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Reversible restriction of electron flow across cytochrome b6f in dark acclimated cells limited for downstream electron sinks

Photosynthetic organisms frequently experience abiotic stresses that restrict their growth and development. Under such circumstances, most absorbed solar energy cannot be used for CO2 fixation and can cause the photoproduction of reactive oxygen species (ROS) that can damage the photosynthetic reaction centers, photosystems I and II (PSI and PSII), resulting in a decline in primary productivity. This work describes a biological ‘switch’ in the green alga Chlamydomonas reinhardtii that reversibly restricts photosynthetic electron transport (PET) at the cytochrome b6f complex when reductant and ATP generated by PET are in excess of the capacity of carbon metabolism to utilize these products; we specifically show a restriction at this switch when sta6 mutant cells, which cannot synthesize starch, are limited for nitrogen (growth inhibition) and subjected to a dark-to-light transition. This restriction, which may be a form of photosynthetic control, causes diminished electron flow to PSI, which prevents PSI photodamage. When electron flow is blocked the plastid alternative oxidase (PTOX) may also become activated, functioning as an electron valve that dissipates some of the excitation energy absorbed by PSII thereby lessening PSII photoinhibition. Furthermore, illumination of the cells following the dark acclimation gradually diminishes the restriction at cytochrome b6f complex. Elucidating this photoprotective mechanism and its modulating factors may offer new insights into mechanisms associated with photosynthetic control and offer new directions for optimizing photosynthesis.


INTRODUCTION
Oxygenic photosynthesis is a fundamental biological process performed by plants, algae, and photosynthetic bacteria that supports most life on Earth. The photosynthetic complexes absorb light, which provides the energy to oxidize water, using the extracted electrons to generate NADPH, reduced ferredoxin (FDX), and energy as ATP and electrochemical gradients. The ATP and NADPH fuel downstream CO2 fixation via the Calvin-Benson-Bassham cycle (CBBC) ( Figure 1A), which produces the carbon (C) backbones required for nutrient assimilation, the generation of essential cellular building blocks, respiratory substrates, and C polymers (e.g., starch and lipids) that serve in energy storage. Coordination between PET and C metabolism involves 'redox control' of CBBC enzymes, which is primarily mediated by FDX and thioredoxin (TRX) (Buchanan, 2016). In the light, as the redox potential in the chloroplast becomes more negative, TRX reduces disulfide bonds on targeted CBBC enzymes leading to an increase in their activity by up to 40-fold, while oxidation of these disulfides when the redox potential becomes more positive results in diminished enzyme activity (Michelet et al., 2013;Wirtz et al., 1982) (light blue box, upper right in Figure 1A).
In nature, environmental change and abiotic stress can cause an imbalance in the activities of PET and the CBBC. Under conditions in which the photosynthetic machinery is unable to effectively balance photosynthetic electron output with utilization, light energy may be converted to toxic ROS that can damage photosystems I (PSI) and II (PSII) through a process known as photoinhibition, and modify other cellular components including DNA and lipids, potentially leading to cell death. Mechanisms have evolved to protect the cell from photodamage through the dissipation of excess absorbed light energy as heat in a process known as nonphotochemical quenching (NPQ) (Müller et al., 2001;Rochaix, 2014). NPQ and other mechanisms that reduce the damaging impact of excess absorbed light energy are highly relevant to both ecological and agricultural issues as anthropogenic activities are escalating the rate of climate/environmental change (Malnoë, 2018;Müller et al., 2001), which can markedly impact photosynthetic function.
An excess of energetic photosynthetic electrons, however, can also be dissipated by photochemical outlets that involve the reduction of oxygen (O2) to regenerate water (H2O-to-H2O cycle) (Curien et al., 2016). In the Mehler reaction (Mehler, 1951), for example, O2 is directly reduced on the acceptor side of PSI (Kozuleva et al., 2021) to superoxide (O2 -), followed by enzymatic conversion of the O2radical to H2O2 and O2 by superoxide dismutase, followed by conversion of H2O2 to H2O by catalase (Asada, 2000). However, if these ROS cannot be rapidly detoxified, they may damage the photosynthetic apparatus and inhibit other cellular processes. An additional reaction that facilitates non-destructive O2 reduction (without ROS production) involves PTOX (plastoquinol:oxygen oxidoreductase; Figure 1A). This enzyme controls the redox state of the plastoquinone (PQ) pool and the chloroplast stroma through chlororespiration, a lightindependent process that involves the NDH complex in plants and the NDA2 protein in green algae (both function as NADPH:O2 oxidoreductases) (Nawrocki et al., 2015;Nixon, 2000;Peltier and Cournac, 2002;Bennoun, 1982). Moreover, when cells are exposed to light and the excitation pressure is higher than the energetic demands for CO2 fixation, PTOX may act as a safety valve by catalyzing a H2O-to-H2O cycle, a reaction that frequently occurs during rapid changes in the stromal redox state (e.g., transitions from dark to light or from low to high light) and during periods of environmental stress (Saroussi et al., 2016;Bailey et al., 2008;Krieger-Liszkay and Feilke, 2016;Saroussi et al., 2019). Other stromal enzymes that facilitate a H2O-to-H2O cycle include flavodiiron proteins (FLV, NADPH:flavin oxidoreductase; homologs have not been reported in angiosperms; Figure 1A), which oxidize NADPH generated by PSI, thereby protecting PSI (and consequently PSII) from oxidative damage during a rapid increase in reducing power that cannot be used by the CBBC (e.g., when CBBC enzymes are not fully active) (Gurrieri et al., 2021;Chaux et al., 2017;Allahverdiyeva et al., 2013;Gerotto et al., 2016).
When ROS causes damage to PSII, it can be rapidly repaired [extensively reviewed (Theis and Schroda, 2016;Komenda et al., 2012;Nath et al., 2013)], while, in contrast, recovery of PSI from oxidative damage is much slower, and in some cases irreversible (Sonoike, 2011;Heinnickel et al., 2016;Shimakawa et al., 2016;Zivcak et al., 2015). PSI photoinhibition can occur when electron flow from PSII exceeds the capacity of oxidized electron acceptors downstream of PSI, including metabolites of the CBBC (i.e., an asymmetry between PET and photosynthetic C metabolism; also known as acceptor side limitation). In this case, PSI reaction centers become reduced, O2may be generated and react with and damage PSI iron-sulfur clusters, which can lead to degradation of PSI core subunits (Trinh et al., 2021;Sonoike, 2011;Terashima et al., 1998;Asada, 2000;Sonoike, 1996); as a consequence, the level of oxidizable P700 dramatically declines (Terashima et al., 1998(Terashima et al., , 1994Hald et al., 2008). Although some aspects of control are known, further elucidation is required to determine how photosynthetic organisms exposed to strong abiotic fluctuations that perturb the balance between PET and CO2 fixation and assimilation, modulate the rate of electron transport to protect PSI from photoinhibition. There is evidence that suggests that the PGR5 protein, which is involved in cyclic electron flow (CEF) around PSI, participates in protecting PSI from damage when plants are exposed to fluctuating and high light by attenuating the rate of electron transport from PSII toward PSI (Leister et al., 2021;Yamori et al., 2016;Kono et al., 2014;Munekage et al., 2002;Suorsa et al., 2012;Munekage et al., 2008;Terashima et al., 1994); however, the mechanism associated with PGR5 function remains speculative. Another recently proposed mechanism, initially reported in cyanobacteria, describes an inhibition of Q-cycle turnover which slows down electron flow from the Cyt b6f complex to PSI [named Reduction-Induced Suppression of Electron flow, RISE ]. This control of Cyt b6f activity is a consequence of the development of a proton motive force (PMF) across the thylakoid membranes (Foyer et al., 2012;Tikhonov 2014, Tikhonov 2015 and contributes to photosynthetic control, the fact that intersystem electron transport is reversibly regulated at Cyt b6f. Supporting evidence of such reversible regulation of intersystem electron transport at Cyt b6f was recently summarized (Johnson and Berry, 2021).
In this study, we created an asymmetry between PET on the one hand, and growth, CO2 fixation and assimilation, on the other, by culturing a Chlamydomonas reinhardtii (designated Chlamydomonas throughout) mutant (sta6) that cannot synthesize starch (Zabawinski et al., 2001;Li et al., 2010) in medium devoid of nitrogen (N). This led to the establishment of a photosensitive regulatory pathway that appears to protect PSI from photodamage by modulating electron flow through the Cyt b6f complex, which may reflect photosynthetic control, while at the same time sustaining PSII activity by redirecting photosynthetic electron flow to the reduction of O2 through PTOX.

Asymmetry between absorbed light energy and carbon metabolism attenuates O2 evolution and electron transport
To disrupt the balance between the absorbed light energy and C metabolism, we deprived Chlamydomonas of N. This treatment, which resulted in the cessation of cell growth, eliminated the main cellular sink (growth) for photosynthetically generated reductant, ATP, and C backbones.
N-deprived Chlamydomonas cells rapidly acclimate to limiting N conditions by activating expression of specific pathways involved in N acquisition and lipid and starch synthesis (Davey et al., 2014;Siaut et al., 2011); the synthesis of starch and triacylglycerides, N-free C polymers, serve as sinks for energy and fixed C generated by PET and the CBBC. To further eliminate potential pathways for assimilating and storing fixed C and utilizing ATP, we combined N deprivation with the use of the Chlamydomonas sta6 mutant (Zabawinski et al., 2001). This mutant is null for the small subunit of ADP-glucose pyrophosphorylase (AGPase), an enzyme that catalyzes the synthesis of ADP-glucose from glucose-1-phosphate and ATP (Figure 1B), the rate-limiting step in starch synthesis. The sta6 mutant, which makes no detectable starch (Figure S1), can be phenotypically rescued with an ectopically expressed, wild type (WT) copy of the STA6 gene; the rescued strain, C6 (sta6::STA6), synthesizes starch; ( Figure S1) (Li et al., 2010). The genome of the C6 strain was recently sequenced and has been used as a reference for evaluating the sta6 phenotype (Siaut et al., 2011;Schmollinger et al., 2014;Blaby et al., 2013;Anderson et al., 2016).
To assess the balance between PET and downstream C metabolism in sta6 and C6, we cultured Chlamydomonas cells for 24 h at 100 µmol photons m -2 s -1 in MOPS-Tris medium (see METHODS) devoid of N and then measured net O2 evolution immediately following a dark-tolight transition (experimental design in Figure 1C). N-deprived C6 maintained high rates of net O2 evolution (Pmaxnet = ~160 μmol O2 mg Chl -1 h -1 , black curve in Figure 1D), similar to those of C6 cultured in the presence of N (~180 μmol O2 mg Chl -1 h -1 , Figure S2). In contrast, net O2 evolution in the N-deprived sta6 mutant was nearly undetectable (i.e., near the O2 compensation point; Figure 1D) under N deprivation conditions, while in the presence of N, net O2 evolution was ~60 μmol O2 mg Chl -1 h -1 ( Figure S2). These results suggest that a depressed ability to use reductant because of the loss of two major electron sinks resulted in strong disruption of the balance between PET and downstream photosynthetic C metabolism.
To further characterize PET with respect to C metabolism, we monitored linear electron flow (LEF) in sta6 and C6 using fluorescence spectroscopy. Following a period of 24 h of N deprivation, the effective PSII quantum yield (ΦPSII) of sta6 was much lower than that of C6 at all tested light intensities, while no strong differences in ΦPSII between the strains were observed when the cells were cultured in the presence of N ( Figure S3). We also measured the re-reduction rate of P700 in the presence of DCMU and hydroxylamine to evaluate the contribution of NDA2dependent reduction of P700 (Saroussi et al., 2016). The rate of P700 re-reduction in N-deprived, WT Chlamydomonas was previously shown to increase as a consequence of elevation of the NDA2-dependent pathway (Saroussi et al., 2016). In this pathway, electrons derived from NADPH are used to reduce the PQ pool. In the absence of N, the re-reduction rate of P700 measured for C6 increased by ~1.5 fold (relative to the re-reduction rate in the presence of N; Figure S4A and B), similar to the observation previously made for WT Chlamydomonas cells (Saroussi et al., 2016).
However, unlike C6, N deprivation of sta6 resulted in a slowing of the rate of P700 re-reduction ( Figure S4A and B). Hence, N-deprived sta6 has both diminished LEF and NDA2-dependent rereduction of PSI.
We also explored the possibility that photosynthetically generated electrons in N-deprived C6 and sta6 were shuttled to the mitochondrial respiratory electron transport chain. N-deprived and N-replete C6 displayed similar rates of dark respiration immediately following the extinction of the light ( Figure S5A). In contrast, while respiratory O2 uptake was observed in sta6 grown in the presence of N, almost no dark respiration was observed in N-deprived sta6 (Figure S5A and B). In sum, in the absence of starch synthesis (sta6) during N deprivation, PET, net O2 evolution, and mitochondrial respiration were all severely diminished. Figure 1: Key aspects of the photosynthetic system and experimental design: A. Schematic representation of the link between PET and downstream C metabolism (CBBC; upper left). Electrons extracted from H2O by PSII flow through the electron transport chain generating a proton (H + ) gradient across the thylakoid membranes and ultimately reducing NADP + to NADPH. The energy of the H + gradient is used to synthesize ATP by the ATP synthase. CBBC utilizes both ATP and NADPH (green lettering and dashed green arrows) to fix CO2 by Rubisco (light blue box). The CBBC includes several enzymes that are activated by redox control (activated enzymes highlighted in orange boxes with red frames). Upper Right: Redox activation of targeted CBBC enzymes by reduction of specific disulfide bonds, which is mediated by FDX and TRX systems. Activation is reversible upon oxidation of the disulfide bonds in the dark. B. Starch biosynthesis: Metabolites of the starch synthesis pathway are in black lettering. Starch metabolismrelated enzymes are highlighted by a light green rectangular background. C. Experimental design: After growing the cells mixotrophically in moderate light (100 µmol photons m -2 s -1 ) to mid log phase, they were transferred to photoautotrophic medium devoid of N and acclimated for 24 h in moderate light. Following a 30 min dark acclimation period, the cells were exposed to light (intensities indicated in text). D. Net O2 evolution of N-deprived C6 (black curve) and sta6 (red curve) cultures. N=3 ± SE. Abbreviations: PSI -photosystem I; PSII -photosystem II; Cyt b6f -cytochrome b6f; ISP -iron sulfur protein; Cyt f -cytochrome f; PC -plastocyanin; PTOX -plastid terminal oxidase; FLV, flavodiiron protein; FDX -ferredoxin; FNR -ferredoxin-NADPH oxidoreductase; FTR -ferredoxin-thioredoxin reductase; Rubisco -ribulose-1-5-bisphosphate carboxylase-oxygenase; GAPDH -glyceraldehyde 3-phosphate dehydrogenase; PGM -phosphoglucomutase; AGPase -ADP-glucose pyrophosphorylase; SS -starch synthase; FBPase -fructose-1,6-bisphosphatase; PRK -phosphoribulokinase. Metabolites: 3PG -3-phospho-glyceraldehyde; F6P -fructose-6-phosphate; G6P -glucose-6-phosphate; G1P-glucose-1-phosphate. PAR -photosynthetically active radiation.
The negligible net O2 production exhibited by N-deprived sta6 cells following a dark-tolight transition ( Figure 1D) can be explained by one or a combination of the following: a.
Feedback on electron transport caused by the extreme imbalance between PET and C metabolism leads to inactivation of PSII which inhibits water oxidation, O2 production, and electron transport; b. PSII remains active (i.e., H2O is oxidized), but the O2 produced is consumed through activation of a H2O-to-H2O cycle associated with alternative outlets for photosynthetic electrons (Curien et al., 2016); c. An electron transport reaction downstream of PSII is restricted that slows the rate of H2O oxidation. To evaluate these hypotheses, we first analyzed the maximal quantum efficiency of PSII (Fv/Fm) and whether the level of the PSII core subunit, D1, was affected under our experimental conditions. We observe a 20% change in the Fv/Fm of sta6 cells deprived of N while no change was observed for C6 (comparing sta6 with C6 cells under -N and +N conditions, Figure   S6A), which does not necessarily reflect the extent of photoinhibition, while the levels of the PSII reaction center protein (determined immunologically) remained approximately the same (normalized to tubulin, Figure S6B). Furthermore, to assess PSII activity, we measured the capacity of C6 and sta6 cells to evolve O2 in the presence of p-benzoquinone (PBQ), an artificial electron acceptor that oxidizes the PQ pool. The rate of O2 evolution in the presence of this compound reflects the maximal capacity of PSII to oxidize H2O and evolve O2. No significant differences in the levels of O2 evolution were observed with the addition of PBQ to C6 and sta6 cells that were cultured in the presence of N ( Figure S6C and D). Under N deprivation conditions, however, the net rate of O2 evolution in sta6 cells was zero (similar results in Figure 1D); most of the activity was regained when the cells were supplemented with PBQ. These results demonstrate that PSII of N-deprived sta6 is still active.
We next monitored the real-time exchange of O2 in N-deprived sta6 cells using membrane inlet mass spectrometry (MIMS) to distinguish between light-dependent production and uptake of O2 (monitoring the 16 O2 signal for production and the 18 O2 signal for uptake; Figure 2). N-deprived sta6, treated as indicated in Figure 1C (i.e., dark-to-light transition), produced O2 at a maximal rate (Pmax) of ~25 µmol O2 mg Chl -1 h -1 soon after being exposed to 300 µmol photons m -2 s -1 (Figure 2A). However, the rate of O2 production and uptake were similar (compare Figure 2A with 2B), which resulted in approximately zero net O2 evolution (Figures 2C and Figure 1D). Net zero O2 evolution was not observed when sta6 cells were cultured in the presence of N (compare  Light-induced gross O2 production ( 16 O2 evolution, green curve), and B. uptake ( 18 O2 consumption, red curve), for N-deprived sta6 following a dark (30 min) to light (300 μmol photons m -2 s -1 ) transition; gas exchange was monitored for 6 min in the light using MIMS. Prior to turning on the light, the cells were maintained for 5 min in the dark (in the MIMS chamber) and then for 3 min in the dark following illumination. C. Net O2 production (blue curve), was calculated according to the formula given in the Methods. Each trace represents an average of at least 3 biological replicates ± SE.

N-deprived sta6 cells have a highly reduced quinone pool
We next explored the mechanism underlying impaired O2 production and electron transport in the sta6 mutant (Figures 2, S3 and S4). To determine if the pool of electron acceptors downstream of PSII were sufficient to support PSII activity, we quantified the photochemical efficiency (qL) of N-deprived C6 and sta6 as a function of light intensity (Figure 3). The photosynthetic parameter qL indicates the redox state of QA, the primary electron acceptor of the PSII reaction center.
Assuming that QA and QB are in equilibrium, qL would reflect the redox status of the PQ pool; a lower qL value indicates a more reduced electron transport chain (Kramer et al., 2004). To create a direct relationship between the qL value and the redox state, we used an inverse function, 1-qL, which positively correlates with the PQ pool redox state. Following dark acclimation, the 1-qL values for sta6 cells were higher at any given light intensity than the 1-qL values measured for C6 cells, suggesting a more negative redox potential (more reduced PQ pool) in sta6 relative to C6 at each of the light intensities. Even when exposed to low light (LL, 15 µmol photons m -2 s -1 ), the 1-qL value was at 70% maximum for sta6 (suggesting a highly reduced PQ pool). In contrast, for C6 at the same light intensity, the 1-qL value attained only 20% of the maximum value ( Figure   3A). Surprisingly, following a 1 h light acclimation of sta6, the 1-qL levels declined and were only slightly higher than those of C6; that is, the 1-qL values following exposure of sta6 to LL indicated a much more highly oxidized PQ pool ( Figure 3B, compare red solid and dashed curves). After exposure to the same light conditions, we did not observe a significant change in 1-qL values for C6 (compare black curves in Figure 3A and 3B). These results suggest that following a period of dark acclimation, the PQ pool of the N-deprived sta6 mutant is maintained in a highly reduced state, although the redox pressure can be significantly relieved by a 1 h exposure to light. These observations led to the hypothesis that N-deprived sta6 experiences a reversible restriction in PET downstream of the PQ pool. 1-qL, used to estimate the redox state of the PQ pool, was calculated for C6 and sta6 (black and red curves, respectively), from a light curve in which the fluorescence output was quantified using a Dual-PAM fluorometer (Walz, Germany) following A, dark acclimation, and B, 1 h of acclimation to 100 µmol photons m -2 s -1 . For comparison, we included the 1-qL data of dark-acclimated sta6, which is shown as the dashed red line in panel B. Each data point reflects an average of at least 3 biological replicates ± SD.

Cyt b6f complex is the site of a reversible restriction in electron transport
The hyper-reduced PQ pool in dark-acclimated, N-deprived sta6 suggested a downstream limitation in electron flow that impairs quinol oxidation. To explore this issue, we measured the oxidation/reduction kinetics of Cyt f in N-replete and deprived sta6 cells; Cyt f oxidation reflects electron flow from Cyt f to PC while re-reduction reflects electron flow from the iron-sulfur cluster of the Rieske iron-sulfur protein (ISP) to Cyt f. Initially, sta6 cells were cultured in the presence of N. After the cells were dark acclimated, they were exposed to a laser flash that oxidized Cyt f, which was then re-reduced with a half time (T1/2) of 24 ms (black curve of Figure 4A and black boxplot in Figure 4C). However, if the cells were light acclimated (put back under standard culture conditions at 100 µmol photons m -2 s -1 for 10 min prior to the measurements), the T1/2 for rereduction of Cyt f was 5 ms (red curve in Figure 4A and red boxplot in Figure 4C); this value is similar to the reported re-reduction rate of Cyt f in WT Chlamydomonas cells (de Vitry et al., 2004). A similar trend for Cyt f re-reduction kinetics was observed when N-replete C6 cells were subjected to the same dark-to-light transition (16 ms and 7 ms for dark-and 10 min lightacclimated cells, respectively; Inset of Figure 4A). Surprisingly, repeating the same analyses with dark-acclimated sta6 cells cultured in a medium devoid of N yielded a T1/2 re-reduction rate for Cyt f of 66 ms, which is much slower than in the presence of N (compare black curves in Figures   4B and Figure 4A and black boxplot in Figure 4D with 4C). However, after exposing N-deprived, dark-acclimated sta6 to light (same as above) for 10 min, the T1/2 for re-reduction of Cyt f was reduced to 28 ms (red curve of Figures 4B and red boxplot in Figure 4D) while after 1 h of light exposure the T1/2 for re-reduction was 17 ms (blue curve of Figures 4B and blue boxplot in Figure   4D). While the N-deprived C6 cells also exhibited some slowing of Cyt f re-reduction kinetics after dark acclimation (T1/2 = 39 ms) (Inset in Figure 4B), unlike sta6 cells, re-reduction of Cyt f was fast (similar to WT nutrient replete) following exposure of the cells to just 10 min of light (3 ms, Inset of Figure 4B). This modulated flow of electrons to Cyt f reflects a reversible restriction of electron transport that appears to be triggered by environmental conditions.
We also sought to determine if the reversible restriction in electron transport was a function of an imbalance between the amounts of Cyt f and PC in the N-deprived sta6 cells. Based on immunodetection, the accumulation of Cyt f and PC were similar in dark-and light-acclimated, Ndeprived sta6 cells. There was also no difference in the accumulation of these proteins in Ndeprived, dark-and light-acclimated C6 cells, although the levels of both Cyt f and PC appeared to be slightly lower in the mutant than in the complemented strain ( Figure S9). Overall, these results suggest that the difference between electron transport in dark-and light-acclimated cells was not affected by changes in the levels of these proteins.
Lastly, we explored the possibility that a low ATP turnover rate in sta6 leads to a compromised chloroplast ATP synthase activity which feeds back on PET (slowing PET) to increase the reduction state of the PQ pool. To test this hypothesis, we repeated the 1-qL measurements shown in Figure 3 for N-deprived sta6 in the presence of a protonophore and ionophore. We used nigericin, a drug that dissipates the H + gradient across the thylakoid membranes, which would eliminate the feedback of ATP synthesis on PET, and valinomycin, a drug that dissipates the electrochemical gradient across the thylakoid membranes. Upon supplementing the cells with 10 µM nigericin, 10 µM valinomycin, or both, 1-qL remained the same as that of non-treated cells (Figure S10), suggesting that PET was not restricted as a consequence of the rate of ATP synthesis. Hence, the H + and electrochemical gradients across the thylakoid membranes do not appear to be required to sustain a restriction in PET across the Cyt b6f complex.

Blocking electron flow sustains oxidized P700 and photoprotection
To validate the reversible restriction in electron transport at the level of Cyt b6f, we analyzed the redox state of P700. Diminished electron flow out of Cyt b6f would result in a mostly oxidized P700 following exposure of the cells to light. As the restriction is removed, the P700 redox status should become similar to that of cells in which no or little restriction was experienced. To examine this, we monitored oxidation/reduction kinetics of P700 in N-deprived sta6 and C6 cells ( Figure   5) that were treated identically to those cells used for the experiment given in Figure 4.
Importantly, the cells were not treated with DCMU, a drug that inhibits electron flow out of PSII, to allow electrons originating from both LEF and CEF to reduce oxidized P700. Following illumination of dark-acclimated C6 cells with non-saturating light (60 µmol photons m -2 s -1 for 10 sec), ~25% of the P700 was oxidized at the end of the light period (black curve, left panels of  Figure 5B), suggesting that electron flow from Cyt b6f to PSI was restricted in N-deprived sta6 cells immediately following the dark period; this restriction would potentially prevent (or diminish) PSI photoinhibition when the cells are limited for downstream electron acceptors. We then exposed N-deprived sta6 to 100 µmol photons m -2 s -1 for 1 h (similar to 1 h treatment of cells used in the measurements of 1-qL and Cyt f re-reduction kinetics; Figures 3B and 4B). Following this light acclimation period, P700 oxidation/reduction kinetics in C6 and sta6 were similar, with the proportion of oxidized P700 just over 25% for both strains at the end of the actinic light exposure (right panel in Figures 5A and right bar graph of Figure 5B), supporting the conclusion that N-deprived sta6 cells experience a reversible restriction in electron flow to Cyt f. Moreover, the total amount of photo-oxidized P700 was identical for the two strains (t-test, p-value = 0.66), suggesting that PSI in sta6 did not experience more oxidative damage than in C6 during the dark-to-light transition, despite the inability of sta6 to store starch, a major sink for assimilating electrons downstream of PSI during N deprivation. sta6 cultured in the presence and absence of N (A and B, respectively). A saturating laser flash (black arrow) was administered at time 0, immediately following the dark incubation, to oxidize Cyt f. Re-reduction kinetics were monitored by absorbance changes (described in Methods) at the indicated times following the flash. Half time (T1/2) of re-reduction was calculated by fitting the re-reduction component of Cyt f kinetics to a first order decay equation (although the kinetics may be more complex). All p-values for these fittings were <0.05. The data are an average of at least five biological replicates ±SE. Inset: Cyt f oxidation reduction kinetics of N-replete (A) and N-deprived (B) C6 cells. For N-replete C6 cells (left panel) the T1/2 for Cyt f re-reduction are 16 ± 3.2 and 7 ms ± 0.9 for the dark-and light-acclimated (10 min) cells, respectively. For N-deprived C6 cells (right panel) the T1/2 for Cyt f re-reduction are 39 ± 4 and 3 ms ± 0.6 for the dark-and light-acclimated (10 min) cells, respectively. Panels C and D show preset boxplots of the Cyt f T1/2 re-reduction rates following the dark incubation, or after 10 min and 1 h in the light following the dark incubation. For C, t-test ** p-value < 0.005. For D, One-way ANOVA, * P-value < 0.05; Post-hoc Tukey, p-value < 0.05. Note the different scales of the y-axes of panels C and D (there is a significant difference in the T1/2 values following 10 min of light exposure; t-test, p-value =0.0018).

PTOX-dependent H2O-to-H2O cycle drives PSII activity
While PSII was still functioning in N-deprived sta6 following a dark-to-light transition (Fv/Fm was >0.5, Figures 2 and S6), net O2 production was approximately zero (Figures 1D and 2C) because the amount of O2 released as a consequence of H2O oxidation was essentially identical to the amount of O2 consumed (Figure 2). Given the restriction in electron flow at the level of Cyt b6f (Figure 4), the most plausible alternative outlet that could facilitate reduction of O2 to H2O (i.e., O2 consumption) upstream of Cyt b6f in the electron transport chain is the alternative oxidase PTOX, which oxidizes a quinol molecule and reduces O2 to H2O (Nawrocki et al., 2015) ( Figure   6A). To evaluate this possibility, we repeated the MIMS experiment shown in Figure 2, but in the presence of 2 mM propyl gallate (PG), a drug that inhibits alternative oxidases such as PTOX.
Following dark acclimation (30 min, as in Figure 1C), the rates of light-dependent O2 production and uptake in N-deprived sta6 were dramatically diminished (300 µmol photons m -2 s -1 ; solid green and red curves in Figure 6B; note the y axis), with PG-treated sta6 reaching a Pmax 15-fold more slowly than the untreated cells (T1/2 = ~ 84 s for PG-treated and ~ 5 s for untreated cells; green bars in Figure 6C and compare solid and dashed green curves in Figure 6B). The O2 uptake rate for PG-treated sta6 was also diminished, reaching maximal uptake (Umax) 10-fold more slowly than the untreated cells (T1/2 = ~ 59 s for PG treated and ~ 5 s for untreated cells; red bars in Figure   6C and compare solid and dashed red curves in Figure 6B). Importantly, O2 production and uptake were directly correlated over the light period and the rate at which Umax and Pmax were obtained were not significantly different (59 ± 7 s, and 84 ± 28 s, respectively; N=3; t-test, p-value = 0.26; compare green and red bars in Figure 6C and green and red curves in Figure 6B); as a result, net O2 production was close to zero (blue curve, Figure 6B). We note that as O2 uptake in the light proceeds, the effect of PG diminishes (>100 sec), which may be a consequence of light inactivation of the inhibitor or a loss of the restriction of electron flow through Cyt b6f because PTOX activity is critical for sustaining the restriction. In addition, we examined the effect of PG on electron transport in N-deprived sta6 cells (Figure S11). While the restriction of electron transport across the Cyt b6f complex was not affected by PG (the switch remains restrictive, Figure S11A), electron flow generated by PSII (i.e., ΦPSII) further decreased in low/moderate light (Figure S11B), congruent with the observation that O2 production is diminished when PTOX is inhibited ( Figure   6B). In contrast, inactivation of PTOX in N-deprived C6 cells did not significantly impact the rate of O2 production or uptake (Pmax = ~120 µmol O2 mg Chl -1 h -1 for both treated and untreated cells; T1/2 to obtain Pmax for treated and untreated cells was 19.4 ± 2.1 s and 15.9 ± 0.23 s, respectively; Figure 6D). Together, these data indicate that following a restriction in electron flow at the Cyt b6f complex in the N-deprived sta6 mutant, inactivation of PTOX by PG strongly inhibits both the production and uptake of O2; PG does not affect O2 production in C6. Therefore, the H2O-to-H2O cycle catalyzed by PTOX appears to be the driving force for O2 production in the N-deprived sta6 mutant, supporting the conclusion that there is a reversible restriction of electron flow across the Cyt b6f complex in N-deprived sta6 cells during a dark-to-light transition. non-restricted pathways, respectively. B. Gross O2 production ( 16 O2 evolution, green curves) and gross O2 uptake ( 18 O2 consumption, red curves) are presented for N-deprived sta6 both in the presence (solid curves) and absence (broken curves) of 2 mM propyl gallate (PG), an inhibitor of PTOX. Net O2 production in the presence of PG is shown by the blue curves. Experimental design was as in Figure 2. Each curve represents an average of at least 3 biological replicates ± SE. C. T1/2 for attaining Pmax (green bars) and Umax (red bars) in N-deprived sta6 in the presence and absence of PG. Rates were calculated after fitting the O2 production and uptake curves to a first order exponential decay. The presented data is an average of at least 3 biological replicates ± SD. D. Gross O2 production ( 16 O2 evolution, green curves) and gross O2 uptake ( 18 O2 consumption, red curves) are presented for N-deprived C6 both in the presence (solid curves) and absence (broken curves) of 2 mM PG. Net O2 production in the presence of PG is shown by the blue curve. Experimental design was as in Figure 2. Each curve represents an average of at least 3 biological replicates ± SE.

DISCUSSION
Abiotic conditions on Earth, including temperature, light, moisture, and levels of available nutrients, are rapidly being altered in response to global climate change. Changing environmental conditions can constrain the growth and development of photosynthetic organisms and lead to an imbalance between the light energy absorbed by the photosynthetic apparatus and its use for the assimilation of CO2 by the CBBC, growth and the storage of starch and lipids. When such imbalances occur, some of the solar energy absorbed by PSI and PSII can be converted to ROS which creates oxidative stress and consequently photoinhibition, degradation of photosynthetic reaction centers, and damage to essential cellular components such as nucleic acids, proteins, and lipids. While an efficient system has evolved to repair damaged PSII, there is no corresponding system associated with PSI; recovery of PSI is slow or irreversible, which can strongly impact plant primary productivity. Hence, photosynthetic organisms have evolved mechanisms to dissipate excess absorbed light energy through NPQ (Müller et al., 2001) and the use of shunts and alternative electron valves (not directly involved in CO2 fixation) for photochemical quenching, many of which consume O2 (Saroussi et al., 2019;Curien et al., 2016;Roach and Krieger-Liszkay, 2014), and the implementation of photosynthetic control, which alters the conductance of electrons through the Cyt b6f complex, similar to a transistor as it was previously described (Johnson and Berry, 2021. The integration of these processes favor optimizing the use of light energy for CO2 fixation, while at the same time protecting reaction centers from damage. In this work, we explored mechanisms by which Chlamydomonas protects PSI from photoinhibition when there is marked asymmetry between the rate at which energy is absorbed and can be used in C metabolism. Chlamydomonas cells exposed to N deprivation undergo rapid metabolic acclimation. The cells stop growing and the light-generated reductant and ATP drive assimilation of CO2, mostly into starch and lipids (Davey et al., 2014;Siaut et al., 2011;Smith and Gilmour 2018). It is not surprising that a severe imbalance between solar energy absorption by reaction centers and C assimilation occurs when Chlamydomonas cells are both deprived of N (limits Ci utilization for growth) and are unable to synthesize starch, which is the case for the sta6 mutant; the imbalance would likely be most severe during dark-to-light transitions, conditions in which the CBBC is not initially fully active (Buchanan, 2016;Michelet et al., 2013;Wirtz et al., 1982; 2019) ( Figure 1A). Consequently, the pool of available electron acceptors downstream of PSI would be severely limited, potentially leading to ROS accumulation and the irreversible photoinhibition of PSI (Asada, 2000;Allahverdiyeva et al., 2015;Sonoike, 1996Sonoike, , 2011. A detailed analysis of electron transport in N-deprived sta6 cells demonstrated that following dark acclimation, Cyt f was readily oxidized by a laser flash (normal Cyt f oxidation; Figure 4B), but that its rate of re-reduction was very slow relative to WT cells or the complemented strain. These results indicated a restriction in electron flow between the PQ pool (or Qo) and Cyt f, raising the possibility that the restriction is located between the Rieske protein and Cyt f (Sarewicz et al., 2017). In accord with this possibility, the mutant exhibited a marked reduction in the rate of quinol oxidation, as indicated by hyper-reduction of the PQ pool ( Figure   3A) and diminished rates of LEF, P700 re-reduction via the NDA2 pathway, and mitochondrial respiration (Figures S3-S5). It was previously shown that when oxidation processes downstream of Cyt b6f are arrested or slowed, the iron-sulfur cluster of the complex adopts a metastable state that maintains electrons at a local energetic minimum making them nonreactive with O2 (Sarewicz et al., 2017). This stable cluster state of the Cyt b6f complex can sustain the block in electron transport without generating ROS, which might reflect altered interactions or accessibility between the iron-sulfur clusters and O2.
The electron transport restriction in dark acclimated, N-deprived sta6 cells resulted in elevated light-dependent P700 oxidization (i.e., P700 mostly in the P700 + state; left panels of Figures 5A and 5B), a state in which PSI is photo-protected since P700 + is an efficient quencher of absorbed excitation energy (i.e., converts it to heat) (Shubin et al., 2008), and the oxidized P700 cannot readily generate O2because of its positive charge. The 'restricted state' could be reversed upon exposure of the dark-acclimated, N-deprived sta6 cells to sub-saturating light. Interestingly, the restricted state could be reversed, resulting in low levels of O2 evolution following exposure of the dark-acclimated, N-deprived sta6 cells to sub-saturating light (Figures 3B and 4B); the time required for reversal in the light requires more than 10 min and is nearly complete in 1 h. These kinetic features may indicate a biochemical modification of the Cyt b6f complex, potentially posttranslational modification(s) and or activation of a pathway that consumes these electrons. Such a pathway might include the synthesis of lipids, which typically occurs in N deprived Chlamydomonas cells (Xu et al., 2008, Siaut et al., 2011, although this would need to be examined in longer term experiments (e.g. transfer to light reversal conditions for several hours instead of just one hour). Additionally, after this light reversal, the kinetics of P700 oxidation/reduction in N-deprived sta6 and C6 were similar (N-deprived C6 does not exhibit a similarly severe restriction in electron transport after incubation in the dark), suggesting that Cyt b6f associated regulation of PET may protect PSI in N-deprived sta6 from oxidative damage as the cells transition to the light.
The regulation of electron conductivity between PSII and PSI according to the cell's capacity to utilize reducing equivalents and ATP for CO2 fixation and in C assimilatory pathways (while protecting PSI) has been designated 'photosynthetic control' (Foyer et al., 1990;Weis et al., 1987). The specific factors that govern photosynthetic control have not been fully detailed, although studies are providing information concerning its mechanistic features and potentially its complexity. As an example, PSI photoinhibition was analyzed in two Nicotiana tabacum knockdown (KD) strains; one was a KD for an enzyme of the CBBC that consumes NADPH (glyceraldehyde 3-phosphate dehydrogenase, GAPDH) and a second was a KD for ferredoxin-NADPH oxidoreductase (FNR) which would be limited for its ability to generate NADPH. Both KD strains maintain a similar level of the total NAPDH + NAD + pool, but the GAPDH KD strain has a WT level of NADPH while the pool of pyridine nucleotides in the FNR KD strain is more oxidized (relatively higher levels of NADP + ). Both KD strains were severely limited for access to oxidized electron acceptors downstream of PSI and exhibited similarly low rates of CO2 fixation.
While the GAPDH KD strain maintained an oxidized P700 and was therefore protected from photoinhibition, the FNR KD strain was unable to protect PSI from photooxidative damage (Hald et al., 2008). It was hypothesized that redox poise, governed to a large extent by the NADPH/NADP + ratio, regulates electron transport; an elevated NADPH/NADP + ratio would downregulate Cyt b6f activity, which would lower the rate of PET and protect PSI from photodamage by favoring oxidized P700 (which is the case in the GAPDH KD) (Hald et al., 2008).
FNR, however, binds to Cyt b6f and probably forms a complex that facilitates CEF around PSI (Iwai et al., 2010). It has been argued (Joliot and Johnson, 2011) that the H + gradient across the thylakoid membranes, mediated by elevated rates of CEF, is a critical component for eliciting downregulation of Cyt b6f activity and preventing PSI photoinhibition. CEF, which competes with LEF and recycles electrons making them unavailable for use by CBBC, would promote the formation of a large H + gradient across the thylakoid membranes (Joliot and Johnson, 2011;Yamamoto and Shikanai, 2019). This large H + gradient could promote the dissipation of excess excitation energy absorbed by PSII as heat (qE component of NPQ), which in turn would diminish the rate of LEF and the generation of ROS. However, some studies suggest that qE may not be an essential component in the protection of PSI from photoinhibition (Joliot and Johnson, 2011;Tikkanen et al., 2015) and Johnson and Berry suggest that the function of qE is to balance the activity of the reaction centers under sub-saturating light conditions for the most effective use of the absorbed light energy in C metabolism (Johnson and Berry, 2021). Additionally, Shimakawa and co-workers (Shimakawa et al., 2018) suggested that the formation of a H + gradient is not sufficient to elicit oxidation (thus protection) of cyanobacterial PSI when CO2 levels are low.
Finally, this restriction cannot be immediately reversed by uncouplers which would relax the PMF across the thylakoid membranes. If the initial switch is triggered by an elevated PMF and/or redox conditions, then enzymatic processes in the light, whether associated with the 'opening' of electron valves downstream of PET and/or the reactivation of the Cyt b6f complex through protein modification, may be required (and require some time). Together, these studies suggest the importance of C metabolism/redox poise and potentially the H + gradient across thylakoid membranes in controlling photosynthetic electron flow and Cyt b6f complex activity. However, as recently stated (Johnson and Berry, 2021), there is still considerable uncertainty regarding the mechanism(s) of photosynthetic control and the reversible restriction of electron flow across the Cyt b6f complex, which may include more than one mechanism for regulating Cyt b6f activity.
Similar to a previous study (Shimakawa et al., 2018), our work suggests that the H + gradient is not necessary to sustain restricted electron flow to Cyt f. The restricted electron flow to Cyt f following dark acclimation of the N-deprived sta6 mutant was not reversed upon the combined addition of nigericin and valinomycin (Figure S10), although as discussed, it was mostly relieved when the mutant was subjected to a short exposure to sub-saturating light.
Despite the restriction in electron transport at the Cyt b6f complex, O2 production in dark acclimated, N-deprived sta6 cells was not completely arrested. Real-time measurements of O2 exchange demonstrated a maximal O2 production rate of 20-25 µmol O2 mg Chl -1 h -1 (this O2 was consumed at approximately the same rate as it was generated; Figure 2), which is 4-5-fold slower than the rate of O2 production by N-deprived C6 cells (~120 µmol O2 mg Chl -1 h -1 ; Figure 6D).
Based on the use of the inhibitor PG, the O2 production and uptake (and electron transfer) in Ndeprived sta6 cells could be almost completely attributed to the function of a PTOX dependent H2O-to-H2O cycle (Figure 6B and C). Importantly, the rates of O2 production and uptake measured when PET was blocked were 5-6 e -PSI -1 s -1 (conversions between O2 production and electron transport rates were performed according to the ratio of 1 e -PSI -1 s - 4 µmol O2 mg Chl -1 h -1 , as calculated by Johnson and Alric (Johnson and Alric, 2012)), which correlates with the maximal rate of the PTOX reaction measured in Chlamydomonas (Houille-Vernes et al., 2011), further supporting our proposal that PTOX is the main electron outlet in dark-acclimated, Ndeprived sta6 cells. Therefore, we suggest that the photochemical quenching process mediated by PTOX is an important component of photosynthetic control and may impact PET. The photochemical quenching activity of PTOX may not only contribute to protection of PSII but may also generate a sufficient H + gradient (through the H2O-to-H2O cycle, which releases 2 H + on the lumenal side of the thylakoid membranes while taking up 2 H + on the stromal side) to enable ATP synthesis and partially fuel the turnover and repair of damaged PSII.
As in previous studies, the work described in this manuscript suggests that photosynthetic control includes the use of an 'activity' gauge associated with the conductivity of electrons through the Cyt b6f complex or as the Cyt b6f was previously described to be functioning like a transistor in an electrical circuit (Johnson and Berry, 2021). Although their work described Cyt b6f in controlling steady-state photosynthesis in vascular plants, these results here presented showcase similarities in responses across different organisms and indicate deeply conserved features of this control.
Our results indicate that this control can be activated in the dark in N-deprived sta6 cells, which generates highly reducing conditions in the chloroplast (and potentially a delta pH across the thylakoid membranes) and be relieved by exposure light (Figure 3) through an unidentified electron outlet that supports O2 evolution. This restriction helps coordinate the production of low potential electrons with the C assimilatory capacity of the cells and would be especially important for protection of PSI as C fixation saturates. Furthermore, its occurrence in the dark, which would allow N-deprived cells unable to perform starch synthesis to limit PSI damage when the cells become exposed to the light. However, preventing electron flow beyond Cyt b6f would increase the reduction state of the PQ pool and possibly cause PSII photodamage. Photosynthetic organisms have evolved mechanisms to rapidly repair PSII and can therefore sustain the 'cost' of PSII photodamage while maintaining the benefit of PSI photoprotection. PTOX may also limit PSII damage when PET is restricted; it would reduce PSII photoinhibition by oxidizing a proportion of the PQ pool while also enabling the formation of a H + gradient that may facilitate NPQ and PSII repair. When the cells are once again able to transport photosynthetic electrons (through alternative electron outlets generated when they are kept in the light or when environmental limitations are altered), the restriction is relieved, and electron transport across the Cyt b6f complex can proceed.

Strains:
The sta6 mutant (CC-4348; sta6 -; rbo1 -) (Zabawinski et al., 2001) and the genetically complemented sta6::STA6 strain, designated C6 (CC-4567; rbo1 -) (Li et al., 2010) were purchased from the Chlamydomonas Resource Center and maintained under low light on solid Tris-Acetate-Phosphate (TAP) medium. The parental strain for sta6 and C6 requires arginine for growth and therefore could not be used with our current experimental design.
Chl concentration: Chl concentrations were determined from methanol extracted pigments according to (Porra et al., 1989). Briefly, cells were pelleted by centrifugation (13,000 x g, 2 min, 21°C) and resuspended in 100% methanol. Cell debris from methanol-extracted samples was pelleted (13,000 x g, 5 min, 21°C) and the supernatant containing the pigments was transferred to plastic cuvettes. Total Chl (in µg mL -1 ) was calculated according to the equation: Chl = 22.12 x A652 + 2.71 x A665, where A652 and A665 are absorbances at 652 and 665 nm, respectively.

O2 gas exchange:
To measure the rates of dark respiration following exposure to different light intensities, the light curves were determined using a Pt-Ag/AgCl polarographic electrode system (ALGI, Littleton, CO, USA) equipped with a temperature-controlled, 1 mL glass water-jacketed reaction chamber, and two YSI 5331A electrodes (Yellow Springs Instruments, Yellow Springs, OH, USA) polarized at -0.8 V. The assay contained 1.5 mL of actively growing cells which were purged with 1% CO2/99% He and supplemented with 12 µL of 0.5 M potassium bicarbonate in 5 mM TRIS (pH 7.5). After injection into the reaction chamber, the samples were exposed to stepped light intensities (Luxeon III Star, Lumileds, San Jose, CA, USA) with intensity increases at 3 min intervals followed by 3 min of darkness. The intensities used were 40, 80, 160, 320, 640, 1280, and 1600 µmol photons m -2 s -1 .
Chl fluorescence analysis: Chl fluorescence was measured using a DUAL-PAM-100 fluorometer (Walz, Germany). For monitoring Chl fluorescence at different light intensities (i.e., a light curve), the cells were supplemented with 2 mM NaHCO3 and exposed to increasing light intensities for 1 min at each intensity. The maximal quantum efficiency (Fv/Fm) and actual quantum efficiency of PSII (ΦPSII) as well as photochemical quenching, qL (and 1-qL (Kramer et al., 2004)), which estimate the redox state of the PQ pool with the assumption that QA and QB are in equilibrium, were directly calculated from the light curves. For dark acclimation, the cells were vigorously shaken in the dark for 30 min before measuring fluorescence (see Figure 1C). O2 production and light-driven O2 photo-reduction: In vivo 16 O2 photoproduction rates and 18 O2 uptake rates were determined using a custom built membrane inlet mass spectrometer (MIMS) with a Pfeiffer quadrupole PrismaPlus QME-220 (Pfeiffer Vacuum, Germany) coupled with a Pt-Ag/AgCl polarographic electrode system (ALGI, Littleton, CO, USA) equipped with a temperature-controlled, water-jacketed 1 mL glass cell, a YSI 5331A electrode (Yellow Springs Instruments, Yellow Springs, OH, USA) polarized at -0.8 V, and an atmospheric barometric pressure sensor (Infineon Technologies Americas Corp, El Segundo, CA, USA). For performing MIMS assays, 1.5 mL of actively growing cells were purged with 1% CO2/99% He that was spiked with 0.5 mL 99% 18 O2 and supplemented with 4 mM potassium bicarbonate suspended in fresh medium. After injection of the sample into the reaction chamber, a light (Luxeon III Star, Lumileds, San Jose, CA, USA) regime was applied as follows: 5 min dark, 6 min light (300 µmol photons m -2 s -1 ) and then 3 min of dark. The electrode and MIMS were calibrated before each measurement using atmospherically equilibrated medium and 1% CO2/99% He purged medium that was spiked with 0.5 mL 99% 18 O2. The light intensity from the LED was calibrated using a Walz US-SQS/L spherical micro quantum sensor (Walz, Germany). Gross O2 production and net uptake were calculated according to the following equations (Beckmann et al., 2009 ΔNet O2 production = ΔGross O2 production -ΔGross O2 uptake To inhibit alternative oxidases, samples were supplemented with 2 mM n-propyl gallate (Sigma) suspended in ethanol. The inhibitor was added just prior to injecting the sample into the glassreaction vessel.
Cyt f oxidation/reduction kinetics: Laser flash-mediated absorbance changes associated with Cyt f oxidation/reduction were monitored using a JTS10 spectrophotometer (Biologic, France) at 554 nm corrected against a base line drawn between the absorption at 546 and 573 nm (de Vitry et al., 2004). N-deprived cells were pelleted by centrifugation (1,600 x g, 2 min, 21°C) and re-suspended to a concentration of 40 µg Chl mL -1 in fresh medium supplemented with 10% ficoll (Sigma) and 2 mM NaHCO3 (Sigma). Cells were dark acclimated for 30 min before the measurements. For light acclimation, dark-acclimated cells were exposed to 100 µmol photons m -2 s -1 for the time periods given in the main text. After the acclimation period, the cells were placed in a cuvette in the dark and then oxidation of Cyt f was induced by a single turnover laser flash. Absorbance data were normalized to the "a" phase values measured by a laser flash induced electrochromic shift at 520 nm. The half times (T1/2) were calculated by fitting the re-reduction phase of the Cyt f kinetics curve to a first order decay equation using Origin 2019.
Oxidation/reduction of P700: Absorbance changes associated with P700 oxidation/reduction were monitored using a JTS10 spectrophotometer (Biologic, France) at 705 nm and corrected using the absorbance at 740 nm (i.e. P700 oxidation/reduction = ΔI/I 705 nm -ΔI/I 740 nm). Cells in logarithmic growth were pelleted by centrifugation (1,600 x g, 2 min, 21°C) and resuspended to a concentration of 25-30 µg Chl mL -1 in fresh medium. P700 oxidation/reduction was monitored over 10 s while the cells were exposed to 60 µmol photons m -2 s -1 , which was followed by a saturating light pulse and a short dark period (as indicated in Figure 5A). For dark acclimation, cells were shaken vigorously in darkness for 30 min. For light acclimation, dark-acclimated cells were exposed to 100 µmol photons m -2 s -1 for 1 h.
Data analysis: Data were analyzed (statistics, data filtering, and curve fitting) using Origin 2018 Pro and 2019 Pro.