Microoxic Niches within the Thylakoid Stroma of
Air-Grown Chlamydomonas reinhardtii Protect
[FeFe]-Hydrogenase and Support Hydrogen
Production under Fully Aerobic Environment1[OPEN]
Oded Liran 2, Rinat Semyatich 2, Yuval Milrad, Haviva Eilenberg, Iddo Weiner, and Iftach Yacoby*
Department of Molecular Biology and Ecology of Plants, The George S. Wise Faculty of Life Sciences, Tel Aviv
University, Ramat Aviv, Tel Aviv 69978, Israel
ORCID IDs: 0000-0002-6163-1469 (R.S.); 0000-0003-0177-0624 (I.Y.).
Photosynthetic hydrogen production in the microalga Chlamydomonas reinhardtii is catalyzed by two [FeFe]-hydrogenase isoforms,
HydA1 and HydA2, both irreversibly inactivated upon a few seconds exposure to atmospheric oxygen. Until recently, it was thought
that hydrogenase is not active in air-grown microalgal cells. In contrast, we show that the entire pool of cellular [FeFe]-hydrogenase
remains active in air-grown cells due to efficient scavenging of oxygen. Using membrane inlet mass spectrometry, 18O2 isotope, and
various inhibitors, we were able to dissect the various oxygen uptake mechanisms. We found that both chlororespiration, catalyzed
by plastid terminal oxidase, and Mehler reactions, catalyzed by photosystem I and Flavodiiron proteins, significantly contribute to
oxygen uptake rate. This rate is considerably enhanced with increasing light, thus forming local anaerobic niches at the proximity of
the stromal face of the thylakoid membrane. Furthermore, we found that in transition to high light, the hydrogen production rate is
significantly enhanced for a short duration (100 s), thus indicating that [FeFe]-hydrogenase functions as an immediate sink for
surplus electrons in aerobic as well as in anaerobic environments. In summary, we show that an anaerobic locality in the chloroplast
preserves [FeFe]-hydrogenase activity and supports continuous hydrogen production in air-grown microalgal cells.
Photosynthetic hydrogen production is limited to
microbial photosynthetic organisms, such as Chlamydomonas reinhardtii, and is absent from terrestrial plants
(Prince and Kheshgi, 2005). The reasons for this absence
are vague, but they could be related to differences in
the growth environments. Microalgae naturally reside
in ponds, wet soils, and lakes (Clowez et al., 2015), and
have a tendency to swim into deeper water layers as a
defense mechanism against intense light. Hence, they
occasionally face fluctuating levels of oxygen in their
1
This work was supported by Israel Science Foundation-iCORE
(757/12, “Comprehensive understanding and modeling of plant responses to multiple abrupt abiotic stresses and to prolonged climatic
changes”).
2
These authors contributed equally to the article.
* Address correspondence to [email protected].
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 ([email protected]).
R.S. discovered the aerobic hydrogen production and performed
the westerns; O.L. and I.Y. conceived the original research plans; O.L.
performed most of the experiments and analyzed all the data; Y.M.
conceived and performed the CO experiments, and participated
in the westerns and oxygen consumption experiments; H.E. conceived the western protocol and provided technical assistance; I.W.
analyzed the RNAseq data; I.Y. conceived the project and wrote the
article with contributions of all the authors.
[OPEN]
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264
proximate liquid or soil surroundings. In contrast, terrestrial plants have a constant atmospheric oxygen concentration in their surroundings. Thus, the difference in
oxygen concentration might be the selective pressure for
preservation or loss of the hydrogen-producing enzyme
[FeFe]-hydrogenase in microalgae and terrestrial plants,
respectively.
Microalgal hydrogen production is catalyzed anaerobically by the oxygen-sensitive [FeFe]-hydrogenase
(Ghirardi et al., 1997; Stiebritz and Reiher, 2012),
which is reduced by ferredoxin, a multipotent electron
carrier, and functions in three pathways. The first takes
place in the dark, via a fermentative route, and involves
pyruvate ferredoxin reductase (Dubini et al., 2009).
The second is the photosynthetic route, which takes
place in light at the stromal side of the photosystem I
complex (PSI; Winkler et al., 2010). PSI is reduced by
linear electron flow (LEF) that originates at the photosystem II complex (PSII). PSII absorbs and uses light
energy to split water molecules into electrons, protons,
and oxygen. Electron transport through the cytochrome
b6f complex (cytb6f) facilitates proton pumping from
the stroma into the lumen, thus maintaining a proton
gradient necessary for ATP production through ATPsyntase. In a third pathway, LEF originating at NDAPH
dehydrogenase complex type II (NDA2) receives electrons from starch breakdown and reduces the plastoquinone pool bypassing water splitting at PSII (Jans
et al., 2008). Here, as in the formers, ferredoxin mediates
electron flow to [FeFe]-hydrogenase.
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The Microalgal Thylakoid Contains Microoxic Niches
Figure 1. Schematic illustration of the oxygen uptake mechanism in the chloroplast. Water splitting at PSII generates molecular
oxygen, protons, and LEF. Electrons are transferred from PSII to the plastoquinone pool (QA; QB) from which the electron flow is
transferred either to oxygen uptake in PTOX or to cytb6f. From cytb6f, electrons are transferred to plastocyanin (PC), which in turn
reduces PSI that consumes oxygen directly via the Mehler reaction. The inhibitor DCMU specifically blocks electron output from PSII
to the plastoquinone pool. Hence, upon addition of DCMU, the chloroplast oxygen uptake is omitted, without affecting mitochondrial
respiration (not shown). The inhibitor DBMIB specifically blocks electron transfer output from cytb6f, inhibiting oxygen consumption
by the Mehler reaction, catalyzed by PSI, without affecting PTOX chlororespiration. n-PG is a specific inhibitor of PTOX.
Chloroplast oxygen uptake through chlororespiration, illustrated in Figure 1, takes place upstream to PSI
and involves plastid terminal oxidases (PTOX1 and
PTOX2; Houille-Vernes et al., 2011). In addition, chloroplast oxygen uptake through the Mehler reaction
takes place directly at PSI (Mehler, 1951a; Asada, 2000)
or via ferredoxin resulting in water release. Recently, an
additional Mehler-like reaction was found. This reaction
is carried out by FLVA and FLVB, two newly isolated
diiron flavoproteins, which are reduced by NADPH and
convert oxygen directly into water (not shown in the
scheme; Jokel et al., 2015).
Until recently it was accepted that active [FeFe]hydrogenase in C. reinhardtii is absent in air-grown
cultures, and its accumulation is induced only upon
anoxia, with a maximal accumulation observed after 3 h
of nitrogen sparging (Happe et al., 1994) or 72 h of
sulfur deprivation (Winkler et al., 2002). Accordingly,
atmospheric levels of oxygen completely inactivate the
catalytic site of [FeFe]-hydrogenase within a few seconds (Ghirardi et al., 2007). In contrast, Kosourov and
Seibert (Kosourov and Seibert, 2009) observed continuous hydrogen production from C. reinhardtii cultures
embedded in films under ambient atmosphere, although
the oxygen levels within the films were not determined.
In further support of this, recent reports have suggested
that [FeFe]-hydrogenase is capable of aerobic activity in
strains of Chlorella. It was shown that Chlorella vulgaris
could maintain hydrogen production under atmospheres of 21% (Hwang et al., 2014) or 15% (Chader et al.,
2009) oxygen. The active pool of hydrogenase was estimated at ;30 units per mg of dry weight (Hwang et al.,
2014). Last, in a recent paper by Godaux et al. (Godaux
et al., 2015), the authors claim that in a transition from
dark anoxia to light, high rate of hydrogen production
decreased to lower rate, before the onset of oxygen
accumulation. Hence, they conclude that a competition
for electrons with downstream processes such as FNR
and CEF govern the hydrogen production rate rather
than oxygen concentration. All of these findings contradict the common view since the oxygen sensitivity of
[FeFe]-hydrogenase is well documented (Ghirardi et al.,
1997). Furthermore, it is unclear why the transcripts of
C. reinhardtii hydrogenase and its maturases, Hyd EF
and G, are present under aerobiosis and increase under anaerobiosis induced by sulfur deprivation by
roughly 20%, as we analyzed from published RNAseq
data (González-Ballester et al., 2010).
The mechanism underlying [FeFe]-hydrogenase activity in air-grown cultures is yet to be resolved. We
have therefore used the model organism C. reinhardtii,
cultivated in air, to examine whether an active pool
of [FeFe]-hydrogenase indeed exists, as well as to explore the mechanism(s) protecting [FeFe]-hydrogenase
under aerobic conditions. We present evidence showing hydrogen production for durations longer than
expected in strictly aerobic cultures of C. reinhardtii. We
further demonstrate the coupling of aerobic hydrogen
production with the photosynthetic electron transport
chain and dissect the mechanisms protecting [FeFe]hydrogenase activity under aerobic conditions.
RESULTS
Aerobic Hydrogen Production
To study steady-state hydrogen production rate, 50-mL
suspensions of wild-type C. reinhardtii (strain CC-124) at
2.5 mg (chl)/mL were cultivated in aerated 100-mL flasks
(Fig. 2A, inset) with constant stirring and under three
light intensities (77, 155, and 600 mE m22 s21; hereafter
mE). To ensure full aerobiosis in these conditions, we
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Liran et al.
Figure 2. Photosynthetic activity and quantification of [FeFe]-hydrogenase in air-cultivated C. reinhardtii cells. A, Steady-state hydrogen
production rates measured by MIMS in wild-type cultures cultivated
under three light intensities: 77, 155, and 600 mE. Inset, Cells were
grown in flasks stoppered with a sponge completely permeable to air,
with constant light and stirring. B, LEF from PSII to hydrogen recorded
upon transition from 77 mE to 1200 mE of either wild-type cells in the
presence (broken green line) or absence (solid green line) of the PSII
donor side inhibitor DCMU. The mutant hydEF-1 was used as negative
control (black). C, Inhibition of hydrogenase by the inhibitor CO shuts
down hydrogen production (green bar) in air-grown cells upon transition from 77 mE to 1200 mE at the onset of the recording. Photosynthetic
oxygen production was not affected by the addition of CO (blue bars).
D, Quantification of [FeFe]-hydrogenase in cells cultivated aerobically
under 77, 155, and 600 mE. Top, An immunoblot performed using antiHydA antibody. Purified [FeFe]-hydrogenase (5 ng of HydA) was used as
marker and reference for band intensity quantification. Equal amounts
of total protein were loaded in each lane. [FeFe]-hydrogenase quantities
were normalized either to ng HydA per mg total protein (brown), shown
in the left y axis, or to ng HydA per 1 million cells (orange), shown in the
right y axis. The cultivation light intensities are shown at the bottom of
each lane. All experiments were carried out in triplicates.
quantified the concentration of dissolved oxygen using
oxygen electrode and Winkler reaction (Winkler, 1888).
Furthermore, we analyzed the headspace gas using a gas
chromatograph. Growth at all irradiances showed
constant atmospheric levels of oxygen in the headspace of the growth vessels as well as ;250 mM dissolved oxygen in the growth media (Supplemental
Fig. S1); the same oxygen concentration was measured
for the positive control—a cell-free aerobic growth
media.
Hydrogen production and oxygen production/
uptake rates were investigated using a membrane
inlet mass spectrometer, which monitors multiple gas
traces in real time and thus can discriminate between
isotopes of the same gas. In this study we simultaneously analyzed the traces of H2 (Mus et al., 2005),
stable oxygen isotopes 16O2 and 18O2, and Ar (Radmer
and Kok, 1976; Kana et al., 1994; Tchernov et al., 1997).
We found that the wild-type strain (Fig. 2, A and B), but
not the negative control, a hydEF-1 mutant lacking
[FeFe]-hydrogenase activity (Fig. 2B), exhibited hydrogen
production rates that were inversely dependent on light
intensity (Fig. 2A).
To ascertain the source of electrons responsible for
hydrogen production, we inhibited the LEF from water,
the electron donor of PSII, to [FeFe]-hydrogenase, using
dichlorophenyl dimethyl urea (DCMU), which blocks
electron output from PSII (schematically illustrated in
Fig. 1). We observed no hydrogen production upon
addition of DCMU at growth light intensity (77 mE) that
was increased to 1200 mE at the onset of measurement,
indicating that LEF was the principal electron source for
hydrogen production (Fig. 2B).
To validate that the observed hydrogen production is
indeed catalyzed by the [FeFe]-hydrogenase enzyme
and that the enzyme pool is at the active, oxygensensitive “ox state” (Adamska-Venkatesh et al., 2014),
we added the [FeFe]-hydrogenase inhibitor CO at
growth light intensity (77 mE) that was increased to
1200 mE at the onset of measurement. CO blocks the
active site of the hydrogenase, forming an Hox-CO state
(Lubitz et al., 2014). A complete inhibition of hydrogen
production was observed, while photosynthetic oxygen
evolution was uninterrupted (Fig. 2C).
Cellular Content of Hydrogenase
Previous studies using Chlamydomonas detected both
[FeFe]-hydrogenase transcripts (Forestier et al., 2003;
González-Ballester et al., 2010) and protein (Forestier
et al., 2003; Hemschemeier et al., 2008) in air-grown
microalgal cells, but did not detect [FeFe]-hydrogenase
activity under these conditions. As shown above, our
measurements revealed a steady rate of hydrogen production in air-grown Chlamydomonas cells. To further
support this observation, we determined the cellular
content and activity of [FeFe]-hydrogenase pools in
microalgal samples directly withdrawn from the aerated
cultivation flasks. [FeFe]-hydrogenase contents were
analyzed by immunoblot using a commercial antibody
against HydA1 and revealed ;6 fmol of enzyme per
1 3 106 cells in the irradiance range tested (Fig. 2D;
Supplemental Fig. S2). Since the content of chlorophyll
and protein per cell is modulated under different irradiance, [FeFe]-hydrogenase content was normalized per
cell. The [FeFe]-hydrogenase pool per cell slightly decreased above irradiance of 77 mE. To further study the
observed [FeFe]-hydrogenase activity independently of
photosynthetic electron transfer, which could be limiting, we lysed the cells and used a chemical electron donor. Cell extracts were exposed to an excess of reduced
methyl viologen (MV), which serves as an artificial
electron donor for [FeFe]-hydrogenase (Meuser et al.,
2012). [FeFe]-hydrogenase activity was observed in all
cultures within the irradiance range tested (Fig. 3A, orange bars). Interestingly, net oxygen production under
steady growth light intensity was observed only when
light irradiance reached 600 mE (Supplemental Fig. S3).
This increase had a direct impact on [FeFe]-hydrogenase,
as indicated by the drastic decrease in its activity (Fig.
3A, orange bars). Still, [FeFe]-hydrogenase activity was
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The Microalgal Thylakoid Contains Microoxic Niches
Figure 3. [FeFe]-hydrogenase activity under fluctuating light in air-cultivated C. reinhardtii cells. A, The measured photosynthetic
(green) versus chemical (orange) hydrogen production rates in cells cultivated aerobically (as shown in the inset of Fig. 2A) under
varying irradiance. B, Recorded traces of net hydrogen (green) and oxygen (blue) kinetics as a function of irradiance. The top x axis
shows the light intensity at each time point. Dissolved gases in the samples were measured simultaneously to illumination for
100 s under 77 mE (phase I), followed by a continuous illumination under 1200 mE. The onset of the transition to 1200 mE, lasting
;100 S, where hydrogen production is maximal, is defined as phase II. The remaining period at 1200 mE is defined as phase III. C,
Hydrogen production under fluctuating light. Photosynthetic hydrogen production rates during light irradiance of 77 mE punctuated each 2 min with 3 min of high irradiance (600, 1200, or 2400 mE). Phase II (yellow) and phase III (pink) are shown for each
high light intensity. All experiments were carried out in triplicates. D, Pie graphs showing the divergence of oxygen consumption
between the various mechanisms under the three phases shown in B. The percentages were calculated using the rates shown in
Table I. Hence, while mitochondrial respiration (blue) was constant, 160 [mmol (O2) mg21 h21], PTOX (green) and Mehler reactions (red) were significantly increased in a transition to high light.
not totally eliminated under 600 mE, suggesting that
mitochondrial respiration and chloroplast O2 uptake can
effectively form a microoxic environment at the stromal
side of the thylakoid membrane, even under continuous
growth under high light.
To estimate the fraction of [FeFe]-hydrogenase driven
by photosynthetic electron transfer from the total active
fraction, chemically measured using cell extract and the
artificial electron donor MV, we plotted the in vivo
photosynthetic hydrogen production rate in the light
(Fig. 3A, green bars) versus its chemically measured rate
(Fig. 3A, orange bars; Meuser et al., 2012). Notably, a
large fraction of the active [FeFe]-hydrogenase pool did
not function as an electron sink in air-cultivated cells
(Fig. 3A, green bars versus orange bars), raising the
question as to its function.
Hydrogenase Functions as an Electron Sink in a Transition
to High Light
A sudden increase in irradiance may create an increased electron pressure on the electron transfer chain,
especially when light-dependent oxygen consumption
does not function properly, e.g. under low oxygen. In
such cases, active [FeFe]-hydrogenase might take part
in a first-line defense to immediately dissipate excess
electrons (White and Melis, 2006; Hemschemeier et al.,
2008). We challenged this assumption aerobically by
exposing cells grown under 77 mE (“phase I” in Fig. 3B)
to a continuous intense light irradiance of 1200 mE
while simultaneously determining the hydrogen and
oxygen kinetics. A short-lived burst of hydrogen production (;100 s; “phase II” in Fig. 3B) preceded the
onset of oxygen evolution. Interestingly, following the
100-s burst, the basal hydrogen production rate was
maintained throughout the exposure to high light irradiance (“phase III” in Fig. 3B). To study whether this
phenomenon can continue for a longer extant, we
recorded the kinetics for additional 30 min. We observed
that both hydrogen and oxygen were accumulating simultaneously even when oxygen concentration was as
high as 350 mM (Supplemental Fig. S4).
To further investigate the role of [FeFe]-hydrogenase
under fluctuating light intensities, we measured the
hydrogen production rates in cells cultivated under
77 mE and then exposed to fluctuating light, consisting
of 3 min at high irradiance of 600, 1200, and 2400 mE,
punctured with relaxation periods of 2 min at 77 mE. As
shown in Figure 3C, the hydrogen production rate at
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Liran et al.
phase II (yellow bars) upon each exposure to high irradiance reached up to 6 fmol H2 cell21 h21, which
clearly indicates that the [FeFe]-hydrogenase pool in
microalgae is in a “standby” state, ready to dissipate
excess electrons upon sudden exposure to high light.
Thus, it seems that [FeFe]-hydrogenase provides an
immediate electron sink that chronologically precedes
oxygen accumulation and gains maximal thrust only at
the onset of transition into high light. In addition, net
oxygen production that was simultaneously measured
showed that PSII was active at all light intensities
(Supplemental Table S1). Hence, the increased net oxygen production rates alongside higher irradiance imply
that a mechanism involving strong photoinhibition is
unlikely.
DBMIB alone, when both respiration and chlororespiration are active (dark controls are shown in
Supplemental Fig. S5). To further validate if the chlororespiration can be attributed to PTOX activity, DCMU
was replaced with n-propyl gallate (n-PG). Hence, we
subtracted the 18O2 uptake rate measured in the presence of DBMIB + n-PG from the rate measured in the
presence of DBMIB. As shown in Table I, chlororespiration and 18O2 uptake rate by PTOX are highly similar,
supporting the notion that PTOX is the principle oxygen consumer upstream to PSI, notably, the measured
PTOX rates under steady state are in agreement with
Cournac et al. (2002). However, the rate of chlororespiration significantly increased upon transition from
medium (77 mE) to high light (1200mE; Table I), compensating for the increased oxygen production by PSII.
Finally, we analyzed the oxygen uptake by Mehler and
Mehler-like reactions. The Mehler contributions were
calculated by subtracting the chlororespiration rate
from the total chloroplast 18O2 uptake rate that was
determined by subtracting mitochondrial respiration in
light (18O2 uptake in the presence of DCMU) from the
gross oxygen consumption in light. Here, as for PTOX, a
significant enhancement in the Mehler reaction rate was
observed upon transition from medium (77 mE) (Table
I, phase I) to high light (1200 mE; Table I, phase II),
compensating for the increased oxygen production
by PSII. At later time points (phase III), the Mehler rate
was decreased, while oxygen consumption by PTOX
remained constant (Table I, phase III), resulting in a net
oxygen release (Table I, phase III). It is notable that in
transition from medium (77 mE) to high light (1200 mE),
as much as 50% of the oxygen produced by PSII was
consumed by the chloroplast oxygen uptake pathways
(Fig. 3D).
Determination of Oxygen Consumption Pathways
The unanticipated hydrogen production in air-grown
microalgae observed under both stable and fluctuating
light intensities suggests efficient oxygen consumption
at the stromal side of PSI, where [FeFe]-hydrogenase
naturally resides. Oxygen produced upon water splitting at PSII may be consumed by mitochondrial respiration and by chloroplast oxygen uptake (illustrated in
Fig. 1). Chloroplast oxygen uptake takes place before
PSI by chlororespiration driven by PTOX, and at PSI
by Mehler and Mehler-like reactions (Mehler, 1951a,
1951b; Badger et al., 1980; Houille-Vernes et al., 2011;
Jokel et al., 2015). To differentiate between these possibilities, we analyzed the oxygen kinetics during the
three phases of hydrogen production shown in Figure
3B. Since 16O2 is continuously produced under light
by the water-splitting activity of PSII, accurate oxygen
consumption was measured using injections of known
amounts of an 18O2 isotope that is not produced in vivo
(Radmer et al., 1978). To calculate the magnitude
of oxygen uptake by chlororespiration, we subtracted
the 18O2 consumption rate measured in the presence
of 2,5-dibromo-6-isopropyl-3-methyl-1,4-benzoquinone
(DBMIB) + DCMU, when only respiration is active,
from the 18O2 uptake rate measured in the presence of
DISCUSSION
In this work we show that [FeFe]-hydrogenase activity in air-grown Chlamydomonas cultures is evident
due to anaerobic niches within the thylakoid membranes. Still, [FeFe]-hydrogenase could be aerobically
Table I. Oxygen kinetics during the three phases observed following a sharp transition from medium to
high light
Dissection of the independent elements of oxygen kinetics in a transition from irradiance of 77 mE to
1200 mE was conducted in three phases using 18O2 isotope in the presence or absence of the inhibitors
DCMU, DBMIB, and n-PG.
Phase
21
I
II
III
21
Rates [mmol (O2) mg h ] 6 SE
Calculated gross
Net O2 release
Thylakoid total
Chlororespiration (DBMIB + DCMU)
PTOX (DBMIB + n-PG)
Calculated Mehler reactions
Mitochondria dark
Mitochondria light (+DCMU)
93
272.64 6 15.91
211.35 6 9.84
28.7
28.8 6 3.4
22.55
302
285
0.42 6 2
20.85 6 5.5
2148.43 6 33.2 2110.62 6 42.19
264.73 6 12.58
272.83 6 16.84
284
246
2162 6 10
2154 6 34
268
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The Microalgal Thylakoid Contains Microoxic Niches
active due to (1) intrinsic tolerance or (2) fast [FeFe]hydrogenase protein synthesis that could not take
place under strictly aerobic conditions (Happe et al.,
1994; Winkler et al., 2002).
Concerning intrinsic tolerance, the known mechanisms
for partial “oxygen-tolerance” of [FeFe]-hydrogenase
were shown only in vitro and involve reversible inactivation (Vincent et al., 2005; Stripp et al., 2009; Stiebritz
and Reiher, 2012). Yet, our data clearly show that the
cellular [FeFe]-hydrogenase pool is active, as evident
from the increased hydrogen evolution rates during the
transition from low to high light. Furthermore, we show
that the entire [FeFe]-hydrogenase pool is at the CO/O2sensitive ox state since the [FeFe]-hydrogenase activity
was completely blocked by the addition of external CO,
ceasing all hydrogen production at the transition from
77 mE to 1200 mE (Fig. 2C). Thus, we did not observe an
inactive population of [FeFe]-hydrogenase, rendering
the possibility of intrinsic oxygen tolerance unlikely. To
understand how the activity of such an oxygen-sensitive
enzyme is possible in air-grown microalgae, we dissected
the oxygen uptake mechanisms, namely, mitochondrial
respiration and gross chloroplast oxygen uptake, which
consists of chlororespiration catalyzed by PTOX, and
Mehler reactions. We found that the combined oxygen
uptake activity of these pathways keeps the oxygen level
within the stromal face of PSI to a minimum, which enables [FeFe]-hydrogenase activity resulting in hydrogen
production in air-grown microalgae. In support of our
findings of microoxic niches in the chloroplast, one can
observe some similarity to other photosynthetic organisms; for example, the protection of another strictly
anaerobic enzyme, nitrogenase, in the cyanobacteria
Anabaena is carried out by an efficient removal of oxygen
by the Mehler-like reaction. Recently, Ermakova et al.
(2014) showed that Flv3B protects nitrogenase by
performing light-induced O2 uptake, which maintains
microoxic conditions inside of the heterocysts.
Interestingly, we found that a major fraction of the
[FeFe]-hydrogenase pool is idle but active. Hence, its
full capacity is revealed only upon transition from low
to high light. The period of full [FeFe]-hydrogenase
activity is short, around 100 s, suggesting that [FeFe]hydrogenase participates in a first line of defense for the
immediate dissipation of the increased electron flux in
fluctuating light environments. Still, hydrogen production accounted for less than 1% of this initial electron
flux. Thus, it appears that under aerobic conditions, the
conversion of oxygen into water, rather than hydrogen
production, is the prime sink for excess electrons generated under sharp repetitive transitions into high light.
Taken together, these findings suggest that higher plants
have probably not preserved this enzyme due to a constant atmospheric oxygen level in their surroundings,
which enables, under high irradiance, efficient dissipation of excess electrons via PTOX and Mehler reactions.
Thus, the fluctuating oxygen concentrations in the
proximity of microalgae could have formed an evolutionary pressure to preserve an additional mechanism,
namely, [FeFe]-hydrogenase, to confront instances such
as under low oxygen, where the chlororespiration catalyzed by PTOX and Mehler reactions cannot function
properly.
CONCLUSION
Due to [FeFe]-hydrogenase oxygen sensitivity, we
exploited [FeFe]-hydrogenase as an oxygen reporter.
The presence of [FeFe]-hydrogenase activity in airgrown microalgae indicates a microoxic locality in the
thylakoid stroma, where the enzyme resides.
We show that in air-cultivated Chlamydomonas cells,
[FeFe]-hydrogenase (1) is active due to efficient oxygen
consumption fed by LEF at the stromal side of PSI, and
(2) functions as a minor alternative sink for excess
electrons, especially upon transition to high light.
In relation to futuristic algal hydrogen production,
it is widely accepted that the main barrier of the engineering efforts toward photosynthetic hydrogen
production is thought to be the oxygen sensitivity of
[FeFe]-hydrogenase. Our findings, of the microoxic locality at the thylakoid stroma, might be used as a platform for reengineering of the natural oxygen-scavenging
pathways. Such modifications, if successful, could facilitate a continuous production of hydrogen in air-grown
microalgae.
MATERIALS AND METHODS
Organisms and Growth Conditions
Chlamydomonas reinhardtii wild-type strain CC-124 (mt- [137C]) and the hydEF1 mutant or the hydA1-hydA2 double mutant were cultivated on 1.5% (w/v) agar
plates with Tris-Acetate-Phosphate (TAP) medium. The cultures were maintained
at 24.5°C under 77 mE cool-white light. For all experiments, 50 mL of microalgal
suspension were cultivated aerobically in aerated 100-mL flasks under constant
stirring using a magnetic bar. Cell density was kept below 2.5 mg (chl)/mL.
Determination of Cell Density and
Chlorophyll Concentration
Cell density was determined using a Neubauer hemocytometer. Chlorophyll
was extracted and measured as previously described (Jeffrey and Humphrey,
1975).
Determination of [FeFe]-Hydrogenase Content by
Western Blot
Soluble proteins were isolated from air-grown, midlog-phase C. reinhardtii
(total of ;3 mg chlorophyll). Samples were withdrawn directly from the cultivation flasks and kept at 4°C to prevent further protein synthesis. Cells at 2.5
mg (chl)/mL were precipitated (3,200g, 5 min, 4°C) and resuspended in 600 mL
of buffer A (50 mM Tris-HCl, pH 8.5, 20 mM sodium dithionite, 60 mM NaCl, and
1 mM protease inhibitor cocktail [Sigma]). The cell suspension was lysed using a
Minilys tissue lyser (Bertin Technologies) at two 5000 rpm cycles of 45 s each in
the presence of glass beads (Sigma). The soluble proteins were separated by a
10-min (14,000g, 4°C) centrifugation and further concentrated using Vivaspin
500 (Sartorius) to yield 180 mL at protein concentration of ;1mg/mL. Increasing
amounts (3–10 mL corresponding to 30–100 mg chl) of soluble proteins were
loaded onto 4-12% Bis-Tris Plus PAGE gels (Novex by Life Technologies) and
analyzed by immunoblotting using rabbit polyclonal HydA1/HydA2 antibodies (Agrisera). A known standard of purified HydA1 (5 ng) was mixed with
a lysate of the hydA1-hydA2 double mutant, to mimic protein interference. The
HydA1/2 bands were analyzed by DNR imager and quantified using GelQuant software. The minimal sensitivity was 1 ng of HydA1/2 per lane.
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Copyright © 2016 American Society of Plant Biologists. All rights reserved.
Liran et al.
Chemical Determination of [FeFe]-Hydrogenase Hydrogen
Production Rate
Statistics
Samples were withdrawn directly from the cultivation flasks and pelleted by
centrifugation for 2 min at 4000 rpm. Then they were perforated by Triton/MV
mix as described previously (Meuser et al., 2012). Dissolved hydrogen production was analyzed using membrane inlet mass spectrometry (MIMS).
Each group within the charts is represented as value 6 SE. Comparison
between rates was examined by Kruscal-Wallis a-parametric ANOVA.
Repeated-measurement ANOVA was used when the dependent variable was
measured three times on the same sample and the same experiment (Fig. 2D).
Significance was set at P , 0.05.
Gas-Exchange Profiles of H2 and O2
Supplemental Data
Samples of cell suspensions at 2.5 mg (chl)/mL were centrifuged for 2 min at
4000 rpm at room temperature, then resuspended in fresh TAP + 50 mM HEPES
pH 7.8 (NaOH) to yield a final concentration of 40 mg chl mL21. For analyses,
5 mL of a concentrated sample was introduced to a sealed quartz cuvette
(Starna Cells). A quadrupole mass spectrometer (QMS 200 M1; Pfeiffer Vacuum) was connected to the inlet probe by a vacuum line. The cuvette construct
was then fitted into a metabolic chamber (optical unit ED-101US/MD; Walz),
which kept the sample thermostated at 24.5°C during the experiment. Light was
guided through the light ports of the chamber with a Schott light (KL1500-T)
covered with light-reducing filters to attain final light intensities of 77 to
2000 mE. Masses of H2, N, 16O2, 18O2, and Ar were repeatedly measured using a
0.5-s dwelling time per mass. The measured traces of oxygen were corrected for
the continuous removal of the measured gas by the vacuum line as described by
Luz and Barkan (2005). H2 trace was analyzed as described by Mus et al. (2005).
The following supplemental materials are available.
Supplemental Figure S1. Quantification of oxygen in cell cultures cultivated aerobically under 77, 155, and 600 mE.
Supplemental Figure S2. Quantification of [FeFe]-hydrogenase in cells
cultivated aerobically under 77, 155, and 600 mE.
Supplemental Figure S3. Net measured oxygen kinetics.
Supplemental Figure S4. Prolonged H2 production under 1200 mE.
Supplemental Figure S5. Modulation of dark respiration by chloroplast
inhibitors.
Supplemental Table S1. Net oxygen production in phase III under intermittent light.
Stable Isotope Analysis
Oxygen unidirectional flux rates were determined as described previously
(Radmer et al., 1978), with a slight modification. A small volume of 99% 18O2
(Sigma-Aldrich) saturated TAP solution was injected at time points of interest.
The trace of the isotope was then corrected for the constant MIMS consumption.
Absolute rates were calculated as follows:
Net oxygen production in light ¼ RL32
Gross oxygen consumption in light ¼
RL36
3 F32;36 3 K
We thank Prof. Michael Gurevitch (Tel Aviv University), Alexandra Dubini
(National Renewable Energy Laboratory), and Dr. Yuval Mazor (Tel Aviv
University) for helpful discussion and critical reading of the manuscript. The
MIMS equipment was bought using the generous donation of the Australian
Friends of Tel Aviv University.
Received July 6, 2016; accepted July 18, 2016; published July 21, 2016.
Gross oxygen production in light ¼ RL32 2 RL36 3 F32;36 3 K
Mitochondrial oxygen consumption in dark ¼ RD
32
Thylakoid oxygen consumption in light ¼
ACKNOWLEDGMENTS
RL36 3 F32;36 3 K 2 RD
32
R is the linear regression of mass 32 or 36 in light or dark (L or D, respectively); F
is the enrichment factor taken as the ratio between masses 32 and 36 at the
maximum peak obtained after injection; and K is an additional correction to the
assumption that respiration in the dark should be the same for both masses
(Radmer et al., 1978; Kana, 1990).
Dissection of Chloroplast Oxygen Uptake
Cells at a concentration of 40 mg chl mL21 were kept in a thermostated
(24.5°C) MIMS cuvette under irradiance of 77 mE with constant stirring while
the cuvette was open to air. DBMIB was added to yield a final concentration of
20 mM. The cells were incubated for 2 min and then the cuvette was sealed for the
MIMS measurement. A small volume of 99% 18O2 (Sigma-Aldrich) solution was
injected, and after reaching a recorded mass spectrometer peak of 18O2, additional injections of DCMU (final concentration 10 mM) or n-PG (final concentration 400 mM) were followed. Then, the sample was incubated for an
additional 3 min in light, after which irradiance was increased to 1200 mE for an
additional 3 min.
CO Inhibition
N2 or CO saturated solution (200 mL) of TAP + 50 mM HEPES pH 7.8 (NaOH)
was injected into the MIMS cuvette, which contained cells at a concentration of
40 mg chl mL21 under irradiance of 77 mE, 24.5°C with constant stirring. To
monitor the injection of CO, mass 12 (corresponding to carbon) was measured.
After 8 min of incubation, the light intensity was increased to 1200 mE, and H2
and 16O2 concentrations were measured simultaneously as described above.
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