Photosynthetic electrons are distributed among many bioenergetics and biosynthetic pathways. Recent re-ports have suggested that the improvement of crop and biofuel yields depends on enhanced photosynthetic ef-ficiency (Ort et al., 2015; Gu et al., 2017; Orr et al., 2017; Rogers et al., 2017). Therefore, deciphering the main electron transfer junctions is a prerequisite for further engineering. An excellent candidate for such studies and engineering attempts is algal photobiological hy-drogen production, which has gained interest during the search for sustainable energy alternatives (Ghirardi, 2015). In the plant kingdom, hydrogen production, catalyzed by the enzyme hydrogenase, is unique to green algae. The enzyme activity is mediated by pho-tosynthetic electron transfer from the main electron hub, ferredoxin (Winkler et al., 2009; Hemschemeier and Happe, 2011). Ferredoxin is the ultimate electron mediator of the photosynthetic apparatus, mediating photosynthetic electron transfer to numerous compet-ing pathways (Hemschemeier and Happe, 2011). Al-though there are several pathways in which ferredoxin

can be reduced, the most prominent is photoreduction by photosystem I (Redding et al., 1999).

It is well established that hydrogenase accumulates under a dark anoxic adaptation (Happe and Kaminski, 2002; Mus et al., 2007). Following such induction, ex-posure of algae to light supports high rates of hydro-gen production. However, it is commonly known that hydrogen production ceases within a few minutes of illumination (Gaffron and Rubin, 1942; Ghirardi, 2015; Noone et al., 2017). The common dogma claims that oxygen, which originates from the light-mediated wa-ter splitting at photosystem II, irreversibly inactivates hydrogenase and is therefore responsible for this fast cessation (Erbes et al., 1979; Ghirardi, 2015). An alter-native hypothesis suggests a complex series of events where hydrogen production stops before the inevitable destruction of hydrogenase by oxygen. It was suggested that the cessation of hydrogen production is caused by competition for reducing equivalents, initially with cyclic electron flow (CEF), oxygen reduction, and ul-timately with carbon dioxide fixation by the Calvin Benson Bassham (CBB) cycle. (Lee and Greenbaum, 2003; Yacoby et al., 2011; Godaux et al., 2015). Still, conclusive evidence that supports either hypothesis is missing (Fig. 1). Deciphering the mechanism of hydro-genase inactivation requires assessing the enzyme’s pool half-life and identifying the prominent competing processes in a transition from dark anoxia to light.

In this report, we aimed to gain unbiased measure-ments of the electron divergence between hydrogen production and competing processes. We studied con-ditions in which the active hydrogenase pool is not a limiting factor, i.e. after anaerobic induction. Measuring the rates of photosynthetically relevant gases upon light onset allowed us to identify two main players—carbon

Green Algal Hydrogenase Activity Is Outcompeted by Carbon Fixation before Inactivation by Oxygen Takes Place1

Yuval Milrad, Shira Schweitzer, Yael Feldman, and Iftach Yacoby2

School of Plant Sciences and Food Security, The George S. Wise Faculty of Life Sciences, Tel Aviv University, Ramat Aviv, Tel Aviv 69978, IsraelORCID ID: 0000‑0003‑0177‑0624 (I.Y.)

Photoproduction of hydrogen by green algae is considered a transitory release valve of excess reducing power and a potential carbon-free source of sustainable energy. It is generally accepted that the transitory production of hydrogen is governed by fast inactivation of hydrogenase by oxygen. However, our data suggest that photosynthetic electron loss to competing processes, mainly carbon fixation, stops hydrogen production, supports hydrogen uptake, and precedes the inevitable inactivation by oxy-gen. Here, we show that when transitioning from dark anaerobiosis to light, hydrogen production ceases within 2 min, regard-less of the presence of oxygen. Simultaneous monitoring of the active hydrogenase pool size shows that it remains entirely intact up to 4 min after illumination and is inactivated only later. Thus, our data reveal a window of 4 min in which the hydrogenase pool is not being degraded by oxygen. Furthermore, we show that electron loss, prominently to carbon fixation, outcompetes hydrogen production and leads to hydrogen uptake. Indeed, supplying additional reducing power to hydrogenase at the cessa-tion point regenerates the accumulation of hydrogen. Our results imply the fast cessation of hydrogen production is governed by electron loss rather than oxygen inactivation, which takes place minutes later.

1This work was supported in part by Israel Science Foundation iCore grant 757/12 and Israel Science Foundation personal grant 1646/16.

2Address correspondence to iftachy@tau.ac.il.The author responsible for distribution of materials integral to

the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Iftach Yacoby (iftachy@tau.ac.il).

Y.M. and I.Y. conducted the experimental design and analysis; Y.M., S.S., and Y.F. conducted the measurements; I.Y. and Y.M. wrote the manuscript with the assistance of S.S. and Y.F.

www.plantphysiol.org/cgi/doi/10.1104/pp.18.00229

918 Plant Physiology®, July 2018, Vol. 177, pp. 918–926, www.plantphysiol.org © 2018 American Society of Plant Biologists. All Rights Reserved.

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dioxide and hydrogen—defining the arena of compe-tition. We show when and under what circumstances hydrogen production ceases and what the content of active hydrogenase is before, at, and after the cessation point of hydrogen production. Furthermore, we deter-mine that the prominent cause for this fast cessation is electron funneling to carbon fixation, which precedes the inactivation of the catalytic site of hydrogenase by oxygen.

RESULTS

Kinetics and Duration of Hydrogen Production

To study the duration of hydrogen production at the onset of light following dark anaerobic incubation, we carried out studies using a membrane inlet mass spec-trometer (MIMS). This apparatus monitors the concen-trations of dissolved hydrogen and other soluble gases of interest in real time (Mus et al., 2005). To establish an anaerobic environment, Chlamydomonas reinhardtii cells were put in a MIMS cuvette and kept in the dark for 1 h, which created conditions that support the accumula-tion of hydrogenase (Mus et al., 2007). Within the initial period of 15 min, the cells respired the entire oxygen

pool (Fig. 2A). Interestingly, fermentative hydrogen production (Klein and Betz, 1978; Gfeller and Gibbs, 1984; Mus et al., 2007) also initiated at this moment (Fig. 2A; -45 min) in the absence of previous anoxic induction, thus reflecting the activity of a basal pool of hydrogenase that was present under air, which agrees with the findings of Liran et al. (2016). Previously, it was reported that under such conditions hydrogenase accumulation continues linearly up to 4 h (Eilenberg et al., 2016). However, the rate of photosynthetic hy-drogen production increases only during the first 2 h of dark anoxic incubation; therefore, the enzymatic activity is saturated (any additional dark incubation time does not increase the rate under irradiance). The full anaerobic induction period was accompanied by a gradual increase in hydrogen concentration (Fig. 2A, green line), resulting from the fermentative route (Klein and Betz, 1978; Gfeller and Gibbs, 1984; Mus et al., 2007). Following 1 h dark incubation, we illuminated the sample with a continuous actinic light (370 µE m−2 s−1) for 15 min (Fig. 2A, time = 0). As expected, hy-drogen evolved rapidly for 2 min, while oxygen levels remained low. Then, hydrogen accumulation ceased (marked with a red arrow) and its concentration started declining because of hydrogen uptake by hydrogenase (Noone et al., 2017). The uptake of hydrogen was con-firmed to be of a biological nature (Supplemental Fig. S1). Simultaneously, oxygen concentration, which is evolved from water splitting by photosystem II in the light, increased. During the measurement, we sampled 50 µL of cells directly from the MIMS cuvette (which holds a total volume of 5 mL) and performed a chemi-cal determination of the active hydrogenase pool. This determination was conducted using an excess of re-duced methyl viologen (Eilenberg et al., 2016; Khanna et al., 2017; Sawyer et al., 2017; Nikolova et al., 2018; Weiner et al., 2018), which provides electrons to hydro-genase for H+ reduction. This determination, in con-trast to immunoblotting, quantifies solely the pool of active hydrogenase. This quantification shows that as expected, hydrogenase quantity rose during the dark incubation (Fig. 2A). Remarkably, the active enzyme pool was intact and did not change pursuant to light exposure during the first 4 min of illumination, at which time oxygen levels increased up to 4 μm. After 4 min of illumination, a decrease of the active hydroge-nase pool is observed in correlation with the increasing oxygen concentration (traces of active enzyme could also be observed after 10 min of illumination). These results indicate that at the cessation points of hydro-gen production, ∼2 min after light onset, the pool of hydrogenase is entirely intact. Therefore, oxygen in-activation of hydrogenase pool cannot explain the fast cessation of photosynthetic hydrogen production.

Kinetics of the Dissolved Gases under Light/Dark Cycles

To investigate the roots of the fast cessation of hy-drogen production at light onset, we examined the ki-netics of hydrogen, oxygen and carbon dioxide. Since

Figure 1. Schematic of possible hydrogenase inhibition mechanisms. Photosynthetic electron flow (orange arrows) is generated by water splitting at photosystem II (PSII) upon illumination (yellow lightning). Oxygen is produced as a by-product at photosystem II (blue arrow). Electron transfer from photosystem II drives proton pumping via cyto-chrome b6f (Cytb6f), after which it is mediated by plastocyanin (PC) to photosystem I (PSI). An additional illumination excites photosystem I, which in turn reduces the main electron hub Fd. The following mech-anisms were suggested as potential inhibitors of hydrogen production: (1) electron loss to carbon fixation through the CBB cycle (Yacoby et al., 2011); (2) electron loss to oxygen reduction within the chloroplast (Lee and Greenbaum, 2003); (3) deactivation of hydrogenase by oxygen (Erbes et al., 1979); and (4) electron loss to cyclic electron flow (Godaux et al., 2015).

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we observed that the hydrogenase pool is intact after 2 min of illumination (at the cessation point of hydrogen production), we exposed the cells to short interval of illumination, which lasted 2 min (at an irradiance of 370 µE m−2 s−1). Following each light exposure, the cells were kept in the dark for 3 min until anaerobiosis re-sumed and then the light was switched on again (Fig. 2B). Each experiment consisted of 10 light/dark cycles. At the onset of each light exposure cycle, we observed a rapid accumulation of hydrogen, while oxygen also accumulated but remained at low concentrations (up to 5 μm). In addition, we observed a sharp increase in the concentration of carbon dioxide probably due to bicarbonate conversion to carbon dioxide and water catalyzed by carbonic anhydrases. This increase lasted a minute, after which the concentration of carbon diox-ide started declining presumably due to operation of the CBB cycle (Fig. 2B). Subsequently, when the light was turned off, hydrogen and oxygen concentrations

dropped, while the concentration of carbon dioxide in-creased again for a short period of time. Interestingly, each light onset triggered a rapid hydrogen accumula-tion, which declined in the same manner (average of 49 µmol H2 mg chlorophyll [Chl]−1 h−1 under mixotropic conditions). However, although the enzyme’s pool size did not change from cycle to cycle (Fig. 2B, columns), the initial rate at the onset of the first cycle (following 1 h of dark induction; 150 µmol H2 mg Chl−1 h−1) was 3-fold higher than each of the sequential exposures. This is also reflected by larger accumulation of hydro-gen in the first cycle compared to the following cycles (Supplemental Fig. S2).

Oxygen Has a Limited Role in the Cessation of Hydrogen Production after 2 min of Illumination

To gain better understanding of these observations, we averaged the concentration data for cycles 2 to 10

Figure 2. Hydrogen production kinetics and quantification of the enzyme pool. C. reinhardtii wild-type strain cc-124 cells were incubated for 1 h in the dark (gray background), after which they were exposed to an irradiance of 370 µE m−2 s−1 (white background) light for 15 min, in which hydrogen evolution was observed for 2 min before it ceased completely (red arrow; A), or cycles of 2 min of illumination (370 µE m−2 s−1; white background) and 3 min of darkness (B). Shown are dissolved concen-trations of oxygen (O2; blue), hydrogen (H2; green), and carbon dioxide (CO2; red) as measured in the MIMS. Simultaneously in both measurements, the cells were sampled for quantification of the active pool of hydrogenase (HydA) using the methyl violo-gen assay (MV; orange columns). Each experiment was repeated using three biological replicates. Error bars indicate se (n = 3).

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(Fig. 3A). We then plotted the rates for each of the gas-es of interest (hydrogen, oxygen, and carbon dioxide) as a function of time (Fig. 3C). Using this analysis, we determined the initial rate of hydrogen production at the light onset. The exposure to light triggered an im-mediate reaction, in which hydrogen was produced at a high rate. Then after 30 s of illumination, the rate started declining gradually and stopped after 2 min of illumination. It should be noted that this phenome-non was observed also in cells that were grown under autotrophic conditions, although in lower rates (Sup-plemental Fig. S3). To study whether the cessation of hy-drogen accumulation relates to oxygen uptake, e.g. by the Mehler reaction or chlororespiration, we repeated the measurements and rate calculations described above under strictly anaerobic conditions (Fig. 3, B and D). Continuous oxygen removal was maintained by Glc oxidase (GOx), which supports a rapid consump-tion of oxygen in the medium. To study whether an efficient removal of intracellular oxygen was occurring in the presence of GOx, we used the active pool of hy-drogenase as an indicator of internal oxygen accumu-lation. Hence, in the presence of GOx, one can expect that the hydrogenase pool will remain intact for longer durations. For testing this, we determined the active enzyme pool using the methyl viologen procedure described earlier. We observed (Supplemental Fig. S4) that in the presence of GOx the enzyme’s pool was

intact for the entire duration of illumination (12 min). Hence, intracellular oxygen that could damage the enzyme pool or compete for electrons was effectively eliminated by the addition of GOx. Interestingly, under these conditions, in the presence of GOx, the kinetics of hydrogen and carbon dioxide did not differ much, and hydrogen production still ceased after 2 min. Any attempt to differentiate between the hydrogen rates or accumulation in presence or absence of GOx resulted in nonsignificant differences (data not shown). There-fore, we concluded that electron loss to oxygen uptake plays a minor role in ceasing hydrogenase activity at this regime of low oxygen concentrations. At the end of the illumination period, the accumulated oxygen was respired in line with the observed positive carbon dioxide efflux, which changes direction rapidly once the culture becomes anaerobic again (Fig. 3, A and C). The lack of dissolved oxygen in the presence of GOx (Fig. 3, B and D) eliminated that emission. This obser-vation is in line with the increase in carbon efflux to the medium at the onset of darkness, governed by oxygen respiration and mitochondrial activity.

Electron Loss to Carbon Fixation Hinders Hydrogen Production

The kinetics of carbon dioxide (a combination of uptake and respiration; Fig. 3, A and C) at light onset

Figure 3. Hydrogen production following light and dark cycles. C. reinhardtii wild-type strain cc-124 cells were incubated for 1 h in the dark, after which they were exposed to cycles of 2 min of illumination (370 µE m−2 s−1; white background) and 3 min of darkness (gray background). Shown are the concentrations of carbon dioxide (CO2; red), hydrogen (H2; green), and oxygen (O2; blue) in the absence inhibitors or treatments (NT; A) or in the presence of GOx (B). Using the concentration data, the rates (production or consumption) are plotted in the absence of inhibitors or treatments (NT; C) or the presence of GOx (D). A to D present an average of cycles 2 to 10. The results show an average of three biological replicates.

Carbon Fixation Outcompetes Hydrogen Production

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show that in the absence of reducing equivalents, car-bon dioxide is not fixed (observed as rapid carbon ac-cumulation). Then, as carbon fixation initiates, the rate decreases and the net rate turns negative. Remarkably, the increase in carbon fixation is in line with the decline in the rate of hydrogen resulting from hydrogen uptake (Fig. 3, A and C), hence implying that diversion of elec-trons toward carbon fixation (expressed here as carbon dioxide uptake) shifts hydrogenase activity toward a mode of hydrogen uptake. These results emphasize the role of the competition between the different metabolic pathways. To confirm the involvement of carbon fixa-tion in the fast cessation of hydrogen production, we added the CBB cycle inhibitor glycolaldehyde (GA), a known inhibitor of phosphoribulokinase (Roberty et al., 2014). In the presence of GA and under 370 µE m−2 s−1, we observed a stable rate of hydrogen production that did not cease at any point (Supplemental Fig. S5). However, at this irradiance, oxygen accumulation was inferior owing to the fact that inhibition of CBB slows down photosynthesis because of acceptor side limita-tion. This fact forms a condition in which the oxygen pool is smaller and cannot be an effective electron sink. To validate that the stable rate of hydrogen production observed in the presence of GA resulted from CBB in-hibition and was not an outcome of a lessened electron loss to oxygen uptake, i.e. Mehler reactions or chloro-respiration, etc., we examined the kinetics of hydrogen evolution in light conditions that feature roughly the same oxygen accumulation. To that aim, we increased the irradiance for the GA treated cells to 1,500 µE m−2 s−1. Then, we compared the accumulation of oxygen (Fig. 4A) and hydrogen (Fig. 4B) between the first cy-cle of either GA-treated cells (GA, 1,500 µE m−2 s−1) or the untreated cells (NT, 370 µE m−2 s−1). While oxygen accumulation was roughly at the same levels (with a slight advantage for the GA treated cells), hydrogen ac-cumulation was linear and did not stop in the presence of GA. This lack of inhibition was reflected by a stable rate of hydrogen production observed only for the GA treated cells (Fig. 4C). Furthermore, the amount of hy-drogen that was accumulated in the presence of GA was higher than in the NT cells (Supplemental Fig. S2), thus reflecting a larger routing of reducing equivalents toward hydrogen production on the account of carbon fixation. Interestingly, in the presence of the CBB cycle inhibitor GA, the hydrogen uptake activity was elim-inated. Finally, since carbon dioxide concentration in-creased substantially following the addition of GA, its emission/fixation rates could not be determined.

Hydrogenase Activity Resumes upon Light Increase

The quantification of hydrogenase showed that its entire pool remains intact following 4 min of illumina-tion (Fig. 2A), after which it declines gradually. We also provide clear evidence that supports the claim that the loss of electrons toward carbon fixation affects the rapid decline of hydrogen production. This led us to

hypothesize that supplying additional reducing equiv-alents to the idle hydrogenase pool would resume its activity. To study this, following 2 min of illumination (at an irradiance of 370 µE m−2 s−1), the irradiance was increased to extremely high light (1320 µE m−2 s−1) for an additional 1 min (Fig. 5A), after which the light was turned off and anaerobiosis was reestablished (4 min of dark acclimation). This scheme was repeated for 10 cycles (Supplemental Fig. S6). The sudden increase in light intensity, which supplied additional electrons, re-generated the accumulation of hydrogen. This observa-tion implies that the hydrogenase pool is indeed active

Figure 4. Inhibition of the CBB cycle supports linear hydrogen produc-tion regardless of oxygen accumulation. C. reinhardtii wild-type strain cc-124 cells were incubated for 1 h in the dark (gray background), after which they were exposed to 2 min of illumination (white background). The concentration of oxygen (A) and hydrogen (B) was measured in the absence of inhibitors or treatments (NT, dark, solid) or the presence of the CBB inhibitor GA (light, dashed). Since the addition of GA hinders electron flow, and therefore oxygen accumulation, the irradiance that was provided was set to an intensity in which the same levels of oxy-gen will be accumulated (370 µE m−2 s−1 for NT versus 1,500 µE m−2 s−1 for GA). C, The rates of hydrogen evolution were plotted as a function of time. The results show an average of three biological replicates.

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but idle. The phenomenon was repeated at each cycle, although the oxygen concentration rose to 12 µm (3-fold higher than the concentration at the point of light increase; Fig. 5A). In addition, chemical determination of the hydrogenase pool confirms that it maintained intact under theses consecutive cycles (Supplemental Fig. S6). Since increasing the light intensity after 2 min at an irradiance of 370 µE m−2 s−1 resumed hydrogenase activity, we extended the initial exposure time (at an irradiance of 370 µE m−2 s−1) to either 4, 7, or 15 min, af-ter which the irradiance was increased to an extremely high light (1320 µE m−2 s−1), this time without cycles (Fig. 5B). Light increase following either 4 or 7 min of exposure resulted in higher oxygen levels (a 10-fold rise following an exposure of 7 min) and, as expected, lowered the hydrogen production rates. However, fol-lowing an illumination period of 15 min (at an irradi-ance of 370 µE m−2 s−1) as aerobiosis was established, hydrogenase activity did not resume (Fig. 5B).

DISCUSSION

Scaling up hydrogen production via hydrogenase is hindered due to two main challenges: oxygen inhibi-tion and inefficient electron transfer to hydrogenase. To study the relations between these two, we exposed cells to a scheme consisting of a dozen light/dark cy-cles. If oxygen accumulation during illumination was indeed destructive to hydrogenase, a gradual decrease in hydrogen production rate from one cycle to another would be expected (Fig. 2B). Nonetheless, it was ev-ident that all the cycles featured a similar increase in hydrogen production. Furthermore, chemical deter-mination of the hydrogenase pool showed that it was intact from cycle to cycle (Fig. 2B). Thus, in this work, we provide conclusive evidence that the fast cessa-tion of hydrogen production due to electron loss for carbon fixation after 2 min of illumination precedes the inactivation of the hydrogenase pool by oxygen after 4 min of illumination. Still, a residual hydroge-nase pool is observed even after 10 min. Accordingly, we found that while hydrogen production comes to a complete halt following 2 min of illumination (Fig. 2A), the active hydrogenase pool remains intact and supports hydrogen uptake (Fig. 2A). This implies that once electron transport to hydrogenase stops, hydro-gen production ceases and shifts toward hydrogen up-take. Indeed, it was previously suggested that at the transition from dark anaerobic environment to light, the functionality of hydrogenase is hindered first due to inefficient electron transfer and only later by inac-tivation due to oxygen (Yacoby et al., 2011; Godaux et al., 2015). However, the identity of the primary path-way that is responsible for this inhibition was vague. Accordingly, in all cycles, the increasing rates of car-bon dioxide uptake coincide with a gradual cessation of hydrogen production (Fig. 3). By blocking the CBB cycle using GA, hydrogen production did not stop and was maintained roughly at a steady rate (Fig. 4C;

Supplemental Fig. S5). Furthermore, our data also showed that during the cycle scheme (Fig. 3B), oxygen was not functioning as a major electron acceptor that can outcompete hydrogenase. Thus, our data indicate that the CBB cycle is the prominent element that out-competes hydrogen production within the transition from dark anoxia to light.

Given the electron loss to carbon fixation, one would expect that resupplying electrons to hydrogenase would renew hydrogen production. Indeed, overflow-ing the system with additional reducing equivalents by suddenly increasing the illumination (from 370 to 1,320 µE m−2 s−1) renewed hydrogen production, al-though only for a short duration (Fig. 5, A and B, light green), supporting the notion that other electron sinks are preferred. In addition, although the active hydro-genase pool size remained constant throughout 4 min of illumination, we see a plasticity in electron transfer to hydrogenase, which shrinks rapidly as the period of low light treatment increases before an increase in illumination, for example, following 2 and 4 min under

Figure 5. Increased irradiance renews hydrogen production. C. reinhardtii wild-type strain cc-124 cells were incubated for 1 h in the dark (gray background). A, Average of 10 cycles. Following an initial period of exposure of 2 min at irradiance of 370 µE m−2 s−1 (white background), after which the irradiance was increased to 1,320 µE m−2 s−1 (yellow background). Shown are dissolved oxygen concentrations (blue) and hydrogen rates (production or consumption, green). B, Following an initial period of exposure at irradiance of 370 µE m−2 s−1 for periods of 2, 4, 7, or 15 min, the irradiance was increased to 1,320 µE m−2 s−1 without cycles. Shown are the initial hydrogen production rates at the onset of light (370 µE m−2 s−1; dark green) and at light increase (1,320 µE m−2 s−1; light green). In each of the above, oxygen concentrations were simultaneously measured at the light increase point (blue). Error bars indicate se (n = 3).

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370 µE m−2 s−1 (Fig. 5B, light-green bars). Interestingly, the partial irreversible inactivation of the hydrogenase pool after 7 min was reflected as a lower hydrogen production rate. Eventually, a complete irreversible in-activation of the hydrogenase pool after 15 min of con-tinuous irradiation at 370 µE m−2 s−1 stopped hydrogen production.

Based on our observations, we can safely state that within 4 min of illumination (following dark anoxia), hydrogen production kinetics studies could elucidate the proportion of electron divergence to the vari-ous competing processes. In addition, hydrogenase activity could be used to monitor fast structural re-modeling of the photosynthetic apparatus. Structural remodeling occurs following an hour of anaerobiosis, as the CEF super complex, consisting of cytochrome b6f and photosystem I, is formed (Iwai et al., 2010). We suggest that the spatial proximity of cytochrome b6f and photosystem I within the CEF supercomplex could support higher electron transfer rate to/from photosystem I, due to a shorter distance that the elec-tron mediator plastocyanin needs to travel. To sup-port this hypothesis, we report here that during the first cycle of illumination, the initial hydrogen pro-duction rate was 3-fold higher than its sequential cy-cles (Fig. 2B; Supplemental Fig. S2). In addition, an enhanced carbon dioxide uptake was also observed only in the first cycle of illumination. Remarkably, the following cycles did not trigger any further changes in the kinetics of these gases, thus indicat-ing that the CEF supercomplex disassembles within less than 2 min of irradiance. More support for this was observed from initial assessments of a mutant having a perturbed CEF complex, i.e. proton gra-dient regulation 5 or 1 (pgr5/pgrl1; Mosebach et al., 2017). There, we observed no difference between the first cycle (20.9 ± 5.5 μmol H2 mg Chl−1 h−1) and the second cycle (19.8 ± 5.4 μmol H2 mg Chl−1 h−1). Still, further biochemical and structural work is needed to provide additional evidence for this hypothesis. Al-ternatively, two additional scenarios can explain this enhanced rate of hydrogen production: (1) Given that ferredoxin NADP+ oxidoreductase (FNR) binding to photosystem I takes place in the wild type (Andersen et al., 1992; and is diminished in the pgr5/pgrl1 mu-tant) and assuming that the majority of the NADPH pool is reduced during the dark anoxia, at light onset, the FNR could act reversibly, oxidizing the NADPH pool while reducing Fd, which in turn supports high-er rates of hydrogen production. (2) The decrease in initial hydrogen production rate agrees with the fact that following an hour of dark incubation most of the downstream sinks, with the exception of hydroge-nase, are deactivated (Terashima et al., 2010; Godaux et al., 2015). Accordingly, the initial exposure to light triggers the activation of various electron acceptors and therefore increases the competition. However, this cannot explain the accelerated carbon dioxide uptake observed in the first cycle.

CONCLUSION

In this work, we provide clear evidence that hydro-genase ceases to produce hydrogen 2 min following light onset, much earlier than damage to its active pool occurs. Using several techniques, we show that the hydrogenase pool is intact up to 4 min in light and some partial pools are observed even after 10 min of illumination. Furthermore, we provide conclusive evidence showing that electron loss to competing processes, prominently carbon fixation, not only elim-inates the positive hydrogen accumulation but also shifts hydrogenase activity toward hydrogen uptake. Therefore, bypassing the electron competition is the preferred path for engineering a continuous process of hydrogen production. Indeed, recent reports show that blocking the CBB cycle either by substrate lim-itation (Nagy et al., 2018) or avoiding its activation (Kosourov et al., 2018) supports continuous hydro-gen production. In addition, preliminary engineering attempts (Nielsen et al., 2013; Wittenberg et al., 2013; Rumpel et al., 2014; Eilenberg et al., 2016; Mellor et al., 2016) provide promising results and in our opinion, should be incorporated into a “super mosaic” single algae clone.

MATERIALS AND METHODS

Cell Growth and Experimental Conditions

Chlamydomonas reinhardtii wild-type strain CC-124 (mt- [137C]) was cul-tivated in 50 mL TAP (Tris/acetate/phosphate) medium kept in Erlenmeyer flasks capped with a silicone sponge. Cells were grown under constant irra-diation of 100 μE m−2 s−1 at 24.5°C and stirring until they reached early log phase (2–5 μg Chl mL−1). Chl was extracted and measured according to Jeffrey and Humphrey (1975). The cells were then centrifuged at 3,300g for 2 min and resuspended to a final concentration of 15 μg Chl mL−1 in a medium containing TAP, 50 mm HEPES and 2 mm Na2CO3 at pH 7.8.

Gas Concentration Measurements

To determine gas concentration within a liquid culture, we transferred the cells into a 5-mL quartz cuvette that was attached to a MIMS, as previously described (Liran et al., 2016). The measured masses were, 2, 32, 40, and 44 m/z, which correlate to the molecules of H2, O2, Ar, and carbon dioxide, respectively. The traces of hydrogen and oxygen were analyzed as described (Liran et al., 2016). The conversion factor of the carbon dioxide trace/concentration was evaluated using a free cell medium purged with carbon dioxide, and its solu-bility was calculated according to Weiss (1974). In order to generate anaerobic conditions, the cells were kept in the sealed MIMS cuvette for a period of 1 h. For each experiment, the cuvette was exposed to red actinic light supplied by a Dual Pulse Amplitude Modulated Fluorometer (DUAL-PAM-100; Walz). The modules used were DUAL-DR and NIR. Light intensity was determined by a Walz light detector (model US-SQS/L) attached to a Li-250A light meter (LI-COR Biosciences). If stated, GA was added 10 min prior to light exposure according to Anderson et al. (2007). When examining the effect of oxygen ac-cumulation on the hydrogen production phenotype, we added to the cuvette: Glc oxidase (200 units mL−1), catalase (200 units mL−1), and Glc (50 mm) accord-ing to Roberty et al. (2014) at the beginning of the measurement.

Rate Analysis

To assess the rate in which the concentration of hydrogen, oxygen, and carbon dioxide was changing during the measurement, we analyzed the de-rivative of the concentration trace in time [μmol gas L−1 min−1] using Microsoft

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Excel software. We used a LINEST function for a frame of four points and implemented it as a rolling fit for all measured points. The derivative was nor-malized to the Chl concentration in order to convert the units to [μmol gas mg Chl−1 h−1]. Each measurement was repeated three times where each time point was averaged. se was calculated using the STDEV function divided by √3. Average of hydrogen rate or oxygen concentration was calculated using a least square analysis for the relevant five points as stated.

Chemical Measurement of the Intrinsic Hydrogenase Activity

Chemical reaction buffer (100 mm Tris-HCl, pH 7.2, 1 m NaCl, 10 mm meth-yl viologen, 20 mm sodium dithionite, and 0.2% [v/v] Triton X-100) was pre-pared in an anaerobic glove box (H2/N2). The buffer was divided into sealed 6-mL vials in a total volume of 1 mL buffer per vial. The sealed vials were then removed from the anaerobic glove box and purged with argon for 3 min in order to remove residual hydrogen traces. Fifty microliters of cell culture was sampled from the MIMS cuvette and injected into a vial and immediately purged with argon for 3 min to remove any leftover hydrogen contamination. The vials were then incubated at 50°C in a water bath for 30 min. The concen-tration of hydrogen in the vial’s headspace was measured by a Hewlett-Packard 5890 Series II gas chromatograph. For each measurement, we sampled 500 µL of gas. The chemical activity of the active hydrogenase was determined in [µmol H2 mg Chl−1 h−1]. The amount of active enzyme was calculated as described by Eilenberg et al. (2016), based on the enzyme’s specific activity that was previ-ously determined in vitro and was reported by Yacoby et al. (2011).

Accession Numbers

Sequence data from this article can be found in the GenBank/EMBL data libraries under the following accession numbers: hydrogenase (HydA 1 and 2), AAL23572.1 and AAL23573.1, respectively; Fd, P07839.2; and FNR, P53991.1.

Supplemental Data

The following supplemental materials are available.

Supplemental Figure S1. Raw data of hydrogen injection into a cell-free medium.

Supplemental Figure S2. Hydrogen accumulation in each light/dark

cycle.

Supplemental Figure S3. Hydrogen traces measured under autotrophic

versus mixotropic conditions.

Supplemental Figure S4. Active hydrogenase pool size in the presence of

Glc oxidase.

Supplemental Figure S5. Hydrogen production rates in the presence of

glycolaldehyde at 370 µE m−2 s−1.

Supplemental Figure S6. Hydrogen and oxygen concentrations with in-creased light intensity.

ACKNOWLEDGMENTS

We thank Iddo Weiner, Meital Avitan, and Marina Kozuleva for carefully reading the manuscript. The MIMS equipment was purchased using a kind donation from the Australian friends of Tel Aviv University.

Received February 21, 2018; accepted May 11, 2018; published May 21, 2018.

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