Proton gradient regulation 5 (PGR5) is involved in the control of photosynthetic electron transfer, but its mechanistic role is not yet clear. Several models have been proposed to explain phenotypes such as a diminished steady-state proton motive force (pmf) and increased photodamage of photosystem I (PSI). Playing a regulatory role in cyclic electron flow (CEF) around PSI, PGR5 contributes indirectly to PSI protection by enhancing photosynthetic control, which is a pH-dependent down-regulation of electron transfer at the cytochrome b6f complex (b6f). Here, we re-evaluated the role of PGR5 in the green alga Chlamydomonas reinhardtii and conclude that pgr5 possesses a dysfunctional b6f. Our data indicate that the b6f low-potential chain redox activity likely operated in two distinct modes — via the canonical Q cycle during linear electron flow and via an alternative Q cycle during CEF, which allowed efficient oxidation of the low-potential chain in the WT b6f. A switch between the two Q cycle modes was dependent on PGR5 and relied on unknown stromal electron carrier(s), which were a general requirement for b6f activity. In CEF-favoring conditions, the electron transfer bottleneck in pgr5 was the b6f, in which insufficient low-potential chain redox tuning might account for the mutant pmf phenotype. By attributing a ferredoxin-plastoquinone reductase activity to the b6f and investigating a PGR5 cysteine mutant, a current model of CEF is challenged.
In linear electron flow (LEF), the two photosystems (PSII and PSI) act in series to ultimately reduce NADP+ via the enzyme ferredoxin (Fd)-NADP(H) oxidoreductase (FNR). The cytochrome b6f complex (b6f) functionally interconnects the two photosystems (reviewed in ), accepting electrons from plastoquinol (PQH2) and donating electrons to plastocyanin (PC). Functional b6f occurs as a homodimer, each monomer consisting of four major subunits (cytochrome b6, subunit-IV, cytochrome f (cyt.f) and the Rieske iron-sulfur protein (ISP)), as well as four minor subunits (PetG, L, M and N). In addition, each monomer includes six cofactors: two b-hemes (bl and bh), two c-hemes (cyt.f and ci), one chlorophyll a and one β-carotene. Light indirectly induces b6f turnover: Upon oxidation by the primary PSI electron donor P700, PC extracts one electron from cyt.f, which is re-reduced by the Rieske ISP. The positively charged Rieske FeS domain moves towards the lumenal Qo-site, where an electron flow bifurcation occurs: PQH2 donates one electron to the Rieske ISP (part of the high-potential chain with a midpoint potential Em = 300–350 mV) and a second electron to bl (low-potential chain; Em = −130 mV). PQ is re-reduced at the stromal Qi-site via bh (Em = −35 mV) and/or ci (Em = 100 mV, flexible as described below). Via the canonical Q cycle, the production of one PQH2 at Qi requires the oxidation of two PQH2 at Qo. The spatial proximity between bh and ci suggests electron sharing between the two and the presence of a membrane potential (ΔΨ) promotes the shared electron to rest on bhred/ciox . Furthermore, the presence of a ΔΨ is a general prerequisite for efficient b-heme oxidation  but it remains enigmatic by which mechanism the b6f senses the ΔΨ. It is of note that heme-ci is unique since it lacks an amino acid axial ligand and thus might ligate with the semiquinone analog NQNO [4,5], which downshifted the heme-ci midpoint potential from 100 mV to ∼−150 mV . Furthermore, heme-ci was proposed to engage in a Qi-site gating function  by either ligating tightly with the phenyl group of F40 in subunit-IV in the oxidized stated (closed Qi-site), or, after transient heme-ci reduction, with (semi-)PQ. A recent cryo-EM structure of the spinach b6f complex contained the native PQ in proximity to heme-ci . Following the ligation-associated midpoint potential downshift of heme-ci , it is not clarified yet whether heme-ci reduces heme-bh or the quinone. Since not more than half of the b-heme population is reduced per Qo-site turnover in uninhibited complexes ([9–11] and references therein), the occurrence of blred/bhred is unlikely. By approaching blred/bhred, i.e. during a Qo-site turnover in the presence of bhred, the strongly reducing redox potential in the low-potential chain injects the first electron into the quinone-ciox ensemble . Thereby, heme-ci could force a quasi-concerted PQ reduction [5,12]. The deprotonation of PQH2 at Qo and the protonation of PQ at Qi couple electron transfer to proton translocation into the thylakoid lumen. The resulting transmembrane electrochemical proton gradient (pmf) fuels ATP synthesis via the chloroplast ATP-synthase. Besides LEF, which produces both NADPH2 and ATP, diverse auxiliary electron flow pathways, including cyclic electron flow (CEF) around PSI, contribute to the pmf and thereby equilibrate the NADPH2 to ATP output ratio of the light reactions (reviewed in ). In addition, the pmf plays an integral photoprotective role, since the chemical component (ΔpH) induces energy-dependent quenching (qE) and modulates the rate-limiting, pH-dependent oxidation of PQH2 at the Qo-site, which is termed photosynthetic control (, reviewed in ). Hence, CEF creates a regulatory feedback loop, linking the stromal redox poise to the efficiency of light harvesting and the rate of electron transfer.
PGR5 (proton gradient regulation 5) has been first identified in Arabidopsis thaliana as a component being involved in the regulation of the pmf via CEF . The corresponding knockout mutant in Chlamydomonas reinhardtii features multi-faceted phenotypes resembling its vascular plant counterpart [17,18]: The algal pgr5 fails to induce qE-dependent NPQ and is extremely susceptible to PSI photodamage in response to high light [19,20] as well as fluctuating illumination . These defects have been attributed to an impaired acidification of the thylakoid lumen due to compromised Fd-PQ reductase-dependent CEF and a resulting lack of photosynthetic control in response to enhanced stromal redox pressure [19,22]. However, the detailed mechanism of this CEF route is still elusive, as is the molecular role of PGR5. In the past, the association of FNR with the b6f [23–25] has been proposed to induce a switch from LEF to CEF: According to this model, FNR would tether reduced Fd in the vicinity of bh and ci, ultimately facilitating PQ reduction via a modified Q cycle that combines lumenal and stromal electrons [26–28]. Our previous work showed less stable binding of algal FNR to the thylakoid membrane in the absence of PGR5 , suggesting a structural or regulatory contribution of PGR5 to this CEF pathway by influencing the localization of FNR. By contributing to photosynthetic control and potentially providing the Fd-PQ reductase activity required for CEF, the b6f seems to be at the core of the phenotypes the absence of PGR5 produces. Therefore, we spectroscopically reinvestigated the impact of PGR5 on photosynthetic electron transfer in C. reinhardtii, with a focus on b6f functionality by measuring the behavior of the high- and low-potential chain as well as the electrogenic efficiency of the photosynthetic machinery. We provide evidence that during CEF, a Fd-assisted Q cycle is active which requires PGR5 for sustained b6f function in the light.
Materials and methods
Strains and cell cultures
As described previously , the C. reinhardtii WT strain t222+, pgr5 and a complemented line, termed C1, were used. The complemented C1 strain accumulated ∼75% of WT PGR5 levels . Cells were cultivated at 20 µmol photons m−2 s−1 on agar-supplemented tris-acetate-phosphate (TAP) plates . When growing cells for experiments, liquid tris-phosphate (TP) medium was devoid of acetate. Stirred cultures were grown at 10 µmol photons m−2 s−1 (16 h light/8 h dark) and were bubbled with sterile air at 25°C. Grown cultures were diluted ∼6-fold at least once after inoculation and grown to a density of ∼2 × 105 cells ml−1 before harvesting (5000 rpm, 5 min, 25°C). For experiments with PGR5-complemented lines that feature zeocin resistance, WT, pgr5 and complemented lines were grown in TAP in the same conditions as TP cells, but without air bubbling. One day before the experiments, cells were diluted in fresh TAP and 5 µg ml−1 zeocin was added to resistant cultures to drive PGR5 expression. Before the measurements, cells were resuspended at 20 µg chlorophyll ml−1 (determined as in ) in TP supplemented with 20% (w/v) Ficoll. Before transferring the samples to an open cuvette, cells were shaken vigorously in dim light. Figure 1 pictures the routine of sample handling and dark adaptation in open cuvettes involved regular mixing of the 2 ml oxic sample. For oxygen-deprived conditions in the dark, cells were supplemented with 50 mM glucose, 10 U glucose oxidase and 30 U catalase in the cuvette, and then overlaid with mineral oil for at least 30 min. Inhibition of mitochondrial respiration by oxygen deprivation results in a strongly reduced chloroplast stroma (reviewed in ). This redox poise is known to promote PGR5-dependent CEF in algae . Independent of PSII photochemistry, these illuminated cells will be referred to as anoxic. In some experiments throughout this study, 1 mM hydroxylamine (HA, from 1 M aqueous stock) and 10 µM 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU, from 10 mM ethanolic stock) were used as PSII inhibitors. Where indicated, oxic control samples were poised for at least 20 min with 10 mM methyl viologen (MV, added from 1 M aqueous stock) to abolish PSI acceptor side limitation and inhibit CEF.
Generation of PGR5 complemented lines using a bicistronic system
The PGR5 gene (Cre05.g242400) was amplified from genomic DNA extracts using forward (5′-GCCCCGAATTCATGCTGGCCTCCAAGCCCGTTGTTG) and reverse primer (5′-CTAGTCTAGATTAAGCCAGGAAGCCCAG), harboring the underlined EcoRI/XbaI restriction sites. The reverse primer was used in combination with a second forward primer (5′-GGACTTCTCCAAGCTGCTTGGC, underlined base alters codon 113 from Cys to Ser) to generate a 107 bp mega primer . The latter served to generate the PGR5C113S template, amplified with the EcoRI/XbaI primer pair above. The digested PGR5 fragments were introduced into a bicistronic expression vector [33,34] under the control of the PSAD promoter. In addition, the construct conferred zeocin resistance, since PGR5 expression was linked to ble via a skipping peptide FMDV2A. The DNA was introduced to the pgr5 nuclear genome by electroporation (25 µF, 1 kV). Transformants were pre-selected on TAP agar plates supplemented with 10 µg ml−1 zeocin. Picked colonies, putatively carrying the bicistronic PGR5 construct, were grown at 10 µmol photons m−2 s−1 and submitted to spot tests performed on TP agar plates.
Chlorophyll fluorescence analysis
Pre-selection of pgr5::PGR5 and pgr5::PGR5 C113S was done with colonies grown on TP plates after 24 h exposure to 200 µmol photons m−2 s−1 (Supplementary Figure S1), using a Maxi-Imaging PAM chlorophyll fluorometer (Walz, Germany). All other experiments were carried out in an LED-based spectrophotometer (JTS-10, BioLogic, France). The device was equipped with a Fluo_59 accessory in fluorescence mode with separate light sources to induce photosynthetic processes and measure the fluorescence yield . Regarding the latter, weak 10 µs pulses (LED435-03, Roithner, Austria) were used for excitation and were placed in the dark and 100 µs after cessation of the actinic light, respectively. The light-detecting photodiodes were protected from scattered actinic light and the measuring pulses by using appropriate 3 mm filters (reference diode: BG39; measuring diode: LPF650+RG665, Schott, Mainz, Germany). Note that the experimental routines in Figure 1 are not in the same order as they are referenced in the following text. After 30 min dark-adaptation (Figure 1 item a), Fo was measured in the dark and Fm was obtained by a 250 ms saturating pulse (520 nm LED, 5000 µmol photons m−2 s−1) to give Fv/Fm ((Fm − Fo)/Fm). The photochemical quantum yield of PSII, ΦPSII, was calculated as (Fm′ − Fs)/Fm′ by measuring maximal (Fm′) and steady-state (Fs) fluorescence in the light. Actinic light exposure (630 nm LEDs emitting ∼150 µmol photons m−2 s−1) was either a brief 10 s (Figure 1 item b) or 30 min to establish a steady-state regime (Figure 1 item d). For the latter, oxic samples were regularly resuspended in an open cuvette and oxygen-deprived cells were not mixed. PSII operation efficiency depends, in part, on the number of open centers. Two different parameters were calculated, aiming to describe the pool of oxidized QA (reviewed in ). The PSII efficiency factor qP was derived from (Fm′ − Fs)/(Fm′ − Fo′) in light-adapted samples . Fo′ was estimated from Fo/(Fv/Fm + Fo/Fm′) . qP is based on a puddle model, but when a lake model for light harvesting is assumed, the redox state of the QA pool in the light can be more accurately assessed by qL, which was derived from qP × Fo′/Fs . Therefore, qL is preferred over qP (reviewed in ), although controversy exists (reviewed in ). We included both parameters and high values of (1 − qL), or (1 − qP), approximate a high proportion of reduced QA− over total QA [41,42].
Time-resolved absorption spectroscopy
All time-resolved absorption measurements are expressed as ΔI/I and were carried out in the JTS-10 with light-adapted cells (630 nm LEDs emitting ∼150 µmol photons m−2 s−1, Figure 1 item d). Using white pulsed LED detection light, the electrochromic shift (ECS) signals were measured as the difference of the absorbance changes at 520 and 546 nm (ΔI/I520nm−546nm, respective interference filters FWHM: 20 nm). The light-detecting diodes were protected from scattered actinic light by 3 mm BG39 filters (Schott, Mainz, Germany). Illumination of the samples was interrupted by short dark intervals (250 µs) during which 10 µs detecting pulses were placed after 200 µs. The light intensity used to establish a steady-state regime was further described by the initial ECS slope during 2 ms when transitioning from darkness to ∼150 µmol photons m−2 s−1 red actinic light (Figure 1 item c). Assuming that the light excited both photosystems equally, the initial photochemical rates (kiPSII ∼ kiPSI) are expressed in charge separations PSI−1 s−1 . The unit of the slope arises from signal normalization (see next paragraph). A saturating pulse (22 ms of 630 nm LED, ∼3000 µmol photons m−2 s−1 resulting in kiPSII ∼ kiPSI ∼ 1700 charge separations PSI−1 s−1) was used to follow the kinetics of the ECS and the b6f redox reactions (Figure 1 item f). The pulse duration was timed to generate a stable ECS level in WT. Using appropriate interference filters (FWHM: 10 nm), the b6f redox reactions were monitored on the level of cyt.f (554 nm) and cytochrome b (563 nm) with a baseline drawn between 546 and 573 nm . At variance with the previous deconvolution , 554 nm signals were corrected with 0.23 × (563 nm–546 nm) for a spectral overlap with reduced cytochrome b, estimated from extinction coefficients . Only this correction abolished cyt.f signal offsets in reducing conditions, thus approximating the dark reference several tens of ms after a flash (Supplementary Figure S2). The kinetics of 520–573 nm signals were obtained from back and forth recordings. Four and nine technical replicates were averaged for each wavelength in the multiple and single turnover setups, respectively (Figure 1).
The functional PSI:PSII ratios under the growth conditions were obtained by comparing the ECS amplitudes that were produced after a saturating ∼6 ns laser flash (a-phase). The flashes were provided by a dye laser (DCM, exciton laser dye) pumped by a frequency doubled Nd:YAG laser (Minilite II, Continuum). The ECS signals developed in less than 1 µs, i.e. before the first detection point in the dark at 300 µs. The a-phase amplitude was measured in oxic samples before and after the addition of PSII inhibitors (Figure 1 items c,h). In the presence of HA and DCMU, PSII fails to promote flash-induced charge separations. Accordingly, since the a-phase corresponded to 1 charge separation PSI−1, it revealed the concentration of active PSI in the sample. At 20 µg chlorophyll ml−1 in the cuvette, ΔI/I520nm−546nma-phase amplitudes of ∼900 to ∼1100 were typically observed (used to normalize all ΔI/I signals). In the absence of PSII inhibitors, ΔI/I520nm−546nma-phase amplitudes of ∼2000 were produced in oxic samples.
The apparent b-phase of single turnover kinetics (∼10 ms of electrogenic b6f contribution to the ECS signal; Figure 1 item i) was deconvoluted to obtain kΔΨ by subtracting the c-phase (ATP-synthase activity that results in ECS decay), which followed a first-order exponential decay [45,46]. To avoid double turnover, subsaturating laser flashes were used in this type of experiments, which generated an a-phase of ∼0.35–0.45 charge separations PSI−1. Like the b- and c-phase, the dark relaxation kinetics of the b6f were fitted with the mono-exponential decay function ExpDec1 of the OriginPro software. For kf-red and kb-ox, the 600 ms phase was used for fitting. However, in multiple turnover experiments (Figure 1 item f), a shorter time window for anoxic kb-ox calculation was necessary for a seamless transition to the fit of kb-red up to 9 s dark.
The overall membrane potential formation rates in saturating light (Figure 1 item f) were determined via a dark pulse-based protocol (, reviewed in ), which yielded a similar parameter as νH+ developed by Kramer et al. . Accordingly, the ECS signals were continuously recorded when changing the light intensities (from weak steady-state light to saturating light and from there to darkness). Therefore, the light-dependent photochemical rates were instantaneously altered, whereas the turnover rates of the ATP-synthase and the b6f stayed initially unchanged. Linear slopes were calculated during the initial 2 ms of transitioning from steady state to saturating light (Sini), which reflected the initial electrogenic efficiency in light-saturating conditions (kini, obtained in light-adapted samples at variance with kiPSII ∼ kiPSI above). In the light-adapted state, the initial electrogenic efficiency depended on the number of electrons in the high-potential chain that was immediately available . These electron pools were exhausted during the 22 ms saturating light pulse, i.e. the absence of acceptors (oxidized PQ or oxidized Fd) or donors (reduced P700) was promoted. The linear slopes at the end of the pulse (SL; final 6 ms) and in subsequent darkness (SD) were calculated and yielded the effective electron transfer after several turnovers in saturating light (kend = SL − SD). The initial dark signals were disregarded since they may include electrogenic PSI charge recombinations , and linear SD was obtained from 2 to 16 ms darkness.
As described previously , P700 redox changes (Figure 1 item e) were measured with short-pulsed detection LEDs, which peaked at 700 nm and were used in combination with interference filters at 705 and 740 nm (FWHM: 10 nm and 15 nm, respectively). The light-detecting photodiodes were protected from scattered actinic light by a 3 mm RG695 filter (Schott, Mainz, Germany). Illumination of the samples was interrupted by short dark intervals (250 µs) during which 10 µs detecting pulses were placed after 200 µs. The presented P700 kinetics originated from light-adapted cells which had a functional PSII. The steady-state kinetics were recorded in the light (Ps) and during a saturating 12 ms pulse, using a 630 nm LED ring (same as above, kiPSII ∼ kiPSI ∼ 1700 charge separations PSI−1 s−1). The pulse served to drive a multiple turnover process which increased the efficiency of photosynthetic control and emptied the immediate donor pools of reduced PC and cytochrome f . Thus, the strong pulse yielded in each condition the maximal P700 oxidation state (Pm), producing a lower ΔI/I705nm−740nm than Ps. By fitting the mono-exponential decay kinetics during the 12 ms pulse (ExpDec1 function of the OriginPro software), the P700 oxidation rate kP-ox was obtained. Prolonging the pulse did not change Pm (not shown). As a reference (Figure 1 item h), PSII inhibitors HA and DCMU were added in light-adapted samples to obtain the maximal P700 oxidation during the saturating pulse (P′m, kinetics not shown). After the 12 ms pulse, dark recovery of P700 was recorded for several seconds and the dark kinetics were corrected for a linear drift that developed in steady-state light-adapted cells, especially in anoxic conditions (Supplementary Figure S3). For each kinetics, the ΔI/I of the P700 redox state in darkness served as zero baseline (P0, only detecting light present). P0 was calculated from the offset of a two-component exponential decay function (OriginPro software, using dark kinetics from 4 to 2550 ms). The first decay rate of this function, k1P-red, describes the fast component of the P700 dark relaxation. Using the P0-corrected values, the yield of PSI (ΦPSI) is defined as (Pm − Ps)/P′m . The PSI acceptor side limitation (YNA, non-photo-oxidizable P700) is defined as (P′m − Pm)/P′m and the PSI donor side limitation (YND, photo-oxidized P700 in steady-state light) is defined as Ps/P′m.
When assessing the function of PGR5 in photosynthetic electron transfer, PSI photodamage has to be anticipated in pgr5 under high to moderate, as well as fluctuating light conditions [19–21]. To avoid such alterations of functional PSI, we conducted experiments at low light conditions under a photo-autotrophic growth regime. In fact, the weak irradiance during growth and measurements provided permissive conditions for the original pgr5 phenotype by avoiding photodamage [19–21]. To re-evaluate the role of PGR5 in electron transfer regulation, we combined several in vivo measurement protocols to assess PGR5-dependent electron transfer under these conditions in C. reinhardtii. For this dataset, we compared the pgr5 strain with WT, and furthermore investigated a partially PGR5-rescued strain C1 . To determine the functional PSI:PSII ratio, the amplitudes of the ECS signals were calculated after a laser flash. Thereby, we obtained a PSI:PSII ratio of 1.13 ± 0.13, 1.18 ± 0.13 and 1.24 ± 0.18 in WT, pgr5 and complemented C1, respectively (N = 3 ± SD). Another indication that permissive conditions were found was provided by the initial photochemical rates (kiPSII ∼ kiPSI) of 216 ± 23, 193 ± 11 and 185 ± 12 charge separations PSI−1 s−1 (N = 3 ± SD) in WT, pgr5 and C1, respectively. In the following sections, we will present redox and electrogenic parameters obtained with cultures that were comparable on the levels of the PSI:PSII ratio and kiPSII ∼ kiPSI.
Absence of PGR5 facilitates PSI oxidation in Chlamydomonas
The P700 redox state was analyzed in steady-state light-adapted cells. The resulting kinetics are shown in Figures 2A,B for WT and pgr5, respectively. The C1 line is shown in Supplementary Figure S4A. At a first glance, the kinetics were very similar, with exception of a strong pulse-induced P700 oxidation signal in anoxic pgr5. To further deconvolute the signals, the corresponding P700 redox states are shown in Figure 2C (see Supplementary Figure S4B for C1). It became evident that the yield of PSI (ΦPSI) was comparable in oxic conditions, also in the presence of methyl viologen (MV). Due to the artificial electron acceptor, the MV treated samples were not limited on the PSI acceptor side (YNA) and showed larger ΦPSI as well as larger donor side limitation (YND). The anoxic WT exhibited an increased YNA, unlike pgr5. Largely owing to YNA, the anoxic WT showed a significant decrease in YND compared with oxic samples (Student's t-test, P = 0.03). On the contrary, YND increased in anoxic pgr5 (P = 0.02).
The electron transfer via P700 is changed in
pgr5 under different cellular redox states, pointing to a more photo-oxidizable PSI.
When comparing the black traces in panels A and B of Figure 2, P700 was oxidized faster during the saturating light pulse in oxic pgr5, since kP-ox was significantly larger than in WT (Figure 2D, see Materials and methods). The oxic C1 sample was intermediary and kP-ox remained similar in the presence of MV, like in pgr5. In WT, kP-ox was faster after MV addition. As expected, kP-ox was increased in all strains due to an increased PSI antenna size in anoxic conditions. After increasing photosynthetic control efficiency during the pulse and thus emptying PSI electron donors, the P700 dark relaxation rate k1P-red was slower in anoxic conditions (Figure 2E, see Materials and methods). An insignificant slowdown was seen in anoxic pgr5 and MV addition had an inhibiting effect on k1P-red in all strains.
b6 f is subjected to enhanced oxidation by PSI in oxic pgr5 and becomes strongly impaired in anoxic pgr5
The b6f redox behavior of the samples in Figure 2 is presented in Figure 3. The kinetics in the high- and low-potential chain will be described first in WT and pgr5 (for C1 see Supplementary Figure S5), followed by calculations of the apparent dark relaxation rates. At variance with the representations in Figure 2, the steady-state signals before the strong pulse were the reference levels. It is of note that the b6f signals, although small in amplitude, displayed specific redox information, since the signals were absent in b6f-lacking mutants (Supplementary Figure S6). When oxic WT in the steady-state light was exposed to the saturating ms-pulse, a net cyt.f oxidation of ∼−0.4 a.u. was observed (hatched box in Figure 3A). In the dark, after the pulse, the fast cyt.f net reduction phase had an amplitude of ∼+0.65 a.u. and finished in ∼50 ms. Both amplitudes of cyt.f net oxidation (−0.2 a.u.) and reduction phase (+0.4 a.u.) were smaller in anoxic samples, compared with oxic conditions. The time during the pulse to reach maximal cyt.f oxidation was shorter compared with oxic cells, which was expected after showing lower ΦPSI (Figure 2C) and thus less injection of positive charges into the high-potential chain. The cyt.f re-reduction kinetics in darkness were faster in these samples as well (Figure 3A). When MV was present, there was almost no cyt.f net oxidation during the pulse. No distinct fast cyt.f reduction phase was observed during 100 ms of darkness. When monitoring redox changes of the hemes bl/bh during the saturating ms-pulse in oxic WT samples (Figure 3B), a net reduction of ∼+0.1 a.u. was observed. The hemes bl/bh net oxidation in the dark was finished within ∼50 ms after the saturating pulse in oxic WT samples. The redox state of hemes bl/bh was slightly more oxidized for several seconds darkness compared with the steady state before the pulse. The net reduction amplitude of the hemes bl/bh during the ms-pulse was slightly more pronounced in the presence of MV. After the pulse, however, there was a significantly larger amplitude of hemes bl/bh net oxidation in the MV treated samples, which transiently reached −0.2 a.u. compared with the steady-state reference. The oxidation was finished after ∼300 ms of darkness. In anoxic samples during the ms-pulse, the hemes bl/bh net reduction amplitude reached a slightly lower plateau earlier. In darkness, the hemes bl/bh net oxidation amplitude was small and the phase finished within ∼25 ms. In contrast with oxic samples, a unique hemes bl/bh redox feature was a net reduction phase, that had started by 100 ms of darkness in anoxic WT.
The electron transfer via
b6 f is changed under different cellular redox states, pointing to a more oxidized high-potential chain in pgr5, especially in anoxic cells, and to a b6 f inhibition in the low-potential chain in the absence of cyclic electron flow.
The net cyt.f oxidation amplitude during the pulse was smaller in oxic pgr5 (−0.2 a.u., Figure 3C), and thus half as large as in WT (cf. Figure 3A). In this sample, the cyt.f reduction phase amplitude was slightly larger (+0.8 a.u.) but the decay kinetics resembled WT. Whereas MV treated samples were indistinguishable from WT, cyt.f redox changes in anoxic pgr5 differed in several aspects. Unlike anoxic WT, fast cyt.f reduction kinetics were absent after the pulse in anoxic pgr5. The net oxidation amplitude was almost non-existent in these cells. As expected from the elevated ΦPSI and YND, which indicated the amount of P700+ before darkness in Figure 2C, the cyt.f reduction amplitude in anoxic pgr5 was indistinguishable from oxic cells (Figure 3C) and contrasted with the difference observed in WT (cf. Figure 3A). The hemes bl/bh redox signals during and after the saturating ms-pulse were like WT with two exceptions in anoxic pgr5 (Figure 3D): The hemes bl/bh oxidation phase after the pulse finished after ∼25 ms, i.e. later than in WT (cf. Figures 3B,D). Moreover, the onset of re-reduction was significantly delayed.
To quantitate the observations after the pulse, the apparent net reduction rate of cyt.f was calculated (kf-red, Figure 3E). With kf-red between ∼40 and ∼45 s−1, oxic samples were comparable and significantly slowed down by ∼75% in the presence of MV. Compared with oxic samples, kf-red was increased by a factor of ∼2 in anoxic WT cells. On the other hand, kf-red was lowered by a factor of ∼2 and ∼8 in anoxic C1 and pgr5, respectively. After the pulse, the apparent net oxidation kinetics in the low-potential chain are expressed as kb-ox (Figure 3F). In WT and C1, kb-ox was ∼50 s−1 and was significantly lowered upon MV addition. Oxic pgr5 controls trended towards lower kb-ox compared with WT and C1, and the inhibitory effect of MV was statistically not significant. The anoxic WT displayed higher kb-ox, whereas the pgr5 mutant and the partially complemented C1 strain did not show this effect. Yet, the C1 line was less severely affected than pgr5. Figure 3G shows the slow dark-reduction rates of hemes bl/bh in anoxia, kb-red. The WT rate was significantly faster than C1 and pgr5.
Using a multiple turnover protocol, it is reasonable to assume that the presented rates also strongly depended on the oxidation level of the PC pool as well as the PQ pool redox state. Regarding the former, the P700 parameters in Figure 2 served as a satisfactory proxy, since PC equilibrates with P700. The next section will present various chlorophyll fluorescence parameters that reflect PSII photochemistry, and to some extent the PQ pool redox state.
The enhanced PSII photochemistry in oxic
pgr5 stands in stark contrast with the anoxic mutant phenotype
During the workflow presented in Figure 1, samples have also been examined for their chlorophyll fluorescence yields. After 30 min dark adaptation, the maximum quantum efficiency, Fv/Fm, was comparable in all strains (Supplementary Table S1). Figure 4A shows the photochemical quantum yield of PSII (ΦPSII) in WT and pgr5, which was significantly lower in anoxic samples as expected. ΦPSII was determined after a short 10 s period of light acclimation as well as in the steady state after 30 min (the parameters for C1 are summarized in Supplementary Figure S7). Only oxic pgr5 showed a significant increase of ΦPSII in the steady state, which was lowered to WT-levels upon MV addition. Figure 4B shows a similar picture of the PSII efficiency factor (qP), yielding higher values in oxic steady-state pgr5. Moreover, qP inhibition in the presence of MV was observed in oxic algae, which has been reported for vascular plants previously . Figure 4B also shows that, in contrast with oxic conditions, the steady-state qP in anoxic WT was higher than in pgr5. Like the closely related qP, the fraction of open PSII centers (qL) in Figure 4C revealed the same phenotypes of 10 s vs. 30 min light adaptation. Furthermore, qL increased significantly only in fully light-acclimated anoxic WT cells.
The electron transfer via PSII is changed under different cellular redox states, pointing to higher PSII efficiency in oxic
pgr5 and to a lower efficiency in reducing conditions.
The initial electrogenic efficiency of the light-adapted photosynthetic chain in Chlamydomonas depends on PGR5
The data in Figures 2–4 display alterations in the mutant electron transfer chain. In a similar fashion to the P700 measurements, in which a superimposed saturating light pulse emptied the immediate donor pool of PSI during several turnovers, we analyzed the charge separation efficiency of the photosynthetic apparatus by recording the ECS. The results are shown in Figure 5 (see Supplementary Figure S8 for C1). The ECS, which serves as intrinsic voltmeter, was recorded in the background light (as reference), during and after the saturating pulse. In the steady state, before the short pulse, the slope of the signal was zero. The additional membrane potential (ΔΨ) that was built up at the very onset of the 22 ms pulse depended on the immediate availability of electrons in the photosynthetic chain, and thus the photochemical yield of both photosystems (which we determined above). The initial rate of this process, kini (see Materials and methods), was calculated from the data shown in green symbols for WT (inset of Figure 5A) and pgr5 (inset of Figure 5B). Oxic samples produced ∼4 additional, stable charge separations during the pulse, also in the presence of MV. Anoxic cells generated less additional ΔΨ during the pulse in general, and WT generated more extra ΔΨ than pgr5. During several seconds of darkness while ATP-synthase was active, the ΔΨ collapsed to ∼−4 units in oxic and anoxic samples, and to ∼−8 units in MV treated samples. At the end of the pulse, a new steady state (zero slope) was established in oxic and anoxic WT only. The progressed ΔΨ production rate at the end of the pulse, kend (see Materials and methods), gave an indication of how electron transfer in the chain was diminished upon exhaustion of electron donors (reduced P700) or acceptors (oxidized PQ or oxidized Fd).
The electrogenic efficiency of the photosynthetic electron transfer chain is compromised in
pgr5 under anoxic conditions.
The kini values are shown in Figure 5C and indicate that oxic controls produced similar rates between ∼630 and ∼730 charge separations PSI−1 s−1. In the presence of MV, with exception of pgr5, kini was less efficient compared with the controls. Nonetheless, the MV trend existed in pgr5 as well, which showed a relatively low kini in oxic controls already. In anoxic samples, kini was compromised in all strains compared with oxic conditions. WT kini in anoxia showed the smallest decrease, followed by C1 and pgr5. As shown in Figure 5D, kend was diminished to a similar extent in oxic samples, and anoxic conditions lowered kend further in all strains.
The phenotype in Chlamydomonas
pgr5 is rescued by overexpressing PGR5 and PGR5Cys113Ser, respectively
It is of note that throughout the study, the partially complemented C1 line, which accumulates ∼75% of WT PGR5 levels under the control of its native promoter , resembled WT in oxic conditions, whereas it tended to partially perform like pgr5 in anoxia. Although P700 of WT and C1 behaved similarly in anoxic conditions (cf. Figure 2C and Supplementary Figure S4B), the pgr5 resemblance was most apparent on the levels of b-hemes oxidation (kb-ox in Figure 3F). For cyt.f reduction (kf-red in Figure 3E) and electrogenicity (kini in Figure 5C), both rates were significantly faster under anoxia as compared with pgr5, but still slower than WT. To eliminate PGR5 titration effects in anoxic C1, we generated independent PGR5-complemented lines which, besides the P700 redox behavior (Figure 6A), also produced WT-like cyt.f reduction rates after the pulse (kf-red in Figure 6B), as well as kb-ox (Figure 6C) and kini (Figure 6D). Figure 6 also includes a mutated version of PGR5, in which the sole Cys at position 113 was replaced with Ser. Interestingly, this mutation did not interfere with restoration of a WT-like phenotype when overexpressing PGR5Cys113Ser. The only exception was an intermediate kb-ox (Figure 6,C) which was significantly faster than pgr5, but not as fast as WT and the pgr5::PGR5 strain, respectively. We noted that hetero-phototrophic cells varied slightly in their rates, compared with photoautotrophic cells (cf. Figures 4–6). Although the pgr5 phenotype was more effectively recovered in the overexpression lines compared with C1, slow photoautotrophic growth at low irradiance was not feasible to maintain controlled zeocin levels. Therefore, we introduced the C1 strain with the native PGR5 promotor in the first part of the manuscript. In the following part, we will focus on single turnover b6f kinetics in WT and pgr5, since the complex appeared to underperform under anoxic conditions in the mutant.
Measurements of various redox parameters under anoxic conditions are shown, which demonstrate the recovery of the
pgr5 phenotype by overexpression of PGR5 and a Cys113Ser variant of the polypeptide.
Redox finetuning of the
b6 f low-potential chain is PGR5-dependent
To rule out possible dark redox equilibration artefacts (owing to different pre-oxidation levels in the light-adapted steady state), this section introduces single turnover measurements. Here, the light-adapted cells have been investigated in the absence of PSII photochemistry and upon a 30 s dark period (Figure 1 item i). This dark period ensured the reduction of primary and secondary PSI donors as well as pmf consumption, especially since the ΔpH governs photosynthetic control (, reviewed in ). Furthermore, as seen from the varying number of open PSII centers in Figure 4C, we intended to mitigate differences in the PQ pool redox state by inhibiting PSII activity and exerting a more homogeneous reducing pressure on the b6f samples. During the single b6f turnover, an electron hole is passed from the oxidized c-heme in cyt.f to the Rieske ISP which, after swapping back the FeS domain closer to the cytochrome b6 subunit, is reduced by PQH2 at the Qo-site. The latter is a bifurcated process that also reduces hemes bl/bh. When heme-bh receives an electron from the Qo-site, a ΔΨ is generated, which we monitored via ECS signals. Redox changes in the b6f (Figure 7A) and the corresponding ECS changes (Figure 7B) were assayed in oxic samples. The ECS kinetics in Figure 7 are relative and are composed of three phases (reviewed in [48,54]). The deconvolution is explained in Materials and methods. We observed no significant differences in the two strains regarding the decay rate of the c-phase, related to ATP synthesis (Supplementary Figure S10). We also measured the b6f redox kinetics and ECS in the presence of MV (Figures 7C,D) as well as in anoxic conditions (Figures 7E,F). On a time scale after injecting an electron hole into the b6f, cyt.f reduction (kf-red) preceded the electrogenic b-phase (kΔΨ). The last phase was the relatively slow oxidation of the hemes bl/bh (kb-ox). A statistical evaluation of kf-red (Figure 7G), kΔΨ (Figure 7H), and kb-ox (Figure 7I) is shown, as well as the amplitude of the b-phase relative to one charge separation per PSI (Figure 7J).
Redox kinetics and electrogenic signals reveal a PGR5-dependent low-potential chain tuning in anoxia as well as an inhibitory effect of methyl viologen on the single
b6 f turnover.
In oxic samples, whether MV was present or not, cyt.f oxidation was finished before the first record at 300 µs after the flash and resulted in an amplitude of ∼−0.1 units compared with the reference signal before the flash (circle symbols in the b6f redox kinetics panels of Figures 7A,C and E). In anoxic samples, oxidation of cyt.f was slowed down slightly, finishing between ∼1 and ∼2 ms after the flash. With exception of the MV treated samples, cyt.f reduction by the FeS domain was initiated at ∼1 ms, yielding similar kf-red values in oxic and anoxic conditions. When MV was added, kf-red was lowered significantly (5% and 9% residual rates of oxic WT and pgr5, respectively) and a delayed onset of reduction became apparent between ∼5 and ∼10 ms.
Before cyt.f was getting reduced in oxic and anoxic samples (during the first ms), net redox changes of the hemes bl/bh were very small (square symbols in b6f redox kinetics panels of Figures 7A,C and E). Only after the onset of cyt.f reduction, a net reduction of the hemes bl/bh became varyingly apparent. In the same timescale, the different hemes bl/bh net reduction amplitudes coincided with the electrogenic b-phase (square symbols in ECS kinetics panels of Figures 7B,D and F). The sequential reduction of cyt.f and hemes bl/bh was expected and was also observed in MV treated samples but there was a significant slowdown of the low-potential chain turnover. The signal amplitude of the hemes bl/bh net reduction was a function of Qo- (kf-red or kΔΨ as proxy) and Qi-site activity (kb-ox as proxy). Accordingly, the amplitude appeared larger in the presence of MV since Qo-site turnover was less slowed down than Qi-site activity. For instance, 23% and 26% residual kΔΨ were measured in oxic WT and pgr5 after adding MV (Figure 7H), compared with 10% and 28% residual kb-ox (Figure 7I). Since the inhibitory MV effect on kb-ox was less pronounced in pgr5, the hemes bl/bh net reduction amplitude was smaller during the first 10 ms after the flash in the mutant (Figure 7C).
When comparing the respective strains under oxic and anoxic conditions, the net reduction amplitudes in the hemes bl/bh redox kinetics differed during the first 10 ms (Figures 7A,E). Both anoxic strains showed a relatively small net reduction of the hemes bl/bh. As mentioned above, this amplitude was a function of Qo- and Qi-site activity. Since the Qo-site, i.e. kf-red (Figure 7G) and kΔΨ (Figure 7H), was not significantly different from oxic conditions in the respective strains, the small amplitude in anoxia was related to Qi-site events, where electrons exited the low-potential chain during the 10 ms phase.
On the other hand, kΔΨ differed between WT and the less efficient pgr5, although the hemes bl/bh net reduction amplitude was comparable. After injection of Qo-site electrons into the low-potential chain finished, kb-ox was faster in anoxic WT only (Figure 7I). The relative amplitude of the b-phase was between 50% and 85% of the a-phase and tended to be slightly higher in pgr5 (Figure 7J). The ΔΨ generated by one b6f turnover in our conditions was close to the values in earlier reports [55,56], which attributed similar fractions of one charge separation when measuring electron transfer ‘within’ the membrane bilayer from hemes bl to bh in the bc1 complex, as opposed to across the whole membrane.
PGR5 is an important regulator of photosynthetic electron transfer, however, its function has not been linked to the operation of the b6f. Our data indicate a dysfunctional b6f in the absence of PGR5 which is manifested in an impaired redox cross-talk between the b6f complex and PSI. The Q cycle of the b6f complex is modified in strongly reducing as compared with oxic conditions. We conclude that the b6f is inhibited when being disconnected from signals downstream of PSI in the absence of PGR5 or in the presence of artificial electron acceptors. Moreover, PGR5 is functionally involved in a modified Q cycle which has access to stromal electrons and operates in WT but less efficiently in pgr5. To facilitate interpretation of pgr5 performance in the steady-state measurements, we will first discuss the single turnover experiments. For simplicity of redox signal discussions, the b6f will be treated as a homogeneous population, although distinct subsets may exist in the sample, e.g. in close vicinity to PSI in CEF-supercomplexes [57–60]. As summarized below, five previous findings are the conceptual framework for the interpretation of the hemes bl/bh signal evolution shown in Figure 7. Moreover, Table 1 is a guide for Figure 8:
The light-induced ΔΨ produces bhred/ciox, which converts to bhox/cired in the dark .
Presence of a ΔΨ is crucial for Qi-site activity, i.e. oxidation of the b-hemes after a Qo-site turnover . How exactly ΔΨ is sensed in the b6f is not yet understood.
As reviewed elsewhere for the cytochrome bc1 complex [12,61], an intrinsic short-circuit-preventing process influences the cyt.f reduction rate by governing the interaction between the Rieske ISP FeS domain and cyt.f. Only after the b-hemes become oxidized, the reduced FeS domain will swap closer to cyt.f to release the ‘trapped’ high-potential chain electron.
To fill the Qi-site with substrate (PQ and/or H+), cired is transiently required. Besides having an effect on other side chains in the cavity , cired likely weakens the interaction with the moiety of F40 in subunit-IV . In fact, slight changes of the F40 aromatic ring position relative to the heme-ci plane were observed depending on the Qi-site occupation in spinach  and cyanobacterial b6f . Considering (i) and the possible gating mechanism, the Qi-site pocket may remain free after a flash and fill with substrate in the dark.
blred/bhred has not been observed in single turnover measurements ([9–11] and references therein). Accordingly, when semi-PQ reduces blox in the presence of bhred, simultaneous transitions of blox → blred and bhred → bhox occur, followed by electrogenic blox/bhred equilibration. The inability to accumulate electrons in the low-potential chain can be linked to the terminal electron acceptor heme-ci. The tight ligation of the heme-ci with PQ (or quinone analog inhibitors) is favored by ciox , and thus by ΔΨ (i). A reducing pressure on the b6f is required for PQH2 formation since, compared with the second step, tight ciox/PQ ligation requires more energy for the first reduction step and prevents semi-PQ accumulation [5,7].
As introduced and in agreement with the general view [1,63], the b6f displayed a canonical Q cycle in oxic conditions (Figure 8A,B, referring to Figure 7). Thus, the b6f low-potential chain harbors blox/bhox/cired after the first (Figure 8A), and blox/bhox/ciox after the second Qo-site turnover that is associated with PQH2 formation at Qi (Figure 8B). The (semi-)PQ in the Qi-site receives the electrons from cired and/or bhred in the presence of ΔΨ, probably in a concerted and closely spaced process. Since the redox signals re-equilibrated near the zero baseline in oxic samples, an electron that entered the low-potential chain at the Qo-site did not reside on either b-heme after the reactions ceased. Note that in the absence of PSII activity in the light the PQ pool is oxidized, so that equilibration of bhred during 30 s dark was unlikely.
Model summarizing the single turnover measurements.
|Sample .||bl/bh after 30 s D (ci at low ΔΨ) .||bl/bh after 10 ms b-phase (PQH2 formed at Qi) .||bl/bh after ∼100 ms (PQH2 formed at Qi) .|
|oxic||ox/ox (red)||ox/red (0 or 0.5)||ox/ox (0.5 or 1)|
|anoxic WT||ox/red (red)||ox/red (1)||ox/ox (2, Fd-assisted ci red)|
|anoxic pgr5||ox/red (red)||ox/red (1)||ox/ox (1, ΔΨ-assisted ci red)|
|Sample .||bl/bh after 30 s D (ci at low ΔΨ) .||bl/bh after 10 ms b-phase (PQH2 formed at Qi) .||bl/bh after ∼100 ms (PQH2 formed at Qi) .|
|oxic||ox/ox (red)||ox/red (0 or 0.5)||ox/ox (0.5 or 1)|
|anoxic WT||ox/red (red)||ox/red (1)||ox/ox (2, Fd-assisted ci red)|
|anoxic pgr5||ox/red (red)||ox/red (1)||ox/ox (1, ΔΨ-assisted ci red)|
ox: oxidized; red: reduced.
The b-heme re-equilibration near the zero baseline was also true for oxic samples in the presence of MV (not shown in Figure 8). However, according to (iii), the b6f inhibition by MV can be attributed to altered kb-ox. Before the laser flash was applied, the FeS domain of the Rieske ISP might rest in an unusual position more distant to cyt.f, thus producing slow kf-red. The observed b6f redox kinetics in the presence of MV showed attenuated resemblance to b6f samples treated with the Qi-site inhibitors MOA-stilbene  and NQNO .
This observation links the Qi-site functionality to a stromal redox poise, which we also examined in the anoxic samples by creating a reduced stroma. Here, the major finding concerns the kb-ox tuning (Figures 7E,I) which was missing in anoxic pgr5 (Figure 8C) and produced high rates in WT (Figure 8D). As expected, the b-heme redox re-equilibration below the zero reference in the anoxic samples can be attributed to the accumulation of electrons in the PQ pool (reviewed in ) and the generation of bhred within several seconds in the dark [65,66]. Thus, the b-heme redox state before the flash differed from oxic conditions by containing blox/bhred (left of dashed line in Figures 8C,D). According to (i) and (ii), equilibration of bhred in the dark is only possible in the presence of cired and the low ΔΨ precluded a Qi-site turnover. The establishment of blred, which requires several minutes darkness , depended on a flash-induced Qo-site turnover after the 30 s of darkness. According to (v), an electrogenic oxidation/reduction of heme-bh occurred within 10 ms after the flash. Meanwhile ((i) and (ii)), blox/bhred/cired converted to blox/bhred/ciox and yielded one PQH2 at the Qi-site in both strains.
The 10 ms phase entails a difference between WT and pgr5 (not shown in Figure 8) that arises from panels H (kΔΨ) and J (b-:a-phase ratio) of Figure 7. Further experiments need to clarify whether less efficient PQH2 formation at the pgr5 Qi-site was responsible for the slightly slower charge separation and higher b-phase amplitudes in the mutant. It could be that the distance between stromal H+ and the heme-ci is larger in pgr5 due to a modified protein interaction in the stromal b6f domain. This may slow down the b-heme redox rates (v), while increasing relative electrogenicity. Involvement of H+ movements in b6f electrogenicity has been discussed recently in the context of heme-ci protonation upon reduction .
The b-heme redox re-equilibration after the flash below the zero reference, i.e. formation of blox/bhox in the ∼100 ms range, was slow in pgr5 (right part in Figure 8C). According to (i), the consumption of ΔΨ was required which yielded a similar kb-ox as oxic samples (cf. right part in Figure 8A). On the other hand, the WT (right part in Figure 8D) showed a faster kb-ox in the same time range which strongly suggests a modified Qi-site turnover as a response to the stromal redox poise. Considering that thylakoid membranes isolated from WT retain more bound FNR than those from pgr5  and that FNR copurifies with the b6f [23–25], it could be an allosteric regulator of the b6f. Thus, in the presence of ΔΨ, b6f-associated FNR overcomes the energetic barrier (i) by using stromal electrons from Fd to produce cired (Em of Fd in Fd:FNR = ∼−500 mV ). Importantly, this modified Q-cycle yields a second PQH2 at the WT Qi-site (right part in Figure 8D), when inducing a single turnover of a strongly pre-reduced WT complex. In turn, such a modified Q-cycle would facilitate more efficient proton pumping into the lumen as compared with the canonical Q-cycle, which would be hampered in pgr5 especially under conditions where the modified Q-cycle is favored (see also below).
Switching to the Fd-assisted Q cycle might (in)directly rely on reduced Fd since FNR membrane recruitment is stimulated in anoxic conditions and requires functional b6f and PSI . Moreover, MV-treated thylakoid membranes do not retain bound FNR in the light . Considering the above-mentioned consequences of MV on the Qi-site functionality upon illumination, the stromal interface of the b6f might be modulated due to events downstream of PSI.
As evidenced by the large b-heme oxidation amplitudes after the pulse (Figures 3B,C), MV addition in the steady-state experiments also led to an electron backup in the b6f low-potential chain due to an underperforming Qi-site (Figure 9A). According to (iii) and in agreement with the low kf-red (Figure 3E), the elevated YND (Figure 2C) and lower k1P-red (Figure 2E), this eventually imposed a bottleneck on LEF. If one target of the MV effect is localized at the b6f, an altered pmf composition with a lower ΔpH component should be expected. At least the larger ECS decay amplitude in Figure 5 indicates a larger ΔΨ component that drives ATP synthesis after the pulse. Moreover, future pmf parsing studies might correlate ΔΨ-induced charge recombination in PSII  with altered chlorophyll fluorescence that we (Figure 4) and others  observed.
Model summarizing the multiple turnover measurements.
In agreement with previous studies [71,72], the chlorophyll fluorescence pattern in oxic pgr5 suggests that the mutant displayed an altered electron flow downstream of PSI (Figure 9B). Accordingly, the higher ΦPSII in light-adapted pgr5, which we (Figure 4A) and others  observed in oxic conditions, indicates that PSI oxidized the high-potential chain more efficiently between PQH2 and PC. This resulted in a stronger pre-oxidation of cyt.f in the steady state (Figure 3C) and a faster PSI oxidation during saturating light pulses (Figure 2D). Accordingly, higher LEF rates via an unthrottled PSI may be expected in the mutant, but not in the WT (Figure 9C). The higher LEF rates in oxic pgr5 are associated with a drain of electron carriers downstream of PSI, at the expense of a PQ pool reduction via CEF. This shortage of immediate electron donors in the photosynthetic chain might account for a lowered initial electrogenic efficiency in light-saturating conditions (Figure 5C). When the photon flux density is higher than in our permissive growth conditions [19,20], it is reasonable to link the PSI photodamage phenotype in pgr5 to a deregulation of events downstream of PSI. As suggested previously, the resulting lower PSI:b6f ratio creates a situation where more electrons are funneled through the remaining PSI centers, which lowers ΦPSI . We prevented this scenario in low light growth since algal mutants of the PGR5-dependent CEF pathway cope with the drained stromal electrons, up to a certain light intensity and time period, by increasing metabolic cooperation with mitochondria [21,73]. Algal photosynthesis features additional electron sinks like flavodiiron proteins, which are absent in angiosperms. Thus, in addition to the weak growth light, flavodiiron proteins might prevent PSI overreduction in oxic pgr5 as opposed to Arabidopsis . Accordingly, introducing flavodiiron proteins in the Arabidopsis pgr5 background alleviates the PSI phenotype .
When depriving oxygen, the substrate for mitochondrial respiration and flavodiiron proteins, the remaining acceptor for the drained stromal electrons is hydrogenase. In fact, the hydrogenase successfully competes for reduced Fd in anoxic pgr5 [75,76]. On the contrary, the ATP-depleted WT is engaged in CEF and thereby generates the pmf, in part, via the Fd-assisted Q cycle (Figure 9D). Thus, YNA is maintained in anoxic WT and the electrons are evenly distributed in the photosynthetic chain. The pgr5 fails to switch to a Fd-assisted Q cycle in anoxic conditions (Figure 9E). Since we did not observe a b6f phenotype in oxic pgr5 during steady-state experiments, the intrinsic Q cycle switch in anoxia could involve posttranslational modifications that impede a b6f operation as under oxic conditions. By failing to efficiently generate a ΔpH upon Qi-site tuning, the b6f in pgr5 did not underlie photosynthetic control imposed by a strong light pulse in the WT. This resulted in an insignificant slowdown of k1P-red in anoxia (Figure 2E). Moreover, probably as response to the reduced PQ pool, more PSII centers were closed (Figure 4C) and the electron donors downstream of the b6f were strongly oxidized (Figures 2C and 3C). According to (iii), the underperforming b6f was responsible for the redox pool imbalance between PQ and PC/P700. This strongly impaired the initial electrogenic efficiency in light-saturating conditions (Figure 5C). Moreover, the slow and delayed b-heme reduction phase in pgr5 (Figure 3G) might be indicative of the underperformance since it could be the sum of a coincident b-heme oxidation phase when the b6f relaxes in the dark (i). The small number of electrons that flow downstream of the b6f over time might promote the unbinding of FNR from PSI and/or b6f. As elaborated above, FNR binding to the membrane might rely on a critical Fd photoreduction rate and was affected in pgr5, thus favoring hydrogen production. An interesting side note of our in vivo results in algae is that they challenge the suggested role of the PGRL1-PGR5 complex, which is believed to be the Fd-PQ reductase in Arabidopsis, where PGR5 supports PGRL1 reduction . The authors demonstrated in vitro that PGR5 forms heterodimers with PGRL1. According to their non-reducing SDS–PAGE, several Cys in PGRL1 were crucial for the dimerization interface. Our results rule out that mixed disulfides play a role in the corresponding algal model. Moreover, they exclude that the only cysteine in PGR5 is involved in PGRL1 reduction or that PGRL1 redox state is relevant for the effects we observed.
Taken together, we propose that the b6f receives a stromal redox feedback from PSI and represents the Fd-PQ reductase that is switched on in ATP-depleted conditions. PGR5 is required for the activation and/or tuning of the Fd-assisted Q cycle.
The authors declare that there are no competing interests associated with the manuscript.
F.B. conceptualization, investigation, formal analysis, visualization, writing — original draft; L.M. formal analysis, writing — review and editing; P.G. resources, formal analysis; M.H. funding acquisition, conceptualization, formal analysis, writing — review and editing.
The b6f-deficient mutants were kindly provided by Sandrine Bujaldon (IBPC, Paris). M.H. acknowledges support from Deutsche Forschungsgemeinschaft (DFG) Grant HI 739/13-1 and Grant HI 739/13-2. M.H. also acknowledges support from the RECTOR program (University of Okayama, Japan).
cytochrome b6f complex
cyclic electron flow
iron sulfur protein
linear electron flow
proton gradient regulation 5
proton motive force