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Keywords:

  • activation;
  • cyanobacteria;
  • diaphorase;
  • Hox genes;
  • hydrogen

Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Cyanobacterial NAD(P)+-reducing reversible hydrogenases comprise five subunits. Four of them (HoxF, HoxU, HoxY, and HoxH) are also found in the well-described related enzyme from Ralstonia eutropha. The fifth one (HoxE) is not encoded in the R. eutropha genome, but shares homology with the N-terminal part of R. eutropha HoxF. However, in cyanobacteria, HoxE contains a 2Fe–2S cluster-binding motif that is not found in the related R. eutropha sequence. In order to obtain some insights into the role of HoxE in cyanobacteria, we deleted this subunit in Synechocystis PCC6803. Three types of interaction of the cyanobacterial hydrogenase with pyridine nucleotides were tested: (a) reductive activation of the NiFe site, for which NADPH was found to be more efficient than NADH; (b) H2 production, for which NADH appeared to be a more efficient electron donor than NADPH; and (c) H2 oxidation, for which NAD+ was a much better electron acceptor than NADP+. Upon hoxE deletion, the Synechocystis hydrogenase active site remained functional with artificial electron donors or acceptors, but the enzyme became unable to catalyze H2 production or uptake with NADH/NAD+. However, activation of the electron transfer-independent H/D exchange reaction by NADPH was still observed in the absence of HoxE, whereas activation of this reaction by NADH was lost. These data suggest different mechanisms for diaphorase-mediated electron donation and catalytic site activation in cyanobacterial hydrogenase.


Abbreviations
BV

benzyl viologen

MIMS

membrane-inlet MS

MV

methyl viologen

Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Hydrogenases are enzymes that catalyze the reversible oxidoreduction of protons into dihydrogen and vice versa. Three types of hydrogenase are known in nature, and are classified according to the metal content of their active site: [FeFe]-hydrogenases, which have been found in bacteria and eukaryotes, [NiFe]-hydrogenases in archaeons and bacteria, and [Fe]-hydrogenases in some methanogenic archaeons [1]. In hydrogenases, H2/H+ oxidoreduction is associated with various donors or acceptors, depending on the organism and the physiological context in which these enzymes are expressed. Oligomeric, reversible, NAD(P)-linked [NiFe]-hydrogenases constitute a group (group 3d in reference [1]) that is found in several bacteria, including cyanobacteria, carrying out the reaction:

  • image

Because the action of these hydrogenases is reversible, they may, in certain environmental conditions, produce reductive equivalents by oxidizing H2 present in the medium, and therefore consume it in conjunction with, for instance, (anaerobic) respiration or photosynthesis. Conversely, they also have the ability to evacuate reducing power and reoxidize cofactors through H+ reduction (resulting in H2 synthesis) in fermentative conditions, especially in the absence of other electron acceptors. Therefore, they can be envisaged for use in H2 production processes, in particular when they are expressed in environmental conditions (and/or genetic backgrounds) in which the reduction of pyridine nucleotide pool is enhanced [2].

The NAD(P)-linked [NiFe]-hydrogenases share in common four subunits that are conserved among this group: HoxH and HoxY constitute the hydrogenase module, whereas HoxF and HoxU constitute the diaphorase module [3,4]. Although these enzymes were originally thought to be tetrameric, additional subunits have been found. A well-known member of the group is the H2ase from Ralstonia eutropha (Cupriavidus necator), an NAD+-reducing hydrogenase in which two additional HoxI subunits have been discovered. These subunits feature a putative cyclic nucleotide-binding motif, and are essential for NADPH-dependent enzyme activation [5]. In purple bacteria and cyanobacteria, a different subunit named HoxE, which is not related to HoxI, is found, and it has been reported to form part of the hydrogenase complex [6].

Most Hox subunits show homology with components of the NADH-oxidizing respiratory complex I [designated Nuo(x) in Escherichia coli]: HoxF shares homology with NuoF, HoxU with the N-terminus of NuoG, HoxH with NuoD, HoxY with NuoB, and HoxE with NuoE [7]. In R. eutropha, which does not have a hoxE gene, the N-terminus of the HoxF sequence bears a NuoE (and thus a cyanobacterial HoxE) homologous region [3] that is not found in cyanobacterial HoxF [8]. However, the NuoE/HoxE-like domain of HoxF in R. eutropha contains only one conserved cysteine, as compared with four cysteines in NuoE, and may have lost the ability to bind 2Fe–2S clusters. Paradoxically, in the cyanobacterium Synechocystis strain PCC6803, the N-terminus of HoxF does bear a 2Fe–2S-binding motif, although this organism also produces a distinct HoxE subunit that comprises a four-cysteine motif [7]. Therefore, the electron transport chain in Synechocystis hydrogenase might include two supplemental 2Fe–2S clusters when compared with that of the Ralstonia bidirectional hydrogenase.

A cyanobacterial-like HoxEFUYH hydrogenase has been described in the purple bacterium Thiocapsa roseopersicina [9]. In this organism, it has been shown to function reversibly in vivo, catalyzing fermentative H2 production in the dark, light-mediated H2 production in the presence of thiosulfate, and H2 consumption in the light (but not H2 uptake in the presence of O2, which relies on membrane hydrogenase) [10]. Deletion of hoxE resulted in an impairment of hydrogenase-dependent H2 evolution in vivo, but the enzyme retained full activity in vitro with benzyl viologen (BV) as an electron acceptor [9]. Recently, mutants with tagged HoxE have been obtained and the bidirectional hydrogenase has been purified [11], confirming the pentameric composition of the enzyme. It was observed that, in these enzymatic preparations, the diaphorase part of the complex (HoxEFU) tended to dissociate from the hydrogenase catalytic part (HoxYH), as was also found in Allochromatium vinosum Hox hydrogenase [12]. HoxE, HoxF and HoxU have been suggested to also take part in the NAD(P)H-binding activity of cyanobacterial complex I, whose pyridine-binding subunits have not yet been formally identified [7]. However, this involvement has been questioned, as hydrogenase-deficient cyanobacteria do not show any impairment in respiratory capacity [13].

In the present study, we investigated the function of the Synechocystis hydrogenase in the presence and absence of HoxE, together with the presence and absence of another major diaphorase component, HoxF. Simultaneous HoxE and HoxF deficiency severely reduced the amount of hydrogenase, and completely prevented interactions with NADH and NADPH. A different picture was observed with HoxE deficiency alone, which moderately decreased the amount of hydrogenase. Indeed, HoxE was found to be essential for electron transfer between NAD(H) and H2/H+, but the process of NADPH-mediated hydrogenase activation of H/D exchange capacity appeared to be largely independent of HoxE.

Results

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Enzyme expression in modified strains

As expected, no trace of HoxH was detected by western blot in the ΔhoxH strain and no trace of HoxF in the ΔhoxEF strain (Fig. 1). Deletion of hoxE alone had a slight effect on the amounts of HoxH (60%) and HoxF (70%) as estimated from western blot band quantification. Deletion of hoxH also induced some decrease in HoxF abundance; however, this subunit continued to be expressed at a relatively high level (∼ 65% of the wild type). Deletion of the hoxE–hoxF cluster had a very strong effect on HoxH abundance: this subunit remained detectable, but at a very low level (∼ 10% of that in the wild type).

image

Figure 1.  Immunological detection of Synechocystis hydrogenase subunits in soluble extracts. M: molecular weight markers. Lane 1: wild type. Lane 2: ΔhoxH. Lane 3: ΔhoxE. Lane 4: ΔhoxEF. Bands corresponding to HoxH and HoxF are indicated by arrows.

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In order to determine whether differences in hydrogenase subunit amounts could be linked to transcriptional polar effects resulting from antibiotic cassette insertion, quantitative real-time PCR was performed to assay hoxE, hoxF and hoxH expression levels for the strains used in this study (Fig. S1). No difference could be seen between wild-type and mutant strains for these genes, respectively located at the beginning (hoxE and hoxF) and at the end (hoxH) of the hox operon. It therefore appears that the observed differences in protein amounts are most likely attributable to post-transcriptional effects, such as differences in protein translation, maturation, or stability.

Hydrogen exchange and hydrogenase activity in vivo

As previously reported [2], H2 production in the dark was observed in anaerobic conditions for the wild-type strain (in the presence of glucose and glucose oxidase), and a transient stimulation of H2 production was observed upon illumination, rapidly followed by H2 uptake (Fig. 2). Conversely, no H2 evolution could be detected for any of the deletion mutants, indicating that all deleted subunits are essential for proton reduction in vivo. However, when hydrogenase activity was assayed with the H/D exchange assay, which detects turnover of the catalytic site independently of electron transfer (H+ reduction or H2 oxidation), a quite different picture emerged (Fig. 3). In this case, the hydrogenase activities of the wild type and of ΔhoxE appeared to be quite comparable; ΔhoxEF retained faint but detectable activity (∼ 5% of that of the wild type), whereas hoxH deletion totally abolished activity, which is expected, as this subunit is responsible for coordination of the active site. These results are in line with the western blot quantifications of HoxH, indicating that, once hydrogenase catalytic units are synthesized, H/D exchange can be activated in vivo, even if electron transfer is impaired.

image

Figure 2.  Time course monitoring by MIMS of H2 production and uptake during anaerobic incubation of Synechocystis cells, first in the dark and then in the light. WT, wild type.

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image

Figure 3.  Time course monitoring of D2, HD and H2 dynamics during anaerobic incubation of Synechocystis cells. The progressive decrease in the deuterium content of the hydrogen gas (decrease in D2, and increase in HD and then in H2) reveals hydrogenase catalytic activity in vivo. The dotted curve represents the sum of all hydrogen species. WT, wild type.

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Hydrogenase function in cell-free extracts

In order to obtain further insights into the mechanisms of hydrogenase activation and electron transfer, studies of hydrogenase activation and hydrogenase-mediated electron transfer capacities in soluble (cell-free) protein extracts were performed. We first assayed H/D exchange capacities. We observed that fresh cell-free extracts, which are inactive in the presence of O2, as previously reported [2], spontaneously reactivated as soon as they were placed in anaerobic conditions, indicating that some soluble compounds were able to mediate this activation (data not shown). In order to investigate which compounds could be responsible for this reactivation, we let the extract oxidize until spontaneous reactivation was inhibited, and then tested how the enzyme could be activated by physiological and artificial electron donors in vitro [2]. For this, cell-free soluble protein extracts were kept at room temperature for ∼ 3 h before measurements. Extracts were then placed in the mass spectrometer vessel, put under anaerobic conditions with the addition of glucose and glucose oxidase, and activated with reductive compounds. In the presence of reduced methyl viologen (MV), all extracts (except from ΔhoxH) exhibited hydrogenase activity as assayed by H/D exchange. However, activity was much lower in extracts from the ΔhoxEF strain (Fig. 4; Table 1). Thus, H/D exchange rates in the presence of reduced MV were in line with protein amounts as detected by western blotting.

image

Figure 4.  Time course monitoring of hydrogenase activity in cell-free extracts by (log) decrease in isotopic labeling of hydrogen, τ = ([D2] + [HD]/2)/([D2] + [HD] + [H2]). Air-oxidized cell-free extracts were placed in the vessel, D2 was bubbled into the suspension, and anoxia was then imposed by injection of glucose and glucose oxidase/catalase at t = 5 min. NADH (N), NADPH (P) or MV (M) were then injected, as indicated by circled letters. Curves have been shifted incrementally (by 0.05 or 0.1) towards the bottom for better legibility; otherwise, initial isotopic contents were the same in all experiments [close to 1, i.e. Ln(τ) ∼ 0]. WT, wild type.

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Table 1.   H/D exchange rates (nmol·min−1·mg protein−1) catalyzed by soluble extracts from Synechocystis strains. Effect of reductants on previously oxidized (and therefore hydrogenase-inactivated) extracts. Results are expressed as mean ± standard deviation, with n = 3–6.
StrainMVredNADHNADPH
Wild type34.9 ± 7.98.6 ± 2.816.0 ± 0.2
ΔhoxE27.1 ± 3.10.2 ± 0.214.2 ± 5.7
ΔhoxEF1.2 ± 0.90.00.0
ΔhoxH 0.00.00.0

We then investigated the ability of hydrogenase to restore activity in the presence of the physiological electron donors NADH and NADPH (Fig. 4; Table 1). Only the wild type and ΔhoxE enzymes could be significantly activated in the presence of these compounds. Whereas the wild type enzyme could be activated by both (although less efficiently with NADH than with NADPH), the ΔhoxE enzyme could only be activated by NADPH.

As the hydrogenase catalytic site was differentially activated by reducing agents, we investigated whether hydrogenase-mediated electron transfer (i.e. reduction of H+ and oxidation of H2) could also be detected in extracts from the different mutants. H2 production was detected in the presence of reduced MV in the wild type, ΔhoxE (60% versus the wild type), and ΔhoxEF extracts (15% versus the wild type), but NADH or NADPH-mediated production could only be detected in the wild type extracts, NADH being much more efficient than NADPH (Table 2). Simultaneous addition of both substrates did not further increase H2 production in the wild type. In the other direction, BV-driven H2 oxidation (measured by monitoring H2 uptake from the medium) could be observed in wild-type, ΔhoxE and ΔhoxEF extracts (62 ± 44, 18 ± 13 and 0.7 ± 0.5 nmol·min−1·mg protein−1, respectively), which is consistent with previous reports on the Thiocapsa enzyme [9]. NAD+-driven H2 uptake could only be detected in the wild type (4.6 ± 1.5 nmol·min−1·mg protein−1). No significant NADP+-driven H2 uptake could be detected in any of the preparations, including extracts from the wild type.

Table 2.   Hydrogen production rates (nmol·min−1·mg protein−1) catalyzed by soluble extracts from Synechocystis strains. Effects of different electron donors.
StrainMVNADHNADPHNADH + NADPH
Wild type5.44 ± 4.120.48 ± 0.240.08 ± 0.080.55 ± 0.36
ΔhoxE3.39 ± 4.310.00.00.0
ΔhoxEF0.92 ± 1.790.00.00.0
ΔhoxH0.00.00.00.0

In complementary experiments, with the objective of checking the integrity of the hydrogenase fractions in the extracts, we tested enzyme activity with colorimetric MV reduction assays after electrophoresis in native gels (Fig. S2). In our hands, activity spots could only be obtained for wild-type and ΔhoxE extracts from cultures grown in nitrogen-depleted medium. Below the main spot, a smear was observed in the wild-type lane, indicative of the presence of lower-mass proteins (possibly degradation products) retaining activity. The spot in ΔhoxE lane was of lower apparent mass than that in the wild-type lane, which was expected. Its intensity was very weak, although the activity checked before gel loading was quite strong (∼ 25% of that of the wild type in the conditions of the experiment). This finding, which might indicate lower stability of the hydrogenase of ΔhoxE during migration, is in favor of a stabilizing role of HoxE. Unfortunately, we did not succeed in further attempts to determine the association of the activity spots with the presence of hydrogenase subunits by immunodetection, which would have yielded a stronger indication of the integrity (or not) of the complex and of the possible occurrence of dissociation between hydrogenase and diaphorase moieties, as previously observed in similar enzymes [12].

Discussion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

The experiments described above indicate that NADH is a much better donor of the Synechocystis hydrogenase for H2 production than NADPH. NAD+ is also a better acceptor than NADP+ for H2 oxidation. NAD(H) is therefore probably the preferential substrate of the diaphorase moiety in this enzyme. Moreover, HoxE appears to be an essential component for the connection between the NAD+/NADH and H+/H2 oxidoreduction reactions.

On investigation of the effect of pyridine nucleotides on the reductive activation of hydrogenase, as assayed by restoration of H/D exchange capacity after O2 exposure, a different picture emerges. Previous reports have shown the ability of the enzyme to be activated by both NADH and NADPH [2,14]. In the present study, we show that NADPH is an efficient activator of the hydrogenase in Synechocystis, in a HoxE-independent manner. NADH can also exert an activating effect on the enzyme, with a lower efficiency than NADPH, but only when HoxE is present. The most straightforward interpretation is that NADH-mediated activation occurs via diaphorase donation to the electron transfer chain up to the NiFe cluster, whereas NADPH activation proceeds via a mechanism that is independent of electron donation through diaphorase or at least that does not involve the HoxE-dependent route. The differences that were observed concerning NADH and NADPH efficiencies between this study and a previous one from our laboratory [2] can be attributed to two essential protocol variations: (a) the special care taken in the current work to remove, as much as possible, membrane debris (ultracentrifugation instead of centrifugation); and (b) the oxidation of the samples before measurements in order to avoid interference with residual intracellular donors.

The process of NADPH-mediated activation remains to be elucidated at the molecular level. No homolog of HoxI [5] is present in cyanobacteria, but the fact that cyanobacterial Hox subunits bear supplemental FeS cluster-binding motifs when compared with those of R. eutropha might open possibilities for alternative electron pathways related to this phenomenon, stemming either from the diaphorase module or from another binding site. Whether NADPH activation proceeds by direct interaction with the enzyme, or indirectly (e.g. via ferredoxin–NADP+ reductase or reduced ferredoxin), is also not clear, and deserves further investigation. A reduced ferredoxin-mediated pathway, for instance, would yield a low-potential donor that could also directly interact with the HoxYH moiety, independently of diaphorase. This might account for a significant part of the hydrogenase activation in ΔhoxE, especially if the stability of the complex is impaired by the subunit deletion, as indicated by the native gel staining experiments. We tried to add ferredoxin–NADP+ reductase and ferredoxin to the cellular extracts in order to check this hypothesis, but did not observe any stimulation of the NADPH-mediated activation rate (Fig. S3), which currently leaves two possibilities: that these compounds are not involved in hydrogenase activation, or that they are already present in sufficient amounts in the cell extracts.

In NADH dehydrogenases, subunit E (NuoE, Nqo2, or 24-kDa subunit, depending on the considered organism) coordinates the 2Fe–2S cluster N1a [15] and is homologous to the thioredoxin-like [2Fe–2S]-bacterial ferredoxins, such as that from Aquifex aeolicus [1]. Interestingly, the localization of N1a in the Thermus thermophilus enzyme structure suggested that this cluster was prone to exchange electrons with the FMN group, but not with the other FeS clusters of the enzyme [15]. Moreover its redox potential is low, making it a poor potential acceptor for electrons coming from NADH, at least in isolated complexes [16]. Despite this, the deletion of the 24-kDa subunit led to total abolition of electron transfer in Neurospora crassa complex I [17]. Our finding that HoxE deletion in Synechocystis hydrogenase suppresses electron transfer with NAD(H) is in line with these previous data. In R. eutropha, the absence of a cysteine motif in the NuoE-homolog sequence at the HoxF N-terminus suggests a structural role for this module, independent of FeS cluster coordination. Because 24-kDa subunit deletion induced a strong decrease in the abundance of the 51-kDa subunit in N. crassa, it was suggested that NuoE could have a stabilizing effect [17].

So far, the conservation of the NuoE-like motif in all types of diaphorase with a NuoF-like NADH-binding module (including, for instance, NAD+-linked formate dehydrogenase [18] and NADP+-linked [FeFe]-hydrogenase [19]) indicates that it is probably part of the constitutive backbone of this reaction, although the precise electron transfer mechanism and the reason for the absolute requirement for NuoE remain to be elucidated, and might depend greatly on the actual redox potential of the cluster in vivo [17,20].

Considering enzyme subunit abundances, deletion of hoxH slightly affects the amount of the diaphorase subunit HoxF, deletion of hoxE slightly impacts on the amounts of HoxF and HoxH, and deletion of both hoxE and hoxF strongly decreases the amount of HoxH, in a manner that is compatible with the (also strong) observed decrease in H/D exchange capacity. The low amount of HoxH in HoxE/HoxF-depleted mutants could result from several factors: an effect on expression through polar transcriptional effects, owing to interruption of the operon; or post-transcriptional effects such as impaired maturation or instability of the enzyme. Quantitative real-time PCR experiments (Fig. S1) indicated that hoxH is expressed at the same level in ΔhoxEF strains as in the wild type, and therefore post-transcriptional effects are most probably responsible for the observed differences.

As NADH is one of the major products of glycolysis, being oxidized at the level of NADH dehydrogenase, and NADPH is produced by the photosynthetic electron transport chain, these results are in line with the observed dynamics of H2 during light–dark transitions. Indeed, in wild-type cells, H2 is produced during anaerobiosis in the dark as a result of NADH accumulation, this production being transiently stimulated by light, which yields NADPH and probably further stimulates hydrogenase. On another hand, when reducing equivalents are reoxidized in the light (CO2 fixation, and respiration of released O2) and before O2 reaches critical levels, H2 is consumed as the result of an activated hydrogenase working in the opposite direction (Fig. 2) [2]). This is also in line with the finding that NAD(P)H dehydrogenase suppression, which impairs NADPH oxidation [21], enhances light-stimulated H2 production [2].

When put together, these results indicate that differential regulation of hydrogenase subunit expression, differential interaction of the enzyme with electron donors and activators and complementary roles of diaphorase subunits might cooperate to adjust enzyme function in response to environmental variations.

Experimental procedures

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Strains and growth conditions

E. coli strains (DH10β, M15Rep4, and BL21) were grown on LB medium at 37 °C. The wild-type Synechocystis strain PCC6803 and mutant strains were grown autotrophically in liquid modified Allen’s medium [22] at 30 °C under continuous illumination at 30 μmol photons·m−2·s−1.

Construction of plasmid for targeted deletion of hoxH, hoxE, and hoxF

In order to perform hoxH deletion, a DNA region containing the coding sequence (positions 1 673 795–1 671 771 of the genomic sequence [23]) was amplified by PCR and inserted in pUC18 between the BamHI and HindIII sites with primers P1 and P2 (Table S1). The chloramphenicol resistance gene was then inserted between the ClaI sites at positions 19 and 1376 of hoxH, with primers P3 and P4 (Table S1).

A fragment of 2478 bp containing the hoxE and hoxF genes (positions 1 676 257–1 678 735 of the genomic sequence) was amplified by PCR and inserted into pUC19 between the BamHI and EcoRI sites with primers P5 and P6 (Table S1). For performance of hoxE interruption, a SnaBI–PshAI fragment of 201 bp in hoxE was deleted and replaced by the chloramphenicol resistance gene with primers P7 and P8 (Table S1).

For performance of simultaneous hoxE and hoxF interruption, a SnaBI–ApaI fragment of 1625 bp comprising regions of hoxE and hoxF was deleted and replaced by the chloramphenicol resistance gene with primers P7–P9 (Table S1).

Wild-type Synechocystis was transformed with these constructions, and transformants were selected on Allen medium agar plates containing 25 μg chloramphenicol·mL−1. Correct segregation of the strains was checked by PCR.

Antibody generation

The DNA region of hoxH was amplified by PCR with primers P10 and P11 (Table S1) and cloned between the BamHI and HindIII sites in the expression vector pQE30 (Qiagen, Courtaboeuf, France), in order to confer His6-tag fusion. The insert was verified by sequencing (genome express). His6-HoxH was overexpressed in E. coli strain M15Rep4 and purified from the soluble protein fraction of isopropyl thio-β-d-galactoside cultures with a His Trap HP column, following the protocols of the manufacturer (GE Healthcare Europe, Vélizy, France).

hoxF was cloned with primers P12 and P13 (Table S1) in the pET151 directional TOPO expression vector (Life Technologies, Cergy Pontoise, France), in order to confer His6-tag fusion. His6-HoxF was overexpressed in E. coli strain BL21. The membranes from isopropyl thio-β-d-galactoside cultures were solubilized in 8 m urea, and centrifuged at 245 000 g for 1 h. The soluble fraction was then purified with a His Trap HP column, following the instructions of the manufacturer (GE Healthcare Europe).

HoxH and HoxF protein fractions were then desalted by dialysis two times against NaCl/Tris and lyophilized. Aliquots of the HoxH and HoxF fractions (each 1–2 mg of protein) were used to generate corresponding polyclonal antibodies from rabbits (Eurogentec, Angers, France).

Extract preparation

Synechocystis cells were harvested by centrifugation (15 min at 5300 g) after 6 days of growth, suspended in 5 mL of phosphate buffer (20 mm, pH 7.2), and broken with ‘one shot’ cell disruptor apparatus (Constant Systems, Daventry, UK; pressure set at 2000 bars). The cell suspension was then centrifuged (60 min at 148 000 g) to remove cell debris, resulting in the cell-free extract. The protein content of the soluble fraction was determined with the Bradford protein assay (Pierce, Rockford, IL, USA). This fraction was used either for MS measurements (cell-free extract) or for enzyme expression assays.

The soluble fraction was then separated by SDS/PAGE and transferred to a nitrocellulose membrane. HoxH and HoxF were identified by using anti-HoxH serum diluted 1 : 1000 and anti-HoxF serum diluted 1 : 5000. IRDye 680 goat anti-(rabbit IgG) was used as secondary antibody. Detection was performed by fluorescence imaging (Li-Cor Odyssey, Lincoln, NE, USA).

Analysis of gas exchange by MS

Hydrogenase activity (H/D exchange rate of labeled dihydrogen) and hydrogen production were determined by membrane-inlet MS (MIMS) on cell suspensions or on cell-free extracts as previously described [2]. For H2 uptake assays, glucose and glucose oxidase were added to 1.5-mL cell extracts in the mass spectrometer measurement vessel, in order to ensure anaerobiosis, and 100 μL of H2-saturated water was then injected, followed by 10 μm reduced MV to activate the enzyme, and finally electron acceptor (5 mm BV, NAD+, or NADP+). The subsequent decrease in H2 concentration was monitored by MIMS.

For measurements on cell suspensions, Synechocystis cultures were harvested as above, and then diluted up to 30 μg chlorophyll·mL−1. Chlorophyll concentration was measured by absorbance at 665 nm after methanol extraction.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

This work was supported by CEA (BioH2 program), ANR (DIVHYDO and HYLIOX projects), and CNRS (REPROGRAMHYDROGEN project). M. Cano received support from Conseil Régional Provence Alpes Côte d’Azur and EDF, and experimental facilities were provided by the HelioBioTec platform, funded by Région Provence Alpes Cote d’Azur and the European Union (Regional Development Fund).

References

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information
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Supporting Information

  1. Top of page
  2. Abstract
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Fig. S1. Quantitative real-time PCR determination of hoxE, hoxF and hoxH expression in the four strains used during this study.

Fig. S2. Detection of hydrogenase activity by colorimetric (MV reduction) assay after electrophoresis in nondenaturing polyacrylamide gels.

Fig. S3. Effects of ferredoxin and ferredoxin–NADP+ reductase (FNR) addition on NADPH-mediated hydrogenase activation in Synechocystis extracts.

Table S1. List of primers used in this study.

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