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.

Introduction

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 [1]), 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 [2]. Furthermore, the presence of a ΔΨ is a general prerequisite for efficient b-heme oxidation [3] 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 [6]. Furthermore, heme-ci was proposed to engage in a Qi-site gating function [7] 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 [8]. Following the ligation-associated midpoint potential downshift of heme-ci [6], 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 [7]. 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 [13]). 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 ([14], reviewed in [15]). 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 [16]. 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 [21]. 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 [20], 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 [19], 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 [19]. Cells were cultivated at 20 µmol photons m−2 s−1 on agar-supplemented tris-acetate-phosphate (TAP) plates [29]. 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 [30]) 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 [31]). This redox poise is known to promote PGR5-dependent CEF in algae [22]. 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.

Figure 1.

The experimental workflow of the steady-state protocol in items a to h is shown, as well as the single turnover experiments in i. The references are in parenthesis. AL, actinic light (∼150 µmol photons m−2 s−1); ECS, electrochromic shift; MV, methyl viologen; HA, hydroxylamine; DCMU, 3-(3,4-dichlorophenyl)-1,1-dimethylurea.

Figure 1.

The experimental workflow of the steady-state protocol in items a to h is shown, as well as the single turnover experiments in i. The references are in parenthesis. AL, actinic light (∼150 µmol photons m−2 s−1); ECS, electrochromic shift; MV, methyl viologen; HA, hydroxylamine; DCMU, 3-(3,4-dichlorophenyl)-1,1-dimethylurea.

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 [32]. 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 [35]. 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 [36]). The PSII efficiency factor qP was derived from (Fm′ − Fs)/(Fm′ − Fo′) in light-adapted samples [37]. Fo′ was estimated from Fo/(Fv/Fm + Fo/Fm′) [38]. 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 [39]. Therefore, qL is preferred over qP (reviewed in [36]), although controversy exists (reviewed in [40]). 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 [43]. 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 [44]. At variance with the previous deconvolution [44], 554 nm signals were corrected with 0.23 × (563 nm–546 nm) for a spectral overlap with reduced cytochrome b, estimated from extinction coefficients [6]. 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 ([47], reviewed in [48]), which yielded a similar parameter as νH+ developed by Kramer et al. [49]. 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 [50]. 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 [51], and linear SD was obtained from 2 to 16 ms darkness.

As described previously [52], 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 [50]. 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 [50]. 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.

Results

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 [19]. 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.

Figure 2.
The electron transfer via P700 is changed in pgr5 under different cellular redox states, pointing to a more photo-oxidizable PSI.

For (A) WT and (B) pgr5, saturating pulse-induced P700 redox changes are shown in the absence and presence of 10 mM methyl viologen (MV), as well as in anoxic cells. The 12 ms pulse (hatched red box) was applied on light-adapted cells in the steady state (red box), followed by several seconds dark measurements (black box). (C) As described in Materials and methods, the different P700 populations were deconvoluted as photo-oxidizable fraction (ΦPSI, yield of PSI), non-oxidizable P700 owing to acceptor side limitation (YNA), and pre-oxidized P700 due to donor side limitation (YND). The electron acceptor MV abolished YNA and increased YND despite PSII activity. In WT, anoxic conditions lowered ΦPSI and increased YNA. The latter effect was absent in anoxic pgr5. (D) As described in Materials and methods, the rate of P700 oxidation during the pulse, kP-ox and (E) the fast P700 dark relaxation rate after the pulse, k1P-red, were calculated from the kinetics in (A) and (B) for WT and pgr5, respectively (N = 3 ± SD; Student's t-test *P < 0.05 and **P < 0.005). The parameters for the partially complemented line C1 are shown as well (see Supplementary Figure S4 for P700 kinetics and redox states). kP-ox was faster in oxic pgr5. Only MV samples in the WT showed faster kP-ox. Anoxic cells also produced faster kP-ox than oxic controls. k1P-red was slowed down in MV samples. The anoxic WT and C1 also showed slower k1P-red.

Figure 2.
The electron transfer via P700 is changed in pgr5 under different cellular redox states, pointing to a more photo-oxidizable PSI.

For (A) WT and (B) pgr5, saturating pulse-induced P700 redox changes are shown in the absence and presence of 10 mM methyl viologen (MV), as well as in anoxic cells. The 12 ms pulse (hatched red box) was applied on light-adapted cells in the steady state (red box), followed by several seconds dark measurements (black box). (C) As described in Materials and methods, the different P700 populations were deconvoluted as photo-oxidizable fraction (ΦPSI, yield of PSI), non-oxidizable P700 owing to acceptor side limitation (YNA), and pre-oxidized P700 due to donor side limitation (YND). The electron acceptor MV abolished YNA and increased YND despite PSII activity. In WT, anoxic conditions lowered ΦPSI and increased YNA. The latter effect was absent in anoxic pgr5. (D) As described in Materials and methods, the rate of P700 oxidation during the pulse, kP-ox and (E) the fast P700 dark relaxation rate after the pulse, k1P-red, were calculated from the kinetics in (A) and (B) for WT and pgr5, respectively (N = 3 ± SD; Student's t-test *P < 0.05 and **P < 0.005). The parameters for the partially complemented line C1 are shown as well (see Supplementary Figure S4 for P700 kinetics and redox states). kP-ox was faster in oxic pgr5. Only MV samples in the WT showed faster kP-ox. Anoxic cells also produced faster kP-ox than oxic controls. k1P-red was slowed down in MV samples. The anoxic WT and C1 also showed slower k1P-red.

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.

The functional b6f 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 b6f is changed under different cellular redox states, pointing to a more oxidized high-potential chain in pgr5, especially in anoxic cells, and to a b6f inhibition in the low-potential chain in the absence of cyclic electron flow.

Figure 3.
The electron transfer via b6f is changed under different cellular redox states, pointing to a more oxidized high-potential chain in pgr5, especially in anoxic cells, and to a b6f inhibition in the low-potential chain in the absence of cyclic electron flow.

(A) Cyt.f redox changes in a steady-state light/pulse/dark regime showed different pulse-induced oxidation amplitudes in WT cells with oxic > anoxic > oxic + MV, compared with the steady-state reference. The addition of the cyclic electron flow inhibitor methyl viologen (MV, 10 mM) caused no significant cyt.f net oxidation during the pulse. Most of the dark relaxation finished in ∼50 ms, except for the slow re-reduction in MV samples. (B) Redox changes of b-hemes in WT were comparable during the pulse and most of the oxidation was finished in ∼50-ms, ∼300-ms, ∼25-ms in oxic, oxic + MV and anoxic samples, respectively. The latter samples showed a slow re-reduction phase with an onset at less than 100 ms darkness. (C) In pgr5, cyt.f showed different pulse-induced oxidation amplitudes with oxic > anoxic = oxic + MV, compared with the steady-state reference. Oxidation amplitudes in oxic controls were smaller than in WT (cf. A). Mutant kinetics in the presence of MV and in anoxia were similarly slowed down. (D) Redox changes of b-hemes in pgr5 were comparable during the pulse and showed a similar trend as WT, except for a slightly longer oxidation phase after the pulse in anoxia (cf. B). Anoxic pgr5 also showed a re-reduction phase which started later and was slower than in WT. (E) After the pulse in (A) and (C), fitting the cyt.f reduction phase yielded a kf-red which was slowed down by MV (N = 3 ± SD; Student's t-test *P < 0.05 and **P < 0.005, see Materials and methods). Anoxic WT showed a faster kf-red, whereas it was much slower in anoxic pgr5, followed by C1 (for intermediary C1, see Supplementary Figure S5). (F) Following the pulse, b-heme oxidation rates were calculated as kb-ox which was slowed down by MV in WT and C1. In anoxic WT, kb-ox was faster compared with the oxic control and the other anoxic strains. (G) The slow b-heme re-reduction rate in anoxia, kb-red, was fastest in WT.

Figure 3.
The electron transfer via b6f is changed under different cellular redox states, pointing to a more oxidized high-potential chain in pgr5, especially in anoxic cells, and to a b6f inhibition in the low-potential chain in the absence of cyclic electron flow.

(A) Cyt.f redox changes in a steady-state light/pulse/dark regime showed different pulse-induced oxidation amplitudes in WT cells with oxic > anoxic > oxic + MV, compared with the steady-state reference. The addition of the cyclic electron flow inhibitor methyl viologen (MV, 10 mM) caused no significant cyt.f net oxidation during the pulse. Most of the dark relaxation finished in ∼50 ms, except for the slow re-reduction in MV samples. (B) Redox changes of b-hemes in WT were comparable during the pulse and most of the oxidation was finished in ∼50-ms, ∼300-ms, ∼25-ms in oxic, oxic + MV and anoxic samples, respectively. The latter samples showed a slow re-reduction phase with an onset at less than 100 ms darkness. (C) In pgr5, cyt.f showed different pulse-induced oxidation amplitudes with oxic > anoxic = oxic + MV, compared with the steady-state reference. Oxidation amplitudes in oxic controls were smaller than in WT (cf. A). Mutant kinetics in the presence of MV and in anoxia were similarly slowed down. (D) Redox changes of b-hemes in pgr5 were comparable during the pulse and showed a similar trend as WT, except for a slightly longer oxidation phase after the pulse in anoxia (cf. B). Anoxic pgr5 also showed a re-reduction phase which started later and was slower than in WT. (E) After the pulse in (A) and (C), fitting the cyt.f reduction phase yielded a kf-red which was slowed down by MV (N = 3 ± SD; Student's t-test *P < 0.05 and **P < 0.005, see Materials and methods). Anoxic WT showed a faster kf-red, whereas it was much slower in anoxic pgr5, followed by C1 (for intermediary C1, see Supplementary Figure S5). (F) Following the pulse, b-heme oxidation rates were calculated as kb-ox which was slowed down by MV in WT and C1. In anoxic WT, kb-ox was faster compared with the oxic control and the other anoxic strains. (G) The slow b-heme re-reduction rate in anoxia, kb-red, was fastest in WT.

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 [53]. 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.

Figure 4.
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.

Panels (AC) show the chlorophyll fluorescence-derived quantum yield of PSII (ΦPSII), the PSII efficiency factor (qP) and the fraction of open PSII centers (qL) for WT and pgr5 (N = 3 ± SD; Student's t-test *P < 0.05 and **P < 0.005). During light adaptation in oxic conditions, all parameters were unaltered in WT after 10 s illumination and in the steady-state after 30 min light. In oxic pgr5, all parameters were like in WT after 10 s light but increased significantly after 30 min light. In the presence of 10 mM methyl viologen (MV), all parameters in pgr5 were lowered significantly to levels as in the WT, in which MV lowered qP only. During light adaptation in anoxic conditions, all parameters were at the same low level after 10 s light. They remained low in anoxic pgr5 after 30 min illumination and qL was significantly changed in anoxic WT only. The light-adapted anoxic WT showed a significantly higher qP and qL than pgr5.

Figure 4.
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.

Panels (AC) show the chlorophyll fluorescence-derived quantum yield of PSII (ΦPSII), the PSII efficiency factor (qP) and the fraction of open PSII centers (qL) for WT and pgr5 (N = 3 ± SD; Student's t-test *P < 0.05 and **P < 0.005). During light adaptation in oxic conditions, all parameters were unaltered in WT after 10 s illumination and in the steady-state after 30 min light. In oxic pgr5, all parameters were like in WT after 10 s light but increased significantly after 30 min light. In the presence of 10 mM methyl viologen (MV), all parameters in pgr5 were lowered significantly to levels as in the WT, in which MV lowered qP only. During light adaptation in anoxic conditions, all parameters were at the same low level after 10 s light. They remained low in anoxic pgr5 after 30 min illumination and qL was significantly changed in anoxic WT only. The light-adapted anoxic WT showed a significantly higher qP and qL than pgr5.

The initial electrogenic efficiency of the light-adapted photosynthetic chain in Chlamydomonas depends on PGR5

The data in Figures 24 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.

Figure 5.
The electrogenic efficiency of the photosynthetic electron transfer chain is compromised in pgr5 under anoxic conditions.

The ability to generate and dissipate an electric field is assessed by measuring the electrochromic shift, ECS. Signals were recorded in steady-state light, during a saturating pulse and in darkness. (A) The ECS kinetics of the WT and (B) pgr5 indicated that the 22 ms pulse led to equilibration of a new membrane potential ΔΨ (for ECS kinetics of C1 refer to Supplementary Figure S8). The initial efficiency to generate a higher ΔΨ level at the onset of the pulse was calculated with data from the green symbols, yielding kini from the linear slope during the first 2 ms of the pulse (inset, see Materials and methods). The apparent ΔΨ generation efficiency at the end of the pulse (kend) was corrected with the ΔΨ-consuming activity of the ATP-synthase. (C) kini was similar in oxic strains and, except for pgr5, slower upon addition of 10 mM methyl viologen (MV; N = 3 ± SD; Student's t-test *P < 0.05,**P < 0.005, ***P < 0.0005 and ****P < 0.00005). In anoxic cells, kini was diminished in all strains but highest in WT, followed by C1 and pgr5. (D) kend was low in the strains due to exhaustion of electron acceptors or donors. Several small differences among treatments and strains were observed.

Figure 5.
The electrogenic efficiency of the photosynthetic electron transfer chain is compromised in pgr5 under anoxic conditions.

The ability to generate and dissipate an electric field is assessed by measuring the electrochromic shift, ECS. Signals were recorded in steady-state light, during a saturating pulse and in darkness. (A) The ECS kinetics of the WT and (B) pgr5 indicated that the 22 ms pulse led to equilibration of a new membrane potential ΔΨ (for ECS kinetics of C1 refer to Supplementary Figure S8). The initial efficiency to generate a higher ΔΨ level at the onset of the pulse was calculated with data from the green symbols, yielding kini from the linear slope during the first 2 ms of the pulse (inset, see Materials and methods). The apparent ΔΨ generation efficiency at the end of the pulse (kend) was corrected with the ΔΨ-consuming activity of the ATP-synthase. (C) kini was similar in oxic strains and, except for pgr5, slower upon addition of 10 mM methyl viologen (MV; N = 3 ± SD; Student's t-test *P < 0.05,**P < 0.005, ***P < 0.0005 and ****P < 0.00005). In anoxic cells, kini was diminished in all strains but highest in WT, followed by C1 and pgr5. (D) kend was low in the strains due to exhaustion of electron acceptors or donors. Several small differences among treatments and strains were observed.

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 [19], 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 46). 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.

Figure 6.
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.

(A) The P700 redox state in TAP-grown WT and pgr5 follows a similar trend as in anoxic samples of Figure 2C. Both overexpression lines had a WT-like P700 redox state. (B) After the pulse (for kinetics, see Supplementary Figure S9A), the reduction rates of cytochrome f (kf-red) were significantly slower in the mutant and were recovered to WT-level in both overexpressors (N = 3 ± SD; Student's t-test *P < 0.05 and ***P < 0.0005). (C) Simultaneously, the oxidation rates of the b-hemes after the pulse (kb-ox) were significantly slowed down in pgr5 and, to a lesser extent, in pgr5::PGR5 C113S (for kinetics, see Supplementary Figure S9B). (D) During the saturating pulse, the high initial electrogenic charge separation rates (kini) in steady-state cells were PGR5-dependent, and were also recovered in the C113S variant.

Figure 6.
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.

(A) The P700 redox state in TAP-grown WT and pgr5 follows a similar trend as in anoxic samples of Figure 2C. Both overexpression lines had a WT-like P700 redox state. (B) After the pulse (for kinetics, see Supplementary Figure S9A), the reduction rates of cytochrome f (kf-red) were significantly slower in the mutant and were recovered to WT-level in both overexpressors (N = 3 ± SD; Student's t-test *P < 0.05 and ***P < 0.0005). (C) Simultaneously, the oxidation rates of the b-hemes after the pulse (kb-ox) were significantly slowed down in pgr5 and, to a lesser extent, in pgr5::PGR5 C113S (for kinetics, see Supplementary Figure S9B). (D) During the saturating pulse, the high initial electrogenic charge separation rates (kini) in steady-state cells were PGR5-dependent, and were also recovered in the C113S variant.

Redox finetuning of the b6f 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 ([14], reviewed in [15]). 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 b6f turnover.

Figure 7.
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 b6f turnover.

(A) Cyt.f and b-heme signals are shown for oxic WT and pgr5. Cyt.f was rapidly (<300 µs) oxidized after the flash and re-reduced within 100 ms darkness (fitted curves). The b-heme net reduction lasted between ∼1 and 10 ms darkness, followed by a slower oxidation phase (fitted curves) for several tens of ms. (B) The corresponding oxic ECS kinetics were normalized to the signal produced by a flash hitting ∼40% of PSI centers (ΔI/I520-546 < 300 µs, a-phase). The following b-phase (fitted curve of squares) resulted from b6f-dependent charge separation activity in the ∼10 ms range. As described in Materials and methods, the b-phase was deconvoluted from raw ECS kinetics by subtracting the c-phase (fitted curve of circles). In this sample, the b-phase developed slower in pgr5 and showed a slightly larger relative amplitude. (C) The addition of 10 mM methyl viologen (MV) slowed down cyt.f reduction and b-heme oxidation in both strains, and the mutant was slightly less affected. The inhibition of b-heme oxidation allowed larger reduction amplitudes compared with (A). (D) Evolution of the b-phase was slowed down by MV but the amplitude was not altered. MV also slowed down the c-phase upon disulfide promotion in the ATP-synthase γ-subunit [78]. (E) The b6f redox kinetics in anoxia showed slightly slower cyt.f oxidation whereas reduction was like in oxic cells. Net reduction of b-hemes in the first 10 ms was of negligible amplitude and a large oxidation phase followed. (F) ECS and b-phase kinetics in anoxia resembled oxia (B). (G) Cyt.f reduction rates kf-red were calculated and showed a significant slowdown in the presence of MV (N = 3 ± SD; Student's t-test *P < 0.05, **P < 0.005 and ***P < 0.0005). (H) The electrogenic b-phase also evolved at slower rates (kΔΨ) in MV samples. Compared with WT, the pgr5 mutant showed slower kΔΨ in oxic and anoxic conditions. (I) After the varyingly apparent b-heme reduction phase, the slow oxidation rates were expressed as kb-ox. MV slowed down kb-ox and the WT showed faster kb-ox in anoxia, whereas the anoxic mutant kb-ox remained unchanged. (J) Compared with the amplitude of the a-phase, the mutant had a slight tendency to produce a larger b-phase, which was significant for pgr5 MV samples.

Figure 7.
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 b6f turnover.

(A) Cyt.f and b-heme signals are shown for oxic WT and pgr5. Cyt.f was rapidly (<300 µs) oxidized after the flash and re-reduced within 100 ms darkness (fitted curves). The b-heme net reduction lasted between ∼1 and 10 ms darkness, followed by a slower oxidation phase (fitted curves) for several tens of ms. (B) The corresponding oxic ECS kinetics were normalized to the signal produced by a flash hitting ∼40% of PSI centers (ΔI/I520-546 < 300 µs, a-phase). The following b-phase (fitted curve of squares) resulted from b6f-dependent charge separation activity in the ∼10 ms range. As described in Materials and methods, the b-phase was deconvoluted from raw ECS kinetics by subtracting the c-phase (fitted curve of circles). In this sample, the b-phase developed slower in pgr5 and showed a slightly larger relative amplitude. (C) The addition of 10 mM methyl viologen (MV) slowed down cyt.f reduction and b-heme oxidation in both strains, and the mutant was slightly less affected. The inhibition of b-heme oxidation allowed larger reduction amplitudes compared with (A). (D) Evolution of the b-phase was slowed down by MV but the amplitude was not altered. MV also slowed down the c-phase upon disulfide promotion in the ATP-synthase γ-subunit [78]. (E) The b6f redox kinetics in anoxia showed slightly slower cyt.f oxidation whereas reduction was like in oxic cells. Net reduction of b-hemes in the first 10 ms was of negligible amplitude and a large oxidation phase followed. (F) ECS and b-phase kinetics in anoxia resembled oxia (B). (G) Cyt.f reduction rates kf-red were calculated and showed a significant slowdown in the presence of MV (N = 3 ± SD; Student's t-test *P < 0.05, **P < 0.005 and ***P < 0.0005). (H) The electrogenic b-phase also evolved at slower rates (kΔΨ) in MV samples. Compared with WT, the pgr5 mutant showed slower kΔΨ in oxic and anoxic conditions. (I) After the varyingly apparent b-heme reduction phase, the slow oxidation rates were expressed as kb-ox. MV slowed down kb-ox and the WT showed faster kb-ox in anoxia, whereas the anoxic mutant kb-ox remained unchanged. (J) Compared with the amplitude of the a-phase, the mutant had a slight tendency to produce a larger b-phase, which was significant for pgr5 MV samples.

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.

Discussion

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 [2].

  • Presence of a ΔΨ is crucial for Qi-site activity, i.e. oxidation of the b-hemes after a Qo-site turnover [3]. 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 [8], cired likely weakens the interaction with the moiety of F40 in subunit-IV [7]. 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 [8] and cyanobacterial b6f [62]. 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 [6], 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.

Figure 8.
Model summarizing the single turnover measurements.

Different operation modes of the b6f are shown which highlight Qi-site alterations between (A,B) the canonical Q cycle in oxic conditions and (C,D) an alternative Fd-assisted Q cycle in anoxic conditions. Referring to Figure 7, the panels display schematics of the developing electrochromic shift (ECS) signals and b-heme redox changes in the 10 ms range after a subsaturating single turnover flash and, separated by the dashed line, beyond the initial phase. Each turnover is induced by a laser flash, which besides an electron hole also generates a membrane potential (ΔΨ) via PSI charge separation. This further influences b6f properties and the figure focuses on events after the reduction of cyt.f and the swapping of the elliptic FeS domain of the Rieske ISP to a cytochrome b6-proximal position. (A) Per PQH2 formed by Qi-site turnover, the Qo-site turns over twice in a canonical Q cycle. After the first full oxidation of PQH2, the b6f produces a ΔΨ by converting the redox couple blred/bhox to blox/bhred. With the decay of ΔΨ upon ATP synthesis, the prevalent bhred/ciox converts to bhox/cired. The reduction of heme-ci opens the Qi-site for the PQ substrate, which ligates with heme-ci. Swapping of the FeS domain closer to cyt.f is linked to b-heme oxidation. (B) The flash-induced ΔΨ converts bhox/cired to bhred/ciox, which tightly ligates heme-ci with the substrate. To accomplish the ci/(semi-)PQ reduction, reducing pressure is accumulated on the low-potential chain upon a second Qo-site turnover by reduction of blox. In the presence of ΔΨ, this drives the quasi-simultaneous bhred oxidation since blred/bhred is unlikely during single turnover measurements. The (semi-)PQ in the Qi-site receives the electrons from cired and/or bhred in a concerted and closely spaced process. (C,D) In anoxic samples, the low-potential chain was partially pre-reduced before the flash since, with the consumption of ΔΨ during 30 s darkness, equilibration of bhred follows that of cired. The initial reaction steps left from the dashed line were very similar in both (C) pgr5 and (D) WT. The pre-flash blox/bhred re-equilibrated, again, to blox/bhred after the electrogenic charge transfer associated with a Qo-site turnover. During this initial phase, one PQH2 is formed at the Qi-site by utilizing the pre-reduced bhred/cired redox couple. (C) The following b-heme oxidation phase is slower in the mutant and may be attributed to poor Qi-site substrate availability, which depends on cired. In the presence of ΔΨ, bhred/ciox is prevalent and a slower transition to bhox/cired follows the ΔΨ decay in pgr5 (gray and violet curves). (D) Unlike pgr5, the WT retains more FNR at the thylakoid membrane, which might be an allosteric modulator by interacting with the b6f. Thus, FNR-bound Fd may drive the transient generation of cired in the presence of ΔΨ. This creates the ‘PQ-accessible’ Qi-site specifically in WT and, due to the bhred/cired redox couple, produces faster b-heme oxidation and a second PQH2 molecule. Compared with pgr5 and possibly important during multiple turnover, the faster b-heme oxidation also accelerates the Rieske FeS domain to swap closer to cyt.f.

Figure 8.
Model summarizing the single turnover measurements.

Different operation modes of the b6f are shown which highlight Qi-site alterations between (A,B) the canonical Q cycle in oxic conditions and (C,D) an alternative Fd-assisted Q cycle in anoxic conditions. Referring to Figure 7, the panels display schematics of the developing electrochromic shift (ECS) signals and b-heme redox changes in the 10 ms range after a subsaturating single turnover flash and, separated by the dashed line, beyond the initial phase. Each turnover is induced by a laser flash, which besides an electron hole also generates a membrane potential (ΔΨ) via PSI charge separation. This further influences b6f properties and the figure focuses on events after the reduction of cyt.f and the swapping of the elliptic FeS domain of the Rieske ISP to a cytochrome b6-proximal position. (A) Per PQH2 formed by Qi-site turnover, the Qo-site turns over twice in a canonical Q cycle. After the first full oxidation of PQH2, the b6f produces a ΔΨ by converting the redox couple blred/bhox to blox/bhred. With the decay of ΔΨ upon ATP synthesis, the prevalent bhred/ciox converts to bhox/cired. The reduction of heme-ci opens the Qi-site for the PQ substrate, which ligates with heme-ci. Swapping of the FeS domain closer to cyt.f is linked to b-heme oxidation. (B) The flash-induced ΔΨ converts bhox/cired to bhred/ciox, which tightly ligates heme-ci with the substrate. To accomplish the ci/(semi-)PQ reduction, reducing pressure is accumulated on the low-potential chain upon a second Qo-site turnover by reduction of blox. In the presence of ΔΨ, this drives the quasi-simultaneous bhred oxidation since blred/bhred is unlikely during single turnover measurements. The (semi-)PQ in the Qi-site receives the electrons from cired and/or bhred in a concerted and closely spaced process. (C,D) In anoxic samples, the low-potential chain was partially pre-reduced before the flash since, with the consumption of ΔΨ during 30 s darkness, equilibration of bhred follows that of cired. The initial reaction steps left from the dashed line were very similar in both (C) pgr5 and (D) WT. The pre-flash blox/bhred re-equilibrated, again, to blox/bhred after the electrogenic charge transfer associated with a Qo-site turnover. During this initial phase, one PQH2 is formed at the Qi-site by utilizing the pre-reduced bhred/cired redox couple. (C) The following b-heme oxidation phase is slower in the mutant and may be attributed to poor Qi-site substrate availability, which depends on cired. In the presence of ΔΨ, bhred/ciox is prevalent and a slower transition to bhox/cired follows the ΔΨ decay in pgr5 (gray and violet curves). (D) Unlike pgr5, the WT retains more FNR at the thylakoid membrane, which might be an allosteric modulator by interacting with the b6f. Thus, FNR-bound Fd may drive the transient generation of cired in the presence of ΔΨ. This creates the ‘PQ-accessible’ Qi-site specifically in WT and, due to the bhred/cired redox couple, produces faster b-heme oxidation and a second PQH2 molecule. Compared with pgr5 and possibly important during multiple turnover, the faster b-heme oxidation also accelerates the Rieske FeS domain to swap closer to cyt.f.

Table 1.
Key events ofFigure 8 .
Samplebl/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) 
Samplebl/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 [10] and NQNO [64].

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 [31]) 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 [65], 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 [6].

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 [20] 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 [67]). 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 [68]. Moreover, MV-treated thylakoid membranes do not retain bound FNR in the light [69]. 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 [70] with altered chlorophyll fluorescence that we (Figure 4) and others [53] observed.

Model summarizing the multiple turnover measurements.

Figure 9.
Model summarizing the multiple turnover measurements.

Except for PC and NADPH2 pools, the redox levels were measured (blue and red stand for oxidized and reduced, respectively). PSI acceptor side limitation (YNA) and photosynthetic control (ΔpH) were established between weakly (transparent) and strongly contributing levels (bold). Modified forward reaction efficiencies are highlighted (Qo: PQH2 oxidation, Rieske ISP/cyt.f interaction; Qi: PQ reduction, bh/ci electron sharing, Fd-dependent ci reduction; NADPH2 formation). (A) Qi-site inhibition by methyl viologen (MV) disturbs the low-potential chain oxidation and thus ISP/cyt.f interaction. MV likely interferes with the redox signal that mediates FNR tethering to the membrane. (B) The pgr5 mutant shows unregulated electron utilization downstream of PSI, (C) unlike WT. Faster electron flow via PSI was sustained without increasing YNA in algal pgr5 since alternative electron acceptors, such as flavodiiron proteins, prevented the reduction of the Fd pool and PSI photodamage. The mutant showed impaired FNR membrane recruitment and the PQ pool and cyt.f were slightly more oxidized. (D) When anoxic WT b6f operates in the Fd-assisted Q cycle mode, b6f electrogenicity is maintained. Moreover, the generated ΔpH protects PSI, which also throttles electron flow via YNA. (E) Inefficient utilization of excess stromal electrons at the Qi-site stalls the pgr5 b6f in anoxic conditions, thus weakening YNA. The backup of low-potential chain electrons could be linked to a disturbed ISP/cyt.f interaction and loss of efficient pmf generation of the photosynthetic chain.

Figure 9.
Model summarizing the multiple turnover measurements.

Except for PC and NADPH2 pools, the redox levels were measured (blue and red stand for oxidized and reduced, respectively). PSI acceptor side limitation (YNA) and photosynthetic control (ΔpH) were established between weakly (transparent) and strongly contributing levels (bold). Modified forward reaction efficiencies are highlighted (Qo: PQH2 oxidation, Rieske ISP/cyt.f interaction; Qi: PQ reduction, bh/ci electron sharing, Fd-dependent ci reduction; NADPH2 formation). (A) Qi-site inhibition by methyl viologen (MV) disturbs the low-potential chain oxidation and thus ISP/cyt.f interaction. MV likely interferes with the redox signal that mediates FNR tethering to the membrane. (B) The pgr5 mutant shows unregulated electron utilization downstream of PSI, (C) unlike WT. Faster electron flow via PSI was sustained without increasing YNA in algal pgr5 since alternative electron acceptors, such as flavodiiron proteins, prevented the reduction of the Fd pool and PSI photodamage. The mutant showed impaired FNR membrane recruitment and the PQ pool and cyt.f were slightly more oxidized. (D) When anoxic WT b6f operates in the Fd-assisted Q cycle mode, b6f electrogenicity is maintained. Moreover, the generated ΔpH protects PSI, which also throttles electron flow via YNA. (E) Inefficient utilization of excess stromal electrons at the Qi-site stalls the pgr5 b6f in anoxic conditions, thus weakening YNA. The backup of low-potential chain electrons could be linked to a disturbed ISP/cyt.f interaction and loss of efficient pmf generation of the photosynthetic chain.

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 [19] 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 [19]. 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 [16]. Accordingly, introducing flavodiiron proteins in the Arabidopsis pgr5 background alleviates the PSI phenotype [74].

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 [77]. 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.

Competing Interests

The authors declare that there are no competing interests associated with the manuscript.

Funding

Author Contribution

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.

Acknowledgements

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).

Abbreviations

     
  • ΔΨ

    membrane potential

  •  
  • b6f

    cytochrome b6f complex

  •  
  • CEF

    cyclic electron flow

  •  
  • cyt.f

    cytochrome f

  •  
  • ECS

    electrochromic shift

  •  
  • Fd

    ferredoxin

  •  
  • FNR

    Fd-NADP(H) oxidoreductase

  •  
  • ISP

    iron sulfur protein

  •  
  • LEF

    linear electron flow

  •  
  • MV

    methyl viologen

  •  
  • PC

    plastocyanin

  •  
  • PGR5

    proton gradient regulation 5

  •  
  • Pmf

    proton motive force

  •  
  • PQ

    plastoquinone

  •  
  • PQH2

    plastoquinol

  •  
  • PSI/PSII

    photosystem I/II

References

References
1
Cramer
,
W. A.
and
Hasan
,
S. S.
(
2016
) Structure-function of the cytochrome b6f lipoprotein complex. In
Cytochrome Complexes: Evolution, Structures, Energy Transduction, and Signaling
(
Cramer
,
W. A.
and
Kallas
,
T.
, eds), pp.
177
207
,
Springer Netherlands
,
Dordrecht
2
Lavergne
,
J.
(
1983
)
Membrane potential-dependent reduction of cytochrome b6 in an algal mutant lacking photosystem I centers
.
Biochim. Biophys. Acta
725
,
25
33
3
Barbagallo
,
R.P.
,
Breyton
,
C.
and
Finazzi
,
G.
(
2000
)
Kinetic effects of the electrochemical proton gradient on plastoquinone reduction at the Qi site of the cytochrome b6f complex
.
J. Biol. Chem.
275
,
26121
26127
4
Zatsman
,
A.I.
,
Zhang
,
H.
,
Gunderson
,
W.A.
,
Cramer
,
W.A.
and
Hendrich
,
M.P.
(
2006
)
Heme–heme interactions in the cytochrome b6f complex: EPR spectroscopy and correlation with structure
.
J. Am. Chem. Soc.
128
,
14246
14247
5
Baymann
,
F.
,
Giusti
,
F.
,
Picot
,
D.
and
Nitschke
,
W.
(
2007
)
The ci/bH moiety in the b6f complex studied by EPR: a pair of strongly interacting hemes
.
Proc. Natl. Acad. Sci. U.S.A.
104
,
519
524
6
Alric
,
J.
,
Pierre
,
Y.
,
Picot
,
D.
,
Lavergne
,
J.
and
Rappaport
,
F.
(
2005
)
Spectral and redox characterization of the heme ci of the cytochrome b6f complex
.
Proc. Natl. Acad. Sci. U.S.A.
102
,
15860
15865
7
de Lavalette
,
A.D.
,
Barucq
,
L.
,
Alric
,
J.
,
Rappaport
,
F.
and
Zito
,
F.
(
2009
)
Is the redox state of the ci heme of the cytochrome b6f complex dependent on the occupation and structure of the Qi site and vice versa?
J. Biol. Chem.
284
,
20822
20829
8
Malone
,
L.A.
,
Qian
,
P.
,
Mayneord
,
G.E.
,
Hitchcock
,
A.
,
Farmer
,
D.A.
,
Thompson
,
R.F.
et al (
2019
)
Cryo-EM structure of the spinach cytochrome b6 f complex at 3.6 A resolution
.
Nature
575
,
535
539
.
9
Furbacher
,
P.N.
,
Girvin
,
M.E.
and
Cramer
,
W.A.
(
1989
)
On the question of interheme electron transfer in the chloroplast cytochrome b6 in situ
.
Biochemistry
28
,
8990
8998
10
Rich
,
P.R.
,
Madgwick
,
S.A.
,
Brown
,
S.
,
von Jagow
,
G.
and
Brandt
,
U.
(
1992
)
MOA-stilbene: a new tool for investigation of the reactions of the chloroplast cytochrome bf complex
.
Photosynth. Res.
34
,
465
477
11
Mulkidjanian
,
A.Y.
(
2010
)
Activated Q-cycle as a common mechanism for cytochrome bc1 and cytochrome b6f complexes
.
Biochim. Biophys. Acta
1797
,
1858
1868
12
Allen
,
J.F.
(
2004
)
Cytochrome b6f: structure for signalling and vectorial metabolism
.
Trends Plant Sci.
9
,
130
137
13
Nawrocki
,
W.J.
,
Bailleul
,
B.
,
Picot
,
D.
,
Cardol
,
P.
,
Rappaport
,
F.
,
Wollman
,
F.A.
et al (
2019
)
The mechanism of cyclic electron flow
.
Biochim. Biophys. Acta
1860
,
433
438
14
Stiehl
,
H.H.
and
Witt
,
H.T.
(
1969
)
Quantitative treatment of function of plastoquinone in photosynthesis
.
Z.Naturforsch. B
24
,
1588
1598
15
Kramer
,
D.M.
,
Sacksteder
,
C.A.
and
Cruz
,
J.A.
(
1999
)
How acidic is the lumen?
Photosynth. Res.
60
,
151
163
16
Munekage
,
Y.
,
Hojo
,
M.
,
Meurer
,
J.
,
Endo
,
T.
,
Tasaka
,
M.
and
Shikanai
,
T.
(
2002
)
PGR5 is involved in cyclic electron flow around photosystem I and is essential for photoprotection in arabidopsis
.
Cell
110
,
361
371
17
Nandha
,
B.
,
Finazzi
,
G.
,
Joliot
,
P.
,
Hald
,
S.
and
Johnson
,
G.N.
(
2007
)
The role of PGR5 in the redox poising of photosynthetic electron transport
.
Biochim. Biophys. Acta
1767
,
1252
1259
18
Suorsa
,
M.
,
Jarvi
,
S.
,
Grieco
,
M.
,
Nurmi
,
M.
,
Pietrzykowska
,
M.
,
Rantala
,
M.
et al (
2012
)
PROTON GRADIENT REGULATION5 is essential for proper acclimation of Arabidopsis photosystem I to naturally and artificially fluctuating light conditions
.
Plant Cell
24
,
2934
2948
19
Johnson
,
X.
,
Steinbeck
,
J.
,
Dent
,
R.M.
,
Takahashi
,
H.
,
Richaud
,
P.
,
Ozawa
,
S.
et al (
2014
)
Proton gradient regulation 5-mediated cyclic electron flow under ATP- or redox-limited conditions: a study of (ATpase pgr5 and (rbcL pgr5 mutants in the green alga Chlamydomonas reinhardtii
.
Plant Physiol.
165
,
438
452
20
Mosebach
,
L.
,
Heilmann
,
C.
,
Mutoh
,
R.
,
Gabelein
,
P.
,
Steinbeck
,
J.
,
Happe
,
T.
et al (
2017
)
Association of Ferredoxin:NADP(+) oxidoreductase with the photosynthetic apparatus modulates electron transfer in Chlamydomonas reinhardtii
.
Photosynth. Res.
134
,
291
306
21
Jokel
,
M.
,
Johnson
,
X.
,
Peltier
,
G.
,
Aro
,
E.M.
and
Allahverdiyeva
,
Y.
(
2018
)
Hunting the main player enabling Chlamydomonas reinhardtii growth under fluctuating light
.
Plant J.
94
,
822
835
22
Alric
,
J.
(
2014
)
Redox and ATP control of photosynthetic cyclic electron flow in Chlamydomonas reinhardtii: (II) involvement of the PGR5-PGRL1 pathway under anaerobic conditions
.
Biochim. Biophys. Acta
1837
,
825
834
23
Clark
,
R.D.
,
Hawkesford
,
M.J.
,
Coughlan
,
S.J.
,
Bennett
,
J.
and
Hind
,
G.
(
1984
)
Association of ferredoxin-NADP+ oxidoreductase with the chloroplast cytochrome b-f complex
.
FEBS Lett.
174
,
137
142
24
Zhang
,
H.
,
Whitelegge
,
J.P.
and
Cramer
,
W.A.
(
2001
)
Ferredoxin:NADP+ oxidoreductase is a subunit of the chloroplast cytochrome b6f complex
.
J. Biol. Chem.
276
,
38159
38165
25
Okutani
,
S.
,
Hanke
,
G.T.
,
Satomi
,
Y.
,
Takao
,
T.
,
Kurisu
,
G.
,
Suzuki
,
A.
et al (
2005
)
Three maize leaf ferredoxin:NADPH oxidoreductases vary in subchloroplast location, expression, and interaction with ferredoxin
.
Plant Physiol.
139
,
1451
1459
26
Joliot
,
P.
,
Béal
,
D.
and
Joliot
,
A.
(
2004
)
Cyclic electron flow under saturating excitation of dark-adapted Arabidopsis leaves
.
Biochim. Biophys. Acta
1656
,
166
176
27
Joliot
,
P.
and
Joliot
,
A.
(
2006
)
Cyclic electron flow in C3 plants
.
Biochim. Biophys. Acta
1757
,
362
368
28
Joliot
,
P.
and
Johnson
,
G.N.
(
2011
)
Regulation of cyclic and linear electron flow in higher plants
.
Proc. Natl. Acad. Sci. U.S.A.
108
,
13317
13322
29
Sueoka
,
N.
(
1960
)
Mitotic replication of deoxyribonucleic acid in Chlamydomonas reinhardi
.
Proc. Natl. Acad. Sci. U.S.A.
46
,
83
91
30
Porra
,
R.J.
,
Thompson
,
W.A.
and
Kriedemann
,
P.E.
(
1989
)
Determination of accurate extinction coefficients and simultaneous-equations for assaying chlorophyll-a and chlorophyll-b extracted with 4 different solvents - verification of the concentration of chlorophyll standards by atomic-absorption spectroscopy
.
Biochim. Biophys. Acta
975
,
384
394
31
Johnson
,
X.
and
Alric
,
J.
(
2013
)
Central carbon metabolism and electron transport in Chlamydomonas reinhardtii: metabolic constraints for carbon partitioning between oil and starch
.
Eukaryot. Cell
12
,
776
793
32
Landt
,
O.
,
Grunert
,
H.P.
and
Hahn
,
U.
(
1990
)
A general method for rapid site-directed mutagenesis using the polymerase chain reaction
.
Gene
96
,
125
128
33
Scholz
,
M.
,
Gabelein
,
P.
,
Xue
,
H.
,
Mosebach
,
L.
,
Bergner
,
S.V.
and
Hippler
,
M.
(
2019
)
Light-dependent N-terminal phosphorylation of LHCSR3 and LHCB4 are interlinked in Chlamydomonas reinhardtii
.
Plant J.
99
,
877
894
34
Rasala
,
B.A.
,
Lee
,
P.A.
,
Shen
,
Z.
,
Briggs
,
S.P.
,
Mendez
,
M.
and
Mayfield
,
S.P.
(
2012
)
Robust expression and secretion of Xylanase1 in Chlamydomonas reinhardtii by fusion to a selection gene and processing with the FMDV 2A peptide
.
PLoS One
7
,
e43349
35
Rappaport
,
F.
,
Beal
,
D.
,
Joliot
,
A.
and
Joliot
,
P.
(
2007
)
On the advantages of using green light to study fluorescence yield changes in leaves
.
Biochim. Biophys. Acta
1767
,
56
65
36
Baker
,
N.R.
(
2008
)
Chlorophyll fluorescence: a probe of photosynthesis in vivo
.
Annu. Rev. Plant Biol.
59
,
89
113
37
Genty
,
B.
,
Briantais
,
J.-M.
and
Baker
,
N.R.
(
1989
)
The relationship between the quantum yield of photosynthetic electron transport and quenching of chlorophyll fluorescence
.
Biochim. Biophys. Acta
990
,
87
92
38
Oxborough
,
K.
and
Baker
,
N.R.
(
1997
)
Resolving chlorophyll a fluorescence images of photosynthetic efficiency into photochemical and non-photochemical components - calculation of qP and Fv′/Fm′ without measuring Fo
.
Photosynth. Res.
54
,
135
142
39
Kramer
,
D.M.
,
Johnson
,
G.
,
Kiirats
,
O.
and
Edwards
,
G.E.
(
2004
)
New fluorescence parameters for the determination of QA redox state and excitation energy fluxes
.
Photosynth. Res.
79
,
209
40
Kalaji
,
H.M.
,
Schansker
,
G.
,
Brestic
,
M.
,
Bussotti
,
F.
,
Calatayud
,
A.
,
Ferroni
,
L.
et al (
2017
)
Frequently asked questions about chlorophyll fluorescence, the sequel
.
Photosynth. Res.
132
,
13
66
41
Schreiber
,
U.
and
Bilger
,
W
. (
1987
) Rapid assessment of stress effects on plant leaves by chlorophyll fluorescence measurements. In
Plant Response to Stress
(
Tenhunen
,
J.D.
,
Catarina
,
F.M.
,
Lange
,
O.L.
and
Oechel
,
W.C.
, eds), pp.
27
53
,
Springer Berlin Heidelberg
,
Berlin, Heidelberg
42
Weis
,
E.
and
Berry
,
J.A.
(
1987
)
Quantum efficiency of photosystem II in relation to ‘energy’-dependent quenching of chlorophyll fluorescence
.
Biochim. Biophys. Acta
894
,
198
208
43
Joliot
,
P.
and
Joliot
,
A.
(
2008
)
Quantification of the electrochemical proton gradient and activation of ATP synthase in leaves
.
Biochim. Biophys. Acta
1777
,
676
683
44
Malnoe
,
A.
,
Wollman
,
F.A.
,
de Vitry
,
C.
and
Rappaport
,
F.
(
2011
)
Photosynthetic growth despite a broken Q-cycle
.
Nat. Commun.
2
,
301
45
Joliot
,
P.
and
Joliot
,
A.
(
2001
)
Electrogenic events associated with electron and proton transfers within the cytochrome b6/f complex
.
Biochim. Biophys. Acta
1503
,
369
376
46
Zito
,
F.
,
Finazzi
,
G.
,
Joliot
,
P.
and
Wollman
,
F.A.
(
1998
)
Glu78, from the conserved PEWY sequence of subunit IV, has a key function in cytochrome b6f turnover
.
Biochemistry
37
,
10395
10403
47
Joliot
,
P.
and
Joliot
,
A.
(
2002
)
Cyclic electron transfer in plant leaf
.
Proc. Natl. Acad. Sci. U.S.A.
99
,
10209
10214
48
Bailleul
,
B.
,
Cardol
,
P.
,
Breyton
,
C.
and
Finazzi
,
G.
(
2010
)
Electrochromism: a useful probe to study algal photosynthesis
.
Photosynth. Res.
106
,
179
189
49
Avenson
,
T.J.
,
Cruz
,
J.A.
and
Kramer
,
D.M.
(
2004
)
Modulation of energy-dependent quenching of excitons in antennae of higher plants
.
Proc. Natl. Acad. Sci. U.S.A.
101
,
5530
5535
50
Klughammer
,
C.
and
Schreiber
,
U.
(
1994
)
An improved method, using saturating light-pulses, for the determination of photosystem-I quantum yield via P700+-absorbance changes at 830 nm
.
Planta
192
,
261
268
51
Brettel
,
K.
(
1997
)
Electron transfer and arrangement of the redox cofactors in photosystem I
.
Biochim. Biophys. Acta
1318
,
322
373
52
Alric
,
J.
,
Lavergne
,
J.
and
Rappaport
,
F.
(
2010
)
Redox and ATP control of photosynthetic cyclic electron flow in Chlamydomonas reinhardtii (I) aerobic conditions
.
Biochim. Biophys. Acta
1797
,
44
51
53
Fan
,
D.Y.
,
Jia
,
H.
,
Barber
,
J.
and
Chow
,
W.S.
(
2009
)
Novel effects of methyl viologen on photosystem II function in spinach leaves
.
Eur. Biophys. J.
39
,
191
199
54
Joliot
,
P.
and
Delosme
,
R.
(
1974
)
Flash-induced 519 nm absorption change in green algae
.
Biochim. Biophys. Acta
357
,
267
284
55
Robertson
,
D.E.
and
Dutton
,
P.L.
(
1988
)
The nature and magnitude of the charge-separation reactions of ubiquinol cytochrome c2 oxidoreductase
.
Biochim. Biophys. Acta
935
,
273
291
56
Glaser
,
E.G.
and
Crofts
,
A.R.
(
1984
)
A new electrogenic step in the ubiquinol:cytochrome c2 oxidoreductase complex of rhodopseudomonas sphaeroides
.
Biochim. Biophys. Acta
766
,
322
333
57
Iwai
,
M.
,
Takizawa
,
K.
,
Tokutsu
,
R.
,
Okamuro
,
A.
,
Takahashi
,
Y.
and
Minagawa
,
J.
(
2010
)
Isolation of the elusive supercomplex that drives cyclic electron flow in photosynthesis
.
Nature
464
,
1210
1213
58
Terashima
,
M.
,
Petroutsos
,
D.
,
Hudig
,
M.
,
Tolstygina
,
I.
,
Trompelt
,
K.
,
Gabelein
,
P.
et al (
2012
)
Calcium-dependent regulation of cyclic photosynthetic electron transfer by a CAS, ANR1, and PGRL1 complex
.
Proc. Natl. Acad. Sci. U.S.A.
109
,
17717
17722
59
Takahashi
,
H.
,
Clowez
,
S.
,
Wollman
,
F.A.
,
Vallon
,
O.
and
Rappaport
,
F.
(
2013
)
Cyclic electron flow is redox-controlled but independent of state transition
.
Nat. Commun.
4
,
1954
60
Steinbeck
,
J.
,
Ross
,
I.L.
,
Rothnagel
,
R.
,
Gäbelein
,
P.
,
Schulze
,
S.
,
Giles
,
N.
et al (
2018
)
Structure of a PSI–LHCI–cyt b6f supercomplex in Chlamydomonas reinhardtii promoting cyclic electron flow under anaerobic conditions
.
Proc. Natl. Acad. Sci. U.S.A.
115
,
10517
10522
61
Mulkidjanian
,
A.Y.
(
2005
)
Ubiquinol oxidation in the cytochrome bc1 complex: Reaction mechanism and prevention of short-circuiting
.
Biochim. Biophys. Acta
1709
,
5
34
62
Yamashita
,
E.
,
Zhang
,
H.
and
Cramer
,
W.A.
(
2007
)
Structure of the cytochrome b6f complex: quinone analogue inhibitors as ligands of heme cn
.
J. Mol. Biol.
370
,
39
52
63
Mitchell
,
P.
(
1975
)
The protonmotive Q cycle: a general formulation
.
FEBS Lett.
59
,
137
139
64
Jones
,
R.W.
and
Whitmarsh
,
J.
(
1988
)
Inhibition of electron transfer and the electrogenic reaction in the cytochrome bf complex by 2-n-nonyl-4-hydroxyquinoline N-oxide (NQNO) and 2,5-dibromo-3-methyl-6-isopropyl-p-benzoquinone (DBMIB)
.
Biochim. Biophys. Acta
933
,
258
268
65
Joliot
,
P.
and
Joliot
,
A.
(
1988
)
The low-potential electron-transfer chain in the cytochrome bf complex
.
Biochim. Biophys. Acta
933
,
319
333
66
Joliot
,
P.
and
Joliot
,
A.
(
1986
)
Mechanism of proton-pumping in the cytochrome b/f complex
.
Photosynth. Res.
9
,
113
124
67
Batie
,
C.J.
and
Kamin
,
H.
(
1981
)
The relation of pH and oxidation-reduction potential to the association state of the ferredoxin (ferredoxin:NADP+ reductase complex
.
J. Biol. Chem.
256
,
7756
7763
PMID:
[PubMed]
68
Takahashi
,
H.
,
Okamuro
,
A.
,
Minagawa
,
J.
and
Takahashi
,
Y.
(
2014
)
Biochemical characterization of photosystem I-associated light-harvesting complexes I and II isolated from state 2 cells of Chlamydomonas reinhardtii
.
Plant Cell Physiol.
55
,
1437
1449
69
Palatnik
,
J.F.
,
Valle
,
E.M.
and
Carrillo
,
N.
(
1997
)
Oxidative stress causes ferredoxin-NADP+ reductase solubilization from the thylakoid membranes in methyl viologen-treated plants
.
Plant Physiol.
115
,
1721
1727
70
Davis
,
G.A.
,
Kanazawa
,
A.
,
Schottler
,
M.A.
,
Kohzuma
,
K.
,
Froehlich
,
J.E.
,
Rutherford
,
A.W.
et al (
2016
)
Limitations to photosynthesis by proton motive force-induced photosystem II photodamage
.
eLife
5
,
e16921
71
DalCorso
,
G.
,
Pesaresi
,
P.
,
Masiero
,
S.
,
Aseeva
,
E.
,
Schunemann
,
D.
,
Finazzi
,
G.
et al (
2008
)
A complex containing PGRL1 and PGR5 is involved in the switch between linear and cyclic electron flow in Arabidopsis
.
Cell
132
,
273
285
72
Takagi
,
D.
and
Miyake
,
C.
(
2018
)
PROTON GRADIENT REGULATION 5 supports linear electron flow to oxidize photosystem I
.
Physiol. Plant.
164
,
337
348
73
Dang
,
K.V.
,
Plet
,
J.
,
Tolleter
,
D.
,
Jokel
,
M.
,
Cuine
,
S.
,
Carrier
,
P.
et al (
2014
)
Combined increases in mitochondrial cooperation and oxygen photoreduction compensate for deficiency in cyclic electron flow in Chlamydomonas reinhardtii
.
Plant Cell.
26
,
3036
3050
74
Yamamoto
,
H.
,
Takahashi
,
S.
,
Badger
,
M.R.
and
Shikanai
,
T.
(
2016
)
Artificial remodelling of alternative electron flow by flavodiiron proteins in Arabidopsis
.
Nat. Plants
2
,
16012
75
Steinbeck
,
J.
,
Nikolova
,
D.
,
Weingarten
,
R.
,
Johnson
,
X.
,
Richaud
,
P.
,
Peltier
,
G.
et al (
2015
)
Deletion of proton gradient regulation 5 (PGR5) and PGR5-Like 1 (PGRL1) proteins promote sustainable light-driven hydrogen production in Chlamydomonas reinhardtii due to increased PSII activity under sulfur deprivation
.
Front. Plant. Sci.
6
,
892
76
Chen
,
M.
,
Zhang
,
J.
,
Zhao
,
L.
,
Xing
,
J.L.
,
Peng
,
L.W.
,
Kuang
,
T.Y.
et al (
2016
)
Loss of algal proton gradient regulation 5 increases reactive oxygen species scavenging and H2 evolution
.
J. Integr. Plant Biol.
58
,
943
946
77
Hertle
,
A.P.
,
Blunder
,
T.
,
Wunder
,
T.
,
Pesaresi
,
P.
,
Pribil
,
M.
,
Armbruster
,
U.
et al (
2013
)
PGRL1 is the elusive ferredoxin-plastoquinone reductase in photosynthetic cyclic electron flow
.
Mol. Cell
49
,
511
523
78
Kramer
,
D.M.
and
Crofts
,
A.R.
(
1989
)
Activation of the chloroplast ATPase measured by the electrochromic change in leaves of intact plants
.
Biochim. Biophys. Acta
976
,
28
41