Cyanobacterial-Type, Heteropentameric, NAD+-Reducing NiFe Hydrogenase in the Purple Sulfur Photosynthetic Bacterium Thiocapsa roseopersicina

Gábor Rákhely; Ákos T Kovács; Gergely Maróti; Barna D Fodor; Gyula Csanádi; Dóra Latinovics; Kornél L Kovács

doi:10.1128/AEM.70.2.722-728.2004

. 2004 Feb;70(2):722–728. doi: 10.1128/AEM.70.2.722-728.2004

Cyanobacterial-Type, Heteropentameric, NAD⁺-Reducing NiFe Hydrogenase in the Purple Sulfur Photosynthetic Bacterium Thiocapsa roseopersicina

Gábor Rákhely ¹, Ákos T Kovács ¹, Gergely Maróti ¹, Barna D Fodor ¹, Gyula Csanádi ¹, Dóra Latinovics ¹, Kornél L Kovács ^1,^*

PMCID: PMC348915 PMID: 14766547

Abstract

Structural genes coding for two membrane-associated NiFe hydrogenases in the phototrophic purple sulfur bacterium Thiocapsa roseopersicina (hupSL and hynSL) have recently been isolated and characterized. Deletion of both hydrogenase structural genes did not eliminate hydrogenase activity in the cells, and considerable hydrogenase activity was detected in the soluble fraction. The enzyme responsible for this activity was partially purified, and the gene cluster coding for a cytoplasmic, NAD⁺-reducing NiFe hydrogenase was identified and sequenced. The deduced gene products exhibited the highest similarity to the corresponding subunits of the cyanobacterial bidirectional soluble hydrogenases (HoxEFUYH). The five genes were localized on a single transcript according to reverse transcription-PCR experiments. A σ⁵⁴-type promoter preceded the gene cluster, suggesting that there was inducible expression of the operon. The Hox hydrogenase was proven to function as a truly bidirectional hydrogenase; it produced H₂ under nitrogenase-repressed conditions, and it recycled the hydrogen produced by the nitrogenase in cells fixing N₂. In-frame deletion of the hoxE gene eliminated hydrogen evolution derived from the Hox enzyme in vivo, although it had no effect on the hydrogenase activity in vitro. This suggests that HoxE has a hydrogenase-related role; it likely participates in the electron transfer processes. This is the first example of the presence of a cyanobacterial-type, NAD⁺-reducing hydrogenase in a phototrophic bacterium that is not a cyanobacterium. The potential physiological implications are discussed.

Hydrogenases catalyze the simple redox reaction H₂ ↔ 2H⁺ + 2e⁻. These enzymes are grouped on the basis of their metal content; some hydrogenases contain only iron atoms (Fe hydrogenases), but the majority of known hydrogen-activating enzymes contain nickel and iron at the active center (some NiFe hydrogenases also contain selenium), and one enzyme that displays hydrogenase activity in methanogens has been shown to contain no redox-active metal at all (29, 38, 40). The protein core of the NiFe hydrogenases is composed of at least two subunits. The small subunit harbors the electron-transferring Fe-S clusters, and the large subunit contains the heterobinuclear NiFe metallocenter. In the active center the Fe is coordinated by one CO and two CN ligands (39). Biosynthesis of an active hydrogenase involves complex posttranslational processing, which includes assembly of the NiFe centers, insertion of the CO and CN ligands, proteolytic cleavage of the C-terminal end of the large subunit by an endoprotease, and biosynthesis and orientation of the Fe-S clusters (9, 23). This maturation process requires the concerted action of several accessory proteins.

Hydrogenases may differ in electron carrier specificity, in cellular localization, and in regulation of expression. Hydrogenases are involved in energy conservation, in the disposal of excess electrons formed during fermentation processes, or in hydrogen sensing as a component of an H₂-dependent molecular signal transduction cascade, which regulates the expression of many hydrogenases (14, 29, 38.)

In a number of species, more than one hydrogenase has been described. For example, in Escherichia coli there are four membrane-associated hydrogenases (two belonging to one group and two belonging to another group) (3, 29, 38). In Ralstonia eutropha one membrane-bound hydrogenase, one heterotetrameric cytoplasmic NAD⁺-reducing hydrogenase, and one regulatory hydrogenase have been described, and each of these enzymes belongs to a distinct NiFe hydrogenase family (14, 29, 38). Cyanobacteria have a special type of NAD⁺-reducing hydrogenases, in which an additional subunit, HoxE, is a member of the enzyme complex (HoxEFUYH) (34, 36). Most microorganisms apparently contain designated hydrogenases for diverse physiological tasks, but cross-reactions may occur as well (29).

Thiocapsa roseopersicina BBS is a purple sulfur photosynthetic bacterium (4). Two membrane-bound hydrogenases (HupSL and HynSL) have been identified in this microorganism (10, 28); both of these enzymes belong to the same group (29, 38). HynSL, which was characterized at the protein and gene levels in detail previously (19, 20, 28), is an unusually stable enzyme. It is remarkably active at high temperatures and is resistant to oxygen, proteases, and detergents (19). The organization of the hynSL genes is also extraordinary, as the genes of the small and large subunits are separated by two open reading frames (isp1 and isp2), which seem to code for the components of a transmembrane redox complex (28). No hydrogenase accessory gene was found in the vicinity of hynSL, while downstream of hupSL several genes homologous to specific accessory genes were identified (10). Seven genes coding for accessory proteins involved in the biosynthesis of hydrogenases were isolated after transposon mutagenesis in the genome of T. roseopersicina (13, 24). These genes are clustered in various loci.

The physiological role of the HynSL hydrogenase is not known, but this enzyme is believed to play a role in the maintenance of the redox balance of the cells. The other hydrogenase (HupSL) has been characterized mainly at the gene level (10), since it is difficult to purify. The putative hupSL gene products resemble the uptake hydrogenases of other microbes, which recycle the hydrogen produced by the nitrogenase during nitrogen fixation.

To establish the physiological functions of the HynSL and HupSL hydrogenases, their structural genes were deleted in the present study. Surprisingly, the cells still had hydrogenase activity, which was localized in the cytoplasm. Characterization of a gene cluster which encodes a heteropentameric NAD⁺-reducing hydrogenase related to the cyanobacterial bidirectional hydrogenases is described below.

(Preliminary results were presented at the Biohydrogen 2002 Conference, Ede-Wageningen, The Netherlands, 21 to 24 April 2002, and have been reviewed by Kovács et al. [20a].)

MATERIALS AND METHODS

Bacterial strains and plasmids.

The strains and plasmids used are listed in Table 1. T. roseopersicina strains were grown photoautotrophically and anaerobically in liquid cultures for 3 to 4 days in Pfennig's mineral medium supplemented with 0.1% NH₄Cl (PC medium) (18). For nitrogen-fixing conditions, the NH₄Cl was omitted. The medium was solidified with 7 g of Phytagel (Sigma) per liter, and acetate (2 g/liter) was also added when transconjugants were selected. Plates were incubated for 2 weeks in anaerobic jars by using the GasPack (BBL) or AnaeroCult (Merck) system. Cultures were illuminated with continuous light at 27 to 30°C (18). E. coli strains were maintained on Luria-Bertani medium (30). Antibiotics were used at the following concentrations: 50 μg of streptomycin per ml, 100 μg of ampicillin per ml, 50 μg of kanamycin per ml, 20 μg of gentamicin per ml, and 50 μg of erythromycin per ml for E. coli; and 20 μg of kanamycin per ml, 5 μg of streptomycin per ml, 5 μg of gentamicin per ml, and 50 μg of erythromycin per ml for T. roseopersicina.

TABLE 1.

Strains and constructs used in this study

Strain or plasmid	Relevant genotype and/or phenotype	Reference or source
Thiocapsa roseopersicina strains
BBS	Wild type	4
M539	hypF::Km^r	13
GB11	hynSLΔ::Sm^r	This study
GB21	hupSLΔ::Gm^r	This study
GB1121	hynSLΔ::Sm^rhupSLΔ::Gm^r	This study
GB112131	hynSLΔ::Sm^rhupSLΔ::Gm^r ΔhoxH::Em^rerm (Em^r) oriented as hox	This study
GB112132	hynSLΔ::Sm^rhupSLΔ::Gm^r ΔhoxH::Em^rerm (Em^r) oriented opposite the hox orientation	This study
HoxEDM	hynSLΔ::Sm^rhupSLΔ::Gm^r ΔhoxE	This study
Escherichia coli strains
XL1-Blue MRF′	Δ(mcrA)183 Δ(mcrCB-hsdSMR-mrr)173 endA1 supE44 thi-1 recA1 gyrA96 relA1 lac [F′ proAB lacI^qZΔM15 Tn10 (Tet^r)]^c	Stratagene
S17-1(λpir)	294 (recA pro res mod) Tp^r Sm^r (pRP4-2-Tc::Mu-Km::Tn7) λpir	15
Plasmids
pBluescript SK (+)	Amp^r, cloning vector, ColE1	Stratagene
pHRP317	Km^r Ω Sm^r/Sp^r	27
pK18mobsacB	Km^rsacB RP4 oriT ColE1 ori	31
pRL271	Cloning vector carrying sacB, Em^r (ermC) Cm^r	GenBank accession no. L05081
p34S-Gm	oriR, Amp^r Gm^r	11
pHOXED2	Upstream and downstream region of hoxE in pK18mobsac; construct for in-frame deletion of hoxE	This study
pHOXHD3	Upstream and downstream region of hoxH separated by the Em^r gene in pK18mobsac; construct for replacing the hoxH genes with Em^r; the gene of Em^r is oriented like hox genes	This study
pHOXHD4	Upstream and downstream region of hoxH separated by the Em^r gene in pK18mobsac; construct for replacing the hoxH genes with Em^r; the gene of Em^r is oriented opposite the orientation of the hox genes	This study
pΔhydSm	Upstream region of hynS and downstream region of hynL separated by the Sm^r gene in pK18mobsac; construct for replacing the hynS, isp1, isp2, and hynL genes with the Sm^r gene	This study
pKUSDSGm	Upstream region of hupS and downstream region of hupL separated by the Gm^r gene in pK18mobsac; construct for replacing the hupSL genes with the Gm^r gene	This study
pTCB4/2	Amp^r; 11,927-bp BgIII fragment; pBluescript SK+ BamHI site	This study

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Molecular biology techniques.

Standard recombinant DNA techniques were performed as described previously (2, 30) or by using the specifications of the manufacturers.

Deletion of the hynS, isp1, isp2, and hynL genes.

The 929-bp PstI-HindIII fragment, which was located 448 bp downstream of the hynL gene, from pAK41 (28) was cloned into pK18mobsacB (31). The resulting construct was designated pTS3′ and carried a 929-bp genomic region from T. roseopersicina. The 956-bp EcoRI-SalI fragment from pTSH2/8, containing a 935-bp homologous fragment 138 bp upstream from hynS, was cloned into pTS3′ (pΔhyd). The streptomycin resistance cassette from pHRP317 (27) was cloned as a polished HindIII fragment into SalI-cleaved, blunted pΔhyd, resulting in pΔhydSm. Thus, a streptomycin resistance cassette was inserted between the homologous genomic regions found upstream and downstream of the hyn operon. This plasmid was conjugated into T. roseopersicina BBS, as described previously (13), and kanamycin-resistant single recombinants were selected. The genotype of a clone was confirmed by PCR and Southern blotting. This recombinant clone was grown in liquid medium with streptomycin and plated on PC medium supplemented with streptomycin and 3% sucrose. Streptomycin- and sucrose-resistant, kanamycin-sensitive colonies were selected, and the genotype was confirmed by Southern blotting and hybridization. The clone in which the hyn operon was replaced by the streptomycin resistance cassette was selected and designated GB11.

Deletion of the hupSL genes.

A 1,100-bp fragment upstream from the hupS gene was amplified from the genome, using primers OHUP3 (5′ CACCGCCTTGCAGCTGTCGGC 3′) and OHUP4 (5′ CTCGAAATCCGGAAAGGCTC 3′), and was cloned into the SmaI site of pBluescript SK+ (pBUS). A 1,230-bp fragment covering the downstream region of the hupL gene was obtained by PCR by using genomic DNA template and primers OHUP5 (5′ TCGCATCAAGGTCCGCTGAA 3′) and OHUP6 (5′ GCCGGGCACCATGAAGCGGG 3′). The PCR product was digested with SalI, and a 1,130-bp fragment was cloned into the EcoRV-SalI site of pBUS (pBUSDS). The pBUSDS vector was cut by PstI and polished, and a blunted 0.9-kb KpnI fragment of p34S-Gm (11) containing the gentamicin resistance gene was inserted into it (pBUSDSGm). This was digested with KpnI, blunted, and cleaved with XbaI, which yielded a 3-kb fragment, which was cloned into the SmaI-XbaI site of pK18mobsacB (pKUSDSGm). The construct was introduced into T. roseopersicina BBS and GB11, and double recombinants were selected as described above for the hynSL deletion. All genotypes were confirmed by Southern blotting and PCR by using appropriate primers. The ΔhynSL ΔhupSL strain was designated GB1121.

Partial purification of the hydrogenase from the soluble fraction of the cells.

After 4 to 5 days of photoautotrophic growth, the cells were centrifuged in a CEPA cross-flow centrifuge, which yielded 60 to 90 g of wet bacterial biomass per 60 liters. Cells were lysed in 20 mM potassium phosphate buffer (pH 7.0) by using an MSE PB-620 ultrasonic disintegrator. Cell debris and sulfur crystals were removed by centrifugation (4°C, 15,000 × g, 10 min). The soluble and membrane fractions were separated by ultracentrifugation of the supernatant at 4°C and 100,000 × g for 3 h. The soluble fraction was heat treated at 60°C for 20 min, and the heat-labile protein precipitate was removed by centrifugation. In the subsequent sequential (NH₄)₂SO₄ precipitation experiments, the activity was collected from the protein fraction precipitates between 25 and 35% (vol/vol) saturated (NH₄)₂SO₄. Concentration and desalting were performed with an Amicon stirred cell with a PM-30 membrane (Millipore) under N₂. Column chromatography was performed by using Bio-Rad DuoFlow liquid chromatography equipment fitted with a Bio-Rad 2128 fraction collector. For the first two separation steps 12-ml Sepharose Q columns were eluted with 50 mM Tris-HCl (pH 8.5)-1 M NaCl and with 20 mM potassium phosphate (pH 7.0)-1 M NaCl. In a third chromatographic separation a 12-ml DEAE column and an elution buffer containing 50 mM Tris-HCl (pH 9.4) and 1 M NaCl were employed. All chromatographic manipulations were performed under air. The active fractions were combined and then concentrated and desalted with an Amicon PM-30 membrane, as described above.

The protein concentration was determined as described by Lowry et al. (22). Protein patterns were analyzed on either a sodium dodecyl sulfate-12% polyacrylamide gel electrophoresis gel or a 5 to 20% linear sodium dodecyl sulfate gradient gel (2), and this was followed by silver staining (35). Western blotting experiments (2) were performed by using antibody raised against HoxH of Synechocystis sp. strain 6803 (kindly provided by Rudiger Schulz, University of Kiel, Kiel, Germany) (1) and R. eutropha (a gift from Bärbel Friedrich and Oliver Lenz, Humboldt University, Berlin, Germany).

Hydrogen uptake activity assay in vitro.

H₂ uptake was assayed spectrophotometrically by using benzyl viologen or methyl viologen as a redox dye. The samples were taken up in 20 mM potassium phosphate buffer (pH 7.0). Two milliliters of a sample was placed into a cuvette, 20 μl of 20 mM benzyl viologen was added, and the cuvettes were sealed with SubaSeal rubber stoppers. The assay was initiated by flushing the gas phase with N₂ and H₂ repeatedly (24).

Hydrogen evolution assay.

The samples were mixed or suspended in 1.2 ml of 20 mM potassium phosphate buffer (pH 7.0) in Hypo-Vials (volume, 10 cm³; Pierce), and 1 ml of 1 mM methyl viologen was added. The gas phase was flushed with N₂ for 10 min, and this was followed by anaerobic addition of 0.5 ml of a 0.1-g ml⁻¹ dithionite solution. Hydrogen evolution was measured by gas chromatography (13).

Hydrogen production in vivo.

The cells were anaerobically grown in Hypo-Vials (Pierce), and the hydrogen content of the gas phase was determined by gas chromatography at various times during growth. In each set of experiments the volume of the cultures and the ratio of the gas phase to the liquid phase were kept constant.

Isolation of the hox gene cluster.

The following primers were designed on the basis of the amino acid sequence data: SHO1 (5′ AGCAYGTSGAIATCGTNTA 3′ and SHO2 (5′ GGGAAGAASACRTCSACC 3′) (I is deoxyinosine, N is A, G, C, or T, R is A or G, S is C or G, and Y is C or T). A 380-bp fragment was amplified, cloned, and sequenced. The deduced amino acid sequence coincided with the protein sequence data. This fragment was used as a probe for isolation of genes coding for the soluble hydrogenase. A partial genomic library was prepared from BglII-digested genomic DNA of T. roseopersicina in the pBluescript SK+ BamHI site. This library was screened with the labeled probe, and a clone giving a positive signal (pTCB4/2) was selected. This clone was subcloned, and finally both strands of an 11,927-bp region were sequenced.

Deletion of the hoxH gene.

From pRL271 (GenBank accession number L05081), a 914-bp blunted EcoRI-SalI fragment containing the erythromycin resistance gene was cloned into the XhoI-digested, polished, vectorial fragment of pTCB4/2 (harboring 404 bp from the 3′ end of the hoxH gene and 843 bp of the downstream region [see Fig. 2]), yielding pHOXHD1/2, in which the erythromycin resistance gene was in the sense direction with respect to the hox genes. The 1,319-bp blunted PstI fragment of pTCB4/2 (1,259-bp upstream region plus 60 bp from the 5′ end of the hoxH gene [see Fig. 2]) was cloned into the SmaI site of pK18mobsac (pHOXHD2). A 2,188-bp NotI-KpnI fragment from pHOXHD1/2 was blunted and ligated into the polished EcoRI site of pHOXHD2, resulting in pHOXHD3, in which the orientations of the upstream and downstream regions of hoxH were the same. This construct was introduced into GB1121, and double recombinants were selected by analogy as described above. The hynSLΔ::Sm hupSLΔ::Gmr ΔhoxH::Emr mutant was designated GB112131. The whole procedure was repeated so that the erythromycin gene was oriented in the direction opposite that of the hox genes, which yielded the pHOXHD1/1 and pHOXHD4 constructs and the hynSLΔ::Smr hupSLΔ::Gmr ΔhoxH::Emr triple mutant strain designated GB112132.

In-frame deletion of the hoxE gene.

The 1,538-bp PstI-EcoRI fragment of pTCB4/2 (downstream region of hoxE [see Fig. 2]) was cloned into PstI-EcoRI-digested pK18mobsac (pHOXED1). The 1,558-bp FspI-PstI fragment of pTCB4/2 (upstream region of hoxE [see Fig. 2]) was inserted into the PstI and HindIII sites (filled up) of pHOXED1, which yielded pHOXED2, which was conjugated into the T. roseopersicina GB1121 strain. Single recombinants were selected on kanamycin-containing plates. One clone was grown under nonselective conditions, and then the double recombinants were screened for kanamycin sensitivity. The genotype of the hynSLΔ::Smr hupSLΔ::Gmr ΔhoxE mutant, designated HoxEDM, was confirmed by sequencing the corresponding genomic region which was amplified by PCR.

RNA isolation and reverse transcription-coupled PCR.

RNA was isolated with the TRIzol reagent (Gibco BRL) by following the manufacturer's recommendations. Prior to reverse transcription, the RNA was treated with DNase I (4 U of RNase-free Dnase I in 40 mM Tris-HCl [pH 7.5]-20 mM MgCl₂-20 mM CaCl₂ at 37°C for 60 min). The mixture was extracted with phenol-chloroform, and after precipitation, the RNA was dissolved in 20 μl of H₂O. Reverse transcription-PCR experiments were carried out as previously described (13). The reverse transcription was initiated at primer TCHO24 (5′ ACTTGGCGTCTTCGACCTCC 3′) located at the 5′ end of the hoxH gene, while the PCR was performed with the following primers: TCHO27 (5′ CTGCAGGAGCGCTACGACAT 3′) in the hoxE gene and TCHO9 (5′ GGATGACTGACAGCTGGCCGCGAGG 3′) in the hoxF gene.

Nucleotide sequence accession number.

The nucleotide sequence of clone pTCB4/2 has been deposited in the GenBank database under accession number AY214929.

RESULTS AND DISCUSSION

Detection of a third hydrogenase activity in T. roseopersicina.

Two gene clusters (hupSL and hynS-isp1-isp2-hynL), each encoding a membrane-associated NiFe hydrogenase, have been found previously in T. roseopersicina (10, 28). The structural genes coding for the two enzymes were deleted in the present study, and much to our surprise, in the absence of the membrane-associated NiFe hydrogenases the cells produced molecular hydrogen and the cultures grew uninhibited. The hydrogen production data for the wild type, GB1121 (ΔhupSL ΔhynSL), and the hypF mutant strains (13) under nitrogen-fixing and nitrogenase-repressed conditions are compared in Fig. 1.

FIG. 1. — Normalized in vivo hydrogen production by *T. roseopersicina* BBS (wild type [wt]) and the GB1121 (Δ*hynSL* Δ*hupSL*) and *hypF*::Tn5 mutant strains cultivated under nitrogenase-repressed or nitrogen-fixing conditions. The cells were inoculated into 15-ml portions of media in 30-ml Hypo-Vials, which were flushed with N₂ and incubated with continuous illumination at 30°C for 5 days. The H₂ concentration in the gas phase was determined by gas chromatography and was normalized for the optical densities of the cultures. The amount of H₂ produced by the wild-type strain divided by the optical density at 600 nm was considered 100%, and the other values were compared to this.

From these experiments the following conclusions were apparent. (i) Under nitrogenase-repressed conditions, an NiFe hydrogenase evolved the unexpected hydrogen, since H₂ production was not observed in the hypF mutant (HypF is a pleiotropic protein that is essential for maturation of all NiFe hydrogenases). The GB1121 strain produced practically the same amount of hydrogen as the wild type produced; hence, the NiFe hydrogenase involved was neither Hyn nor Hup. (ii) Under nitrogen-fixing conditions, when hydrogen was generated predominantly by the nitrogenase, much less hydrogen was produced by the GB1121 mutant (ΔhupSL ΔhynSL) than in the absence of all NiFe hydrogenases (hypF mutant). It is reasonable to assume that the hydrogen production capacity of the nitrogenase complex was the same in both cases; therefore, an NiFe hydrogenase must have been responsible for H₂ recycling (i.e., it accounted for the difference between the values for GB1121 and M539 in Fig. 1). The NiFe enzyme carrying out this function is a truly bidirectional NiFe hydrogenase, which produces and recycles H₂ depending on the physiological requirements of the cell. It is shown below (see Fig. 4) that this hydrogenase is indeed the Hox enzyme.

FIG. 4. — Characterization of the hydrogenase activities of various *T. roseopersicina* mutants in vivo and in vitro. The genotypes of the mutants used are as follows: GB1121, *hynSL*Δ::Sm^r *hupSL*Δ::Gm^r; GB112131, *hynSL*Δ::Sm^r *hupSL*Δ::Gm^r Δ*hoxH::*Er^r; HoxEDM, *hynSL*Δ::Sm^r *hupSL*Δ::Gm^r Δ*hoxE.* (A) Relative hydrogen production by the cells grown under nitrogenase-repressed, photosynthetic conditions after 5 days. (B) Relative in vitro uptake hydrogenase activities of the same cells with benzyl viologen as an artificial electron acceptor. In both cases the values obtained for the GB1121 strain were defined as 100%.

Purification of an NiFe hydrogenase from the soluble fraction.

Although the enzyme purification protocol has not been fully optimized yet, each chromatographic step resulted in a considerable (i.e., >20-fold) increase in the specific activity without a loss of a significant amount of the total activity. In the best sample, more than 800-fold purification was achieved, although the protein yield was rather low (1 mg of partially purified hydrogenase from 3,000 mg of starting protein).

According to our preliminary characterization, the stability of this hydrogenase of T. roseopersicina is outstanding with regard to both oxygen inactivation and heat inactivation.

In many cases, the soluble hydrogenases catalyze the hydrogen-dependent reduction of NAD⁺ or NADP⁺ (29, 38). The fact that we could detect hydrogen-dependent NAD⁺ reduction in the soluble fraction raised the possibility that our new hydrogenase is a Hox type of enzyme.

The presence and location of the HoxH-type polypeptide were further confirmed by Western hybridization experiments in which the HoxH antibody of Synechocystis sp. strain 6803 was used (1). However, no signal was obtained with the antibody raised against R. eutropha HoxH. The corresponding protein band was excised for sequence analysis with a Pulsar hybrid tandem mass spectrometer with QqTOF geometry. A detailed description of the purification and biochemical properties of the Hox enzyme awaits a better purification yield and will be published elsewhere.

Isolation of the gene cluster coding for a heteropentameric hydrogenase in T. roseopersicina.

Primers were designed on the basis of the protein sequence data and were used to amplify a probe for Southern and colony hybridization experiments. A 12-kb chromosomal locus was isolated with this probe, and numerous ORFs were found (Fig. 2). Five of these ORFs (designated hoxEFUYH) coded for putative proteins showing significant homology to the bidirectional NAD⁺-reducing hydrogenases of cyanobacteria (16, 17, 32) (Table 2). The HoxF and HoxU subunits are responsible for diaphorase activity, while HoxY and HoxH represent the NiFe hydrogenase small and large subunits, respectively. Upstream, close to the hoxF gene, a hoxE-like gene was recognized. Interestingly, there was considerable resemblance between the HoxEFU subunits and the diaphorase subunits of the Fe-only hydrogenases of Desulfovibrio fructosovorans (12) and the hyperthermophilic bacterium Thermotoga maritima (25). The function of the putative hoxE gene product is unclear, but its similarity to the NADH:ubiquinone oxidoreductase 24-kDa NuoE subunit is remarkable. It should be noted that HoxF and HoxU exhibit similarity to NuoF and NuoG, respectively, and the N terminus of HoxF also resembles the NuoE subunit. These subunits are at the NAD-reactive end of the NADH:ubiquinone oxidoreductase complex (26). The homology suggests that there may be a respiration-related function. A lower level of relatedness to the NAD⁺-reducing hydrogenase of R. eutropha (37) was found for the deduced HoxFUYH subunits. These results coincide well with the data obtained in the Western hybridization experiments (data not shown).

TABLE 2.

Levels of amino acid identitiy of the subunits of the T. roseopersicina NAD⁺-reducing hydrogenase and other hydrogenases

Organism	Amino acid identity (%)					Refer- ence
Organism	HoxE	HoxF	HoxY	HoxU	HoxH	Refer- ence
A. variabilis		56	59	49	56	32
Nostoc sp. strain PCC 7120	54	56	59	48	57	17
Synechocystis sp. strain PCC 6803	57	56	54	51	56	16
T. maritima	44	50	37			25
D. fructosovorans	42	51	36			12
R. eutropha		31	33	43	40	37

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Additional ORFs were identified upstream from the hoxE gene, but none of them was related to known genes implicated in hydrogenase biosynthesis (Fig. 2). A short ORF (orf2) fit perfectly between the hoxY and hoxH genes, but the putative gene product had no homologous counterpart in the databanks. Downstream of the hoxH gene orf3 was found, which may be related to the genes encoding hydrogenase-processing proteases. However, the sequence homology is low; therefore, the assumed function of the putative gene product needs additional experimental verification. Site-directed mutagenesis is in progress to determine whether Orf3 is involved in NiFe hydrogenase biosynthesis.

Organization of the hox genes.

Apart from the presence of orf2, the hox genes form a contiguous gene cluster. The cyanobacterial counterpart genes are arranged in a more irregular fashion. In Anabaena sp. strain PCC 7120 or Synechococcus sp. strain PCC 6301 the hoxF and hoxU genes are separated by 8.8 kb and at least 16 kb, respectively (36). Often additional ORFs are inserted between the hox genes; e.g., in Synechocystis sp. strain PCC 6803 one ORF is located between hoxF and hoxU, and two ORFs were found between hoxY and hoxH (36). The role of the putative gene products and the physiological relevance of the numerous insertions are not known. Therefore, we tested whether the hoxEFUYH genes are on one transcript by performing reverse transcription-PCR experiments. For reverse transcription, a primer in the hoxH gene was used, and for PCR separate primers in the hoxE and hoxF genes were used. Figure 3A shows the cotranscription of the hox genes. The results did not eliminate the possibility that there is an additional internal promoter, although this possibility is unlikely. Cotranscription of the entire hox gene cluster raises the possibility that the putative protein product of orf2 may also participate in the Hox enzyme complex. A single transcriptional unit was described in Anabaena variabilis ATCC 29413, but two independent transcripts were detected in an Anacystis nidulans strain (6). In T. roseopersicina, upstream from the hoxE gene a typical σ⁵⁴-type promoter was identified (Fig. 3B), suggesting that expression of the hox genes is regulated (8). It has been shown that in Synechocystis sp. strain PCC 6803 expression of the hox genes is directed by the circadian rhythms (33). In R. eutropha a hydrogen-sensing signal transduction cascade controls the biosynthesis of the soluble NAD⁺-reducing hydrogenase (21).

FIG. 3. — (A) Reverse transcription-PCR analysis of the *hox* operon. Reverse transcription was initiated at the 5′ end of the *hoxH* gene, while the PCR was performed with primers located in the *hoxE* and *hoxF* genes. Lane RT+, reverse transcriptase present in the reverse transcription reaction; lane RT−, reverse transcriptase not present in the reverse transcription reaction; lane M, marker; lane gC, genomic control. (B) Promoter region of the *hoxE* gene. The elements of the σ⁵⁴-type promoter are underlined; the ribosomal binding site is indicated by boldface italic type.

Hox hydrogenase is responsible for in vivo hydrogen evolution activity under nitrogenase-repressed conditions.

Attempts to delete the entire hox gene cluster failed for unknown reasons. Hence, deletion of the large subunit-encoding gene was used to inactivate Hox. Around 1 kb of the hoxH gene was removed and replaced by the erythromycin resistance gene in the GB1121 (ΔhynSL ΔhupSL) mutant strain. As shown in Fig. 4A, the level of in vivo hydrogen production by the GB112131 (GB1121 ΔhoxH) triple mutant was less than 1% of the value measured for the GB1121 strain. The in vitro uptake hydrogenase activity of the triple mutant was also practically zero (Fig. 4B). These experiments provide unequivocal evidence that the Hox hydrogenase is responsible for hydrogen consumption in the GB1121 double mutant (ΔhynSL ΔhupSL) under nitrogen-fixing conditions and for H₂ production under nitrogenase-repressed conditions (Fig. 1). It should be noted that neither the in vivo hydrogen production nor the in vitro hydrogenase activity disappeared completely; very weak (less than 0.5% of the Hox enzyme activity) but reproducible hydrogen production and hydrogenase activity could be measured. Similar data were obtained with the GB112132 (ΔhynSL ΔhupSL ΔhoxH) strain, in which the erythromycin resistance gene was inverted. The origin of this hydrogenase activity is unclear, and the possibility that it is due to a unknown NiFe hydrogenase cannot be eliminated, since this activity cannot be detected in the hypF::Tn5 mutant strain (Fig. 1).

HoxE is required for in vivo hydrogen production but not for in vitro hydrogenase activity.

Very little is known about the possible function of the HoxE subunit. In Synechococcus sp. strain PCC 6301, HoxE copurified with the hydrogenase activity, suggesting that it has a hydrogenase-related function (34). In T. roseopersicina, the genomic and transcriptional context suggests that the HoxE subunit has a Hox hydrogenase-linked function as well. An in-frame deletion was introduced into the hoxE gene to determine the effect on the hydrogen production of the cells in vivo and on the in vitro activity of the Hox hydrogenase with redox dyes as artificial electron acceptors. Figure 4 clearly shows that specific disruption of the HoxE protein had deleterious effects on hydrogen production in the intact cells under nitrogenase-repressed conditions, while the in vitro hydrogenase activity remained unaltered. In the NAD⁺-reducing hydrogenases, the catalytic domains for H₂/H⁺ and NAD⁺/NADH redox reactions are located in the HoxH and HoxF subunits, respectively. Moreover, HoxE is not a component of the R. eutropha type of NAD⁺-reducing hydrogenase; hence, HoxE may not have a direct role in the catalytic function. It is therefore likely to be involved in electron transfer. The presence of a 2Fe-2S cluster binding domain in HoxE and the fact that its counterpart (NuoE) in the Nuo complex has an electron transport role (26) support this assumption. HoxE may also anchor the complex to the photosynthetic membrane under certain conditions, and/or it may simply stabilize the heteropentameric complex or at least its diaphorase part. Remarkably dissimilar results were obtained for Synechococcus sp. strain PCC 6301, in which inactivation of the HoxE protein had a dramatic effect on the hydrogenase activity in vitro as well (Gudrun Boison, University of Cologne, Cologne, Germany, personal communication). A comparison of the activities of HoxE mutants might provide deeper insight into the physiological function of the whole pentameric enzyme complex and its individual subunits.

Possible function of the bidirectional hydrogenase in T. roseopersicina.

The functions of the NAD⁺-reducing hydrogenase are dissimilar in R. eutropha (heterotetrameric NAD⁺-reducing enzymes) and in cyanobacteria. In R. eutropha, this enzyme is related to an energy-conserving mechanism (14, 29, 38), while in cyanobacteria the data obtained for various strains have led to inconsistent conclusions. First, based on experiments done with A. nidulans, it was suggested that HoxEFU act as the missing components (NuoEFG) of the Nuo complex in cyanobacteria, and these subunits were assumed to be used by both respiratory complex I and the bidirectional hydrogenase (5). Later, this hypothesis was revised since a mutation in HoxU and/or HoxF did not impair respiration. In A. nidulans and Nostoc sp. strain PCC 73102, both of which lack the hoxFU and nuoEFG genes, the respiratory rate was comparable to that in other cyanobacterial strains (5, 7). Appel and coworkers mutated the large subunit of the bidirectional hydrogenase in Synechocystis sp. strain PCC 6803 and observed impaired oxidation of photosystem I (1). Taking into account these and other experiments, the bidirectional hydrogenase was presumed to function as an electron valve during photosynthesis in this strain. This ambiguity is the basis of the dissimilar classifications of the cyanobacterial soluble heterotetrameric-heteropentameric NiFe hydrogenases (29, 38).

Since it is a purple sulfur photosynthetic bacterium, T. roseopersicina does not have photosystem II. It probably has respiratory complex type I, because the Nuo complex has been detected in the close relative Allochromatium vinosum (41). T. roseopersicina utilizes reduced sulfur compounds, which provide electrons for the cells. Taking these considerations together, one might speculate that the bidirectional hydrogenase has an electron valve role in this bacterium as well, but the excess electrons are derived from the reduced sulfur compounds. Preliminary results suggest that under non-nitrogen-fixing conditions, the in vivo hydrogen production of T. roseopersicina cells depends on the light-dark conditions and on the reduced sulfur compounds present in the cells (data not shown), which is consistent with the electron valve hypothesis. It might be important to note that this type of hydrogenase has been identified only in photosynthetic organisms (so far only in cyanobacteria), and based on this an interaction between photosynthesis and Hox hydrogenase seems to be logical. If the electron valve model is true, there is still the question of how the Hox hydrogenase is associated with the photosynthetic and/or other bioenergetic processes.

Acknowledgments

This work was supported by EU 5th Framework Programme projects QLK5-1999-01267, QLK3-2000-01528, QLK3-2001-01676, and ICA1-CT-2000-70026 and by OTKA, FKFP, OMFB, OM KFHÁT, and NKFP.

We thank Rudiger Schulz (University of Kiel, Kiel, Germany), as well as Bárbel Friedrich and Oliver Lenz (Humboldt University, Berlin, Germany), for Hox antibodies. We gratefully acknowledge Rózsa Verebély for excellent technical assistance.

REFERENCES

1.Appel, J., S. Phunpruch, K. Steinmüller, and R. Schulz. 2000. The bidirectional hydrogenase of Synechocystis sp. PCC 6803 works as an electron valve during photosynthesis. Arch. Microbiol. 173:333-338. [DOI] [PubMed] [Google Scholar]
2.Ausubel, F. M., R. Brent, R. E. Kingston, D. D. Moore, J. G. Seidman, J. A. Smith, and K. Struhl (ed.). 1996. Current protocols in molecular biology. John Wiley & Sons, New York, N.Y.
3.Bagramyan, K., N. Mnatsakanyan, A. Poladian, A. Vassilian, and A. Trchounian. 2002. The roles of hydrogenases 3 and 4, and the F0F1-ATPase, in H2 production by Escherichia coli at alkaline and acidic pH. FEBS Lett. 516:172-178. [DOI] [PubMed] [Google Scholar]
4.Bogorov, L. V. 1974. About the properties of Thiocapsa roseopersicina BBS, isolated from estuaria of White Sea. Mikrobiologija (Belgrade) 43:326-332. [PubMed] [Google Scholar]
5.Boison, G., O. Schmitz, B. Schmitz, and H. Bothe. 1998. Unusual gene arrangement of the bidirectional hydrogenase and functional analysis of its diaphorase subunit HoxU in respiration of the unicellular cyanobacterium Anacystis nidulans. Curr. Microbiol. 36:253-258. [DOI] [PubMed] [Google Scholar]
6.Boison, G., H. Bothe, and O. Schmitz. 2000. Transcriptional analysis of hydrogenase genes in the cyanobacteria Anacystis nidulans and Anabaena variabilis monitored by RT-PCR. Curr. Microbiol. 40:315-321. [DOI] [PubMed] [Google Scholar]
7.Boison, G., H. Bothe, A. Hansel, and P. Lindblad. 1999. Evidence against a common use of the diaphorase subunits by the bidirectional hydrogenase and by the respiratory complex I in cyanobacteria. FEMS Microbiol. Lett. 174:159-165. [Google Scholar]
8.Buck, M., M. T. Gallegos, D. J. Studholme, Y. Guo, and J. D. Gralla. 2000. The bacterial enhancer-dependent σ⁵⁴ (σ^N) transcriptional factor. J. Bacteriol. 182:4129-4136. [DOI] [PMC free article] [PubMed] [Google Scholar]
9.Casalot, L., and M. Rousset. 2001. Maturation of the [NiFe] hydrogenases. Trends Microbiol. 9:228-237. [DOI] [PubMed] [Google Scholar]
10.Colbeau, A., K. L. Kovács, J. Chabert, and P. M. Vignais. 1994. Cloning and sequencing of the structural (hupSLC) and accessory (hupDHI) genes for hydrogenase biosynthesis in Thiocapsa roseopersicina. Gene 140:25-31. [DOI] [PubMed] [Google Scholar]
11.Dennis, J. J., and G. J. Zylstra. 1998. Plasposons: modular self-cloning minitransposon derivatives for rapid genetic analysis of gram-negative bacterial genomes. Appl. Environ. Microbiol. 64:2710-2715. [DOI] [PMC free article] [PubMed] [Google Scholar]
12.Dermoun, Z., G. De Luca, M. Asso, P. Bertrand, F. Guerlesquin, and B. Guigliarelli. 2002. The NADP-reducing hydrogenase from Desulfovibrio fructosovorans: functional interaction between the C-terminal region of HndA and the N-terminal region of HndD subunits. Biochim. Biophys. Acta 1556:217-225. [DOI] [PubMed] [Google Scholar]
13.Fodor, B., G. Rákhely, Á. T. Kovács, and K. L. Kovács. 2001. Transposon mutagenesis in purple sulfur photosynthetic bacteria: identification of hypF, encoding a protein capable of processing [NiFe] hydrogenases in α, β, and γ subdivisions of the proteobacteria. Appl. Environ. Microbiol. 67:2476-2483. [DOI] [PMC free article] [PubMed] [Google Scholar]
14.Friedrich, B., and E. Schwartz. 1993. Molecular biology of hydrogen utilization in aerobic chemolithotrophs. Annu. Rev. Microbiol. 47:351-383. [DOI] [PubMed] [Google Scholar]
15.Herrero, M., V. Lorenzo, and K. N. Timmis. 1990. Transposon vectors containing non-antibiotic resistance selection markers for cloning and stable chromosomal insertion of foreign genes in gram-negative bacteria. J. Bacteriol. 172:6557-6567. [DOI] [PMC free article] [PubMed] [Google Scholar]
16.Kaneko, T., S. Sato, H. Kotani, A. Tanaka, E. Asamizu, Y. Nakamura, N. Miyajima, M. Hirosawa, M. Sugiura, S. Sasamoto, T. Kimura, T. Hosouchi, A. Matsuno, A. Muraki, N. Nakazaki, K. Naruo, S. Okumura, S. Shimpo, C. Takeuchi, T. Wada, A. Watanabe, M. Yamada, M. Yasuda, and S. Tabata. 1996. Sequence analysis of the genome of the unicellular cyanobacterium Synechocystis sp. strain PCC6803. II. Sequence determination of the entire genome and assignment of potential protein-coding regions. DNA Res. 3:109-136. [DOI] [PubMed] [Google Scholar]
17.Kaneko, T., Y. Nakamura, Y. C. P. Wolk, T. Kuritz, S. Sasamoto, A. Watanabe, M. Iriguchi, A. Ishikawa, K. Kawashima, T. Kimura, Y. Kishida, M. Kohara, M. Matsumoto, A. Matsuno, A. Muraki, N. Nakazaki, S. Shimpo, M. Sugimoto, M. Takazawa, M. Yamada, M. Yasuda, and S. Tabata. 2001. Complete genomic sequence of the filamentous nitrogen-fixing cyanobacterium Anabaena sp. strain PCC 7120. DNA Res. 8:205-213. [DOI] [PubMed] [Google Scholar]
18.Kovács, Á. T., G. Rákhely, and K. L. Kovács. 2003. Genes involved in the biosynthesis of photosynthetic pigments in the purple sulfur photosynthetic bacterium Thiocapsa roseopersicina. Appl. Environ. Microbiol. 69:3093-3102. [DOI] [PMC free article] [PubMed] [Google Scholar]
19.Kovács, K. L., and C. Bagyinka. 1990. Structural properties, functional states and physiological roles of hydrogenase in photosynthetic bacteria. FEMS Microbiol. Rev. 87:407-412. [Google Scholar]
20.Kovács, K. L., G. Tigyi, L. T. Thanh, S. Lakatos, Z. Kiss, and C. Bagyinka. 1991. Structural rearrangements in active and inactive forms of hydrogenase from Thiocapsa roseopersicina. J. Biol. Chem. 266:947-951. [PubMed] [Google Scholar]
20a.Kovács, K. L., B. Fodor, Á. T. Kovács, G. Csanádi, G. Maróti, J. Balogh, S. Arvani, and G. Rákhely. 2002. Hydrogenases, accessory genes and the regulation of [NiFe] hydrogenase biosynthesis in Thiocapsa roseopersicina. Int. J. Hydrogen Energy 27:1463-1469. [Google Scholar]
21.Lenz, O., and B. Friedrich. 1998. A novel multicomponent regulatory system mediates H₂ sensing in Alcaligenes eutrophus. Proc. Natl. Acad. Sci. 95:12474-12479. [DOI] [PMC free article] [PubMed] [Google Scholar]
22.Lowry, O. H., N. J. Rosenbourgh, A. L. Farr, and R. J. Randall. 1951. Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193:265-275. [PubMed] [Google Scholar]
23.Maier, T., and A. Böck. 1996. Nickel incorporation into hydrogenases, p. 173-192. In R. Hausinger, G. L. Eichlorn, and L. G. Marzilli (ed.), Advances in inorganic biochemistry. VHC Publishers Inc., New York, N.Y.
24.Maróti, G., B. D. Fodor, G. Rákhely, Á. T. Kovács, S. Arvani, and K. L. Kovács. 2003. Accessory proteins functioning selectively and pleiotropically in the biosynthesis of [NiFe] hydrogenases in Thiocapsa roseopersicina. Eur. J. Biochem. 270:2218-2227. [DOI] [PubMed] [Google Scholar]
25.Nelson, K. E., R. A. Clayton, S. R. Gill, M. L. Gwinn, R. J. Dodson, D. H. Haft, E. K. Hickey, J. D. Peterson, W. C. Nelson, K. A. Ketchum, L. McDonald, T. R. Utterback, J. A. Malek, K. D. Linher, M. M. Garrett, A. M. Stewart, M. D. Cotton, M. S. Pratt, C. A. Phillips, D. Richardson, J. Heidelberg, G. G. Sutton, R. D. Fleischmann, O. White, S. L. Salzberg, H. O. Smith, J. C. Venter, and C. M. Fraser. 1999. Evidence for lateral gene transfer between Archaea and bacteria from genome sequence of Thermotoga maritima. Nature 399:323-329. [DOI] [PubMed] [Google Scholar]
26.Nicholls, D. G., and S. J. Ferguson. 2001. Bioenergetics 3. Academic Press, New York, N.Y.
27.Parales, R. E., and C. S. Harwood. 1993. Construction and use of a new broad-host-range lacZ transcriptional fusion vector, pHRP309, for gram− bacteria. Gene 133:23-30. [DOI] [PubMed] [Google Scholar]
28.Rákhely, G., A. Colbeau, J. Garin, P. M. Vignais, and K. L. Kovács. 1998. Unusual organization of the genes coding for HydSL, the stable (NiFe) hydrogenase in the photosynthetic bacterium Thiocapsa roseopersicina BBS. J. Bacteriol. 180:1460-1465. [DOI] [PMC free article] [PubMed] [Google Scholar]
29.Robson. R. 2001. Biodiversity of hydrogenases, p. 9-32. In R. Cammack, M. Frey, and R. Robson (ed.), Hydrogen as a fuel. Taylor & Francis, London, United Kingdom.
30.Sambrook, J., T. Maniatis, and E. F. Fritsch. 1989. Molecular cloning: a laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
31.Schafer, A., A. Tauch, W. Jager, J. Kalinowski, G. Thierbach, and A. Puhler. 1994. Small mobilizable multi-purpose cloning vectors derived from the Escherichia coli plasmids pK18 and pK19: selection of defined deletions in the chromosome of Corynebacterium glutamicum. Gene 145:69-73. [DOI] [PubMed] [Google Scholar]
32.Schmitz, O., G. Boison, R. Hilscher, B. Hundeshagen, W. Zimmer, F. Lottspeich, and H. Bothe. 1995. Molecular biological analysis of a bidirectional hydrogenase from cyanobacteria. Eur. J. Biochem. 233:266-276. [DOI] [PubMed] [Google Scholar]
33.Schmitz, O., G. Boison, and H. Bothe. 2001. Quantitative analysis of expression of two circadian clock-controlled gene clusters coding for the bidirectional hydrogenase in the cyanobacterium Synechococcus sp. PCC7942. Mol. Microbiol. 41:1409-1417. [DOI] [PubMed] [Google Scholar]
34.Schmitz, O., G. Boison, H. Salzmann, H. Bothe, K. Schutz, S. H. Wang, and T. Happe. 2002. HoxE—a subunit specific for the pentameric bidirectional hydrogenase complex (HoxEFUYH) of cyanobacteria. Biochim. Biophys. Acta 1554:66-74. [DOI] [PubMed] [Google Scholar]
35.Shevchenko, A., M. Wilm, O. Vorm, and M. Mann. 1996. Mass spectrometric sequencing of proteins from silver-stained polyacrylamide gels. Anal. Chem. 68:850-858. [DOI] [PubMed] [Google Scholar]
36.Tamagnini, P., R. Axelsson, P. Lindberg, F. Oxelfelt, R. Wunschiers, and P. Lindblad. 2002. Hydrogenases and hydrogen metabolism of cyanobacteria. Microbiol. Mol. Biol. Rev. 66:1-20. [DOI] [PMC free article] [PubMed] [Google Scholar]
37.Tran-Betcke, A., U. Warnecke, C. Böcker, C. Zaborosch, and B. Friedrich. 1990. Cloning and nucleotide sequences of the genes for the subunits of NAD-reducing hydrogenase of Alcaligenes eutrophus H16. J. Bacteriol. 172:2920-2929. [DOI] [PMC free article] [PubMed] [Google Scholar]
38.Vignais, P. M., B. Billoud, and J. Meyer. 2001. Classification and phylogeny of hydrogenases. FEMS Microbiol. Rev. 25:455-501. [DOI] [PubMed] [Google Scholar]
39.Volbeda, A., M.-H. Charon, C. Piras, E. C. Hatchikian, M. Frey, and J. C. Fontecilla-Camps. 1995. Crystal structure of the nickel-iron hydrogenase from Desulfovibrio gigas. Nature 373:580-587. [DOI] [PubMed] [Google Scholar]
40.Wu, L.-F., and M. A. Mandrand. 1993. Microbial hydrogenases: primary structure, classification, signatures and phylogeny. FEMS Microbiol. Rev. 104:243-270. [DOI] [PubMed] [Google Scholar]
41.Zanoni, D. 1995. Aerobic and anaerobic electron transport chains in anoxygenic phototrophic bacteria, p. 949-971. In R. E. Blankenship, M. T. Madigan, and C. E. Bauer (ed.), Anoxygenic photosynthetic bacteria. Kluwer Academic Press, Dordrecht, The Netherlands.

[r1] 1.Appel, J., S. Phunpruch, K. Steinmüller, and R. Schulz. 2000. The bidirectional hydrogenase of Synechocystis sp. PCC 6803 works as an electron valve during photosynthesis. Arch. Microbiol. 173:333-338. [DOI] [PubMed] [Google Scholar]

[r2] 2.Ausubel, F. M., R. Brent, R. E. Kingston, D. D. Moore, J. G. Seidman, J. A. Smith, and K. Struhl (ed.). 1996. Current protocols in molecular biology. John Wiley & Sons, New York, N.Y.

[r3] 3.Bagramyan, K., N. Mnatsakanyan, A. Poladian, A. Vassilian, and A. Trchounian. 2002. The roles of hydrogenases 3 and 4, and the F0F1-ATPase, in H2 production by Escherichia coli at alkaline and acidic pH. FEBS Lett. 516:172-178. [DOI] [PubMed] [Google Scholar]

[r4] 4.Bogorov, L. V. 1974. About the properties of Thiocapsa roseopersicina BBS, isolated from estuaria of White Sea. Mikrobiologija (Belgrade) 43:326-332. [PubMed] [Google Scholar]

[r5] 5.Boison, G., O. Schmitz, B. Schmitz, and H. Bothe. 1998. Unusual gene arrangement of the bidirectional hydrogenase and functional analysis of its diaphorase subunit HoxU in respiration of the unicellular cyanobacterium Anacystis nidulans. Curr. Microbiol. 36:253-258. [DOI] [PubMed] [Google Scholar]

[r6] 6.Boison, G., H. Bothe, and O. Schmitz. 2000. Transcriptional analysis of hydrogenase genes in the cyanobacteria Anacystis nidulans and Anabaena variabilis monitored by RT-PCR. Curr. Microbiol. 40:315-321. [DOI] [PubMed] [Google Scholar]

[r7] 7.Boison, G., H. Bothe, A. Hansel, and P. Lindblad. 1999. Evidence against a common use of the diaphorase subunits by the bidirectional hydrogenase and by the respiratory complex I in cyanobacteria. FEMS Microbiol. Lett. 174:159-165. [Google Scholar]

[r8] 8.Buck, M., M. T. Gallegos, D. J. Studholme, Y. Guo, and J. D. Gralla. 2000. The bacterial enhancer-dependent σ⁵⁴ (σ^N) transcriptional factor. J. Bacteriol. 182:4129-4136. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r9] 9.Casalot, L., and M. Rousset. 2001. Maturation of the [NiFe] hydrogenases. Trends Microbiol. 9:228-237. [DOI] [PubMed] [Google Scholar]

[r10] 10.Colbeau, A., K. L. Kovács, J. Chabert, and P. M. Vignais. 1994. Cloning and sequencing of the structural (hupSLC) and accessory (hupDHI) genes for hydrogenase biosynthesis in Thiocapsa roseopersicina. Gene 140:25-31. [DOI] [PubMed] [Google Scholar]

[r11] 11.Dennis, J. J., and G. J. Zylstra. 1998. Plasposons: modular self-cloning minitransposon derivatives for rapid genetic analysis of gram-negative bacterial genomes. Appl. Environ. Microbiol. 64:2710-2715. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r12] 12.Dermoun, Z., G. De Luca, M. Asso, P. Bertrand, F. Guerlesquin, and B. Guigliarelli. 2002. The NADP-reducing hydrogenase from Desulfovibrio fructosovorans: functional interaction between the C-terminal region of HndA and the N-terminal region of HndD subunits. Biochim. Biophys. Acta 1556:217-225. [DOI] [PubMed] [Google Scholar]

[r13] 13.Fodor, B., G. Rákhely, Á. T. Kovács, and K. L. Kovács. 2001. Transposon mutagenesis in purple sulfur photosynthetic bacteria: identification of hypF, encoding a protein capable of processing [NiFe] hydrogenases in α, β, and γ subdivisions of the proteobacteria. Appl. Environ. Microbiol. 67:2476-2483. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r14] 14.Friedrich, B., and E. Schwartz. 1993. Molecular biology of hydrogen utilization in aerobic chemolithotrophs. Annu. Rev. Microbiol. 47:351-383. [DOI] [PubMed] [Google Scholar]

[r15] 15.Herrero, M., V. Lorenzo, and K. N. Timmis. 1990. Transposon vectors containing non-antibiotic resistance selection markers for cloning and stable chromosomal insertion of foreign genes in gram-negative bacteria. J. Bacteriol. 172:6557-6567. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r16] 16.Kaneko, T., S. Sato, H. Kotani, A. Tanaka, E. Asamizu, Y. Nakamura, N. Miyajima, M. Hirosawa, M. Sugiura, S. Sasamoto, T. Kimura, T. Hosouchi, A. Matsuno, A. Muraki, N. Nakazaki, K. Naruo, S. Okumura, S. Shimpo, C. Takeuchi, T. Wada, A. Watanabe, M. Yamada, M. Yasuda, and S. Tabata. 1996. Sequence analysis of the genome of the unicellular cyanobacterium Synechocystis sp. strain PCC6803. II. Sequence determination of the entire genome and assignment of potential protein-coding regions. DNA Res. 3:109-136. [DOI] [PubMed] [Google Scholar]

[r17] 17.Kaneko, T., Y. Nakamura, Y. C. P. Wolk, T. Kuritz, S. Sasamoto, A. Watanabe, M. Iriguchi, A. Ishikawa, K. Kawashima, T. Kimura, Y. Kishida, M. Kohara, M. Matsumoto, A. Matsuno, A. Muraki, N. Nakazaki, S. Shimpo, M. Sugimoto, M. Takazawa, M. Yamada, M. Yasuda, and S. Tabata. 2001. Complete genomic sequence of the filamentous nitrogen-fixing cyanobacterium Anabaena sp. strain PCC 7120. DNA Res. 8:205-213. [DOI] [PubMed] [Google Scholar]

[r18] 18.Kovács, Á. T., G. Rákhely, and K. L. Kovács. 2003. Genes involved in the biosynthesis of photosynthetic pigments in the purple sulfur photosynthetic bacterium Thiocapsa roseopersicina. Appl. Environ. Microbiol. 69:3093-3102. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r19] 19.Kovács, K. L., and C. Bagyinka. 1990. Structural properties, functional states and physiological roles of hydrogenase in photosynthetic bacteria. FEMS Microbiol. Rev. 87:407-412. [Google Scholar]

[r20] 20.Kovács, K. L., G. Tigyi, L. T. Thanh, S. Lakatos, Z. Kiss, and C. Bagyinka. 1991. Structural rearrangements in active and inactive forms of hydrogenase from Thiocapsa roseopersicina. J. Biol. Chem. 266:947-951. [PubMed] [Google Scholar]

[r20a] 20a.Kovács, K. L., B. Fodor, Á. T. Kovács, G. Csanádi, G. Maróti, J. Balogh, S. Arvani, and G. Rákhely. 2002. Hydrogenases, accessory genes and the regulation of [NiFe] hydrogenase biosynthesis in Thiocapsa roseopersicina. Int. J. Hydrogen Energy 27:1463-1469. [Google Scholar]

[r21] 21.Lenz, O., and B. Friedrich. 1998. A novel multicomponent regulatory system mediates H₂ sensing in Alcaligenes eutrophus. Proc. Natl. Acad. Sci. 95:12474-12479. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r22] 22.Lowry, O. H., N. J. Rosenbourgh, A. L. Farr, and R. J. Randall. 1951. Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193:265-275. [PubMed] [Google Scholar]

[r23] 23.Maier, T., and A. Böck. 1996. Nickel incorporation into hydrogenases, p. 173-192. In R. Hausinger, G. L. Eichlorn, and L. G. Marzilli (ed.), Advances in inorganic biochemistry. VHC Publishers Inc., New York, N.Y.

[r24] 24.Maróti, G., B. D. Fodor, G. Rákhely, Á. T. Kovács, S. Arvani, and K. L. Kovács. 2003. Accessory proteins functioning selectively and pleiotropically in the biosynthesis of [NiFe] hydrogenases in Thiocapsa roseopersicina. Eur. J. Biochem. 270:2218-2227. [DOI] [PubMed] [Google Scholar]

[r25] 25.Nelson, K. E., R. A. Clayton, S. R. Gill, M. L. Gwinn, R. J. Dodson, D. H. Haft, E. K. Hickey, J. D. Peterson, W. C. Nelson, K. A. Ketchum, L. McDonald, T. R. Utterback, J. A. Malek, K. D. Linher, M. M. Garrett, A. M. Stewart, M. D. Cotton, M. S. Pratt, C. A. Phillips, D. Richardson, J. Heidelberg, G. G. Sutton, R. D. Fleischmann, O. White, S. L. Salzberg, H. O. Smith, J. C. Venter, and C. M. Fraser. 1999. Evidence for lateral gene transfer between Archaea and bacteria from genome sequence of Thermotoga maritima. Nature 399:323-329. [DOI] [PubMed] [Google Scholar]

[r26] 26.Nicholls, D. G., and S. J. Ferguson. 2001. Bioenergetics 3. Academic Press, New York, N.Y.

[r27] 27.Parales, R. E., and C. S. Harwood. 1993. Construction and use of a new broad-host-range lacZ transcriptional fusion vector, pHRP309, for gram− bacteria. Gene 133:23-30. [DOI] [PubMed] [Google Scholar]

[r28] 28.Rákhely, G., A. Colbeau, J. Garin, P. M. Vignais, and K. L. Kovács. 1998. Unusual organization of the genes coding for HydSL, the stable (NiFe) hydrogenase in the photosynthetic bacterium Thiocapsa roseopersicina BBS. J. Bacteriol. 180:1460-1465. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r29] 29.Robson. R. 2001. Biodiversity of hydrogenases, p. 9-32. In R. Cammack, M. Frey, and R. Robson (ed.), Hydrogen as a fuel. Taylor & Francis, London, United Kingdom.

[r30] 30.Sambrook, J., T. Maniatis, and E. F. Fritsch. 1989. Molecular cloning: a laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.

[r31] 31.Schafer, A., A. Tauch, W. Jager, J. Kalinowski, G. Thierbach, and A. Puhler. 1994. Small mobilizable multi-purpose cloning vectors derived from the Escherichia coli plasmids pK18 and pK19: selection of defined deletions in the chromosome of Corynebacterium glutamicum. Gene 145:69-73. [DOI] [PubMed] [Google Scholar]

[r32] 32.Schmitz, O., G. Boison, R. Hilscher, B. Hundeshagen, W. Zimmer, F. Lottspeich, and H. Bothe. 1995. Molecular biological analysis of a bidirectional hydrogenase from cyanobacteria. Eur. J. Biochem. 233:266-276. [DOI] [PubMed] [Google Scholar]

[r33] 33.Schmitz, O., G. Boison, and H. Bothe. 2001. Quantitative analysis of expression of two circadian clock-controlled gene clusters coding for the bidirectional hydrogenase in the cyanobacterium Synechococcus sp. PCC7942. Mol. Microbiol. 41:1409-1417. [DOI] [PubMed] [Google Scholar]

[r34] 34.Schmitz, O., G. Boison, H. Salzmann, H. Bothe, K. Schutz, S. H. Wang, and T. Happe. 2002. HoxE—a subunit specific for the pentameric bidirectional hydrogenase complex (HoxEFUYH) of cyanobacteria. Biochim. Biophys. Acta 1554:66-74. [DOI] [PubMed] [Google Scholar]

[r35] 35.Shevchenko, A., M. Wilm, O. Vorm, and M. Mann. 1996. Mass spectrometric sequencing of proteins from silver-stained polyacrylamide gels. Anal. Chem. 68:850-858. [DOI] [PubMed] [Google Scholar]

[r36] 36.Tamagnini, P., R. Axelsson, P. Lindberg, F. Oxelfelt, R. Wunschiers, and P. Lindblad. 2002. Hydrogenases and hydrogen metabolism of cyanobacteria. Microbiol. Mol. Biol. Rev. 66:1-20. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r37] 37.Tran-Betcke, A., U. Warnecke, C. Böcker, C. Zaborosch, and B. Friedrich. 1990. Cloning and nucleotide sequences of the genes for the subunits of NAD-reducing hydrogenase of Alcaligenes eutrophus H16. J. Bacteriol. 172:2920-2929. [DOI] [PMC free article] [PubMed] [Google Scholar]

[r38] 38.Vignais, P. M., B. Billoud, and J. Meyer. 2001. Classification and phylogeny of hydrogenases. FEMS Microbiol. Rev. 25:455-501. [DOI] [PubMed] [Google Scholar]

[r39] 39.Volbeda, A., M.-H. Charon, C. Piras, E. C. Hatchikian, M. Frey, and J. C. Fontecilla-Camps. 1995. Crystal structure of the nickel-iron hydrogenase from Desulfovibrio gigas. Nature 373:580-587. [DOI] [PubMed] [Google Scholar]

[r40] 40.Wu, L.-F., and M. A. Mandrand. 1993. Microbial hydrogenases: primary structure, classification, signatures and phylogeny. FEMS Microbiol. Rev. 104:243-270. [DOI] [PubMed] [Google Scholar]

[r41] 41.Zanoni, D. 1995. Aerobic and anaerobic electron transport chains in anoxygenic phototrophic bacteria, p. 949-971. In R. E. Blankenship, M. T. Madigan, and C. E. Bauer (ed.), Anoxygenic photosynthetic bacteria. Kluwer Academic Press, Dordrecht, The Netherlands.

PERMALINK

Cyanobacterial-Type, Heteropentameric, NAD+-Reducing NiFe Hydrogenase in the Purple Sulfur Photosynthetic Bacterium Thiocapsa roseopersicina

Gábor Rákhely

Ákos T Kovács

Gergely Maróti

Barna D Fodor

Gyula Csanádi

Dóra Latinovics

Kornél L Kovács