Báo cáo khoa học: Electron-transfer subunits of the NiFe hydrogenases in Thiocapsa roseopersicina BBS pptx

Electron-transfer subunits of the NiFe hydrogenases in Thiocapsa roseopersicina BBS Lı ´via S. Pala ´ gyi-Me ´ sza ´ ros 1 , Judit Maro ´ ti 2 ,Do ´ ra Latinovics 1 ,Tı ´mea Balogh 1 ,E ´ va Klement 3 , Katalin F. Medzihradszky 3 ,Ga ´ bor Ra ´ khely 1,2 and Korne ´ l L. Kova ´ cs 1,2 1 Department of Biotechnology, University of Szeged, Hungary 2 Institute of Biophysics, Biological Research Centre, Hungarian Academy of Sciences, Szeged, Hungary 3 Proteomics Research Group, Biological Research Centre, Hungarian Academy of Sciences, Szeged, Hungary Hydrogenases are metalloenzymes that catalyse the reversible oxidation of molecular hydrogen according to the reaction: H 2 M 2H + +2e ) . They can catalyse the reaction in both directions in vitro, but usually either evolve or oxidize (take up) H 2 in vivo. The hydrogenases can be classified according to the metal content of their active centre: NiFe, FeFe or Fe hydrogenases [1]. The core of an NiFe hydrogenase consists of a small electron-transfer subunit and a large catalytic subunit. Additional proteins are required for post-translational maturation of the hydrogenase polypeptides and for connection of the core dimer to other bioenergetic ⁄ redox processes of the cells. These accessory hydrogenase-related proteins typically partici- pate in metallocentre assembly and the transcriptional regulation of the hydrogenases, and some seem to have an electron-transfer function [1,2]. The accessory genes are often located in the close vicinity of hydrogenase structural genes, but may also be found scattered in the genome. Numerous microorganisms contain more Keywords electron transfer; haem, cytochrome b; iron–sulfur protein; NiFe hydrogenase; Thiocapsa roseopersicina Correspondence K. L. Kova ´ cs, Department of Biotechnology, University of Szeged, H-6726 Szeged, Ko ¨ ze ´ pfasor 52, Hungary Fax: +36 62 544352 Tel: +36 62 544351 E-mail: kornel@brc.hu (Received 31 August 2008, revised 6 October 2008, accepted 29 October 2008) doi:10.1111/j.1742-4658.2008.06770.x Thiocapsa roseopersicina BBS contains at least three different active NiFe hydrogenases: two membrane-bound enzymes and one apparently localized in the cytoplasm. In addition to the small and large structural subunits, additional proteins are usually associated with the NiFe hydrogenases, con- necting their activity to other redox processes in the cells. The operon of the membrane-associated hydrogenase, HynSL, has an unusual gene arrangement: between the genes coding for the large and small subunits, there are two open reading frames, namely isp1 and isp2. Isp1 is a b-type haem-containing transmembrane protein, whereas Isp2 displays marked sequence similarity to the heterodisulfide reductases. The other membrane- bound (Hup) NiFe hydrogenase contains the hupC gene, which codes for a cytochrome b-type protein that probably plays a role in electron transport. The operon of the NAD + -reducing Hox hydrogenase contains a hoxE gene. In addition to the hydrogenase and diaphorase parts of the complex, the fifth HoxE subunit may serve as a third redox gate of this enzyme. The physiological functions of these putative electron-mediating subunits were studied by disruption of their genes. The deletion of some accessory pro- teins dramatically reduced the in vivo activities of the hydrogenases, although they were fully active in vitro. The absence of HupC resulted in a decrease in HupSL activity in the membrane, but removal of the Isp1 and Isp2 proteins did not have any significant effect on the location of HynSL activity. Through the use of a tagged HoxE protein, the whole Hox hydrogenase pentamer could be purified as an intact complex. Abbreviation tat, twin arginine transport. 164 FEBS Journal 276 (2009) 164–174 ª 2008 The Authors Journal compilation ª 2008 FEBS than one hydrogenase. Each enzyme has a specific phys- iological function, e.g. NAD + reduction, electron removal, H 2 recycling for energy conservation, etc. [3]. The electrons derived from H 2 oxidation are used for the reduction of the central quinone pool or terminal electron acceptors, such as fumarate, NO 3 ) or SO 4 2) .It is noteworthy that, in spite of their specific expression and physiological role, one enzyme can take over the function of another to some extent [4]. Thiocapsa roseopersicina BBS belongs to the family of purple sulfur photosynthetic bacteria, the Chromati- aceae [5]. During anoxygenic photosynthesis, this bac- terium requires reduced sulfur compounds (e.g. S 2) ,S 0 or S 2 O 3 2) ) as electron sources for CO 2 fixation. T. roseopersicina produces at least three NiFe hydro- genases (Hyn, Hup and Hox) and contains the genes of the so-called regulatory hydrogenase (HupUV) [6]. However, their physiological roles are still unclear. Both the HynS and HupS subunits have a ‘tat’-type (‘twin arginine transport’) signal sequence; they are therefore transported through the membrane by the ‘tat’ system [7] and are anchored to the membrane on the periplasmic side. The Hox enzyme has no signal for transport across the membrane. Hyn hydrogenase (formerly Hyd [8]) is a membrane-bound bidirectional enzyme which has remarkable stability under extreme conditions; it is extracted from the photosynthetic membrane as the catalytically active HynSL dimer [9]. The gene arrangement of the hyn operon is unusual: the genes of the small and large subunits are separated by a 2-kbp intergenic region. In this section, two open reading frames, isp1 and isp2, have been recognized [8]. The putative Isp1 and Isp2 gene products exhibit remarkable similarity to the DsrK and DsrM subunits, respectively, of the dissimilatory sulfite reductase com- plex [10]. Isp1 harbours few transmembrane domains, and a putative b-type haem-binding site has been pre- dicted by in silico analysis. In contrast, the putative Isp2 is a cytoplasmic enzyme resembling the hetero- disulfide reductases [8]. Similar gene structures can be found in only a few bacteria, e.g. in Chromatium vinosum [10] (Accession No. U84760), Aquifex aeolicus [11], Aquifex pyrophilus [12] and an Archaeon, Acidi- anus ambivalens [13], but their physiological role has not been clarified so far. The other membrane-bound hydrogenase of T. roseopersicina, HupSL, is encoded in the hupSLCD- HIR operon [14]. It belongs to the group of uptake NiFe hydrogenases which recycle H 2 produced by the nitrogenase complex [15]. As a consequence of the periplasmic location of the NiFe hydrogenases [16], H 2 oxidation leads to the formation of a proton gradient which is used for ATP synthesis [9]. Next to the hupSL genes encoding for the small and large hydrogenase subunits, the operon contains the hupC gene. In Wolli- nella succinogenes, strong evidence has been provided that HupC, a b-type cytochrome [1], can transfer electrons from the NiFe hydrogenase to the quinones [17]. Hence, HupC may link the electron transfer from Hup hydrogenases to the quinone pool. The third (Hox) hydrogenase has been partially purified from the soluble fraction of the cells [18]. The genomic structure of the hox operon suggests a hetero- pentameric enzyme (HoxEFUYH). The HoxFU subunits are usually the NAD + -reducing part of the complex, and the HoxYH subunits are responsible for hydrogenase activity [19]. Recently, a similar enzyme has been purified and partially characterized from a closely related strain, Allochromatim vinosum [20]. Hox hydrogenases are composed of at least four subunits; the HoxYH and HoxFU dimers form the hydrogenase and diaphorase catalytic cores, respectively [19]. In sev- eral cases, additional subunits have also been identi- fied. In the Hox enzyme of Ralstonia eutropha (which was purified as a heterotetrameric enzyme for many years), a new subunit was discovered, and the compo- sition HoxFUYHI 2 was suggested [21]. In cyanobacte- ria and the phototrophic bacteria T. roseopersicina and A. vinosum, the heterotetrameric Hox enzyme is sup- plemented by a HoxE subunit, which is unrelated to the HoxI protein [18,20,22]. In T. roseopersicina, it has been shown previously that in-frame deletion of the hoxE gene impairs Hox activity in vivo, although the remaining part of the complex (HoxFUYH) still shows unaltered H 2 -dependent NAD + -reducing activity in vitro [18]. However, the roles of HoxE and the Hox complex are still not fully understood. In this article, we show that the various hydrogenas- es use distinct electron-transfer subunits and routes. Deletion of the HupC, Isp1,2 and HoxE proteins clearly reveals their physiological relationships to their respective hydrogenases. Affinity purification of the HoxE-tagged protein under mild conditions confirms the heteropentameric structure of this complex. Results Isp1 and Isp2 are expressed proteins The in silico analysis of the intergenic region of the hynS and hynL genes indicated two open reading frames. It has been established that the hynS-isp1-isp2- hynL region is cotranscribed [23]. In order to confirm that isp1 and isp2 are really coding regions, the hynS- isp1-isp2-hynL* genes were cloned behind a T7 promoter (see Experimental procedures). The genes of L. S. Pala ´ gyi-Me ´ sza ´ ros et al. Electron-transfer subunits of NiFe hydrogenases FEBS Journal 276 (2009) 164–174 ª 2008 The Authors Journal compilation ª 2008 FEBS 165 the construct were expressed in the Escherichia coli BL21(DE3) host, and the bands corresponding to the calculated molecular masses of Isp1 (24.6 kDa) and Isp2 (48.4 kDa) could be clearly identified (data not shown). The small and large subunits were also detected. This means that all the translational signals necessary for the expression of the HynSL and Isp subunits are functionally present in the construct, and are recognized by the translational apparatus of E. coli. The coexpression of the Hyn and Isp subunits suggests that they probably form a functional complex. Isp1 and Isp2 are required for the in vivo function of Hyn hydrogenase The solubilized and purified Hyn hydrogenase con- tained only the HynSL subunits [23]. The role of the Isp proteins in T. roseopersicina is unknown, but com- putational analysis has shown that Isp1 is a b-type haem-containing transmembrane electron carrier, whereas Isp2 seems to be a redox Fe–S-containing pro- tein. If these subunits are involved in the electron flow from ⁄ to the hydrogenase, their removal would abolish the hydrogenase activity in vivo, where the endogenous electron donors ⁄ acceptors must be used. Therefore, a double isp1-isp2 in-frame mutant was constructed in the T. roseopersicina GB2131 (DhoxH, DhupSL) strain (ISP12M, see Experimental proce- dures). The hydrogenase activities were measured both in vivo (without the addition of an artificial electron carrier) and in vitro (in the presence of redox viologen dyes). The data in Table 1 unequivocally prove that the in vivo H 2 -producing activity of the isp1,2 mutant strain is completely lost and the in vivo H 2 uptake activity is dramatically decreased relative to the control GB2131 (DhoxH, DhupSL) strain containing all the functional gene products of the Hyn operon. A single Isp1 in-frame deletion mutant was also constructed (ISP1M). Mutation of the Isp1 protein brings about the same phenotype as the deletion of both Isp1 and Isp2 (Table 1). Some remaining in vivo H 2 uptake activity of the Hyn hydrogenase can be detected in both mutants, which suggests an alternative, less effec- tive electron-transfer pathway. In the in vitro measurements, in which benzyl-violo- gen was used as an artificial electron acceptor, the H 2 uptake activity was not influenced by the lack of Isp1 or Isp1,2 proteins (Table 1). On the one hand, these and the in silico results confirm that the Isp proteins play an essential role in the H 2 reduction and oxida- tion ability of Hyn hydrogenase in its natural environ- ment, but the lack of these subunits has no effect on the hydrogenase activity in the artificial assay. On the other hand, this also means that the Isp1,2 proteins do not affect the post-translational maturation and expression level of the Hyn enzyme. A trivial rationali- zation of these observations is that the lack of Isp1 or Isp1,2 proteins results in blockade of the electron flow from ⁄ to Hyn hydrogenase under physiological condi- tions. As the computational analysis implies that Isp1 is an integrated membrane protein, it is plausible to assume that the HynSL dimer is anchored to the mem- brane through the Isp1 protein. Accordingly, we inves- tigated the localization of Hyn hydrogenase in the Isp mutant strains. Unexpectedly, the in vitro H 2 uptake measurements on the various cellular fractions indi- cated that a similar proportion of Hyn hydrogenase remained in the membrane fraction in the presence and absence of the Isp proteins (Table 2). This is surprising, as our protein purification experiments demonstrated that the HynSL subunits are only loosely associated with the membrane and can be easily Table 1. Activities of Hyn hydrogenase in vivo and in vitro in the presence and absence of the Isp1 and Isp2 proteins. The results are given as percentages of the level for GB2131. The cultures were grown on Pfennig’s medium with 4 gÆL )1 of Na 2 S 2 O 3 . The val- ues are normalized to bacteriochlorophyll content. The GB112131 strain (DhupSL, DhoxH, DhynS-isp1-isp2-hynL) and the M539 strain (hypF mutant) containing no active NiFe hydrogenase served as negative controls. Strain Relative H 2 production in vivo Relative H 2 uptake in vivo Relative H 2 uptake in vitro GB2131 (DhupSL, DhoxH) 100 ± 10.2 100 ± 13.7 100 ± 10.0 ISP1M (DhupSL, DhoxH, Disp1) 0.00 29.5 ± 9.8 100.4 ± 9.2 ISP12M t (DhupSL, DhoxH, Disp12) 0.0 35.6 ± 5.9 116.2 ± 8.3 Table 2. Location of Hyn hydrogenase with and without the Isp1,2 proteins (see description in Table 1). Strain Relative uptake activity in vitro Membrane fraction Soluble fraction GB2131 (DhupSL, DhoxH) 100 ± 23.2 100 ± 7.7 ISP1M (DhupSL, DhoxH, Disp1) 106.2 ± 33.4 102.7 ± 4.0 ISP12M (DhupSL, DhoxH, Disp12) 112.4 ± 0.3 113.9 ± 6.8 Electron-transfer subunits of NiFe hydrogenases L. S. Pala ´ gyi-Me ´ sza ´ ros et al. 166 FEBS Journal 276 (2009) 164–174 ª 2008 The Authors Journal compilation ª 2008 FEBS washed off [23]. The strength of the HynSL–membrane interaction apparently does not depend on the presence or absence of the Isp1 protein. The expression of the Hup enzyme depends on the thiosulfate content of the medium With a view to examining the function of the HupC protein in T. roseopersicina,aDhynSL, DhoxH (GB1131) strain was created (see Experimental proce- dures). This strain is suitable for the measurement of Hup hydrogenase activity alone, without the contribu- tions of Hyn and Hox hydrogenases. However, under standard growth conditions, i.e. in the presence of 4gÆL )1 Na 2 S 2 O 3 , only very low HupSL hydrogenase activity was detected in the DhynSL, DhoxH (GB1131) strain. It was postulated that this concentration of thiosulfate resulted in a redox potential in the cells, which downregulated the activity of HupSL hydro- genase (as an uptake, electron-donating enzyme). To test this hypothesis, the expression level and in vitro activity of the Hup hydrogenase were measured in cells grown in the presence of various amounts of thiosul- fate. The data in Table 3 clearly illustrate that the lower the thiosulfate concentration in the medium, the higher the Hup hydrogenase activity both in vivo and in vitro. The effects of the thiosulfate content on the expression level of the hupSL genes were additionally monitored by quantitative RT-PCR. The data in Table 4 reveal that a decrease in the thiosulfate con- tent of the medium from 4 to 2 gÆL )1 resulted in a dra- matic (> 16-fold) increase in the hupSL mRNA level. These data suggest that, when Hup is the only active hydrogenase in the cell, its activity strongly depends on the thiosulfate content of the medium, and changes in the activity primarily correlate with the expression level of the enzyme. Hence, as a practical consequence, the subsequent experiments on Hup activity were per- formed with samples grown in the presence of 2 gÆL )1 thiosulfate. HupC is an electron-transfer subunit of Hup hydrogenase To establish the function of HupC, its gene was deleted in-frame in the DhynSL, DhoxH (GB1131) strain, and the HupSL activities were compared both in vivo and in vitro. The in vivo H 2 uptake activity of Hup hydrogenase was substantially decreased in the DhupC (HCMG4) strain. At the same time, the in vitro activity was twice as high as that of the strain harbouring HupC (Table 5). A comparison of the hupSL mRNA levels of the cells containing or lacking the hupC gene per- ceptibly revealed that a loss of the hupC gene had a positive effect on the transcription level of the hupSL genes (Table 4). To check that the effect was really linked to the loss of HupC, a complementation experi- ment was performed by introducing an expression Table 3. In vivo and in vitro H 2 uptake activities of the GB1131 (DhynSL, DhoxH) strain grown photoautotrophically (Pfennig’s) at various Na 2 S 2 O 3 concentrations. The results are given as percent- ages of that for the sample grown with 1 gÆL )1 of Na 2 S 2 O 3 . Concentration of Na 2 S 2 O 3 (gÆL )1 ) Relative H 2 uptake activity In vivo In vitro 4 0.0 0.0 2 45.0 ± 2.6 83.6 ± 34.2 1 100.0 ± 5.6 100.0 ± 11.1 Table 4. Relative mRNA levels of the hup operon in the presence (GB1131) and absence (HCMG4) of the hupC gene at various Na 2 S 2 O 3 concentrations. The cultures were grown on Pfennig’s medium with 2 or 4 gÆL )1 of Na 2 S 2 O 3 . The mRNA levels were determined by quantitative RT-PCR and the results are given as percentages of the level for GB1131. The values are normalized to the total RNA content. Strain 4gÆL )1 Na 2 S 2 O 3 2gÆL )1 Na 2 S 2 O 3 GB1131 (DhynS-isp1-isp2-hynL, DhoxH) 100.0 ± 0.0 1650.0 ± 44.5 HCMG4 (DhupC, DhynS-isp1-isp2-hynL, DhoxH) 300.0 ± 20.0 2700.0 ± 102.8 Table 5. Activities of Hup hydrogenase in vivo and in vitro in the presence (GB1131, pMHE6C HCMG4) and absence (HCMG4) of the HupC protein. The cultures were grown on Pfennig’s medium with 2gÆL )1 of Na 2 S 2 O 3 . The hydrogenase activity values are normalized to the bacteriochlorophyll content. The results are given as a percent- age of the level for GB1131. The GB112131 strain (DhupSL, DhoxH, DhynS-isp1-isp2-hynL) and the M539 strain (hypF mutant) containing no active NiFe hydrogenase served as negative controls. Strain Relative H 2 uptake activity In vivo In vitro GB1131 (DhynS-isp1-isp2-hynL, DhoxH) 100.0 ± 2.6 100.0 ± 14.5 HCMG4 (DhupC, DhynS-isp1-isp2-hynL, DhoxH) 40.4 ± 5.5 198.9 ± 5.5 pMHE6C HCMG4 (DhupC, DhynS-isp1-isp2-hynL, DhoxH, pMHE6C) 68.3 ± 10.3 231.2 ± 33.5 L. S. Pala ´ gyi-Me ´ sza ´ ros et al. Electron-transfer subunits of NiFe hydrogenases FEBS Journal 276 (2009) 164–174 ª 2008 The Authors Journal compilation ª 2008 FEBS 167 cassette containing the hupC gene driven by the crt promoter (see Experimental procedures). Table 5 shows that the plasmid-borne HupC (pMHE6C ⁄ HCMG4 in Table 5) partially ( 50%) restored the Hup hydrogenase activity in vivo. It is plausible to assume that HupC serves as a membrane anchor for Hup hydrogenase [24]. In con- trast with the findings on Hyn hydrogenase, the lack of HupC substantially reduced the Hup hydrogenase activity in the membrane fraction, i.e. 98% of the activity was lost in the hupC deletion mutant. The hydrogenase activity in the soluble fraction was also decreased; therefore, it is unlikely that HupSL was released from the membrane and accumulated in the cytoplasm. The lower total activity in the HupC-minus cell fractions might be explained by the lower stability of the HupSL enzyme in the absence of HupC in the disrupted and fractionated cells relative to the wild- type (Table 6). It is noteworthy that the HupSL activ- ity was significantly higher in the soluble than in the membrane fraction in the pMHE6C ⁄ HCMG4 (HupC complementing) strain (Table 6). The plasmid-borne HupC could possibly restore the stability of HupSL, although the majority of the activity remained in the soluble fraction. These data suggest that HupC has no role in the maturation process of HupSL hydrogenase, but influ- ences the in vivo activity and the expression level of the HupSL enzyme. Taken together with the findings of computational analysis, the HupC protein serves an electron-transport role in T. roseopersicina and proba- bly forms a functional complex with the small and large hydrogenase subunits in vivo. Purification of Hox hydrogenase In T. roseopersicina, the cytoplasmic Hox hydrogenase is coded by the hoxEFUYH operon. The enzyme contains hydrogenase (HoxYH) and diaphorase (Ho- xEFU) subunits [18]. The diaphorase subunits of the Hox-type hydrogenases exhibit significant sequence similarities to three subunits (NuoEFG) of NADH:ubiquinone oxidoreductase [18,25]. The in-frame deletion of the hoxE gene led to the complete loss of Hox activity in vivo, whereas the enzyme was fully active in vitro [18]. This suggests that HoxE may function in vivo as an electron-transfer protein. Thus, HoxE would offer a third channel for the electrons in addition to the hydrogenase and diaphorase catalytic centres. To test whether the HoxE protein forms a functional complex with the HoxFUYH subunits, its FLAG-tagged form was expressed from a pMHE6 expression vector [26] under the control of the T. roseopersicina crt promoter (pMHE6HoxE Table 7). HoxE was purified by affinity chromatography via the FLAG-tag under very mild conditions in order to pre- serve the protein–protein interactions (see Experimen- tal procedures). The proteins eluted from the affinity column were separated on SDS-polyacrylamide gel and analysed by MALDI-TOF-MS. Each subunit of the HoxEFUYH enzyme complex was easily identified, indicating that HoxE is physically associated with the other (HoxFUYH) subunits (Fig. 1). Discussion Hydrogenases are widespread in the microbial world. The actively expressed hydrogenases must have a dedi- cated physiological role within the cells. For the in vivo function, the catalytic dimers of NiFe hydrogenases must be connected to other oxidoreductases directly or via electron-transfer subunits. In this study, attempts were made to identify the redox partners and the elec- tron-channelling subunits of all three hydrogenases in the cells. In T. roseopersicina, there are at least three NiFe hydrogenases (HynSL, HupSL and HoxYH) with distinct properties and different functions. The HypF accessory protein is required for the maturation of every NiFe hydrogenase, and disruption of the hypF gene therefore results in the hydrogenase-minus pheno- type [27]. However, the hydrogenase-less cells showed virtually identical growth properties as the wild-type under standard growth conditions. Special growth con- ditions, i.e. photoautotrophic in the presence of H 2 and only 0.005% Na 2 S, were identified, in which the presence of each hydrogenase was important, including the Hup enzyme being essential for H 2 -dependent growth (data not shown). This indicates that the Hup enzyme has a direct connection to the central redox, i.e. quinone, pool. Nonetheless, the real redox partners Table 6. Location of HupSL hydrogenase with (GB1131, pMHE6C HCMG4) and without (HCMG4) the HupC subunit (see description in Table 5). Strain In vitro relative uptake activity Membrane fraction Soluble fraction GB1131 (DhynS-isp1-isp2-hynL, DhoxH) 100.0 ± 12.9 36.0 ± 8.5 HCMG4 (DhupC, DhynS-isp1-isp2-hynL, DhoxH) 1.8 ± 0.24 11.3 ± 1.8 pMHE6C HCMG4 (DhupC, DhynS-isp1-isp2-hynL, DhoxH, pMHE6C) 47.3 ± 0.9 115.0 ± 7.1 Electron-transfer subunits of NiFe hydrogenases L. S. Pala ´ gyi-Me ´ sza ´ ros et al. 168 FEBS Journal 276 (2009) 164–174 ª 2008 The Authors Journal compilation ª 2008 FEBS and ⁄ or electron channels of each hydrogenase remain poorly understood. The genes coding for the HynSL enzyme are sepa- rated by two open reading frames, which have been shown to code for real proteins, Isp1 and Isp2. Both proteins have been demonstrated to be important for the function of the HynSL enzyme in vivo, but neither for its in vitro activity or expression. Therefore, they probably play an electron-transfer role from ⁄ to the Hyn enzyme. The heterodisulfide reductase homologue Isp2 is probably an oxidoreductase; its redox substrate still remains to be identified. We conclude that the Hyn enzyme is indirectly linked to the central redox ⁄ bioenergetic processes via the Isp1,2 proteins and an unknown redox substrate. A direct coupling of the HupC protein to the uptake HupSL hydrogenase was demonstrated in this study. Deletion of the hupC gene resulted in reduced and enhanced activities of HupSL in vivo and in vitro, respectively. As HupC is supposed to react with qui- nones directly [17], the reduced in vivo activity stems from the obstruction of the electron flow from the hydrogenase. Consequently, HupC is suggested to be the third subunit of the Hup complex, catalysing the H 2 -dependent reduction of quinones. This is in line with the observation that HupSL hydrogenase is essential for H 2 -dependent growth under the above- mentioned growth conditions. The expression level of the HupSL enzyme was upregulated both by disrupting the HupC subunit and by decreasing the thiosulfate content of the medium. It is assumed that both processes lead to a more oxidized quinone pool, as the disrupted HupC cannot transfer the electrons from HupSL, and thiosulfate serves as reducing power for the photosynthetic carbon fixation via the central quinone pool [28]. The redox status of the quinone pool may influence HupSL expression: the increased electron requirement is reflected in a higher expression level of the electron-donating Hup hydro- genase. Interestingly, removal of the transmembrane elec- tron-transfer subunits of the Hyn and Hup enzymes gave rise to distinct effects on the locations of their corresponding hydrogenases. The lack of Isp1 did not change the membrane association of the Hyn enzyme, whereas the elimination of HupC led to detachment of Table 7. Strains and plasmids. Indicated strains and plasmids are from Stratagene, La Jolla, CA, USA. Strain or plasmid Relevant genotype or phenotype Reference or source Thiocapsa roseopersicina BBS Wild-type [5] GB2131 hupSLD::Gm, hoxHD::Er [18] GB1121 hynSLD_::Smr, hupSLD_::Gmr [18] GB1131 hynSLD_::Smr, hoxHD::Er This work GB112131 hynSLD::Smr, hupSLD::Gm, hoxHD::Er [18] M539 hypFD::Km [27] ISP1M hupSLD::Gm, hoxHD::Er, isp1D This work ISP12M hupSLD ::Gm, hoxHD::Er, D isp1, D isp2 This work HCMG4 hynSLD_::Smr, hoxHD::Er, DhupC This work pMHE6C ⁄ HCMG4 hynSLD_::Smr, hoxHD::Er, DhupC, pMHE6C This work pMHE6HoxE ⁄ GB2131 hupSLD::Gm, hoxHD::Er, pMHE6HoxE This work Escherichia coli XL1-Blue MRF ¢ D (mcrA)183, D (mcrCB-hsdSMR-mrr)173, endA1, supE44, thi-1, recA1, gyrA96, relA1 lac [F¢ proAB lacIqZDM15 Tn10 (Tet r )] c Stratagene BL21 (DE3) F ) ompT gal dcm lon hsdS B (r B ) m B ) ) k(DE3) [lacI lacUV5-T7 gene 1 ind1 sam7 nin5] Stratagene Plasmids pBluescript SK+ Amp r , cloning vector, ColE1 ori Stratagene pK18mobSacB Km r sacB RP4 oriT ColE1 ori [36] pUC19 Amp r , cloning vector, ColE1 ori [37] pMHE6crtKm Km r , mob + , expression vector containing the promoter region of crtD gene [26] pMHE6C Km r , mob + , hupC gene cloned after the crtD gene This work pMHE6HoxE Km r , mob + , hoxE gene cloned after the crtD gene This work pISP1M Km r , in-frame up- and downstream homologous regions of isp1 in pK18mobsacB This work pISP12M Km r , in-frame up- and downstream homologous regions of isp1-isp2 in pK18mobsacB This work pHCD2 Km r , in-frame up- and downstream homologous regions of hupC in pK18mobsacB This work pTSH2 ⁄ 8 Cosmid clone containing hyp operon [8] L. S. Pala ´ gyi-Me ´ sza ´ ros et al. Electron-transfer subunits of NiFe hydrogenases FEBS Journal 276 (2009) 164–174 ª 2008 The Authors Journal compilation ª 2008 FEBS 169 the HupSL enzyme from the membrane. The absence of HupC also resulted in a destabilization of HupSL. A similar phenomenon has been described for HupC in Rhodobacter capsulatus [24]. In T. roseopersicina, the plasmid-borne HupC restored the stability and mem- brane association of HupSL, but a significant amount of enzyme remained in the soluble fraction. Controver- sial data have been published in the literature with regard to the membrane anchoring role of HupC. Deletion of HoxW, the HupC homologous protein in R. eutropha, resulted in detachment of the hydrogenase from the membrane [29]. In contrast in Pseudomo- nas hydrogenovora, the location of the HupSL dimer in the hupC mutant strain did not change [30]. It has been shown previously that HoxE is required for the in vivo function of the Hox hydrogenase [18]. Here, we have demonstrated that HoxE fulfills this role in association with the other subunits of the Hox hydrogenase. Purification of the affinity-tagged HoxE under mild conditions resulted in copurification of the four other (HoxFUYH) subunits. We observed that the hydrogenase dimer dissociated from the HoxEFU trimer relatively easily (data not shown). A similar finding has been published recently for the A. vinosum Hox hydrogenase [20]. This means that the enzyme com- plex has three gates for electron flow: one for H 2 oxid- ation ⁄ proton reduction, one for the NAD + ⁄ NADH redox reaction and one functioning as an electron channel via the HoxE subunit. This makes the potential physiological function of this hydrogenase more complex, as the Hox hydrogenase has a potential to be associated with various metabolic pathways involving redox changes. In cyanobacteria, the Hox hydrogenase was initially suggested to have a relationship to the respiratory complex [25], but evidence challenging this idea was later published [31]. A valve role of the Hox enzyme was suggested for the low-potential electrons generated during photosynthesis [32]. The three gates for electron flow are in line with the valve hypothesis. However, depending on the sulfur source, the Hox enzyme is able to produce H 2 either under illumination or in the dark [33], and thus its physiological function cannot be restricted to photosynthetic electron flow, but the respiratory and fermentative processes should also be considered. Experimental procedures Bacterial strains and plasmids The strains and plasmids are listed in Table 7. The T. roseo- persicina strains were grown photoautotrophically in Pfennig’s medium under anaerobic conditions in liquid cultures with continuous illumination at 27–30 °C for 4–5 days [27]. The acetate-supplemented (2 gÆL )1 ) plates were solidified with 7 gÆL )1 of Phytagel (Sigma, St Louis, MO, USA) [34]. The plates were incubated in anaerobic jars by means of the AnaeroCult (Merck, Darmstadt, Germany) system for 2 weeks. The E. coli strains were maintained on LB-agar plates. Antibiotics were used in the following con- centrations (mgÆL )1 ): for E. coli, ampicillin (100), kanamycin (25), tetracyclin (20); for T. roseopersicina, kanamycin (25), streptomycin (5) and gentamycin (5). Expression of the hynS-isp1-isp2-hynL* genes of T. roseopersicina in E. coli using the T7 promoter ⁄ RNS polymerase system The hynS-isp1-isp2-hynL* gene products were produced from pTSH2 ⁄ 8 [8] in the E. coli BL21(DE3) strain. This construct contains the native promoter ⁄ regulatory region and, additionally, the complete hynS, isp1 and isp2 genes and truncated hynL (denoted by *). The incomplete hynL did not interfere with the outcome of the experiments. Expression of the genes was induced by isopropyl thio-b-d- P T7 P crtD RBS HoxE FLAG-StrepII kDa 120 100 321M 85 70 60 50 40 HoxF HoxU HoxE HoxE-TAG HoxH 30 25 20 Phenylalanyl t-RNA synthetase β subunit Phenylalanyl t-RNA synthetase α subuni t Fig. 1. The HoxFUYH subunits copurifiy with the tagged HoxE sub- unit during the course of affinity chromatography. (A) Scheme of the cassette used to express tagged HoxE in T. roseopersicina (P crtD , carotenoid promoter; P T7 , T7 promoter; RBS, ribosome-bind- ing site; M, marker). (B) Protein patterns of the elution fractions separated by SDS-PAGE. The soluble fraction of T. roseopersicina cells expressing tagged HoxE was loaded onto an Anti-Flag affinity column, the resin was washed, and the bound proteins were eluted three times by Flag peptide (for details, see Experimental proce- dures) (1, 2, 3 indicate the elution fractions). The bordered bands were cut and analysed by mass spectrometry. Electron-transfer subunits of NiFe hydrogenases L. S. Pala ´ gyi-Me ´ sza ´ ros et al. 170 FEBS Journal 276 (2009) 164–174 ª 2008 The Authors Journal compilation ª 2008 FEBS galactoside, and monitored by the incorporation of l-[ 35 S]methionine into the proteins synthesized [35]. The samples were separated in an SDS-polyacrylamide gel and analysed by a Phosphor Imager (Phosphor Imager 445 SI, Molecular Dynamics, Uppsala, Sweden). Conjugation Conjugation was carried out as described previously [27]. Deletion of the isp1,2 genes The in-frame deletion constructs were derived from the pK18mobsacB vector [36]. The upstream region of the isp1,2 genes was amplified with the otsh14r (5¢-GAT CGCGATATTGAACATC-3¢) and trhydo3 (5¢-CATA TGGCTGCCCGTAACCCCACTGAT-3¢) primers. The product was cloned into the polished BamHI site of pUC19 [37], yielding pUNSBamHI. To clone the downstream region, another PCR was per- formed with the isp1o7 (5¢-TCGCACGCTGGTACAA CGGG-3¢) and isp2o2 (5¢-ACCAGGTGCTCGGCGAT CAT-3¢) primers. This fragment was cloned into the XbaI- digested and blunted pUNSBamHI vector (pUS2). The 2502 bp EcoRI fragment of pUS2 was ligated with the EcoRI fragment of pK18mobSacB, yielding pISP12M. The plasmid was transformed into the E. coli S17-1(kpir) strain, and then conjugated into the T. roseopersicina GB2131 strain as described previously [27]. The single recombinants selected through their kanamycin resistance were grown in liquid medium. The double recombinants were selected on 3% sucrose-containing plates. The sucrose-resistant and kanamycin-sensitive colonies were selected, and the genotype was confirmed by Southern blotting and hybridization (ISP12M). Deletion of the isp1 gene The upstream homologous region was taken from the pUNSBamHI vector. The downstream homologous region was amplified with the isp1o8 (5¢-AGCTGACGCACATCT TCACG-3¢) and isp2o7 (5¢-GGTGAGACCGACCACCG GGA-3¢) primers. The product was cloned into the BamHI- cleaved and polished pUNSBamHI construct (pUS3). The EcoRI fragment of pUS3 was cloned into pK18mobSacB (pISM1 ⁄ 3). The construct was conjugated into the GB2131 strain and the double recombinants were selected as described below. Deletion of the hupC gene For deletion of the hupC gene, the pHCD1 and pHCD2 in-frame deletion constructs were created as follows. The upstream region of hupC was amplified with the ohup20 (5¢-CGAGCAGGCCAAGTATTC-3¢) and ohup19 (5¢-TGT TGGTCAGGCGGATCT-3¢) primers, and the 836 bp PCR product was cloned into the SmaI-digested pK18mobsacB (pHCD1). The downstream region was amplified with the ohup21 (5¢-GGCGGATGTTCAAGGACG-3¢) and ohup22 (5¢-TCGACCACGACACTGAAG-3¢) primers. The 800 bp fragment obtained was cloned into the PstI-digested polished pHCD1 (pHCD2). This construct was conjugated into the T. roseopersicina GB1131 strain, yielding the HCMG4 strain. The double recombinants were selected and the genotypes were confirmed as described above. Construction of HupC-expressing plasmid The hupC gene was amplified with the ohupc1 (5¢-CATAT GTCGCGAGCTGCGTCGCG-3¢) and ohupc2 (5¢-AAGCT TTGGCCGATCGTCCTTGAACAT-3¢) primers containing NdeI and HindIII recognition sites. The 777 bp PCR prod- uct was inserted into the EcoRV-digested pBluescripSK+ (pBtC). The 777 bp NdeI-HindIII-digested fragment was ligated into the corresponding sites of pMHE6crtKm [26], resulting in pMHE6C. RNA isolation For RNA isolation, T. roseopersicina was grown in 60 mL of liquid medium in a hypovial to A 600 nm = 1–1.5; 15 mL of culture was centrifuged at 15 000 g for 2 min, the pellet was suspended in 300 lL of SET buffer [20% sucrose, 50 mm EDTA (pH 8.0) and 50 mm Tris ⁄ HCl (pH 8.0)] and 300 lL of SDS buffer was added [20% SDS, 1% (NH 4 ) 2 SO 4 , pH 4.8]; 500 lL of saturated NaCl was added next, the sample was centrifuged at 20 000 g for 10 min and the clear supernatant was transferred into a new tube. 2-Propanol (70% of the total volume of the supernatant) was added to the solution and the mixture was centrifuged at 20 000 g for 20 min. The pellet was washed twice with 1 mL of 70% ethanol. The dried pellet was suspended in 20 lL of diethylpyrocarbonate-treated water. DNase I treatment DNase treatment took place in the presence of 10· reaction buffer with MgCl 2 (Fermentas, Burlington, Canada) and DNase I (RNase-free, Fermentas) at 37 °C for 1 h. The reaction was inactivated by heat at 65 °C for 10 min in the presence of EDTA (Fermentas). Reverse transcription and quantitative real-time PCR For reverse transcription, the OmniscriptÒ Reverse Trans- criptase Kit (Qiagen, Hilden, Germany) was used according L. S. Pala ´ gyi-Me ´ sza ´ ros et al. Electron-transfer subunits of NiFe hydrogenases FEBS Journal 276 (2009) 164–174 ª 2008 The Authors Journal compilation ª 2008 FEBS 171 to the manufacturer’s instructions. One microgram of DNase I-treated total RNA was added to a master mix [10· buffer RT, dNTP mix (0.5 mm of each dNTP), reverse primer (0.2 lm), RNase inhibitor (10 units ⁄ reaction), Omniscript Reverse Transcriptase (2 units ⁄ reaction) in diethylpyrocarbonate-treated water] on ice in a final volume of 20 lL. The reaction mixture was then incubated at 37 °C for 60 min. Reverse transcription was initiated from the huprto2 (5¢-CGCTTGAGCCGATTCTGAACAT-3¢) primer specific for the hupL gene. The cDNA produced during reverse transcription was used as a template for quantitative PCR, which was performed using the ohupSRT1 (5¢-GGA CAAGGGCAGCTTCTATCA-3¢) and ohupSRT2 (5¢-CG CATTGGCCTCGATACC-3¢) primers located in the hupS gene. PCR was carried out and the products were measured with an Applied Biosystems (Foster City, CA, USA) 7500 real-time PCR instrument. PCR was performed in a total volume of 25 lL, including 1 lL of cDNA, 12.5 lLof Power SYBR Green PCR Master Mix (Applied Bio- systems), forward and reverse primers (12.5 pmol of each) and 9 lL of nuclease-free water. The following programme was applied: 95 °C for 10 min; 95 °C for 15 s and 60 °C for 1 min for 40 cycles; 95 °C for 15 s; 60 °C for 1 min; 95 °C for 15 s; 60 °C for 15 s. A calibration curve was gen- erated using sixfold dilutions of pKK48 plasmid DNA (containing the sequence of the hupS gene) in the 100 to 0.001 ngÆlL )1 concentration range. Activity measurements The hydrogenase activities of the various mutants were measured both in vivo and in vitro. In all experiments, the HypF mutant (lacking any NiFe hydrogenase activity) and the GB112131 strain (DhoxH, DhupSL, DhynS-isp1-isp2- hynL) were used as negative controls. In vitro H 2 uptake activity measurements The samples were suspended in 2 mL of 20 mm potassium phosphate buffer containing 0.4 mm of oxidized benzyl- viologen. The cuvettes were closed with SubaSeal rubber stoppers. The gas phase was flushed with H 2 and the H 2 uptake activity was measured spectrophotometrically at 600 nm and 60 °C. In vivo hydrogen evolution activity measurements Cultures (60 mL) were grown in 100 mL hypovials; the gas phase was then flushed with N 2 after inoculation and the H 2 produced was measured gas chromatographically [27] on day 6. In vivo H 2 -uptake activity measurements Medium (60 mL) was inoculated into 100 mL hypovials; the gas phase was flushed with N 2 and 5 mL of pure H 2 was injected into the bottles. The cultures were grown under illumination and the H 2 content of the gas phase was measured gas chromatographically on day 6. Preparation of membrane and soluble fractions of T. roseopersicina The membrane fractions were prepared from 50 and 110 mL cultures for Hyn and Hup measurements, respec- tively. The cells were harvested by centrifugation at 7000 g, suspended in 1 mL of 20 mm potassium phosphate buffer (pH 7.0) and broken by sonication [Bandelin Sonopuls (Berlin, Germany) HD2070 ultrasonic homogenizer; at 85% amplitude six times for 10 s]. The broken cells were centri- fuged at 15 000 g for 10 min. The debris (sulfur globules and intact cells) was discarded and the supernatant was centrifuged at 100 000 g for 1.5 h. The pellet was washed, resuspended in 800 lL of potassium phosphate buffer (pH 7.0) and used as membrane fraction. The supernatant was regarded as the soluble fraction. Measurements of bacteriochlorophyll content The bacteriochlorophyll content was estimated using a methanol extraction procedure, as described previously [38]. The absorption of the samples was measured at 772 nm; the extinction coefficient was 8.41 g )1 ÆLÆcm )1 . The in vivo and in vitro activities were normalized to the bacteriochlo- rophyll content of the samples. Construction of the double-tagged hoxE gene For the construction of an expression system capable of producing the HoxE protein of T. roseopersicina fused with tandem FLAG-tag-Strep-tag II at the C-terminus, a 501-bp fragment was amplified from the pTCB4 ⁄ 2 clone [8] using the TCHO32 (5¢-CATATGAGTCTGCAGCAAGCCA-3¢) and TCHO33 (5¢-AAGCTTGGTCAGCTCCTCGAGC-3¢) primers and cloned into the SmaI site of pBluescript SK+ (pBtHoxE). The 494 bp NdeI-HindIII fragment of pBtHoxE was ligated into the NdeI-HindIII-digested pMHE6crtKm vector (pMHE6HoxE). The construct was confirmed by sequencing and conjugated into the T. roseo- persicina GB1121 strain. Purification of Hox hydrogenase Four grams of cell paste from a GB1121 ⁄ pMHE6HoxE- Km culture were suspended in 5 mL of NaCl ⁄ Tris [50 mm Electron-transfer subunits of NiFe hydrogenases L. S. Pala ´ gyi-Me ´ sza ´ ros et al. 172 FEBS Journal 276 (2009) 164–174 ª 2008 The Authors Journal compilation ª 2008 FEBS Tris (pH 7.4) and 150 mm NaCl]. The sample was sonicated with a Bandelin Sonopuls HD2070 ultrasonic homogenizer (at medium mode, amplitude 2.4 times for 10 s). The cell debris and sulfur crystals were removed by centrifugation (27 000 g, 10 min). The supernatant was incubated with 300 lL of ANTI-FLAG M2 affinity resin (Sigma) at 4 °C for 2 h with gentle shaking. The matrices were washed seven times with 1.5 mL of NaCl ⁄ Tris. For elution, the slurry was washed twice with 100 lL and once with 50 lL of NaCl ⁄ Tris with FLAG-peptide (200 lgÆmL )1 ). Aliquots were collected and the samples were analysed by SDS- PAGE. SDS-PAGE and protein staining SDS-PAGE and silver staining of proteins were performed as described by Ausubel et al. [39]. Identification of proteins by MALDI-TOF-MS Coomassie blue-stained gel bands were cut out and analy- sed by MALDI-TOF-MS, as described previously [26]. Bioinformatics tools Protein sequences in the various databases were compared with the blast (P, X) programs (http://www.ncbi.nih.nlm. gov), the peptide mass fingerprints and the power spectral density spectra; a database search was performed using the National Center for Biotechnology Information protein database with Protein Prospector MS-Fit and MS-Tag, respectively (http://prospector.ucsf.edu/). Acknowledgements The contribution of Drs B. D. Fodor and A ´ . T. 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