 93

114

144

128

173

167

145

160

149

h

c

0

PsaD

24 48 72 96 120 144 168 192 h -S

0

24 48 72 96 120 144 168 192 h -N

Cyt f PetF SDS PAGE

Fig. 7 Efficiency of PSII photochemistry (UPSII = (Fm0 -Ft)/Fm0 ) in illuminated N- or S-depleted algal cells in the absence or presence of 1 mM methyl viologen (a, b) and immunoblot analyses of PsaD, cytochrome f and ferredoxin (c). a Chlorophyll fluorescence was first determined in the absence of methyl viologen in TAP-S (black circles) and TAP-N (gray circles). Then, cells were treated with 1 mM methyl viologen and kept in the dark for 5 min, before the same sequence of light-conditions were applied (TAP-S black

triangles, TAP-N gray triangles). In a, the results obtained from methyl viologen-treated cells as well as the original values already shown in Fig. 4a are depicted. The values represent the average of biological triplicates. In b, the percental increase of relative fluorescence after addition of methyl viologen relative to the value of untreated cells is shown. c Immunoblotting using polyclonal PsaD and cytochrome f as well as anti-spinach ferredoxin antibodies. Coomassie stained SDS gels are shown as loading control

downstream of PSII, PAM chlorophyll fluorescence measurements as described above were repeated using the same culture aliquots, but treated with methyl viologen, which is an efficient electron acceptor of PSI (Dodge 1971; Hiyama and Ke 1971) (Fig. 7a, b). It could be observed that the addition of methyl viologen to S-depleted Chlamydomonas cells resulted in a 172 to 288% (243% on average) enhancement of the efficiency of PSII photochemistry (UPSII) in the hypoxic phase (24 to 192 h after transfer to TAP-S medium). Instead, UPSII of methyl viologen-treated N-starved cells was increased by 128% to maximally 173% (154% on average) during the H2-producing phase (72 to 192 h of incubation in TAP-N) (Fig. 7a, b). These data implicated that a bottleneck downstream of PSII limited linear photosynthetic electron flow in N-deprived cells. It has been described before that the cytochrome b6f complex is selectively degraded upon nitrogen deprivation (Bulte and Wollman 1992). Immunoblot analyses using PsaD and cytochrome f specific as well as antispinach-ferredoxin antibodies showed that PsaD, cytochrome f and ferredoxin amounts stayed rather constant in S-deprived C. reinhardtii cultures over a time period of

192 h (Fig. 7c). In N-starved cells, PsaD levels remained constant, too, but both cytochrome f and ferredoxin levels decreased after 48 and 96 h, respectively, of incubation in TAP-N medium and stayed on this lowered level (Fig. 7c).

Discussion Nitrogen deprivation is a severe stress condition for all organisms, as nitrogen is a major constituent of proteins and nucleic acids and a highly abundant atomic species in living cells. C. reinhardtii has been shown to respond to N deprivation by both altering central metabolic processes (Martin and Goodenough 1975; Plumley and Schmidt 1989; Peltier and Schmidt 1991; Bulte and Wollman 1992; Wang et al. 2009; Miller et al. 2010; Work et al. 2010) and inducing gametogenesis, initiating sexual reproduction (Sager and Granick 1954; Martin and Goodenough 1975; Abe et al. 2004). One of the predominant metabolic changes observed in the microalga when starved for nitrogen is the accumulation of starch (Martin and Goodenough 1975; Ball et al. 1990; Hicks et al. 2001) as well

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as lipid reserves (Wang et al. 2009; Work et al. 2010; Siaut et al. 2011), making Chlamydomonas a good model for analyzing the molecular basics of starch and lipid body biosynthesis both with regard to basic and to applied research, the latter aiming at biodiesel production by microalgae. The large starch reserves accumulated by N-starved C. reinhardtii cells made this nutritional condition interesting with regard to H2 production, since the catabolism of starch is proposed to be one major electron source for photosynthetic H2 production in S-starved wild type C. reinhardtii cells (Melis et al. 2000; Fouchard et al. 2005; Hemschemeier et al. 2008a; Chochois et al. 2009). Under the conditions applied in this study, N-deprived Chlamydomonas cultures did accumulate about twice as much starch as S-deprived cells and they did develop a H2 metabolism. However, N-starved cultures did hardly consume their starch reserves, and the amount of H2 produced upon N starvation was only 50% of the H2 concentrations observed in the gas phase of S-deprived cells. Furthermore, the onset of intracellular hypoxic conditions as concluded from hydrogenase activity and H2 evolution was about 2 days later when compared with S-depleted cells. The delay in establishing hypoxic conditions observed in N-starved algal cultures was apparently due to a prolonged PSII activity. Both nutritional conditions caused a gradual decrease in photosynthetic O2 evolution rates and PSII photochemical efficiency (UPSII). This has been reported before in studies about S-deprived Chlamydomonas cultures both aerated (Wykoff et al. 1998) and sealed (Melis et al. 2000; Antal et al. 2003). The decrease of PSII activity has been explained by an accumulation of QB-non reducing PSII centers (Wykoff et al. 1998), by transition to state II (Wykoff et al. 1998), which, in Chlamydomonas, can result in a complete uncoupling of PSII from the photosynthetic electron transport chain (Finazzi et al. 1999), by an over-reduced plastoquinone pool (Antal et al. 2003) and by impaired turnover of the D1 protein due to a lack of S-containing amino acids (Zhang et al. 2002; Melis and Chen 2005). In S-deprived cells, a reduced photosynthetic activity can be observed after about 1 day, while a similar low level of photosynthesis is observed only after 4 days of P starvation (Wykoff et al. 1998; Melis et al. 2000). Studies on the C. reinhardtii sac1 mutant, which is deficient for a major regulator of the inducible response to S deficiency (Davies et al. 1994, 1996; Ravina et al. 2002) showed that the down-regulation of PSII activity as a response to S starvation is at least partially under SAC1 control, and hence a regulated process: O2 evolution rates of the sac1 mutants do not decrease in the first 24 h of S deprivation, and the cells die within 2 days, a fate which can be prevented by incubating the mutant cells either in the dark or upon addition of the

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PSII inhibitor DCMU in the light (Davies et al. 1996). It was shown in an elaborate study about photosynthetic parameters of sulfur- and phosphorous-deprived Chlamydomonas cells that the sac1 mutant did not accumulate QB-non reducing PSII centers in the absence of sulfur, but did so upon phosphorous starvation (Wykoff et al. 1998). In N-starved cells, a decline in photosynthetic O2 evolution and the maximum quantum yield (Fv/Fm) of PSII has been observed, too (Peltier and Schmidt 1991; Bulte and Wollman 1992; Work et al. 2010). Impaired photosynthetic activity is accompanied or caused by a decrease in light harvesting complex proteins as well as photosystem subunits and Rubisco (Plumley and Schmidt 1989; Peltier and Schmidt 1991; Bulte and Wollman 1992). According to a recent deep-sequencing transcriptome analysis, the decrease in photosynthetic proteins is in many cases due to lowered amounts of the respective transcripts (Miller et al. 2010). During the experiments shown in this study, PSIIactivity of N-starved Chlamydomonas cultures did decrease, but not as fast as observed in S-depleted cultures. This might be explained by three reasons: First, the PQ-pool in N-depleted cells might have been less reduced than in S-starved cells, allowing higher electron transfer rates from QA to QB and finally re-oxidation of QB. Though it has been shown that the PQ-pool of N-starved cells is strongly reduced (Peltier and Schmidt 1991; Bulte and Wollman 1992), and though in this study, cells from both nutritional conditions switched to state 2 conditions, an indicator of a reduced PQ-pool (Lemeille and Rochaix 2010) the 685 nm:715 nm ratio stayed on a slightly higher level in N-depleted cells. Therefore, both nutritional stress conditions result in a reduced PQ-pool and transition to state 2, but these developments seem to be moderately less pronounced in nitrogen-starved cells. The reason for a more oxidized PQ-pool in N- versus S-depleted algal cells might be a higher activity of the chlororespiratory pathway under N deprivation. A study on N-limited cells grown in constantly low ammonium concentrations (0.15 vs. 7.5 mM as present in full medium) showed that these cells had higher NADH-menadione oxidoreductase activity (Peltier and Schmidt 1991), indicating a higher activity of non-photochemical PQ-reduction by the class-II type NAD(P)H dehydrogenase Nda2, which was characterized on the genetic and biochemical level only recently (Jans et al. 2008; Desplats et al. 2009). Enhanced activity of Nda2 alone would result in a more reduced PQ-pool. However, chlororespiratory O2 uptake activity resulting from plastid terminal oxidase (PTOX) (Cournac et al. 2000, 2002) was estimated to be about 30% higher in N-limited than in control cells, while the total respiratory activity was about twice as high (Peltier and Schmidt 1991). In S-deprived, hydrogen-producing Chlamydomonas cells, the activity of PTOX was estimated

Planta (2012) 235:729–745

based on the respiratory rate left after potassium cyanide (KCN) and salicyl hydroxamic acid (SHAM) addition, which inhibit mitochondrial cytochrome oxidase and alternative oxidase, respectively. The data did not show an enhanced PTOX activity, but rather a moderate decrease when compared with nutrient-replete cells (Antal et al. 2003). During the nutrient-deprivation experiments conducted in this study, respiratory O2 consumption rates in the dark did not differ strongly in N- or S-depleted C. reinhardtii cultures. However, it should be noted that aliquots of the cell suspension were withdrawn from the incubation flasks and transferred to the measuring chamber of the oxygen electrode. This transfer resulted in the entry of O2 from air. Consequently, O2 consumption rates measured in the culture aliquots might rather reflect the respiratory capacity in the dark than actual O2 uptake rates in the sealed and illuminated incubation flasks. In contrast to the O2 concentration in the headspace of S-deficient algal cultures, which decreased to non-detectable values after about 50 h of incubation, significant amounts of O2 (about 20% of the air value) remained in the gas phase above N-starved cells. The latter might therefore still use some O2 as an electron acceptor either for mitochondrial or for plastid respiration in situ. The differences in pigment content observed in C. reinhardtii cells transferred to S- or N-free medium might be a second explanation for the observed differences in PSII photochemical activity. N-starved cells showed lower cellular chlorophyll contents as well as higher carotenoid to chlorophyll and Chl a:Chl b ratios. The latter might be explained by a preferential loss of LHCII antennae (Green and Durnford 1996; Elrad et al. 2002). Lower Chl b and LHCII levels in N-limited Chlamydomonas cells have been reported before (Plumley and Schmidt 1989). The differences in pigmentation and antennae composition probably have an impact on light energy transfer to the photosystems, resulting in less excited and thus more open PSII reaction centers. A third reason for the higher PSII quantum yield of Nrelative to S-depleted cells might be the early regulatory ‘‘closure’’ of PSII upon S starvation, which is under control of the SAC1 protein as outlined above. In principal, similar negative feedback events ultimately limiting PSII activity are given in both nutritional stress conditions, which are both marked by a strong decrease in CO2 assimilation (Martin and Goodenough 1975; Zhang et al. 2002; Hemschemeier et al. 2008a) and other anabolic pathways such as protein biosynthesis (Grossman 2000). However, these responses might be accelerated in S-deficient cells due to the limited intracellular resources of sulfur (Wykoff et al. 1998; Grossman 2000). The fast and SAC1-controlled accumulation of QB-non reducing PSII centers seems to be a specific response of S-deprived cells (Davies et al. 1996;

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Wykoff et al. 1998), and might have caused the differences observed in this study. Despite of the 2-day delay of the onset of hydrogen evolution observed in nitrogen- versus sulfur-depleted cells, hypoxic conditions were finally established, as shown by the induction of [FeFe]-hydrogenase biosynthesis and activity as a sensitive marker for low oxygen tensions (Ghirardi et al. 1997; Happe and Kaminski 2002; Goldet et al. 2009; Stripp et al. 2009). However, N-starved cells accumulated much less H2 than S-deprived cells, despite of their large starch reserves. One simple reason for that might be the 2-day delay of the onset of H2-producing activity. S-depleted Chlamydomonas cultures exhibit a stagnation of H2-production after 4–5 days of nutrient starvation, in spite of the fact that the cells do still contain considerable amounts of starch at this time-point. It has been proposed that this slow-down of H2 evolution is rather due to a general impairment of metabolism caused by starvation than to a depletion of energy reserves (Timmins et al. 2009). A similar exhaustion will occur in N-starved cells, resulting in a reduced capacity to photo-evolve H2 after several days of nutrient starvation. However, lower H2 yields of N-deficient Chlamydomonas cultures might be due to N starvation specific changes of the photosynthetic apparatus. Probably due to initially higher PSII activity, the O2 concentration in the gas phase of N-depleted cells remained on a significant level throughout the experiment. This might have resulted in constant destruction of hydrogenase enzymes and be responsible for the slower increase of in vitro hydrogenase activity upon N deprivation. This, in turn, could be one reason for the lower H2 yields of N-deficient cells, though the hydrogenase enzyme is probably not the limiting factor during photosynthetic H2 generation (Winkler et al. 2009). The presence of O2 would also favor respiratory activity, diverting electrons away from H2 production and preventing a Pasteur effect. Indeed, the rate of starch degradation in N-deprived cells transferred to darkness was much higher than in illuminated cells in the same medium, and furthermore in a similar range as observed in shaded S-starved algae. This indicates that starch degrading and downstream pathways are not generally affected upon N starvation, but are inhibited in illuminated cells. Notably, treatment of illuminated N-deprived cells with the PSII inhibitor DCMU did not significantly influence H2 yields. This indicates that electrons provided by wateroxidation at PSII do not contribute to linear electron transport towards the hydrogenase, and moreover, it raises the question about the electron source for the comparably low, but still significant H2 evolution activity of N-depleted algal cells. H2 production by S-deficient Chlamydomonas cultures are reduced in the presence of DCMU, though to varying extents (Fouchard et al. 2005; Hemschemeier et al.

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2008a) and depending on the time point within the sulfur deprivation experiment (Laurinavichene et al. 2004; Hemschemeier et al. 2008a). It has been proposed that residual H2-producing activity of DCMU-treated S-starved cells depends on non-photochemical PQ-reduction by stromal reductants originating from starch breakdown, which, however, is slow in N-deprived cultures. Another source of electrons might be acetate, which is abundant in the medium of photoheterotrophically cultivated Chlamydomonas cells. However, the acetate contents of both media did not decrease further after 48 h (TAP-S) or 72 h (TAP-N) under the experimental conditions applied in this study, an observation which is consistent with findings using N-starved, non-sealed algal cultures (Work et al. 2010) and H2-producing S-depleted cells (Melis et al. 2000). Results from another study analyzing the fate of 13C from labeled acetate indicated that C. reinhardtii cells subjected to N starvation for 48 h did not convert external acetate into glucose via the gluconeogenetic pathway or assimilated it via the glyoxylate cycle, but probably used acetate to build up triacylglycerol (Miller et al. 2010). However, oxidative degradation of other cell components might contribute to the pool of stromal reducing equivalents in both S- and N-starved cultures. The starchless C. reinhardtii mutant sta6 (Zabawinski et al. 2001) produces H2 in the presence of DCMU, though considerably less than the C. reinhardtii wild type under the same condition (Chochois et al. 2009). Still, this result indicates that there are other oxidative pathways contributing to nonphotochemical PQ-reduction and PSII-independent photosynthetic H2 production, respectively. These pathways might include protein and amino acid oxidation, respectively. The protein content of C. reinhardtii cells has been shown to increase in the first 24–48 h of sulfur deprivation and to decrease thereafter, a pattern similar to the build-up and subsequent degradation of starch (Melis et al. 2000; Zhang et al. 2002). Melis et al. (2000) proposed that protein might be a primary substrate for non-photochemical PQ-reduction under these conditions. The protein content of N-starved Chlamydomonas cells has been shown to decrease (Hipkin et al. 1982), so that the resulting free amino acids might constitute a considerable source of electrons for N-deficient algae. The results of dark fermentation analyses presented here furthermore indicate that N-deficient cells have a higher potential for light-independent H2 evolution. It has been proposed that pyruvate ferredoxin oxidoreductase (PFR1) catalyzed ferredoxin reduction might be one source of electrons for H2-production in the dark (Grossman et al. 2011; Philipps et al. 2011). Dark-incubated N-starved cells did produce similar amounts of ethanol, but less formate than S-starved algae, which might indicate a higher PFR1 activity, resulting in higher levels of reduced ferredoxin

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and H2-generation. More elaborate metabolic network analyses would be required to address this question, but a higher activity of fermentative H2-generating pathways might be a further explanation for the observation that PSII was no major electron source for H2 evolution in N-deprived illuminated C. reinhardtii cultures. The impairment of photosynthetic H2-production in N-starved algae might be due to a bottleneck downstream of the PQ-pool. It has been shown that the cytochrome b6f complex is specifically degraded in N-limited Chlamydomonas cells incubated photoheterotrophically (Bulte and Wollman 1992). In S-depleted cells, the amounts of functional cytochrome b6f complexes and PSI decrease in time, too, but much slower than the amount of PSII, as shown spectrophotometrically (Melis et al. 2000) and in analyses of partial photosynthetic reactions using artificial electron donors and acceptors in crude cell membrane preparations (Wykoff et al. 1998). Both phenomena were reflected in this study by a stronger decrease of cytochrome f levels in N- versus S-depleted C. reinhardtii cultures. Furthermore, the addition of methyl viologen to culture aliquots of sulfur- or nitrogen-deficient C. reinhardtii cultures resulted in a significant increase of the efficiency of PSII photochemistry (UPSII) in S-depleted, but only moderately in N-depleted cells. This indicates that artificially optimized oxidation of PSI allows enhanced linear electron transport in S-deficient, but hardly in N-depleted cells. This could be due to a general lower reduction state of the photosynthetic electron transport chain in N-deprived cells or to PSII-derived electrons being diverted away at the site of the PQ-pool and transferred to O2, as discussed above. The latter hypothesis might be in line with the lacking effect of DCMU on H2-yields in N-starved cells. In view of the results of this work and previous studies regarding the different changes of components of the photosynthetic electron transport chain in nitrogenversus sulfur starvation (Plumley and Schmidt 1989; Peltier and Schmidt 1991; Bulte and Wollman 1992; Wykoff et al. 1998; Melis et al. 2000), another hypothesis would explain the minor effect of methyl viologen on UPSII in N-starved cells by a bottleneck generated by the degradation of the cytochrome b6f complex. In addition, a decrease in ferredoxin amounts was observed in N-deficient algae, but not upon sulfur starvation (Jacobs et al. 2009). A bottleneck between the PQ-pool and the hydrogenase, either due to limited cytochrome b6f complex or PetF amounts or both would be one explanation for the inability of N-deficient C. reinhardtii cells to channel electrons resulting from oxidative starch degradation into the H2 evolution pathway. This might also have an impact on attempts to optimize photo-hydrogen evolution by C. reinhardtii. The low hydrogen yields despite higher starch reserves observed in N-starved cells indicate that higher carbohydrate deposits

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are not by default converted in higher hydrogen yields, unless the electrons derived from oxidative pathways are transferred to the PQ-pool and photosynthetic electron transport matches the electron transfer capacity. Furthermore, the effect of methyl viologen on PSII photochemistry in S-depleted Chlamydomonas cultures suggests that electron transfer reactions at PSI might be optimized to result in higher hydrogen evolution rates. Acknowledgments Financial support by the BMBF (H2-Designzelle), the DAAD (Project Based Personnel Exchange Programme with Spain, contract D/06/12793), the European Union (FP7, SolarH2 Consortium) and the VW foundation (LigH2t) is gratefully acknowledged. We are furthermore very thankful for fruitful suggestions and discussions with Emilio Ferna´ndez (University of Co´rdoba, Spain), Laurent Cournac (CEA Cadarache, France) and Ga´bor Berna´t (Ruhr-University of Bochum, Germany) as well as for gifts of antibodies from Francis-Andre´ Wollman (CNRS—University of Paris 6, France), Jean-David Rochaix (University of Geneva, Switzerland) and Sungsoon Park (Aurora Biofuels, USA).

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