GENETICALLY MODIFIED PHOTOTROPHIC CELL FOR IN-VIVO PRODUCTION OF HYDROGEN

Abstract

A genetically modified phototrophic cell for in-vivo production of hydrogen. The phototrophic cell has been genetically modified to the effect that a) at least one of the native photosystem I components has been deleted, b) the native hydrogenase has been deleted, and c) at least one fusion protein is expressed, comprising i. a hydrogenase or hydrogenase component and ii. at least one PSI component, with the proviso that the PSI is complemented by expression of the at least one fusion protein, and the hydrogenase component itself, or together with at least one further hydrogenase component expressibly introduced into the cell, has hydrogenase activity.

Claims

1. A phototrophic cell, which in the wild type has a native photosystem I, PSI, with PSI components and a native hydrogenase, wherein the phototrophic cell has been genetically modified to the extent that a) at least one of the native PSI components has been deleted, b) the native hydrogenase has been deleted, and c) at least one fusion protein is expressed which comprises i. a hydrogenase or hydrogenase component and ii. at least one PSI component, with the proviso that the PSI is complemented by expression of the at least one fusion protein and the hydrogenase component exhibits hydrogenase activity alone or together with at least one further hydrogenase component which has been expressibly introduced into the cell.

2. The phototrophic cell as claimed in claim 1, wherein the phototrophic cell has a higher photobiological hydrogen formation compared with a cell which has not been genetically modified in accordance with a), b) and c) above or compared with a wild type cell.

3. The phototrophic cell as claimed in claim 1, wherein the hydrogenase or hydrogenase component in the fusion protein is inserted into the PSI component or is disposed behind the PSI component or in front of the PSI component in the N-C direction, wherein the hydrogenase or hydrogenase component is optionally linked to the PSI component via a peptide linker.

4. The phototrophic cell as claimed in claim 1, wherein at least one of the native PSI components PsaB, PsaC, PsaD or PsaE has been deleted, and the fusion protein comprises this at least one PSI component, or a functional portion thereof.

5. The phototrophic cell as claimed in claim 1, wherein the native hydrogenase is a Hox hydrogenase, and the native hydrogenase components HoxE, HoxF, HoxU, HoxY and HoxH have been deleted.

6. The phototrophic cell as claimed in claim 5, wherein the phototrophic cell is genetically modified in a manner such that the hydrogenase components HoxY and HoxH are expressed, and wherein the at least one fusion protein comprises the hydrogenase component HoxY or HoxH.

7. The phototrophic cell as claimed in claim 3, wherein a) the hydrogenase or hydrogenase component is fused with PsaB or a functional portion thereof in a manner such that the hydrogenase or hydrogenase component is orientated towards the stroma, and/or b) the hydrogenase or hydrogenase component is fused with PsaC, or a functional portion thereof, in a manner such that the hydrogenase or hydrogenase component is orientated towards the stroma, and/or c) the hydrogenase or hydrogenase component is fused with the C-terminus of PsaD, or of a functional portion thereof, in a manner such that the hydrogenase or hydrogenase component is orientated towards the stroma, and/or d) the hydrogenase or hydrogenase component is fused with the N-terminus of PsaE, or of a functional portion thereof, in a manner such that the hydrogenase or hydrogenase component is orientated towards the stroma.

8. The phototrophic cell as claimed in claim 1, wherein the cell is capable of dividing.

9. The phototrophic cell as claimed in claim 1, wherein the phototrophic cell is a cyanobacterial cell, preferably a cyanobacterial cell of the genus Synechocystis.

10. The phototrophic cell as claimed in claim 1, wherein the hydrogenase component is a component of the NiFe hydrogenase from Synechocystis sp.

11. A nucleic acid coding for a fusion protein produced from i. a hydrogenase or hydrogenase component and ii. a PSI component, with the proviso that the hydrogenase component has hydrogenase activity alone or together with at least one further hydrogenase component.

12. The nucleic acid as claimed in claim 11, wherein the PSI component is PsaB, PsaC, PsaD or PsaE, pre, or a functional portion thereof.

13. The nucleic acid as claimed in claim 11, wherein a) the coding sequence for the hydrogenase or hydrogenase component is fused with the coding sequence for PsaB, or a functional portion thereof, in a manner such that the hydrogenase or hydrogenase component of the fusion protein expressed in-vivo after introduction of the nucleic acid into a phototrophic cell is orientated towards the stroma, and/or b) the coding sequence for the hydrogenase or hydrogenase component is fused with the coding sequence for PsaC, or a functional portion thereof, in a manner such that the hydrogenase or hydrogenase component of the fusion protein expressed in-vivo after introduction of the nucleic acid into a phototrophic cell is orientated towards the stroma, and/or c) the coding sequence for the hydrogenase or hydrogenase component is fused with the coding sequence for PsaD, or a functional portion thereof, in a manner such that the hydrogenase or hydrogenase component of the fusion protein expressed in-vivo after introduction of the nucleic acid into a phototrophic cell is fused with the C-terminus of PsaD, or of a functional portion thereof, and/or d) the coding sequence for the hydrogenase or hydrogenase component is fused with the coding sequence for PsaE, or a functional portion thereof, in a manner such that the hydrogenase or hydrogenase component of the fusion protein expressed in-vivo after introduction of the nucleic acid into a phototrophic cell is fused with the N-terminus of PsaE, or of a functional portion thereof.

14. The nucleic acid as claimed in claim 13, coding for a fusion protein from a Hox hydrogenase component HoxY and PsaD, or a functional portion of PsaD, wherein the coding sequence of HoxY is fused with the coding sequence of PsaD in a manner such that in the expressed fusion protein, HoxY is fused with the C-terminus of PsaD or the C-terminus of a functional portion of PsaD, and wherein the nucleic acid additionally codes for the Hox hydrogenase component HoxH.

15. A vector comprising a nucleic acid as claimed in claim 11.

16. (canceled)

17. A method for the production of hydrogen, wherein at least one phototrophic cell as claimed in claim 1 is cultured in an aqueous medium and/or in a biofilm under light.

18. The method as claimed in claim 17, wherein the phototrophic cell is cultured under anaerobic conditions.

19. The phototrophic cell as claimed in claim 1, wherein at least one of the native PSI components PsaB, PsaC or PsaD has been deleted, and the fusion protein comprises this at least one PSI component, or a functional portion thereof.

20. The phototrophic cell as claimed in claim 5, wherein the phototrophic cell is genetically modified in a manner such that the hydrogenase components HoxY and HoxH are expressed, and wherein the at least one fusion protein comprises the hydrogenase component HoxY.

Description

[0074] The invention will now be described in more detail purely by way of illustration with the aid of exemplary embodiments and the drawings.

[0075] FIG. 1: NAD(P)H/NAD(P)* ratios were determined at the same time as fermentative H.sub.2 production by Synechocystis cells which were cultured on either nitrate or arginine.

[0076] FIG. 2: Hydrogen production of wild type (WT) and Δhox/hoxYH mutants. (A) The hydrogenase assay in the presence of reduced methylviologen (MV) shows that the Δhox/hoxYH mutants express an active hydrogenase. (B) Fermentative and (C) photo-H.sub.2 production from WT and Δhox/hoxYH mutants.

[0077] FIG. 3: The Hox-H.sub.2ase (HoxYH) from Synechocystis was fused in-vivo to its photosystem I (PSI), in which the hydrogenase units hoxYH were fused with the PSI subunit psaD. The H.sub.2ase was fused with a short section of PsaD subunit truncated C-terminally by 35 amino acids, which in its wild type version surrounds PsaC in a clamp-like manner in the immediate vicinity of the 4Fe-4S cluster F.sub.B (44), in order to facilitate electron transfer between F.sub.B and the 4Fe-4S cluster of HoxY. The assumed electron transfer from plastocyanin (PC) via the 4Fe-4S cluster F.sub.X, F.sub.A and F.sub.B of PSI to HoxY and further on to the NiFe centre of HoxH is shown by arrows.

[0078] FIG. 4: Growth of wild type (WT), ΔhoxΔpsaD and psaD-hoxYH (=ΔhoxΔpsaD/psaD-hoxYH mutants) at 10 and 100 μE.Math.m2.Math.s.sup.−1 under photoautotrophic conditions with constant lighting.

[0079] FIG. 5: Hydrogenase activity in wild type (WT) and psaD-hoxYH mutants. (A) A hydrogenase test in the presence of reduced methylviologen (MV) shows that psaD-hoxYH expresses an active hydrogenase. (B) The major portion of the H.sub.2ase activity of WT cell homogenates was detected in the soluble fraction, whereas the largest portion of the H.sub.2ase activity of psaD-hoxYH homogenates was in the membrane fraction as expected, confirming the successful binding of HoxYH to the thylakoid membrane via PSI. It should be noted that some activity was lost upon separation of the fractions. Thus, the values do not add up to 100%.

[0080] FIG. 6: Short-term in-vivo H.sub.2 and O.sub.2 production in wild type (WT) and psaD-hoxYH in darkness and illumination under anaerobic conditions in the absence and presence of DCMU. Anaerobic conditions were obtained by adding glucose, glucose oxidase and catalase. (A) Fermentative and photo-H.sub.2 production in the WT with subsequent H.sub.2 take-up. (B) Photo-H.sub.2 production in psaD-hoxYH. The psaD-hoxYH is deficient in fermentative H.sub.2 production and H.sub.2 pickup. (C) Average amount of photo-H.sub.2 production in WT and psaD-hoxYH. (D) Average rate of photo-H.sub.2 production in WT and psaD-hoxYH. Note the different scales in the two figures.

[0081] FIG. 7: In-vivo photo-H.sub.2 production in WT and psaD-hoxYH based either on oxygenic photosynthesis with H.sub.2O splitting or alternatively on anoxygenic photosynthesis, for example with glucose as the electron donor.

[0082] FIG. 8: Long-term photo-H.sub.2 production in-vivo in wild type (WT) and psaD-hoxYH under anaerobic conditions with constant lighting in the presence and absence of DCMU. The anaerobic conditions were obtained by adding glucose, glucose oxidase and catalase. (A) Long-term photo-H.sub.2 production in WT and psaD-hoxYH. (B) Highest quantities of photo-H.sub.2 produced (under long-term conditions) by WT and psaD-HoxYH. (C) Average quantity of photo-H.sub.2 which was produced by WT and psaD-HoxYH (under long-term conditions). (D) Average rate of photo-H.sub.2 produced (under long-term conditions) by WT and psaD-hoxYH.

[0083] FIG. 9: Light dependency of long-term in-vivo photo-H.sub.2 production in wild type (WT) and psaD-hoxYH under anaerobic conditions in constant lighting in the absence of DCMU. Anaerobic conditions were obtained by the addition of glucose, glucose oxidase and catalase. The light was switched off, shown by the dark bars, and then switched on again. The decay in oxygen concentration in darkness was induced by the ending of H.sub.2O splitting at PSII. While the wild type still produced H.sub.2 by transferring to fermentative H.sub.2 production, H.sub.2 production stopped completely with psaD-hoxYH in darkness.

[0084] FIG. 10: Long-term photo-H.sub.2 production in wild type (WT) and psaD-hoxYH in presence of DCMU with or without glucose. The black bars show a dark phase with fermentative H.sub.2 production in the WT at the beginning of the experiment. Anaerobic conditions were obtained before beginning the measurements by flushing the cultures with argon for 10 minutes. The different scales in the two figures should be noted.

[0085] FIG. 11. Table 1 with primers employed.

[0086] FIG. 12. Example of a vector construct for genetic modification of a cyanobacterial cell in accordance with the invention.

EXAMPLE

[0087] The in-vivo photo-H.sub.2 production in cyanobacteria and green algae was restricted by (1) the sensitivity of the enzyme to oxygen, (2) the rapid uptake of photo-H.sub.2 in light and (3) the transfer of electrons from the photosystem I (PSI) to competing metabolism processes such as nitrogen assimilation and CO.sub.2 fixation, for example (5, 9, 11). In order to circumvent the aforementioned restrictions and to maximize the photo-H.sub.2 production in-vivo, the bidirectional NiFe-H.sub.2ase (Hox hydrogenase) from Synechocystis was fused in-vivo with the endogenous PSI in the cyanobacterium Synechocystis sp PCC 6803 (hereinafter Synechocystis). In this example, HoxYH was fused with the PSI component PsaD.

[0088] It was previously shown that the turnover frequency (TOF) of this hydrogenase in-vivo is large enough to maintain the supply of electrons at the PSI step (25). Bidirectional NiFe hydrogenase (Hox hydrogenase) consists of a diaphorase portion (HoxEFU) and a hydrogenase portion (HoxYH). The diaphorase contains several 2Fe2S and 4Fe4S clusters, which mediate electron transfer from soluble electron donors such as ferredoxin, flavodoxin, NADPH and NADH to the 4Fe4S cluster of HoxY, which then reduces the active centre of NiFe into HoxH for the production of H.sub.2 (5, 7, 26-28). It has already been reported that the hydrogenase unit alone forms a hydrogenase (HoxYH) which, in the presence of the synthetic electron donor methylviologen (MV) can develop H.sub.2 in-vitro (28, 29). The deletion of diaphorase subunits in-vivo leads, however, to a reduced hydrogen production, which is probably caused by a reduced electron transfer to the active centre as well as unstable hydrogenase complexes (26, 30). Thus, in a preliminary experiment, a test was carried out as to whether the hydrogenase (HoxYH) alone could be expressed in a stable and active manner in-vivo, with the aim of then fusing only HoxYH to PSI.

[0089] Materials and Methods

[0090] Construction of Plasmids

[0091] All of the primers used are listed in Table 1 (see FIG. 11). PCR products of the upstream and downstream regions of the Hox operon and psaD were produced together with those of kanamycin and gentamycin-resistance cassettes (“Km cassette”, “Gm cassette”). The three products were then linked by means of a “Gibson assembly” (52) with pUC19 to form pDHOX and pDPSAD. In order to produce an expression construct for hoxYH PCR products from upstream and downstream regions of the Hox operon, the trc promoter (trp-lac promoter) and hoxYH with a 5′-extension, which coded for the Strep-Tag, were amplified and composed into a Gibson Assembly with pBluescript SK-, which had been cut with KpnI and SacI, into a plasmid. In the case of the construct for the fusion of hoxY an psaD, four PCR products were produced: psaD up to the desired fusion site, the hydrogenase gene (hoxY, ORF6, ORF7 and hoxH) from the second codon of hoxY, the erythromycin-resistance cassette (“Em cassette”) and a downstream region of psaD (see FIG. 12). These four fragments were assembled in a single step by TAR cloning (53) in pMQ80 (54), which had been cut by EcoRI and NaeI. In contrast to the original protocol, the fragments only had 40 bp overlaps. For the cloning process itself, the protocol of (55) was employed. In brief, a single colony of the yeast strain BJ5464 was cultured overnight at 30° C. on a rotary shaker with 200 UpM in 50 mL of YPD (1% yeast extract, 2% peptone, 2% glucose). The next day, the culture medium was diluted to a density of OD.sub.600=0.5. The cells had grown to a density of OD.sub.600=2 in approximately 4 h and were harvested at 3000×g for 5 min. The cells were washed twice in 25 mL of sterile water and once in 1 mL of sterile water and then pelletized for 30 seconds at 13000×g in a 1.5 mL beaker. Finally, the pellet was re-suspended in 1 mL of sterile water and 100 μL of cells were divided per transformation into separate beakers. A mixture of 240 μL 50% (wt./vol.) PEG 3350, 36 μL 1 M lithium acetate, 50 μl 2 mg/mL denatured salmon sperm DNA and 34 μL DNA was added to the cells, and the mixture was vigorously stirred. 2 μg of plasmid was used and the various fragments were added to the plasmid in a molar ratio of 1:1. After incubating for 40 minutes at 42° C., the cells were pelletized for 30 seconds at 13000×g, re-suspended in 500 μL of sterile water and plated out onto yeast drop out medium without uracil. All of the colonies which had grown after 3 to 4 days incubation at 30° C. on a single plate were washed from the plate with sterile water. The cells were pelletized and the DNA was isolated using standard protocols. With this DNA, chemically competent Escherichia coli DH5a was transformed and selected on the respective antibiotic. Plasmids were purified out of the resulting colonies. All of the constructs obtained were tested by sequencing.

[0092] Production and Growth of Strains

[0093] Wild type cells from Synechocystis sp. PCC 6803 were transformed in accordance with (56) one after the other with pDHOX and pDPSAD, in order to delete all Hox genes and psaD. The wild type cells and Δhox cells were cultured at 50 μE.Math.m2.Math.s.sup.−1 on BG-11 agar plates and in bubble culture or in shaking culture. Cells transformed with pDPSAD were cultured at 5 μE.Math.m2.Math.s.sup.−1 on plates with 10 mM glucose. After segregation, the ΔhoxΔpsaD-strain was transformed at 10 μE.Math.m2.Math.s.sup.−1 in bubble culture without glucose and transformed with construct I. In this case, the cells were cultured in shaking culture (100 UpM) for approximately three days at 5 μE.Math.m2.Math.s.sup.−1 with 10 mM glucose, before they were plated out onto BG-11 with 10 mM glucose and 20 μg/mL erythromycin. The plates were kept for a few weeks at 5 μE.Math.m2.Math.s.sup.−1, before colonies appeared. The segregation of all of the resulting strains was either checked using PCR or Southern blot. The successful transformation of the Δhox and fusion strains was also checked by measurement of their hydrogenase activity.

[0094] Hydrogen Measurements

[0095] The hydrogenase activity was measured in whole cells in the presence of 10 mM dithionite and 5 mM of methylviologen with a hydrogen electrode as described in (25). The short and long-term hydrogen conversion was measured with the MicroRespiration System (Unisense, Aarhus, Denmark). Cells which had been cultured at 10 μE.Math.m2.Math.s.sup.−1 under a constant stream of air bubbles were harvested by centrifuging for 10 minutes at 5000 U/min and re-suspended in fresh BG-11. The chlorophyll content was determined (25) and adjusted to 20 μg/mL. Cell suspensions were placed in MicroRespiration double chamber and glucose, glucose oxidase and catalase were added in final concentrations of 10 mM, 16 U/mL or 20 U/mL. After inserting an oxygen and a hydrogen electrode into each chamber, the measurement was begun.

[0096] Determination of NAD (P)/NAD(P) Ratios

[0097] The NAD(P)H/NAD(P)+ ratios were determined with the colorimetric NADPH/NADP and NADH/NAD quantification kits from BioVision (California, USA). The cultures were grown under fermentative conditions either in BG11 medium or in BG110 medium, which contained 5 mM arginine and 10 mM glucose. An equivalent of approximately 1.0×10.sup.9 cells/mL (10 mL culture with an OD.sub.750 of 1) was harvested at 3500 g for 10 minutes at −9° C. The pellets were resuspended in 1 mL 20 mM cold PBS-Puffer and centrifuged for 1 min at −9° C. at 12000 g. The pellet was resuspended in 50 μL of extraction buffer (BioVision, California, USA) and pre-cooled glass beads (diameter 0.18 mm) were added close to the surface (0.5-1 mm) of the liquid. The mixture was swirled 4 times for 1 min at 4° C., cooling in between for 1 min on ice. A fresh 150 μL of extraction buffer was added and the mixture was centrifuged at 3500 g, −9° C. The liquid phase was, as far as possible, transferred into a new reaction vessel and centrifuged for 30 minutes at −9° C. at maximum speed. The residue was transferred into pre-cooled 10 kD spin columns (BioVision, California, USA). The sample was centrifuged for 10 min at 10000 g, −9° C. The flux through the liquid was divided into two 30 μL portions. One was incubated with 30 μL NADH extraction buffer (BioVision, California, USA) for 30 minutes at 60° C. and immediately cooled on ice, and was then rapidly centrifuged in order to remove any precipitate, in order to determine the quantity of NADH in the cells. For the other, 30 μL of NADH extraction buffer (BioVision, California, USA) was placed on ice. Two 25 μL heated and two 25 μL unheated samples were placed in four wells of a 96-well mictotitre plate. For the concentration control, 2.5 μL of all of these samples was placed in wells with 22.5 μL of NADH extraction buffer. 50 μL NAD “Cycling Mix” (BioVision, California, USA) was mixed well and pipetted into each well and incubated for 5 minutes at room temperature. Next, 10 μL of NADH developer (BioVision, California, USA) was added to each well and incubated for 1 to 4 hours before the absorption was measured at 450 nm using a TECAN-GENios instrument (TECAN Group Ltd., Austria). The reactions were stopped after adding 5 μL of stop solution (BioVision, California, USA) to each well. The colour should have been made stable for 48 h with the stop solution and was sealed with film at 4° C. The plates were then read at OD 450 nm. The values were collected using Magellan (TECAN) software. 10 μL of 1 nmol/μL of NADH standard was diluted with 990 μL of NADH/NAD extraction buffer in order to produce 10 pmol/μL of standard NADH. The addition of 0, 1, 2, 3, 4, 5 μL of the diluted standard to labelled 96-well plates, in duplicate, produced 0, 20, 40, 60, 80, 100 pmol/Well standards. A final volume of 25 μL was obtained after adding NADH/NAD extraction buffer. All of the diluted NADH standards were treated in the same way as the aforementioned samples. NADPH/NADP quantifications were carried out in a similar manner with the NADPH/NADP “Quantification Colorimetric Kit” from BioVision (BioVision, California, USA).

[0098] Results

[0099] Validation of in-vivo electron donor for the bidirectional cyanobacteria-NiFe hydrogenase FeFe-H.sub.2ases from green algae have long been known to produce hydrogen with the aid of reduced ferredoxin, while for several decades it was assumed that the bidirectional cyanobacteria NiFe hydrogenase formed hydrogen via NAD(P)H. The latter enzyme was therefore seen to be less attractive for biotechnological applications because with reduced ferredoxin compared with NAD(P)H, much higher H.sub.2 concentrations could be produced.

[0100] Based on experimental data and theoretical assumptions, it is now maintained that the bidirectional cyanobacterial NiFe hydrogenase also produces hydrogen, to the detriment of the reduced ferredoxin (5). However, these statements were then called into question because of contradicting data which claimed that NAD(P)H (albeit employed in non-physiologically high concentrations) rather than reduced ferredoxin was confirmed as the electron donor for the cyanobacterial enzyme (6). Thus, fresh investigations to find the electron donor for bidirectional NiFe-H.sub.2ase were undertaken, this time in an in-vivo approach.

[0101] Cells were cultured on either nitrate (NO.sub.3.sup.−) or alternatively on arginine as the source of nitrogen. Nitrate is reduced by ferredoxin in cyanobacteria (31). Thus, it can be assumed that the ferredoxin pool in cells which are cultured with arginine is reduced more strongly. Correspondingly, fermentative H.sub.2 production under these conditions was raised seven-fold (3.2 μM H.sub.2 (NO.sub.3.sup.−); 21.2 μM H.sub.2 (Arg)) (see Table 2), which indirectly again indicates that reduced ferredoxin is the electron donor (FIG. 1). Until now, however, there have not been any methods to hand with which the Fdx.sub.ox/Fdx.sub.red ratio could be determined in-vivo. Therefore, NADH/NAD.sup.+- and NADPH/NADP.sup.+ ratios were determined at the same time as H.sub.2 production. In order to produce 20 μM H.sub.2 at a pH of 7 (measured on arginine), NAD(P)H/NAD(P)* ratios of 50 or instead a Fdx.sub.red/Fdx.sub.ox ratio of 0.001 was necessary (32). These values are far higher than the measured ratios for NADH/NAD* of 0.039 (NO.sub.3.sup.−) and 0.3 (Arg) (see Table 2, FIG. 1) and NADPH/NADP* of 0.058 (NO.sub.3.sup.−) and 0.08 (Arg) (see Table 3, FIG. 1).

[0102] Table 2: NADH/NAD* ratios for fermentative H.sub.2 production by means of Synechocystis during culture on nitrate or arginine as the source of nitrogen. Measured values compared with theoretical values obtained from thermodynamic considerations.

TABLE-US-00001 NO.sub.3.sup.− /Arg NADH/NAD.sup.+ μM H.sub.2 theor. μM H.sub.2 measured NO.sub.3.sup.− 0.039 0.01 3.2 Arg 0.3 0.13 21.2

[0103] Table 3: NADPH/NADP* ratios for fermentative H.sub.2 production by means of Synechocystis during culture on nitrate or arginine as the source of nitrogen. Measured values compared with theoretical values obtained from thermodynamic considerations.

TABLE-US-00002 NO.sub.3.sup.− /Arg NADH/NAD.sup.+ μM H.sub.2 theor. μM H.sub.2 measured NO.sub.3.sup.− 0.058 0.025 3.2 Arg 0.08 0.03 21.2

[0104] The measured NAD(P)H/NAD(P)* ratios agree well with data from the literature, which give NADH/NAD* ratios in E. coli of 0.5 and 0.7, 0.25 and 0.05 in Synechocystis and NADPH/NADP* ratios of 0.05 in Synechococcus under fermentative conditions (33-36). According to thermodynamic calculations, H.sub.2 could be produced at a NADH/NAD* ratio of 0.3 (Ag) 0.13 μM H.sub.2 and at a NADPH/NADP* ratio of 0.08 (Arg) 0.03 μM (Table 3, FIG. 1) (32). The measured NAD(P)H/NAD(P)* ratios are therefore clearly not responsible for the observed H.sub.2 production of 20 μM, whereas the required Fdx.sub.red/Fdx.sub.ox ratio of 0.001 in cells should be readily obtainable. It may therefore be that NAD(P)H is responsible for the production of small quantities of H.sub.2, exerts an activating action on hydrogenase and takes part in a bifurcation reaction together with ferredoxin (5, 7, 37). The above results clearly show, however, that reduced ferredoxin must be responsible for the production of large quantities of H.sub.2 and that the bidirectional cyanobacterium NiFe hydrogenase is therefore a biotechnologically attractive enzyme.

[0105] Expression of Hydrogenase (HoxYH) Unit In-Vivo

[0106] In order to test whether hydrogenase (HoxYH) can be expressed in-vivo in a stable and active manner, HoxYH was overexpressed under the control of a trc promoter in a Δhox deletion strain of Synechocystis. The corresponding Δhox/hoxYH mutant expressed an active hydrogenase in-vivo, as could be shown by methylviologen-based enzyme tests (FIG. 2A). However, the mutants were not able to produce fermentative hydrogen with their natural electron donors (FIG. 2B) and in-vivo exhibited a reduced capacity for photo-H.sub.2 production (FIG. 2C).

[0107] The latter results were in line with expectations, because diaphorase is responsible for an effective transfer of electrons from the redox pairs to the hydrogenase. Because HoxYH could be expressed actively and in a stable manner in-vivo, however, an approach was developed whereby only these subunits were fused with PSI.

[0108] Construction of a PSI-H.sub.2Ase Fusion Product

[0109] The efficiency of the electron transfer between Fe—S clusters depends primarily on the distance between the redox centres, which should ideally be less than 14 Å (38). The vicinity of redox centres is more decisive than its mean potential (39). Shortening the distances from the 14 Å limit enables increasingly endergonic electron transfer steps (38). In contrast to in-vitro tests, the hydrogenase at the PSI in-vivo has to compete successfully with ferredoxin for electrons. The PSI is composed of several subunits, of which PsaA and PsaB are the two largest, which form a heterodimeric reaction centre with the chlorophyll pair P700 and the electron transfer components chlorophyll A.sub.0, phylloquinone A.sub.1 and the 4Fe-4S cluster F.sub.X (40). Three smaller soluble subunits PsaC, PsaD and PsaE are bonded to the reaction centre on the cytoplasm side. PsaC contains two 4Fe-4S clusters F.sub.A and F.sub.B. In the photosynthetic electron transport chain, plastocyanin releases electrons directly to the reaction centre and P700. The electrons are transferred from P700 via A.sub.0, A.sub.1, F.sub.X, F.sub.A and F.sub.B to the soluble electron carrier ferredoxin (see FIG. 7 as well). The binding site of ferredoxin is in a pouch formed by PsaE, PsaC and PsaD (FIG. 3) (40-42). In their in-vitro assay, Ihara et al. (2006) bound the NiFe hydrogenase of Ralstonia to the C-terminus of PsaE which is remote from the ferredoxin binding site. This may be a reason for the preferred electron transfer onto ferredoxin instead of the hydrogenase in this fusion construct (20). In order to place the 4Fe-4S cluster of HoxY in the immediate vicinity of the 4Fe-4S cluster F.sub.B of PsaC, HoxY was fused with the C-terminus of the PsaD shortened by 35 amino acids up to glutamate 103, which is orientated in the F.sub.B direction, in which it surrounds PsaC in the manner of a clamp (FIG. 3) (43, 44).

[0110] The N-terminus of HoxY was fused directly to the glutamate 103 of PsaD without a linker. In order to coarsely estimate the distance between the 4Fe-4S cluster F.sub.B in PsaC and the 4Fe-4S cluster of HoxY in the fusion construct, the distances of the 4Fe-4S clusters to the amino acids on the surface of the respective proteins was determined and added. The largest separation within HoxY was approximately 33 Å to its N-terminus and the shortest separation was 13 Å to Tyr155 (45). Within PsaC, the largest separation was 12.2 Å to Pro29 and the shortest separation was 10.7 Å to Val28 or 10.3 Å to Glu26 (44). As a function of the in-vivo orientation of HoxY with respect to PsaC, it can be estimated that the 4Fe-4S cluster has a distance of approximately 23.3 Å (13+10.3) to 45.2 Å (33+12.2), which is well over the desired limit of 14 Å (38).

[0111] In-Vivo Assembly of PsaD-HoxYH onto PSI

[0112] In order to obtain a Synechocystis mutant into which the psaD-hoxYH fusion could finally be introduced, both the complete Hox gene cluster (hoxEFUYH) as well as the PSI subunit psaD were deleted, which produced a ΔhoxΔpsaD mutant (hereinafter: ΔhoxΔpsaD). The respective mutant was segregated in low lighting conditions (5 μE.Math.m2.Math.s.sup.−1) in the presence of 10 mM glucose. Next, a fusion construct (see FIG. 12) produced from psaDhoxYH in ΔhoxΔpsaD was introduced in order to complement PsaD and to obtain the mutant ΔhoxΔpsaD/psaD-hoxYH. The mutant ΔhoxΔpsaD/psaD-hoxYH will be abbreviated here to psaD-hoxYH. The aim was to develop a mutant which preferentially transferred electrons from the photosynthesis electron transport chain to the hydrogenase (HoxYH) without completely blocking the PSI for other electron acceptors which are responsible for fixing CO.sub.2 and for photoautotrophic growth. In earlier studies, it was shown that the deletion or modification of the PSI subunit PsaD results in mutants with reduced PSI stability, reduced ferredoxin affinity and NADP* photoreduction. However, these mutants could still grow under photoautotrophic conditions, albeit in some cases at extremely low rates (46, 47).

[0113] FIG. 12 is diagrammatic and shown as an example of the structure of the plasmid for introducing a fusion of hoxYH onto PsaD. The sequence corresponds to the sequence in accordance with SEQ ID NO: 31. The shortened sequence of psaD borders directly onto hoxY, followed by two open reading frames (ORF6, ORF7), which are located in the operon of the Hox gene (hoxEFUYH) and the function of which is unclear. The hoxH is adjacent, followed by a resistance cassette against the antibiotic erythromycin (EM cassette), which is used in order to build up selection pressure. In order to be able to integrate the construct into the correct position in the genome by homologous recombination, the sequence was integrated into the vector once downstream of PsaD (down PsaD) and upstream of PsaD (hatched).

[0114] Characterization of psaD-hoxYH

[0115] Photoautotrophic Growth of psaD-hoxYH

[0116] We tested the photoautotrophic growth of WT, ΔhoxΔpsaD and psaD-hoxYH in the absence of external carbohydrate sources in low (10 μE.Math.m2.Math.s.sup.−1) and medium (100 μE.Math.m2.Math.s.sup.−1) lighting.

[0117] ΔhoxΔpsaD could grow slowly under poor lighting conditions, but barely grew at the higher light intensity (FIG. 4), which agrees well with earlier reports regarding the growth of a ΔpsaD mutant of Synechocystis (46). Complementing ΔhoxΔpsaD with psaD-hoxYH resulted in a mutant which grew better under low lighting and furthermore exhibited a photoautotrophic growth at 100 μE m2.Math.s.sup.−1 lighting, albeit at lower rates than the wild type (FIG. 4).

[0118] Hydrogenase Activity of psaD-hoxYH

[0119] In order to test whether the psaD-hoxYH fusion mutant expressed a functional hydrogenase, the H.sub.2 production was measured in the presence of the artificial reduced electron donor methylviologen (MV). With this assay, functional hydrogenases could be quantified in cells. The hydrogen production in psaD-hoxYH was raised compared with the wild type, which was expected, because HoxYH was expressed under the control of a promoter of the PSI subunit (psaD) which should be stronger than the promoter for the hydrogenase operon (FIG. 5A).

[0120] In order to test whether the hydrogenase (HoxYH) in psaD-HoxYH is bonded via PSI to the thylakoid membrane as desired, cells were ruptured and the hydrogenase activity was initially measured in whole cell homogenates and then both in the soluble and also in the membrane fraction. In the wild type, the hydrogenase activity was virtually limited to the soluble fraction (97%) with little activity in the membrane fraction (3%) (FIG. 5B). In contrast to this, in psaD-hoxYH the most hydrogenase activity was found in the membrane fraction (67%), and lower activities (16%) were measured in the soluble fraction, which confirmed successful binding of HoxYH to the thylakoid membrane via PSI (FIG. 5B). Because of the loss of activity upon the separation of the two fractions, the values do not add up to 100%.

[0121] In-Vivo Hydrogen Production

[0122] Fermentative H.sub.2 production and short-lived photo-H.sub.2 production upon irradiation In vivo H.sub.2 production was monitored with H.sub.2 sensors (Unisense) in the wild type and psaD-hoxYH. In contrast to the wild type, with psaD-hoxYH under dark anaerobic conditions, as expected, no fermentative H.sub.2 production was observed, because in psaD-hoxYH, diaphorase (HoxEFU), which is responsible for the electron transfer from the in-vivo electron donor to the active centre of the hydrogenase, is missing (FIG. 6) (26). Because of binding to the PSI, it can also be considered that the 4Fe-4S cluster of HoxY is difficult for electrons other than those from PSI to access.

[0123] Under illumination, photo-H.sub.2 was produced both in the wild type (1.8 μM H.sub.2±0.7; n=17) as well as in psaD-HoxYH (1.1 μM H.sub.2±0.5; n=22) (FIG. 6A-C). The mutants had a much slower photo-H.sub.2 production rate (11.5 μM H.sub.2/h±4.9) compared to the wild type (260 μM H.sub.2/h±143), which indicates that the transfer of electrons between PSI and HoxYH in the case of psaD-hoxYH is less effective than in the wild type and could be optimized further (FIG. 6D). The poorer electron transfer could be due to the estimated distance of 23 Å to 45 Å between the 4Fe4S clusters of HoxY and F.sub.B, which is over the target limiting value of 14 Å (38). However, while the wild type consumed the photo-H.sub.2 within a few minutes, the psaD-hoxYH culture maintained a high photo-H.sub.2 level for several hours (FIG. 6A-B). This is in agreement with the concept that the hydrogenase in the WT takes up electrons for the photo-H.sub.2 production, then catalyses the reverse reaction and possibly feeds electrons back via the diaphorase and the photosynthesis complex 1 (NDH-1) into the photosynthetic electron chain (6, 7, 48). The immobility of HoxYH in psaD-HoxYH together with the lack of diaphorase is therefore highly probably responsible for the lack of photo-H.sub.2 pickup. The oxygen contents in the cultures were kept low by adding glucose and glucose oxidase and catalase to consume the oxygen, in order to prevent inactivation of the hydrogenase. Electrons dissipated from the glucose oxidation can enter the photosynthetic inter-system chain (for example via the photosynthetic complex I, NDH1) and finally reach the PSI for photo-H.sub.2 production (see FIG. 7). This process contains exclusively PSI without the formation of oxygen and therefore enables photo-H.sub.2 production on the basis of an anoxygenic photosynthesis. In order to test whether the observed photo-H.sub.2 is based on either oxygenic or anoxygenic photosynthesis or on both, DCMU was added, which specifically inhibits electron transfer from photosystem II (PSII) into the plastoquinone pool (PQ) (7). The addition of DCMU reduced the photo-H.sub.2 production both in the wild type and also in psaD-hoxYH, but did not cancel this completely, which confirms that PSII probably delivers electrons from the H.sub.2O oxidation for photo-H.sub.2 production in addition to another source.

[0124] Long-Term Photo-H.sub.2 Production Under Anaerobic Conditions with Constant Lighting

[0125] When WT and psaD-hoxYH cultures were left for several days under anaerobic conditions in constant lighting, a second photo-H.sub.2 production phase was observed in which both in WT (average of 15 μM H.sub.2±8.3; n=8) and also, most impressively, in psaD-hoxYH (average 139 μM H.sub.2±146; n=16), much higher quantities of photo-H.sub.2 were formed (FIG. 8). The H.sub.2 concentrations reached fluctuated relatively severely and in WT were between 6 μM H.sub.2 and 29 μM H.sub.2 and in psaD-hoxYH between 32 μM H.sub.2 and 500 μM H.sub.2. The comparison of the maximum H.sub.2 concentrations which were obtained from the WT (23 μM H.sub.2) and psaD-hoxYH (500 μM H.sub.2) exhibited a 22-fold rise in the level of H.sub.2 in the mutant (FIG. 8B). Interestingly, in contrast to the situation with short-term photo-H.sub.2 production, the mutants exhibited significantly higher rates of H.sub.2 production (12.5 μM H.sub.2/h±5.8; n=16) compared with WT (2.1 μM H.sub.2/h±1.8; n=6) (FIG. 8C). Typically, photo-H.sub.2 production in the mutant began sooner than compared with WT (FIG. 8A and FIG. 10). To the knowledge of the inventors, the highest in-vivo concentration measured for Synechocystis cultures until now was approximately 50 μM H.sub.2 (11). By far the best cyanobacterial H.sub.2 producer identified up to now is the filamentary cyanobacterium Lyngbya aestuarii, which under fermentative conditions in-vivo, in the absence of externally added reducing agents, reached concentrations of 160 μM H.sub.2 (49).

[0126] In order to test whether the observed H.sub.2 production was genuinely light-dependent, WT and psaD-hoxYH were initially left under anaerobic conditions with constant lighting until H.sub.2 production began. The light was switched off as soon as the H.sub.2 production reached high rates. The oxygen concentration dropped immediately because of the interrupted H.sub.2O splitting at PSII in darkness (FIG. 9).

[0127] The wild type still produced H.sub.2 in darkness (even at higher rates), probably by switching to fermentative H.sub.2 production. In contrast, PsaD-hoxYH stopped H.sub.2 production entirely in darkness. Both cultures recommenced H.sub.2 production at the earlier rates as soon as the light was switched on again (FIG. 9). The long-term photo-H.sub.2 production is thus clearly light-dependent, at least with psaD-HoxYH.

[0128] Next, the possible dependency of the observed long-term photo-H.sub.2 production on the oxygenic photosynthesis with a contribution from PSII was investigated. To this end, DCMU was added to the cultures; this specifically inhibits electron transfer from PSII to plastoquinone and therefore ends H.sub.2O splitting at PSII (see FIG. 7). The observed results were not always consistent, because in some cases, DCMU reduced the quantity of photo-H.sub.2 produced (for example FIG. 8A), whereas in most cases the photo-H.sub.2 level rose even in the presence of DCMU (FIG. 8C). Because DCMU did not clearly inhibit the long-term photo-H.sub.2 production, however, this process is clearly not primarily dependent on H.sub.2O oxidation at PSII.

[0129] Because glucose, glucose oxidase and catalase were present in the measured cultures, in order to keep the oxygen content low, we tested the influence of glucose on the observed H.sub.2 production further. Unfortunately, in this case there were only two possibilities for keeping cultures anaerobic for several days in constant lighting: (1) Adding DCMU in order to block O.sub.2 development at PSII, or alternatively (2) adding glucose, glucose oxidase and catalase. Flushing cultures with N.sub.2 or argon in order to obtain anaerobic conditions would also have driven H.sub.2 out of the cultures and therefore rendered H.sub.2 measurements impossible. Thus, DCMU was added to the cultures in order to inhibit O.sub.2 production at PSII and to observe the influence of external glucose on the H.sub.2 production. In order to induce anaerobic conditions in the absence of glucose, prior to beginning the O.sub.2 and H.sub.2 measurements, all of the cultures were flushed with argon for 10 min. In WT, similar H.sub.2 levels were produced in the presence and absence of glucose, while the production rates with glucose were much higher (FIG. 10). In the absence of glucose, psaD-hoxYH produced similar quantities of H.sub.2 to the wild type, while in the presence of glucose, concentrations were obtained which were 10 times higher (FIG. 10).

[0130] Sequences (see also Table 1 in FIG. 11):

TABLE-US-00003 Primer SEQ ID NO: hoxout1 01 hoxin1Km 02 Km1 03 Km2 04 hoxin2Km 05 hoxout2 06 pB-Trc-fw 07 pB-Tre-rev NEU 08 pB-NStrep-YH-fw NEU 09 pB-NStrep-YH-rev 10 pB-Gm-fw 11 pB-Gm-rev NEU 12 Hox-out1pB 13 hoxin1pB 14 hoxin2pB 15 hoxout2pB 16 PsaDout1 17 psaDin1 18 Gm1 19 Gm2 20 psaDin2 21 PsaDout2 22 Eco-PsaDout1 23 HoxY-PsaD 24 PsaD-HoxY 25 Em-HoxH 26 PsaD-Em 27 HoxH-Em 28 Em-PsaD 29 NaeI-PsaDout2 30 hoxYH-psaD-Plasmid 31

REFERENCES

[0131] 1. D. C. Catling, C. R. Glein, K. J. Zahnle, C. P. McKay, Why O2 Is Required by Complex Life on Habitable Planets and the Concept of Planetary “Oxygenation Time”. Astrobiology 5, 415-438 (2005). [0132] 2. P. G. Falkowski, L. V. Godfrey, Electrons, life and the evolution of Earth's oxygen cycle. Philosophical Transactions of the Royal Society B-Biological Sciences 363, 2705-2716 (2008). [0133] 3. E. A. Peden et al., Identification of Global Ferredoxin Interaction Networks in Chlamydomonas reinhardtii. Journal of Biological Chemistry 288, 35192-35209 (2013). [0134] 4. A. Sawyer, M. Winkler, Evolution of Chlamydomonas reinhardtii ferredoxins and their interactions with [FeFe]-hydrogenases. Photosynthesis Research 134, 307-316 (2017). [0135] 5. K. Gutekunst et al., The Bidirectional NiFe-hydrogenase in Synechocystis sp. PCC 6803 Is Reduced by Flavodoxin and Ferredoxin and Is Essential under Mixotrophic, Nitrate-limiting Conditions. Journal of Biological Chemistry 289, 1930-1937 (2014). [0136] 6. I. Dutta, W. F. J. Vermaas, The electron transfer pathway upon H.sub.2 oxidation by the NiFe bidirectional hydrogenase of Synechocystis sp. PCC 6803 in the light shares components with the photosynthetic electron transfer chain in thylakoid membranes. International Journal of Hydrogen Energy 41, 11949-11959 (2016). [0137] 7. L. Cournac, G. Guedeney, G. Peltier, P. M. Vignais, Sustained Photoevolution of Molecular Hydrogen in a Mutant of Synechocystis sp. Strain PCC 6803 Deficient in the Type I NADPH-Dehydrogenase Complex. J. Bacteriol. 186, 1737-1746 (2004). [0138] 8. J. Appel, S. Phunpruch, K. Steinmnller, R. Schulz, The bidierctional hydrogenase of Synechocystis sp. PCC 6803 works as an electron valve during photosynthesis. Arch Microbiol 173, 333-338 (2000). [0139] 9. V. Nagy et al., Water-splitting-based, sustainable and efficient H.sub.2 production in green algae as achieved by substrate limitation of the Calvin-Benson-Bassham cycle. Biotechnology for Biofuels 11, 69 (2018). [0140] 10. A. Melis, L. Zhang, M. Forestier, M. L. Ghirardi, M. Seibert, Sustained Photobiological Hydrogen Gas Production upon Reversible Inactivation of Oxygen Evolution in the Green Alga Chlamydomonas reinhardtii. Plant Physiology 122, 127-136 (2000). [0141] 11. F. Gutthann, M. Egert, A. Marques, J. Appel, Inhibition of respiration and nitrate assimilation enhances photohydrogen evolution under low oxygen concentrations in Synechocystis sp. PCC 6803. Biochimica et Biophysica Acta (BBA)—Bioenergetics 1767, 161-169 (2007). [0142] 12. D. C. Ducat, G. Sachdeva, P. A. Silver, Rewiring hydrogenase-dependent redox circuits in cyanobacteria. Proceedings of the National Academy of Sciences 108, 3941-3946 (2011). [0143] 13. E. Reisner, D. J. Powell, C. Cavazza, J. C. Fontecilla-Camps, F. A. Armstrong, Visible Light-Driven H.sub.2 Production by hydrogenases Attached to Dye-Sensitized TiO2 Nanoparticles. Journal of the American Chemical Society 131, 18457-18466 (2009). [0144] 14. M. B. Wilker et al., Electron Transfer Kinetics in CdS Nanorod-[FeFe]-Hydrogenase Complexes and Implications for Photochemical H.sub.2 Generation. Journal of the American Chemical Society 136, 4316-4324 (2014). [0145] 15. M. Hambourger et al., [FeFe] Hydrogenase-Catalyzed H.sub.2 Production in a Photoelectrochemical Biofuel Cell. Journal of the American Chemical Society 130, 2015-2022 (2008). [0146] 16. D. Mersch et al., Wiring of Photosystem II to Hydrogenase for Photoelectrochemical Water Splitting. Journal of the American Chemical Society 137, 8541-8549 (2015). [0147] 17. K. P. Sokol et al., Bias-free photoelectrochemical water splitting with photosystem II on a dye-sensitized photoanode wired to hydrogenase. Nature Energy 3, 944-951 (2018). [0148] 18. I. Yacoby et al., Photosynthetic electron partitioning between [FeFe]-hydrogenase and ferredoxin:NADP-oxidoreductase (FNR) enzymes in-vitro. Proceedings of the National Academy of Sciences 108, 9396 (2011). [0149] 19. C. E. Lubner et al., Solar hydrogen-producing bionanodevice outperforms natural photosynthesis. Proceedings of the National Academy of Sciences of the United States of America 108, 20988-20991 (2011). [0150] 20. M. Ihara et al., Light-driven Hydrogen Production by a Hybrid Complex of a [NiFe] Hydrogenase and the Cyanobacterial Photosystem I. Photochemistry and Photobiology 82, 676-682 (2006). [0151] 21. H. Krassen et al., Photosynthetic Hydrogen Production by a Hybrid Complex of Photosystem I and [NiFe] Hydrogenase. ACS Nano 3, 4055-4061 (2009). [0152] 22. C. E. Lubner, R. Grimme, D. A. Bryant, J. H. Golbeck, Wiring Photosystem I for Direct Solar Hydrogen Production. Biochemistry 49, 404-414 (2010). [0153] 23. L. M. Utschig, S. R. Soltau, D. M. Tiede, Light-driven hydrogen production from photosystem I-catalyst hybrids. Current Opinion in Chemical Biology 25, 1-8 (2015). [0154] 24. M. Ihara, H. Nakamoto, T. Kamachi, I. Okura, M. Maedal, Photoinduced Hydrogen Production by Direct Electron Transfer from Photosystem I Cross-Linked with Cytochrome c3 to [NiFe] Hydrogenase. Photochemistry and Photobiology 82, 1677-1685 (2006). [0155] 25. K. Gutekunst et al., In-vivo turnover frequency of the cyanobacterial NiFehydrogenase during photohydrogen production outperforms in-vitro systems. Scientific Reports 8 (2018). [0156] 26. E. C. Aubert-Jousset, Melissa; Guedeney, Genevieve; Richaud, Pierre; Cournac, Laurent, Role of HoxE subunit in Synechocystis PCC6803 hydrogenase. FEBS Journal 10.111/j.1742-4658.2011.08208.x (2011). [0157] 27. O. Schmitz et al., Molecular biological analysis of a bidirectional hydrogenase from cyanobacteria. European Journal of Biochemistry 233, 266-276 (1995). [0158] 28. L. T. Serebryakova, M. E. Sheremetieva, Characterization of catalytic properties of hydrogenase isolated from the unicellular cyanobacterium Gloeocapsa alpicola CALU 743.

[0159] Biochemistry-Moscow 71, 1370-1376 (2006). [0160] 29. K. Schneider, R. Cammack, H. G. Schlegel, Content and localization of FMN, F3-S clusters and nickel in the NAD-linked hydrogenase of Nocardia opaca 1B. European Journal of Biochemistry 142, 75-84 (1984). [0161] 30. C. Eckert et al., Genetic Analysis of the Hox hydrogenase in the Cyanobacterium Synechocystis sp. PCC 6803 Reveals Subunit Roles in Association, Assembly, Maturation, and Function. Journal of Biological Chemistry 287, 43502-43515 (2012). [0162] 31. E. Flores, J. E. Frfas, L. M. Rubio, A. Herrero, Photosynthetic nitrate assimilation in cyanobacteria. Photosynthesis Research 83, 117-133 (2005). [0163] 32. K. Gutekunst, R. Schulz, “CHAPTER 4 The Physiology of the Bidirectional NiFe Hydrogenase in Cyanobacteria and the Role of Hydrogen Throughout the Evolution of Life” in Microalgal Hydrogen Production: Achievements and Perspectives. (The Royal Society of Chemistry, 2018), 10.1039/9781849737128-00107, pp. 107-138. [0164] 33. J. Biggins, Respiration in Blue-Green Algae. Journal of Bacteriology 99, 570-575 (1969). [0165] 34. M. R. de Graef, S. Alexeeva, J. L. Snoep, M. J. Teixeira de Mattos, The Steady-State Internal Redox State (NADH/NAD) Reflects the External Redox State and Is Correlated with Catabolic Adaptation in Escherichia coli. Journal of Bacteriology 181, 2351-2357 (1999). [0166] 35. G. K. Kumaraswamy et al., Reprogramming the glycolytic pathway for increased hydrogen production in cyanobacteria: metabolic engineering of NAD+− dependent GAPDH. Energy & Environmental Science 6, 3722-3731 (2013). [0167] 36. K. McNeely, Y. Xu, N. Bennette, D. A. Bryant, G. C. Dismukes, Redirecting Reductant Flux into Hydrogen Production via Metabolic Engineering of Fermentative Carbon Metabolism in a Cyanobacterium. Appl. Environ. Microbiol. 76, 5032-5038 (2010). [0168] 37. W. Buckel, R. K. Thauer, Energy conservation via electron bifurcating ferredoxin reduction and proton/Na+ translocating ferredoxin oxidation. Biochim. Biophys. Acta-Bioenerg. 1827, 94-113 (2013). [0169] 38. C. C. Page, C. C. Moser, X. Chen, P. L. Dutton, Natural engineering principles of electron tunnelling in biological oxidation-reduction. Nature 402, 47-52 (1999). [0170] 39. J. M. Hudson et al., Electron Transfer and Catalytic Control by the Iron-Sulfur Clusters in a Respiratory Enzyme, E. coli Fumarate Reductase. Journal of the American Chemical Society 127, 6977-6989 (2005). [0171] 40. I. Grotjohann, P. Fromme, Structure of cyanobacterial photosystem I. Photosynthesis Research 85, 51-72 (2005). [0172] 41. H. Kubota-Kawai et al., X-ray structure of an asymmetrical trimeric ferredoxin-photosystem I complex. Nature Plants 4, 218-224 (2018). [0173] 42. P. Fromme, H. Bottin, N. Krauss, P. Sdtif, Crystallization and electron paramagnetic resonance characterization of the complex of photosystem I with its natural electron acceptor ferredoxin.

[0174] Biophysical Journal 83, 1760-1773 (2002). [0175] 43. P. Fromme, P. Jordan, N. KrauB, Structure of photosystem I. Biochim Biophys Acta 1507, 5-31 (2001). [0176] 44. P. Jordan et al., Three-dimensional structure of cyanobacterial photosystem I at 2.5 Å resolution. Nature 441, 909-917 (2001). [0177] 45. Y. Shomura et al., Structural basis of the redox switches in the NAD-reducing soluble [NiFe]-hydrogenase. Science 357, 928 (2017). [0178] 46. V. P. Chitnis, A. Ke, P. R. Chitnis, The PsaD Subunit of photosystem I (Mutations in the Basic Domain Reduce the Level of PsaD in the Membranes). Plant Physiology 115, 1699-1705 (1997). [0179] 47. B. Lagoutte, J. Hanley, H. Bottin, Multiple Functions for the C Terminus of the PsaD Subunit in the cyanobacterial photosystem I Complex. Plant Physiology 126, 307 (2001). [0180] 48. J. M. Schuller et al., Structural adaptations of photosynthetic complex I enable ferredoxin-dependent electron transfer. Science 363, 257 (2019). [0181] 49. A. Kothari, R. Potrafka, F. Garcia-Pichel, Diversity in hydrogen evolution from bidirectional hydrogenases in cyanobacteria from terrestrial, freshwater and marine intertidal environments.

[0182] Journal of Biotechnology 162, 105-114 (2012). [0183] 50. D. Hoffmann, J. Maldonado, M. F. Wojciechowski, F. Garcia-Pichel, Hydrogen export from intertidal cyanobacterial mats: sources, fluxes and the influence of community composition.

[0184] Environmental Microbiology 17, 3738-3753 (2015). [0185] 51. L. C. Burow et al., Identification of Desulfobacterales as primary hydrogenotrophs in a complex microbial mat community. Geobiology 12, 221-230 (2014). [0186] 52. D. G. Gibson et al., Enzymatic assembly of DNA molecules up to several hundred kilobases.

[0187] Nature Methods 6, 343-U341 (2009). [0188] 53. V. N. Noskov et al., Defining the minimal length of sequence homology required for selective gene isolation by TAR cloning. Nucleic Acids Research 29 (2001). [0189] 54. R. M. Q. Shanks, N. C. Caiazza, S. M. Hinsa, C. M. Toutain, G. A. O'Toole, Saccharomyces cerevisiae-based molecular tool kit for manipulation of genes from gram-negative bacteria. Applied and Environmental Microbiology 72, 5027-5036 (2006). [0190] 55. R. D. Gietz, R. H. Schiestl, High-efficiency yeast transformation using the LiAc/SS carrier DNA/PEG method. Nature Protocols 2, 31-34 (2007). [0191] 56. J. G. K. Williams, CONSTRUCTION OF SPECIFIC MUTATIONS IN PHOTOSYSTEM-II PHOTOSYNTHETIC REACTION CENTER BY GENETIC-ENGINEERING METHODS IN SYNECHOCYSTIS-6803. Methods in Enzymology 167, 766-778 (1988).