Evolution and diversification of Group 1 [NiFe] hydrogenases. Is there a phylogenetic marker for O2-tolerance?

Biochimica et Biophysica Acta 1817 (2012) 1565–1575

Contents lists available at SciVerse ScienceDirect

Biochimica et Biophysica Acta journal homepage: www.elsevier.com/locate/bbabio

Evolution and diversification of Group 1 [NiFe] hydrogenases. Is there a phylogenetic marker for O2-tolerance? Maria-Eirini Pandelia a,⁎, Wolfgang Lubitz a, Wolfgang Nitschke b a b

Max-Planck-Institut für Bioanorganische Chemie, Stiftstrasse 34‐36, D45470, Mülheim a.d. Ruhr, Germany Laboratoire de Bioénergétique et Ingénierie des Protéines, CNRS, FR3479, Aix-Marseille University, France

a r t i c l e

i n f o

Article history: Received 11 March 2012 Received in revised form 21 April 2012 Accepted 24 April 2012 Available online 1 May 2012 Keywords: [NiFe] hydrogenases Phylogenetic tree O2-tolerance Bioenergetics Iron–sulfur cluster

a b s t r a c t Group 1 hydrogenases are periplasmic enzymes and are thus strongly affected by the “outside world” the cell experiences. This exposure has brought about an extensive heterogeneity in their cofactors and redox partners. Whereas in their majority they are very O2-sensitive, several enzymes of this group have been recently reported to be O2-tolerant. Structural and biochemical studies have shown that this O2-tolerance is conferred by the presence of an unusual iron–sulfur cofactor with supernumerary cysteine ligation (6 instead of 4 Cys, hence called ‘6C cluster’). This atypical cluster coordination affords redox plasticity (i.e. two-redox transitions), unprecedented for this type of cofactors and likely involved in resistance to O2. Genomic screening and phylogenetic tree reconstruction revealed that 6C hydrogenases form a monophyletic clade and are unexpectedly widespread among bacteria. However, several other well-defined clades are observed, which indicate early diversification of the enzyme into different subfamilies. The various idiosyncrasies thereof are shown to comply with a very simple rule: phylogenetic grouping of hydrogenases directly correlates with their specific functions and hence biochemical characteristics. The observed variability results from gene duplication, gene shuffling and subsequent adaptation of the diversified enzymes to specific environments. An important factor for this diversification seems to have been the emergence of molecular oxygen. Hydrogenases appear to have dealt with oxidative stress in various ways, the most successful of which, however, was the innovation of the 6C-cluster conferring pronounced O2-tolerance to the parent enzymes. © 2012 Elsevier B.V. All rights reserved.

1. Introduction Chemiosmotic energy conversion, life's penultimate way to harvest energy, taps the ΔG available in redox disequilibria between reducing and oxidizing substrates [1,2]. Whereas the nature of the ancestral electron sink exploited by nascent life is still a matter of debate [3], molecular hydrogen almost certainly was the first electron source [1,2]. [NiFe] hydrogenases, extant enzymes using the H +/H2 couple for bioenergetic ends, are found in a wide variety of phyla in Archaea and Bacteria. Molecular phylogeny suggests them to fall into four distinct groups, the so-called Group 1 to 4 hydrogenases, a distinction supported by phylogenetic markers such as subunit composition and operon structure [4,5].

Abbreviations: DMKn, demethylmenaquinone n, where n refers to the number of the prenyl units present in the side-chain; UQn, ubiquinone n, where n refers to the number of the prenyl units present in the side-chain; MK, menaquinone; Hdr, heterodisulfide reductase; Dsr, dissimilatory sulfate reductase; CoB, coenzyme B (full chemical name 7-mercaptoheptanoylthreoninephosphate); CoM, coenzyme M is the anion with the formula HSCH2CH2SO3− ⁎ Corresponding author at: 104 Chemistry Building, Pennsylvania State University, PA 16803, USA. E-mail address: [email protected] (M-E. Pandelia). 0005-2728/$ – see front matter © 2012 Elsevier B.V. All rights reserved. doi:10.1016/j.bbabio.2012.04.012

Group 1 hydrogenases are generally designated as membraneassociated enzymes coupling the H +/H2 couple to quinones 1 in metabolic pathways with a variety of redox substrates. They are exported to the periplasmic space via an N-terminal signal peptide located in the small subunit that is known as the twin-arginine translocation motif (i.e. RRXFXK) [7], whereas the large subunit lacking this signaling motif is co-translocated with the small subunit. They represent the both phylogenetically most widespread and best studied group. A subclass – the so-called O2-tolerant Group 1 hydrogenases – has recently gained extensive attention, due to their potential biotechnological importance. Only a few cases of O2-tolerant enzymes have been studied so far, e.g. the enzymes from Aquifex aeolicus (Hase I) [5,8], Ralstonia eutropha (MBH) [9], Hydrogenovibrio marinus [10] and Escherichia coli (Hyd-1) [11,12]. Whereas Group 1 hydrogenases are in general typified via the extensively studied Desulfovibrio enzymes [13] (Fig. 1), the O2-tolerant ones are often considered as the exotic outliers of the group. Very recently, crystal structures of three O2-tolerant enzymes [10,12,14] showed the catalytic [NiFe] site and number of iron–sulfur 1 The archaeal hydrogenases from Methanosarcinales do not contain the usual quinones, but methanophenazines (MP), which are functional menaquinone analogs [6] (see also below and SI).

1566

M-E. Pandelia et al. / Biochimica et Biophysica Acta 1817 (2012) 1565–1575

this strategy to cope with O2 appeared several times independently or may it be a relatively ancient trait? To better understand the origin(s) and evolutionary pathways of the O2-tolerant 6C-hydrogenases, we have re-analyzed the topology of the phylogenetic tree of the Group 1 family with specific attention to structural marker characteristics of the complete hydrogenase enzyme complexes. The obtained results suggest surprising evolutionary pathways not only for the oxygen-tolerant subgroup but also for the whole ensemble of Group 1 enzymes. 2. Materials and methods

Fig. 1. Localization of the metal cofactors in the H2-reduced state of the O2-sensitive periplasmic hydrogenase from Desulfovibrio vulgaris Miyazaki F (pdb id: 1H2R, top) and of the membrane-bound O2-tolerant hydrogenase from Hydrogenovibrio marinus (pdb id: 3AYX, bottom). The large subunit (dark grey ribbon) contains the heterobimetallic [NiFe] site and the small subunit (white ribbon) contains the three iron– sulfur centers in an almost linear arrangement. The glutamate-rich surface near the distal [4Fe4S] (1His3Cys) cluster situated close to the surface is shown with red color. Note the difference in the structure of the cluster proximal to the [NiFe] site between the two enzymes (6C cluster in the H. marinus hydrogenase).

cofactors to be conserved, albeit with nuances. The most prominent modification is that the [4Fe4S] cluster proximal to the [NiFe] site (Fig. 1) has been replaced by a novel [4Fe3S] center, coordinated by a total of six cysteine residues (6C cluster), instead of the canonical four. The [4Fe3S] is able to perform two functionally meaningful singleelectron transitions [15], unlike traditional [4Fe4S] clusters. Its unique redox plasticity is promoted by redox-dependent structural changes resulting from its surplus cysteine coordination. Site-directed mutagenesis indeed showed that O2-tolerance of 6C-containing hydrogenases is dominantly linked to the presence of the [4Fe3S] cluster [16,17]. R. eutropha is a β-proteobacterium belonging to the so-called crown-group of bacterial prokaryotes. A. aeolicus, by contrast, is a member of one of the earliest branching bacterial phyla, the Aquificales. Branching-off of the Aquificales is considered to have occurred prior to the advent of oxygenic photosynthesis and the resulting rise in O2 levels in the biosphere. The simultaneous presence of the oxygen-tolerant 6C-hydrogenases in these two species therefore raises a number of evolutionary questions: Does it represent an evolutionary product that marks transition to a more oxygenic lifestyle? Has the trait of the 6C cluster been transferred laterally? Has

BLAST (Basic Local Alignment Search Tool) searches were performed using web tools from the National Center for Biotechnology Information (NCBI) (http://blast.ncbi.nlm.nih.gov/Blast.cgi) [18]. Protein BLAST analyses used the amino acid sequences of the small [NCBI Reference Sequence: NP_213454.1, Locus tag: aq_660], large [NCBI Reference Sequence: NP_213455.1, Locus tag: aq_662], cytb [NCBI Reference Sequence: NP_213456.1, Locus tag: aq_665] subunits of the A. aeolicus Hydrogenase (Hase) I as queries. Multiple sequence analysis and alignments were performed with the programs ClustalX (version 2), MUSCLE (http://www.ebi.ac.uk/Tools/msa/muscle/) and further refined (taking advantage of structural information) using Seaview (version 4). The trees were constructed with the ClustalX (version 2) and the MEGA 5 [19] software using the Bootstrap NJ algorithm without excluding protein gaps. An exhaustive and up-todate inventory of homologs was carried out on current sequence databases; the alignments were checked to remove highly divergent sequences, gaps, and ambiguously aligned positions. For this purpose, also the GUIDANCE web-server (http://guidance.tau.ac.il/overview. html) was additionally used as a comparative tool for assigning a confidence score for each residue, column, and sequence in an alignment and for projecting these scores onto the multiple sequence alignments. For the purpose of rooting our phylogenetic trees, we have tested the robustness of our trees by modifying the precise outgroup enzymes and varying the number of ingroup representatives. In the present study, we constructed our trees using as a single outgroup the soluble hydrogenase from A. vinosum DSM 180 (YP_003443825.1), since it gave essentially the same results. The phylogenetic trees were constructed using the sequences of the small subunits (iron–sulfur cluster containing) and the large subunits ([NiFe]-site containing) of the respective enzymes. Both provided essentially the same tree topology. Therefore, only trees constructed based on the sequences of the large subunit module are presented, which, due to their higher number of informative sites, yielded branching order supported by higher bootstrap values. Trees were visualized and edited further with the open source program TreeView (version 1.6.6). All structures were obtained from the RCSB PDB protein database (www.pdb.org) [20]. Redox potentials of b-cytochromes in the main manuscript are given in Table 2. The redox potentials for the NarI cytochrome in nitrate reductases have been measured in two different organisms in E. coli [21] and in P. denitrificans [22]. These potentials are significantly more positive compared to those measured for the cytbI, occurring in the majority of the Group I hydrogenases studied up to date as well as Formate Dehydrogenases [23–25]. Lists of all the organisms that have been used for the construction of the phylogenetic trees have been attached as a separate Excel Spreadsheet. 3. Results and discussion 3.1. Group 1 heterogeneity; gene cluster organization, subunit composition and nature of iron–sulfur clusters Group 1 hydrogenases are generally considered to consist of the large and the small subunit (Fig. 1) and to be connected to the

M-E. Pandelia et al. / Biochimica et Biophysica Acta 1817 (2012) 1565–1575

1567

Fig. 2. (Left) Gene cluster arrangements of Group 1 hydrogenases. The types of the three iron–sulfur clusters situated in the small subunit are distinguished for each case ([4Fe] as cubic and [3Fe] cubic with missing corner pictograms). The cluster proximal to the [NiFe] site shows extensive variations between different species in its primary coordination motif. The characteristic primary motifs are given in the beginning of each line (a) to (g), the special coordinating amino acids and the pictograms of the clusters are highlighted in the respective colors. (Right) Structure of the classical four-cysteine ligated cluster typically encountered in the proximal position in Group 1 enzymes (pdb id: 1H2R, H2-reduced D. vulgaris MF). The six-cysteine coordinated (6C) cluster in the O2-tolerant H. marinus hydrogenase (pdb id: 3AYX, H2-reduced state); the two extra cysteine residues are colored in red.

quinone pool via a di-heme cytochrome b (cytb) that, together with the hydrophobic C-terminus of the small subunit, anchors the enzyme to the membrane. These prototypical Group 1 hydrogenases are encoded by clustered genes arranged in the order small-largecytb I-subunit (Fig. 2a). In the small subunit, a [3Fe4S] cluster is located in the medial position between two [4Fe4S] centers. The cluster proximal to the [NiFe] site is an all-cysteine (4Cys) ligated [4Fe4S] cluster and the one most distant from the [NiFe] site (distal) is coordinated by three cysteines and one histidine (3Cys1His) (Table 1, Fig. 1). However, both genome surveys [4] and biochemical studies [5,11,26] have shown numerous members of the group featuring substantially different make-ups with respect to all these characteristics. Our comparative genome screening approach for Group 1 hydrogenases came up with 7 basic arrangements differing in gene cluster architecture, presence of additional/alternative subunits and structure of the iron sulfur clusters (Fig. 2). Group 1 thus starts to look more like a mixed crowd than a homogeneous group. Membership to Group 1 was initially based on sequence markers [4] but later shown to correspond to phylogenetic clustering. In the following, we will show that the seven variants shown in Fig. 2 actually correspond to a phylogenetic subclustering and that the subclades most likely arose in response to specific energetic requirements and environmental constraints.

3.2. Phylogeny orders the apparent Group 1 heterogeneity Phylogenetic analysis has been carried out using 20 archaeal and 93 bacterial sequences, based on sequence homology of the large catalytic and the small electron-transferring subunits. The number of sequences retrieved in Blast searches was equilibrated to avoid disproportionate trees resulting from species biases in genomic databases. An unrooted radial phylogram based on the aligned sequences of the large subunits is shown in Fig. 3. For rooting of the tree, the Group 3 soluble hydrogenase from Allochromatium vinosum was used as outgroup (Fig. 4). We have tested the robustness of our trees by modifying the precise outgroup enzymes and varying the number of ingroup representatives. Two observations are noteworthy: (a) The clades discernible were strictly robust, that is, never mixed or coalesced. These clades are further supported by evolutionary marker traits, i.e. specific idiosyncrasies with respect to gene cluster organization, nature of the membrane-anchor subunit and ligands to redox centers. (b) Branching topologies in the archaeal domain were relatively stable. This was also true for the lowest two phyla in the bacterial part of the tree, i.e. Actinobacteria and δ-proteobacteria (Fig. 3). Both these two phyla also figure prominently at the base of recent 16S rRNA-based species trees [27]. Considering the rapid sequence of radiations (Figs. 3, 4) and the low bootstrap values for the

Table 1 Binding motifs of the three iron–sulfur clusters occurring in the membrane bound hydrogenases of Group 1 (Figs. 1, 2). Proximal is the cluster closest to the [NiFe] site, medial the middle one and distal the most distant one (Fig. 1). The two extra cysteines of the proximal 6C cluster occurring in known O2-tolerant hydrogenases are indicated in bold. Classes

Proximal

Medial

Distal

6C Hases Prototypical/HybA CxxN (bacteria) CxxD (bacteria) sulfate reducers (bacteria) Bacteria Isp Crenarchaea Isp (CxxN) Crenarchaea Isp (CxxD) Euryarchaea/Actino 3x[4Fe] Euryarchaea DxxC 3x[4Fe]

CXCC X93–94 CX4CX28–30C CXGC X91–96 CX33–38C CXGN X94–96 CX39–40C CXGD X92–94 CX40C CXGC X94 CX32–35C CXGC X92–96 CX33C CXGN X114–138 CX80–82C CXGD X118–178 CX81–83C CXGC X92–107 CX37–49C DXGC X107 CX49C

CX11 PX6CX2C CX11 PX6CX2C CX11 PX6CX2C CX11 PX6CX2C CX11–13 PX6CX2C CX11 PX6CX2C CX13 PX6CX2C CX13 PX6CX2C CX11 CX6CX2C CX11 CX6CX2C

HX2C X24CX5C HX2C X24CX5C HX2C X24CX5C HX2C X24CX5C HX2C X24CX5C HX2C X24CX5C HX2C X21CX5C HX2C X21CX5C HX2C X24CX5C HX2C X24CX5C

1568

M-E. Pandelia et al. / Biochimica et Biophysica Acta 1817 (2012) 1565–1575

Fig. 3. Radial phylogram of the Group 1 hydrogenases based on the sequence homology and comparison of the large subunit (contains the catalytic [NiFe] site). The Group 3 soluble hydrogenase from A. vinosum DSM 180 (YP_003443825.1) has been included as outgroup. The tree has been built by the neighbor-joining (NJ, distance-matrix) method considering gap positions and accounting for multiple substitutions. The tree has been built using 1000 bootstrap samplings.

corresponding nodes (Fig. 4), it is not surprising that the branching order of the higher clades varied substantially as a function of the whole sequence ensemble.

Most of these higher clades predominantly contain members from more recently branched phyla with only a few isolated cases of obvious lateral gene transfer into specific members of Actinobacteria and δ-proteobacteria.

3.3. Bioenergetics and molecular O2; an intricate interplay driving molecular evolution

3.3.2. Hydrogenases with low oxygen-tolerance

3.3.1. Oxygen-sensitive hydrogenases, the likely ancestral type In the bacterial domain, the earliest branching part consists of Actinobacteria, sulfate-reducing and ε-proteobacteria. These organisms are strict anaerobes and contain the simplest structures of [NiFe] gene operons (Fig. 2, lines c, d, g) as well as a four or threecysteine ligated proximal tetranuclear (most likely) [FeS] cluster. The same types of hydrogenases (disregarding deviant proximal cluster ligation, see below) occur in many Archaea (Fig. 2 lines c, d). It is therefore likely that these come close to the ancestral Group 1 hydrogenases having existed in the Last Universal Common Ancestor (LUCA). These hydrogenases are extremely oxygen-sensitive and typical representatives are those from Desulfovibrio (periplasmic) and Wolinella species (membrane-attached and reducing menaquinone (MK)). The [NiFe] enzymes from Desulfovibrio are the most thoroughly studied and are classified as Group 1 despite their interaction with a soluble cytochrome c3 (cytc3) rather than with quinones (Fig. 5). The properties of the higher radiations (Fig. 3) suggest rapid diversification from this ancestral structure into new subfamilies, based on gene duplication events and recruitment of additional subunits according to the Redox-protein-construction-kit principle [28].

3.3.2.1. HybA-hydrogenases. HybA-enzymes contain in between the small and large subunit genes two other additional genes, hybA and hybB (Fig.2, line f). The HybA protein contains 16 conserved cysteines probably ligating four [4Fe4S] clusters and belongs to the family of tetracubane electron-transfer proteins [25,28]. HybB codes for a membrane-integral protein, predicted to have 9–10 transmembrane helices and frequently annotated as “putative cytochrome subunit”. However, there are no conserved canonical binding sites for hemes. Its high similarity with a class of membrane-associated proteins including PsrC (polysulfide reductase), DmsC (dimethyl sulfoxide reductase) and NrfD (assimilatory nitrite reductase) strongly suggests that HybB serves as a quinone reduction site [29] and, similar to PsrC, is devoid of b-hemes (Fig. 5). HybA-hydrogenases are clustered into a monophyletic clade (Figs. 3, 4) with mainly γ- and very few α- and β-proteobacterial members. The best characterized example is Hydrogenase-2 (Hyd-2) from E. coli [11,30,31] and this enzyme provides important clues for deducing the metabolic function of HybA-hydrogenases. Hyd-2 is bidirectional in vitro and tolerates limited concentrations of O2 under very reducing conditions [11]. It shows an increased tendency for hydrogen production. The observation that Hyd-2 can perform hydrogen catalysis at low

M-E. Pandelia et al. / Biochimica et Biophysica Acta 1817 (2012) 1565–1575

1569

Fig. 4. Rooted phylogram of Group 1 hydrogenases constructed from the aligned sequences of the large subunit gene. The soluble Group 3 hydrogenase from A. vinosum has been used as outgroup. A detailed description of the organisms used is contained in the SI. Selected organisms have been annotated: the crenarchaeal hydrogenases from Thermoproteus tenax (CCC80712.1) and Thermoproteus uzoniensis 768–20 (YP_004338974.1), the MBH from R. eutropha (NP_942644.1), MBH from H. marinus (provided by Higuchi and coworkers)8, Hyd-1 from Escherichia coli (NP_415492.1), Hase I from A. aeolicus (NP_213455.1), MBH from A. vinosum DSM 180 (YP_003443990.1), HynL from Thiocapsa roseopersicina (AAC38282.1), the membrane associated hydrogenase from Alteromonas macleodii str. ‘Deep ecotype’ (YP_004425482.1), the MBH from Paracoccus denitrificans PD1222 (YP_916875.1). Full and open squares denote the bootstrap values of high and small confidence, respectively. The tree was built using 1000 bootstraps repetitions. Different colors have been used to indicate the respective clades (see text).

oxygen concentrations (~1% O2) [11] may indicate that these enzymes encounter, and thus have to cope with, oxygen in vivo, albeit at relatively low concentrations. These hydrogenases, therefore, likely emerged at a point in the Earth's past when oxygen appeared in the environment. The present species distribution of HybA-hydrogenases suggests them to have emerged prior to the radiation of α-, β- and γ-proteobacteria. Precisely this region of the proteobacterial subtree has been suggested previously, based on quinone content, to mark the emerging impact of oxygen in the biosphere [32]. Hyd-2 from E. coli couples periplasmic hydrogen oxidation (−420 mV) to cytoplasmic fumarate reduction (+30 mV) [11,30,33]. The redox potentials of the catalytic [NiFe] site and the iron–sulfur clusters have not been determined so far. By comparison of their electrochemical properties, the reduction potentials may be inferred as close to those of oxygen-sensitive hydrogenases [15,34] (Fig. 6).

A striking property of the HybA-hydrogenases is the absence of bhemes in their membrane-anchor subunit which likely couples H2 oxidation to quinone reduction. For Hyd-2, absence of b-hemes finds a natural rationalization a) in the low ΔG available for electron transfer from H2 to fumarate and b) localization of H2 and fumarate catalytic sites with respect to the cytoplasmic membrane. H2 oxidation releases protons into the periplasm and builds up a proton gradient. Shuttling the derived electrons across the membrane for quinone reduction and hence proton uptake from the cytoplasm further increases this gradient. However, transferring electrons against the 150 mV electrostatic potential of an energized membrane consumes commensurable redox energy. The succinate/fumarate couple is substantially less oxidizing than most common terminal acceptors (nitrogen oxides, Fe(III), O2, etc.), making the ΔE of the donor/acceptor couple insufficient for sending electrons against the membrane

1570

M-E. Pandelia et al. / Biochimica et Biophysica Acta 1817 (2012) 1565–1575

Fig. 5. The four redox partner modules of Group 1 hydrogenases. (a) cytbI features the two-helix motif (the 4 His ligands of the 2 hemes come from two helices). The structural model shown is that of the apo-cytbI from A. aeolicus constructed based on the γ subunit of E. coli Formate Dehydrogenase-N (FdnI, pdb id: 1KQF). Hemes were included after structural alignment. (b) cytbII (or Isp1) has the three-helix motif (the 4 His ligands of the 2 hemes come from three helices). The structural model shown is that of the apo-cytbII from Rhodobacter sp. SW2 (ZP_05843854.1) constructed based on the γ subunit of E. coli nitrate reductase (NarI, pdb id: 1Q16). (c) The cytochrome-devoid HybB protein. It belongs to the superfamily typified by the γ subunit of polysulfide reductase, for which the crystal structure with bound menaquinone is shown (PsrC-MK7, pdb id: 2VPZ). (d) The soluble cytc3 (4 hemes) from D. vulgaris MF (pdb id: 1J0O), is the redox partner of [NiFe] hydrogenases from sulfate-reducing bacteria.

potential and, as a result, the b-hemes are disposed of as in the case of polysulfide reductase [29]. The conserved presence of hybB suggests that throughout this clade, the parent electron transfer chains use weakly oxidizing terminal acceptors. This scenario might, at first sight, seem in conflict with the occurrence of one α- and two β-proteobacterial representatives, i.e. species that usually are devoid of low potential menaquinones. The redox potential of ubiquinone, however, would be too high to operate such “short” chains toward weak oxidants. It is intriguing that one out of these three cases belongs to a group known to produce rhodoquinone [35], a low-potential quinone proposed to be derived from ubiquinone [36]. It seems likely to us that low-potential quinones (likely rhodoquinone) also exist in the two remaining α- and β-proteobacterial species. 3.3.2.2. Isp-hydrogenases. In Isp-hydrogenases two additional open reading frames, called isp1 and isp2, are positioned in between the small and large subunit genes (Fig. 2, line e). Homology modeling and hydrophobicity analyses predict that Isp1 has five transmembrane helices and binds two hemes via the so-called two-helix motif. By virtue of sequence similarity, Isp1 belongs to the b-heme superfamily (Fig. 5, cytb II cytochrome) including NarI [6,37–39], HdrE from methanogenic Archaea [40], DsrM from A. vinosum [38] and HmeC 2 from A. fulgidus [37], whose role, when known, is to oxidize the quinone pool [37,38]. All of these isp1-like genes are followed in their gene clusters by the isp2 gene. Isp2 has high sequence similarity to the catalytic subunit HdrD of heterodisulfide reductase from Methanosarcina barkeri [26,40]. Hdr proteins typically harbor two conventional [4Fe4S] clusters and two tetranuclear clusters with a unique CX31–38CCX33–34CXXC motif (5C) [6,40,42]. The C-terminal copy of these 5C clusters is thought to mediate the reductive cleavage of a disulfide bond [40] and Isp2 contains only this C-terminal copy. Isp-hydrogenases form two separate subclades, one in Archaea and one in Bacteria (Fig. 3), leaving us confronted with an evolutionary conundrum, which can be rationalized by two distinct scenarios. Either the isp1–isp2-module, present in other enzyme systems [37,38,40] was inserted twice independently into an ancestral small-large subunit gene cluster or the tree resolution for very deep evolutionary times may be insufficient. The “actual tree” may in that case correspond to two separate subtrees for prototypical and Isp2 HmeC is more properly referred to as DsrM [41], though when it was isolated from Archaeglobus fulgidus it was given the name HmeC.

hydrogenases, respectively, which would have both co-existed in LUCA. Weak sequence divergence may preclude resolving the two subtrees and thus cause the apparent collapse into a single tree. At this point, both (or still further) scenarios are possible. Gleaning the metabolic role of Isp-hydrogenases is more difficult than for HybA-enzymes, since for no member an unambiguous function has been elucidated so far. Typical representatives are the membrane-attached enzymes from Thiocapsa roseopersicina and A. vinosum. In their purified state they only contain the catalytic heterodimer (HynSL) and lack Isp1–Isp2. Mutational studies showed that Isp1–Isp2 are critical for in-vivo H2 cycling; in their absence hydrogen production is abolished, whereas hydrogen oxidation is ~ 70% decreased [26]. This strong link between hydrogenase function and presence of the isp genes together with their central localization in the operon (rendering co-regulation and co-expression very likely) demonstrate that the two Isp proteins are an integral part of the hydrogenase complex in vivo. The best characterized Isp-hydrogenase, A. vinosum HynSL [43,44], is oxygen-sensitive arguing for a role in an anaerobic bioenergetic chain. The redox potentials of its catalytic [NiFe] states are close to the H2/H + couple (− 420 mV) [45], the [3Fe4S] cluster titrates at +10 mV [46] and the two [4Fe4S] clusters at ≤−300 mV [43], i.e. rather similar to those reported for the oxygen-sensitive hydrogenases from the Desulfovibrio anaerobes [15,34] (Fig. 6). Unfortunately, the redox properties of the associated Isp1–Isp2 modules are not known. Other homologous biochemically characterized cases are the heme/iron–sulfur complex (HmeCD) 3 from A. fulgidus [37], the DsrMK complex from Desulfovibrio desulfuricans ATCC 27774 [41], proposed to be involved in disulfide reduction (linking quinones to dissimilatory sulfite reduction) with so-far unknown thiol substrates. Similar to Isp2, HmeD and DsrK contain only the C-terminal 5C cluster. Their midpoint potentials in the substrate-bound state is +90 mV [37] and +130 mV [41], respectively and thus more positive than that of HdrE from Methanothermobacter marburgensis (−185 mV for CoM-HDR, −30 mV CoB-HDR) [40]. The reduction potentials of cytb II-type hemes (Isp1) will depend on the chemical nature of the quinone pool and the substrate of the Hdr-type protein (Isp2). With the exception of HdrE from M. thermophila, linked to methanophenazine (E = −165 mV) [6,47], the redox potentials of cytb II-type hemes are more positive than those of cytb I

3

HmeCD is more properly referred to as DsrMK [41].

M-E. Pandelia et al. / Biochimica et Biophysica Acta 1817 (2012) 1565–1575

1571

Fig. 6. Redox scheme of the different Group 1 enzymes constructed from spectroscopic and biochemical data. 6C-hydrogenases: the [NiFe], iron–sulfur clusters' and cytbI redox potentials are those measured for A. aeolicus Hase I. Hydrogenases from sulfate reducers (SR): redox potentials given are for D. vulgaris MF and D. gigas. Isp-hydogenases: redox potentials are those measured for A. vinosum. The redox potentials of the Isp1–Isp2 complex were assumed similar to those of HmeCD/DsrMK (Table 2). HybA-hydrogenases: redox potentials were deduced from electrochemical data on E. coli Hyd-2. Redox potentials of the HybA iron–sulfur clusters were assumed to lie close to those of DmsB/NarH. MKreactive hydrogenases: redox potentials were considered similar to those of SR and HybA-hydrogenases (based on the W. succinogenes cytbI redox potentials and phylogenetic position of these enzymes). On the right the redox potentials of electron acceptors are given. Note that sulfate is a much weaker oxidant than O2; linking of the respective enzymes to these two different electron acceptors thus dictates their different redox properties.

[22,23,25,48] (Fig. 6, Table 2). Methanophenazine in methanogens is a functional analog of menaquinone [47,49]. It is a small hydrophobic quinone-like redox cofactor with an isopropenoid side-chain (see Supplementary information). Its role in the energy metabolism of methanogens is similar to that of quinones in mitochondria and bacteria. The final electron acceptor of the HynSL–Isp1–Isp2 supercomplex is unknown. However, it seems to us that the presence of cytb II and the heterodisulfide-related Isp2 protein strongly indicates a link to a sulfur-dependent metabolic pathway [26]. This hypothesis is supported by the observation that HynSL from T. roseopersicina is expressed in the presence of thiosulfate [26]. Which kind of redox reaction between sulfur compounds and the H2/H+ couple might the Isp-hydrogenases catalyze? The gene cluster and observed targeting sequences suggest an enzyme consisting of a periplasmic hydrogenase module (small-large subunit), a cytoplasmic redox protein converting sulfur compounds (most likely S\S bonds) and a b-cytochrome serving as transmembrane electron wire. This structure allows for several bioenergetically sensible electron-transfer scenarios. The simplest one might stipulate periplasmic H2 oxidation (liberating two protons), followed by transmembrane electrontransfer (against the membrane potential) and ultimately reducing unspecified disulfide bonds (consumption of two protons in the cytoplasm). This ultra-short electron-transfer chain can generate a proton

gradient and thus be energetically viable under certain growth conditions. More complicated electron-transfer models can be thought of, if involvement of quinones is envisaged. Electrons issued both from hydrogen and sulfur oxidation to disulfides might merge to reduce quinone at the cytoplasmic face of the membrane. The quinol would then go downstream toward higher potential terminal acceptors (or even pass through the Rieske/cytb complex). Finally, even the reverse scenario is possible, despite its apparent counterintuitive aspects: the Isp-complex might act as a terminal electron acceptor from reduced quinol via electron bifurcation to hydrogen pulled by reduction of a higher potential acceptor (the S\S bond). Recently, the heterodimeric (SL) hydrogenase from the γ-proteobacterium Alteromonas macleodii was reported to tolerate the presence of 1–3% oxygen [50]. This enzyme shares a strong homology (60–69%) with Isp-hydrogenases [50] but lacks isp1–isp2. Phylogenetic analysis places the A. macleodii enzyme in the midst of Isp-hydrogenases (Figs. 3, 4), whereas absence of Isp1–Isp2 likely indicates coupling to altered electron acceptors. Although the T. roseopersicina HynSL was initially reported to be relatively O2-resistant, recent studies showed the activity to be lost at ~ 1% O2 [50]. Similarly, the A. vinosum HynSL is considered to be very oxygen-sensitive. In this context, the decreased O2-sensitivity of the A. macleodii enzyme is intriguing. This difference may be

Table 2 Binding motifs of the two b-cytochrome types encountered in Group 1 hydrogenases. cytbI belongs to the FdnI superfamily. It adopts the so-called three-helix motif, i.e. the histidine ligands to the hemes come from three helices. cytbII or more commonly Isp1 belongs to the NarI superfamily. It adopts the so-called two-helix motif, i.e. the histidine ligands to the hemes come from two helices. The last column contains the midpoint redox potentials for the different di-heme cytochromes quoted versus the standard hydrogen electrode potential.

a

Subgroup

cytbI

CXCC (bacteria)

HX40–46H HX13H

DSGC (Archaea) CXGC (bacteria) CXGD (bacteria) CXGN (bacteria) CXGC (Archaea/actinobacteria [4Fe]) CXGN (bI‐like Crenarchaea) Formate dehydrogenase (W. succinogenes pdb id: 1QLB) CXGC (Isp bacteria) CXGN (Isp Crenarchaea) CXGD (Isp Crenarchaea) Nitrate reductase (E. coli pdb id: 1Q16) Nitrate reductase (P. denitrificans) DsrM (A. vinosum)

HX32H HX13H HX37–42H HX13H HX38–41H HX13H HX38–41H HX13H HX30–33H HX13H HX58H HX14H HX38H HX13H

[19].

b

[44]. c [17].

d

[18]. e [34].

cytbII (Isp1)

Redox potential, mV − 20, − 110 (A. aeolicus)a + 130, − 60 (A. aeolicus)a − 100, − 240 (W. succinogenes)b

− 20, − 190b HX12H HX17H HX12–13H HX17H HX13H HX17H HX9H HX17H HX9H HX17H HX9H HX17H

− 10, + 125c + 95, + 210d + 60, + 110e

1572

M-E. Pandelia et al. / Biochimica et Biophysica Acta 1817 (2012) 1565–1575

explained considering that O2-tolerance of enzymes from the Ispclade may crucially depend on their in vivo operating potential and consequently their precise metabolic role. 3.3.3. Oxygen-tolerant 6C-hydrogenases The most intriguing clade is perhaps that containing the oxygentolerant hydrogenases. Already more than 30 years ago, an atypical EPR signature was observed in hydrogenases from R. eutropha and P. denitrificans [51,52], never seen before in those from sulfate reducers [34,53]. These signatures, suggestive of an additional paramagnetic center, subsequently resurfaced in the spectra of other Group 1 hydrogenases that are members of this clade (A. aeolicus [5,15], Azotobacter vinelandii [54], Hydrogenophaga sp.AH-24 [55], Cupriviadus metallindurans [46] and E. coli [11]). The identity of the unknown cofactor remained enigmatic for almost two decades and was elucidated only recently [15]. EPR electrochemical and Mössbauer studies identified this paramagnetic center to a second, higher potential redox transition of the proximal cluster [15]. The diamagnetic “oxidized” proximal cluster can be reduced to a state that resembles spectroscopically [4Fe4S]1 + ferredoxins (S = 1/2). However, in the O2-tolerant hydrogenases, it can furthermore be super-oxidized to a form reminiscent of the paramagnetic 3+ state of HiPIP centers [15]. The paramagnetism of the super-oxidized proximal cluster is responsible for the atypical complex spectroscopic signatures of these hydrogenases [8,11,56,57], which result from magnetic interactions with the neighboring paramagnetic [NiFe] and [3Fe4S] 1 + centers [15]. The recent crystallographic structures from two members of this family [10,12,14] revealed that the proximal cluster is actually a [4Fe3S] cluster with in-total six coordinating cysteine residues (Figs. 1, 2 line b). These six cysteines are fully conserved in this family, hence called 6C clade. The sulfur atom of the first extra cysteine (C) in the primary binding motif CTCC replaces one of the inorganic sulfides and thus becomes an intrinsic cluster ligand [10,14], whereas the second cysteine terminally coordinates one of the Fe atoms [10,14] (Fig. 1, bottom). In line with the dual redox character of the 6C cluster, the structures of the super-oxidized and reduced forms show unique features. The CC tandem cysteine motif appears to be critical for its structural and redox plasticity, stabilizing the super-oxidized form by offering an additional coordinating ligand via deprotonation of the CC peptide amide nitrogen [10]. These redox-dependent structural changes promoted by the surplus cysteine coordination, allow the 6C [4Fe3S] cluster to carry out two single-electron transitions. Interestingly, a direct correlation of O2-tolerance and the presence of the 6C cluster could be demonstrated via cysteine deletion mutants in R. eutropha MBH [16]. These were found to show growth limitation for oxygen concentrations higher than 2%, the O2-sensitivity of the purified enzyme was significantly increased [16], whereas the atypical EPR features due to the presence of the super-oxidized proximal cluster were absent. Our phylogenetic trees demonstrate that the notion of O2-tolerant 6C hydrogenases being an oddity among Group 1 enzymes is misguided (Figs. 3, 4). The 6C cluster is not restricted to enzymes from (micro)aerophilic organisms, but is also found in those from strict anaerobes (e.g. Heliobacteria, Chlorobi). The presence of the complete gene sequences in these operons and the wide distribution of 6Cenzymes in the Chlorobi/Firmicutes phyla, shows that they encode functional hydrogenases rather than representing pseudogenes. In the light of the proposed rescue function for this cofactor against oxidative stress [16], this was unexpected. The clade of 6C-hydrogenases is monophyletic and all members share the 6C characteristics (i.e. likely the structure of the [4Fe3S] proximal cluster). 6C-type enzymes are found in the crown-group α-, β-, γproteobacteria but also in δ-proteobacteria, Bacteroidetes, Aquificales, Chlorobiaceae and Firmicutes. Since the 6C cluster confers a substantial fitness advantage in environments at least occasionally exposed to O2,

a wide species distribution driven by lateral gene transfer appears likely. However, the topology of the 6C-clade strongly argues for a different scenario. The 6C tree in fact splits into two well-separated parts; (a) Bacteroidetes, Chlorobiaceae and Firmicutes (Heliobacteria and Desulfitobacteria) and (b) α-, β- and γ-proteobacteria. In the upper part (a) of the 6C subtree, low bootstrap values hamper conclusions on vertical or lateral gene transfer. The cleavage, however, between Proteobacteria and Chlorobiaceae/Firmicutes/Bacteroidetes, is strongly bootstrap-supported and reflects 16S rRNA species trees [27] (Fig. 4). Aquificales appear as a subgroup branching at the base of the proteobacterial sequences. As shown previously, a high percentage of their genomes was imported laterally from ε-proteobacteria [58] and Aquificales may even serve as repositories for ε-proteobacterial genes that are difficult to recognize in extant ε-proteobacteria. It therefore appears likely that the group of Aquificales actually represents sequences arising from ε-proteobacteria, making this “side” of the 6C tree closely resemble the proteobacterial species phylogeny (Figs. 3, 4). After the branching-off of the Aquificales, the next higher node splits into two distinct clades, both of which contain α-, β- and γ‐proteobacteria (Fig. 4). The upper clade is characterized by high bootstrap values and reproduces the cleavage into α-, β-, γ-proteobacteria characteristic for species trees [27]. The lower clade features very low bootstrap values and an extreme mixture of α-, β-, γ- and δ-proteobacterial representatives. It cannot be decided whether the disordered topology is due to pervasive lateral gene transfer or whether it reflects insufficient phylogenetic signal. Irrespective of this ambiguity, a gene duplication event in the ancestor of α-, β-, γ-proteobacteria appears to have given rise to these two distinct micro-families of 6C-hydrogenases. The tree topologies shown in Figs. 3 and 4 suggest a provocative conclusion: 6C-hydrogenases appear to have emerged prior to the divergence of Chlorobia/Heliobacteria on one and α-, β- and γ-proteobacterial radiations on the other side. In the traditional thinking that Heliobacteria and Firmicutes have diverged prior to the origin of oxygenic photosynthesis, the presence of 6C-enzymes in these species would indicate that their appearance pre-dates the emergence of light-driven O2 production. Therefore, either the 6C cluster appeared without an apparent link to O2-tolerance or we need to rethink our evolutionary history of oxygenic photosynthesis. The problem we are facing is not entirely new and has been raised previously based on the presence of an additional cofactor also suspected to confer O2-tolerance in the Rieske/cytb complexes of Heliobacteria and Firmicutes [59]. Indeed, molecular phylogenies of several O2-dependent or O2-tolerant enzymes are in favor of a deeper than previously thought ancestry of oxygenic photosynthesis (manuscript in preparation). A more detailed discussion of this topic, however, is beyond the scope of the present article. The expression of 6C-hydrogenases is now commonly thought to be linked to hydrogen oxidation in the presence of O2 or even necessarily with O2 acting as terminal electron acceptor, i.e. to the Knallgas reaction. In addition to the special proximal cluster, however, the 6C-enzymes studied so far stand out by high redox potentials of all three iron sulfur clusters [15,46] (Fig. 6), a property often correlated to O2 serving as terminal acceptor. In several species, by contrast, 6C-hydrogenases have been described to work under anaerobic or microaerobic conditions using high-potential acceptors other than O2, e.g. nitrogen oxides. The 6C Hyd-1 from E. coli is expressed in anaerobic cultures, whereas there is evidence that its presence is of importance when the cells experience microaerobic conditions [11,60]. E. coli deletion mutants containing only Hyd-1 were active when grown on high-potential electron acceptors such as nitrate and O2 [11,60]. The 6C-enzyme from T. roseopersicina (HupSL) is induced under anaerobic conditions and upon decreasing the thiosulfate concentration during growth, which will lead to an increase in cellular redox potential [26] although the ultimate electron acceptor is not known. In the obligate phototrophic Chlorobiaceae and

M-E. Pandelia et al. / Biochimica et Biophysica Acta 1817 (2012) 1565–1575

Heliobacteria, electrons produced from 6C-hydrogenases need to end up reducing the (relatively high-potential) photo-oxidized reaction center pigment. From the presently available data it seems likely to us that enzymes from the 6C clade are not only characterized by their special proximal cluster, but also by a common redox up-shifted electrontransfer FeS chain and correspondingly high-potential redox partners (down the line to the terminal acceptor) (Fig. 6). Whereas the emergence of the 6C cluster was possibly driven by the appearance of O2, the redox upshift is certainly due to the use of high-potential terminal acceptors such as oxygen, nitrogen oxides or photosynthetic reaction centers. Whether the common ancestor of all 6C-hydrogenases originated in a “Knallgas”‐type, a denitrifying or a photosynthetic bioenergetic chain remains an open question. Another open issue is how this ancestor was able to synthesize the sophisticated 6C [4Fe3S] cluster. For [NiFe] hydrogenases no specific maturation genes for the assembly of the iron–sulfur clusters have been identified so far suggesting that biosynthesis of these cofactors occurs via standard biosynthetic pathways (i.e. ISC, SUF) [61,62]. So, is the 6C cluster the result of a more specialized assembly apparatus? Recent studies on the “Knallgas” β-proteobacterial hydrogenase from Rhizobium leguminosarum employing deletion of accessory genes suggested that hupGIJ (homologous to hoxORT in R. eutropha) [63] are involved in the maturation of the iron–sulfur cluster subunit [64]. More precisely, the hoxR gene of R. eutropha MBH was proposed to be important for oxygen-tolerance and presumably involved in the formation of the [4Fe3S] proximal cluster [63]. Intuitively, this proposal is tempting but fails to explain the absence of the hoxR gene from all species having 6C-hydrogenases outside the α-, β-, γ-proteobacterial family. HoxR therefore cannot be essential for maturation of the 6C cluster. HoxR has an all-cysteine ligated rubredoxin site, indicative of its possible role as a redox protein against oxidative stress conditions (electron safety valve to protect easily oxidizable centers). Since adaptation to higher oxygen levels is probably a multi-step process, it seems more likely to us that HoxR is rather involved in accrued protection of the 6C cluster (if linked to the 6C cluster at all). The exclusive occurrence of hoxR in the most recent branching orders of the proteobacteria corroborates this scenario. 3.4. The proximal cluster, a pivotal redox center in [NiFe]-hydrogenases The previous section illustrates the outstanding role of the proximal cluster in modulating the overall properties of [NiFe]-hydrogenases. Due to its close spatial proximity to the [NiFe] center, the proximal cluster has in fact been discussed in the past as being to some degree part of the catalytic core of the enzyme [65]. From this point of view it is not astonishing that hydrogenases extensively experimented with its coordination pattern (Fig. 2, Table 1). The most frequent case of the primary CxxC motif is the CTGC pattern. Two cases of archaeal hydrogenases from Methanosarcinales have replaced the leading coordinating cysteine with an aspartate (DSGC). A second, more common modification is the replacement of the second coordinating cysteine by an aspartate (i.e. CAGD, CTGD) or an asparagine (i.e. CEGN, CAGN). For these cases, there is no direct experimental evidence that a [4Fe4S] cluster is still formed. However, aspartate may act via its carboxylic oxygen either as a mono- or a bi-dentate ligand and aspartate ligation indeed occurs in the [4Fe4S] ferredoxin from Pyrococcus furiosus [66] and the oxygen-sensing [4Fe4S] cluster of Bacillus subtilis [67]. Asparagine ligation is rarer, although this residue may also ligate through its side-chain amide oxygen. The sole spectroscopically characterized hydrogenase with a proximal cluster having an asparagine instead of a cysteine, is the cyanobacterial-like enzyme from Acidithiobacillus ferrooxidans [68]. The aerobically isolated A. ferrooxidans enzyme shows signals reminiscent of a proximal [4Fe4S] cluster magnetically

1573

interacting with the [NiFe] site. The fact that the cluster is already paramagnetic (reduced) in the as-purified enzyme suggests a higher reduction potential induced by the putative asparagine ligation. However, these signals were absent in spectra from whole cells, suggesting the possibility of cluster damage during purification [68]. The fact that this hydrogenase shows optimal activity with relatively high-potential electron acceptors indicates that the redox potentials of the iron–sulfur clusters are more positive than those of standard hydrogenases [15,34]. A CEGN motif for the proximal cluster may therefore be correlated with a higher redox potential (similar to P. furiosus ferredoxin) [66]. There are, so far, no biochemical, biophysical or enzymatic data suggesting specific roles for all these variants in Archaea and Bacteria. From the experiences with the 6C cluster, however, we would predict that these deviant ligation patterns are functionally meaningful and bench-characterizations are thus much needed. Group 1 hydrogenases are expected to still have a few more surprises in stock. 4. Conclusions Group 1 hydrogenases are periplasmically oriented, in contrast to members of the other groups, and thus are exposed to the “outside world” with its variable oxidizing substrates and specific environmental stress factors, the most obvious of which certainly being O2. Specific adaptations to these varying conditions rationalize Group 1's extensive diversification into subclades optimizing the enzyme for diverse chemiosmotic chains. To modulate proton-coupled electron-transfer processes and decrease O2-lability, Group 1 enzymes particularly experiment with the ligation of the proximal cluster, the cofactor closest to the catalytic site and at the interface of the two subunits. Their most successful evolutionary invention to this end is the 6C cluster, shown by mutation studies to be crucial for O2-tolerance. Its most notable peculiarity is a second redox transition promoted by redox-dependent structural changes related to its surplus cysteine coordination. The most logical role of the 6C cofactor's third redox state, rather than participating in catalysis, is that of an electron-safety valve helping to efficiently neutralize reactive oxidative species by releasing an additional electron. The 6C cluster correspondingly is widely distributed in hydrogenases from many more recent phyla, demonstrating that this cofactor is not an odd phenomenon but a wide-spread means to cope with increasing oxidative conditions and most likely direct exposure to O2. Acknowledgements We would like to thank Pascale Infossi and Marie-Therese Giudici Orticoni for helpful comments and Ines Pereira for stimulating discussions. This work has been supported by the Max-Planck Society and the Solar-H2 program. Appendix A. Supplementary data Supplementary data to this article can be found online at http:// dx.doi.org/10.1016/j.bbabio.2012.04.012. References [1] N. Lane, J.F. Allen, W. Martin, How did LUCA make a living? Chemiosmosis in the origin of life, Bioessays 32 (2010) 271–280. [2] W. Nitschke, M.J. Russell, Hydrothermal focusing of chemical and chemiosmotic energy, supported by delivery of catalytic Fe, Ni, Mo/W, Co, S and Se, forced life to emerge, J. Mol. Evol. 69 (2009) 481–496. [3] A.L. Ducluzeau, R. van Lis, S. Duval, B. Schoepp-Cothenet, M.J. Russell, W. Nitschke, Was nitric oxide the first deep electron sink? Trends Biochem. Sci. 34 (2009) 9–15. [4] P.M. Vignais, B. Billoud, Occurrence, classification, and biological function of hydrogenases: an overview, Chem. Rev. 107 (2007) 4206–4272. [5] M. Brugna-Guiral, P. Tron, W. Nitschke, K.O. Stetter, B. Burlat, B. Guigliarelli, M. Bruschi, M.T. Giudici-Orticoni, [NiFe] hydrogenases from the hyperthermophilic

1574

[6]

[7] [8]

[9]

[10]

[11]

[12]

[13]

[14]

[15]

[16]

[17]

[18] [19]

[20]

[21]

[22]

[23]

[24]

[25]

[26]

[27]

[28]

[29]

[30]

[31]

M-E. Pandelia et al. / Biochimica et Biophysica Acta 1817 (2012) 1565–1575 bacterium Aquifex aeolicus: properties, function, and phylogenetics, Extremophiles 7 (2003) 145–157. R.K. Thauer, A.K. Kaster, H. Seedorf, W. Buckel, R. Hedderich, Methanogenic archaea: ecologically relevant differences in energy conservation, Nat. Rev. Microbiol. 6 (2008) 579–591. G. Voordouw, Evolution of hydrogenase genes, Adv. Inorg. Chem. 38 (1992) 397–422. M.E. Pandelia, V. Fourmond, P. Tron-Infossi, E. Lojou, P. Bertrand, C. Leger, M.T. Giudici-Orticoni, W. Lubitz, Membrane-Bound hydrogenase I from the hyperthermophilic bacterium Aquifex aeolicus: enzyme activation, redox intermediates and oxygen tolerance, J. Am. Chem. Soc. 132 (2010) 6991–7004. G. Goldet, A.F. Wait, J.A. Cracknell, K.A. Vincent, M. Ludwig, O. Lenz, B. Friedrich, F.A. Armstrong, Hydrogen production under aerobic conditions by membranebound hydrogenases from Ralstonia species, J. Am. Chem. Soc. 130 (2008) 11106–11113. Y. Shomura, K.S. Yoon, H. Nishihara, Y. Higuchi, Structural basis for a [4Fe–3S] cluster in the oxygen-tolerant membrane-bound [NiFe]-hydrogenase, Nature 479 (2011) 253–256. M.J. Lukey, A. Parkin, M.M. Roessler, B.J. Murphy, J. Harmer, T. Palmer, F. Sargent, F.A. Armstrong, How Escherichia coli is equipped to oxidize hydrogen under different redox conditions, J. Biol. Chem. 285 (2010) 3928–3938. A. Volbeda, P. Amara, C. Darnault, J.M. Mouesca, A. Parkin, M.M. Roessler, F.A. Armstrong, J.C. Fontecilla-Camps, X-ray crystallographic and computational studies of the O2-tolerant [NiFe]-hydrogenase 1 from Escherichia coli, Proc. Natl. Acad. Sci. U. S. A. 109 (2012) 5305–5310. A. Volbeda, M.H. Charon, C. Piras, E.C. Hatchikian, M. Frey, J.C. FontecillaCamps, Crystal-structure of the nickel–iron hydrogenase from Desulfovibrio gigas, Nature 373 (1995) 580–587. J. Fritsch, P. Scheerer, S. Frielingsdorf, S. Kroschinsky, B. Friedrich, O. Lenz, C.M.T. Spahn, The crystal structure of an oxygen-tolerant hydrogenase uncovers a novel iron–sulphur centre, Nature 479 (2011) 249–252. M.E. Pandelia, W. Nitschke, P. Infossi, M.T. Giudici-Orticoni, E. Bill, W. Lubitz, Characterization of a unique [FeS] cluster in the electron transfer chain of the oxygen tolerant [NiFe] hydrogenase from Aquifex aeolicus, Proc. Natl. Acad. Sci. U. S. A. 108 (2011) 6097–6102. T. Goris, A.F. Wait, M. Saggu, J. Fritsch, N. Heidary, M. Stein, I. Zebger, F. Lendzian, F.A. Armstrong, B. Friedrich, O. Lenz, A unique iron–sulfur cluster is crucial for oxygen tolerance of a [NiFe]-hydrogenase, Nat. Chem. Biol. 7 (2011) (p. 310-U87). M.J. Lukey, M.M. Roessler, A. Parkin, R.M. Evans, R.A. Davies, O. Lenz, B. Friedrich, F. Sargent, F.A. Armstrong, Oxygen-tolerant [NiFe]-hydrogenases: the individual and collective importance of supernumerary cysteines at the proximal Fe–S cluster, J. Am. Chem. Soc. 133 (2011) 16881–16892. S.F. Altschul, W. Gish, W. Miller, E.W. Myers, D.J. Lipman, Basic local alignment search tool, J. Mol. Biol. 215 (1990) 403–410. K. Tamura, D. Peterson, N. Peterson, G. Stecher, M. Nei, S. Kumar, MEGA5: molecular evolutionary genetics analysis using maximum likelihood, evolutionary distance, and maximum parsimony methods, Mol. Biol. Evol. 28 (2011) 2731–2739. H.M. Berman, J. Westbrook, Z. Feng, G. Gilliland, T.N. Bhat, H. Weissig, I.N. Shindyalov, P.E. Bourne, The protein data bank, Nucleic Acids Res. 28 (2000) 235–242. N.R. Hackett, P.D. Bragg, The association of 2 distinct b-cytochromes with the respiratory nitrate reductase of Escherichia coli, FEMS Microbiol. Lett. 13 (1982) 213–217. A.L. Ballard, S.J. Ferguson, Respiratory nitrate reductase from Paracoccus denitrificans — evidence for 2 b-type hemes in the gamma-subunit and properties of a water-soluble active enzyme containing alpha-subunit and beta-subunit, Eur. J. Biochem. 174 (1988) 207–212. P. Infossi, E. Lojou, J.P. Chauvin, G. Herbette, M. Brugna, M.T. Giudici-Orticoni, Aquifex aeolicus membrane hydrogenase for hydrogen biooxidation: role of lipids and physiological partners in enzyme stability and activity, Int. J. Hydrogen Energy 35 (2010) 10778–10789. R. Gross, J. Simon, F. Theis, A. Kroger, Two membrane anchors of Wolinella succinogenes hydrogenase and their function in fumarate and polysulfide respiration, Arch. Microbiol. 170 (1998) 50–58. M. Jormakka, S. Tornroth, B. Byrne, S. Iwata, Molecular basis of proton motive force generation: structure of formate dehydrogenase-N, Science 295 (2002) 1863–1868. L.S. Palagyi-Meszaros, J. Maroti, D. Latinovics, T. Balogh, E. Klement, K.F. Medzihradszky, G. Rakhely, K.L. Kovacs, Electron-transfer subunits of the NiFe hydrogenases in Thiocapsa roseopersicina BBS, FEBS J. 276 (2009) 164–174. F.D. Ciccarelli, T. Doerks, C. von Mering, C.J. Creevey, B. Snel, P. Bork, Toward automatic reconstruction of a highly resolved tree of life, Science 311 (2006) 1283–1287. F. Baymann, E. Lebrun, M. Brugna, B. Schoepp-Cothenet, M.T. Giudici-Orticoni, W. Nitschke, The redox protein construction kit: pre-last universal common ancestor evolution of energy-conserving enzymes, Philos. Trans. R. Soc. Lond. B Biol. Sci. 358 (2003) 267–274. M. Jormakka, K. Yokoyama, T. Yano, M. Tamakoshi, S. Akimoto, T. Shimamura, P. Curmi, S. Iwata, Molecular mechanism of energy conservation in polysulfide respiration, Nat. Struct. Mol. Biol. 15 (2008) 730–737. A. Dubini, R.L. Pye, R.L. Jack, T. Palmer, F. Sargent, How bacteria get energy from hydrogen: a genetic analysis of periplasmic hydrogen oxidation in Escherichia coli, Int. J. Hydrogen Energy 27 (2002) 1413–1420. N.K. Menon, C.Y. Chatelus, M. Dervartanian, J.C. Wendt, K.T. Shanmugam, H.D. Peck, A.E. Przybyla, Cloning, sequencing, and mutational analysis of the Hyboperon encoding Escherichia coli hydrogenase-2, J. Bacteriol. 176 (1994) 4416–4423.

[32] B. Schoepp-Cothenet, C. Lieutaud, F. Baymann, A. Vermeglio, T. Friedrich, D.M. Kramer, W. Nitschke, Menaquinone as pool quinone in a purple bacterium, Proc. Natl. Acad. Sci. U. S. A. 106 (2009) 8549–8554. [33] T.V. Laurinavichene, A.A. Tsygankov, The involvement of hydrogenases 1 and 2 in the hydrogen-dependent nitrate respiration of Escherichia coli, Microbiology 72 (2003) 654–659. [34] M. Teixeira, I. Moura, A.V. Xavier, J.J.G. Moura, J. LeGall, D.V. Der Vartanian, H.D. Peck, B.H. Huynh, Redox intermediates of Desulfovibrio gigas [NiFe] hydrogenase generated under hydrogen — Mössbauer and EPR characterization of the metal centers, J. Biol. Chem. 264 (1989) 16435–16450. [35] H. Miyadera, A. Hiraishi, H. Miyoshi, K. Sakamoto, R. Mineki, K. Murayama, K.V.P. Nagashima, K. Matsuura, S. Kojima, K. Kita, Complex II from phototrophic purple bacterium Rhodoferax fermentans displays rhodoquinol-fumarate reductase activity, Eur. J. Biochem. 270 (2003) 1863–1874. [36] B.C. Brajcich, A.L. Iarocci, L.A.G. Johnstone, R.K. Morgan, Z.T. Lonjers, M.J. Hotchko, J.D. Muhs, A. Kieffer, B.J. Reynolds, S.M. Mandel, B.N. Marbois, C.F. Clarke, J.N. Shepherd, Evidence that ubiquinone is a required intermediate for rhodoquinone biosynthesis in Rhodospirillum rubrum, J. Bacteriol. 192 (2010) 436–445. [37] G.J. Mander, E.C. Duin, D. Linder, K.O. Stetter, R. Hedderich, Purification and characterization of a membrane-bound enzyme complex from the sulfate-reducing archaeon Archaeoglobus fulgidus related to heterodisulfide reductase from methanogenic archaea, Eur. J. Biochem. 269 (2002) 1895–1904. [38] F. Grein, I.A.C. Pereira, C. Dahl, Biochemical characterization of individual components of the Allochromatium vinosum DsrMKJOP transmembrane complex aids understanding of complex function in vivo, J. Bacteriol. 192 (2010) 6369–6377. [39] M.G. Bertero, R.A. Rothery, M. Palak, C. Hou, D. Lim, F. Blasco, J.H. Weiner, N.C.J. Strynadka, Insights into the respiratory electron transfer pathway from the structure of nitrate reductase A, Nat. Struct. Biol. 10 (2003) 681–687. [40] R. Hedderich, N. Hamann, M. Bennati, Heterodisulfide reductase from methanogenic archaea: a new catalytic role for an iron–sulfur cluster, Biol. Chem. 386 (2005) 961–970. [41] R.H. Pires, S.S. Venceslau, F. Morais, M. Teixeira, A.V. Xavier, I.A.C. Pereira, Characterization of the Desulfovibrio desulfuricans ATCC 27774 DsrMKJOP complex — a membrane-bound redox complex involved in the sulfate respiratory pathway, Biochemistry 45 (2006) 249–262. [42] N. Hamann, G.J. Mander, J.E. Shokes, R.A. Scott, M. Bennati, R. Hedderich, Cysteine-rich CCG domain contains a novel [4Fe–4S] cluster binding motif as deduced from studies with subunit B of heterodisulfide reductase from Methanothermobacter marburgensis, Biochemistry 46 (2007) 12875–12885. [43] J.M.C.C. Coremans, C.J. Vangarderen, S.P.J. Albracht, On the redox equilibrium between H-2 and hydrogenase, Biochim. Biophys. Acta 1119 (1992) 148–156. [44] H. Ogata, P. Kellers, W. Lubitz, The crystal structure of the [NiFe] hydrogenase from the photosynthetic bacterium Allochromatium vinosum: characterization of the oxidized enzyme (Ni-A state), J. Mol. Biol. 402 (2010) 428–444. [45] B. Bleijlevens, F.A. van Broekhuizen, A.L. De Lacey, W. Roseboom, V.M. Fernandez, S.P.J. Albracht, The activation of the [NiFe]-hydrogenase from Allochromatium vinosum. An infrared spectro-electrochemical study, J. Biol. Inorg. Chem. 9 (2004) 743–752. [46] K. Knüttel, K. Schneider, A. Erkens, W. Plass, A. Muller, E. Bill, A.X. Trautwein, Redox properties of the metal centers in the membrane-bound hydrogenase from Alcaligenes eutrophus CH34, Bull. Pol. Acad. Sci., Chem. 42 (1994) 495–511. [47] M. Tietze, A. Beuchle, I. Lamla, N. Orth, M. Dehler, G. Greiner, U. Beifuss, Redox potentials of methanophenazine and CoB-S-S-CoM, factors involved in electron transport in methanogenic archaea, Chem. Biochem. 4 (2003) 333–335. [48] R. Gross, R. Pisa, M. Sanger, C.R.D. Lancaster, J. Simon, Characterization of the menaquinone reduction site in the diheme cytochrome b membrane anchor of Wolinella succinogenes NiFe-hydrogenase, J. Biol. Chem. 279 (2004) 274–281. [49] U. Beifuss, M. Tietze, S. Baumer, U. Deppenmeier, Methanophenazine: structure, total synthesis, and function of a new cofactor from methanogenic archaea, Angew. Chem. Int. Ed. 39 (2000) 2470. [50] W.A. Vargas, P.D. Weyman, Y.K. Tong, H.O. Smith, Q. Xu, [NiFe] Hydrogenase from Alteromonas macleodii with unusual stability in the presence of oxygen and high temperature, Appl. Environ. Microbiol. 77 (2011) 1990–1998. [51] K. Schneider, D.S. Patil, R. Cammack, Electron-spin-resonance properties of membrane-bound hydrogenases from aerobic hydrogen bacteria, Biochim. Biophys. Acta 748 (1983) 353–361. [52] K. Knüttel, K. Schneider, H.G. Schlegel, A. Muller, The membrane-bound hydrogenase from Paracoccus denitrificans — purification and molecular characterization, Eur. J. Biochem. 179 (1989) 101–108. [53] R. Cammack, V.M. Fernandez, E.C. Hatchikian, Nickel–iron hydrogenase, Inorg. Microb. Sulfur Metabol. 243 (1994) 43–68. [54] J.H. Sun, D.J. Arp, Aerobically purified hydrogenase from Azotobacter vinelandii — activity, activation, and spectral properties, Arch. Biochem. Biophys. 287 (1991) 225–233. [55] K.S. Yoon, N. Tsukada, Y. Sakai, M. Ishii, Y. Igarashi, H. Nishihara, Isolation and characterization of a new facultatively autotrophic hydrogen-oxidizing betaproteobacterium, Hydrogenophaga sp AH-24, FEMS Microbiol. Lett. 278 (2008) 94–100. [56] K.S. Yoon, K. Fukuda, K. Fujisawa, H. Nishihara, Purification and characterization of a highly thermostable, oxygen-resistant, respiratory [NiFe]-hydrogenase from a marine, aerobic hydrogen-oxidizing bacterium Hydrogenovibrio marinus, Int. J. Hydrogen Energy 36 (2011) 7081–7088. [57] M. Saggu, I. Zebger, M. Ludwig, O. Lenz, B. Friedrich, P. Hildebrandt, F. Lendzian, Spectroscopic insights into the oxygen-tolerant membrane-associated [NiFe] hydrogenase of Ralstonia eutropha H16, J. Biol. Chem. 284 (2009) 16264–16276.

M-E. Pandelia et al. / Biochimica et Biophysica Acta 1817 (2012) 1565–1575 [58] B. Boussau, L. Gueguen, M. Gouy, Accounting for horizontal gene transfers explains conflicting hypotheses regarding the position of aquificales in the phylogeny of bacteria, BMC Evol. Biol. 8 (2008) 272. [59] W. Nitschke, R. van Lis, B. Schoepp-Cothenet, F. Baymann, The “green” phylogenetic clade of Rieske/cytb complexes, Photosynth. Res. 104 (2010) 347–355. [60] T.V. Laurinavichene, A.A. Tsygankov, H-2 consumption by Escherichia coli coupled via hydrogenase 1 or hydrogenase 2 to different terminal electron acceptors, FEMS Microbiol. Lett. 202 (2001) 121–124. [61] E.M. Shepard, E.S. Boyd, J.B. Broderick, J.W. Peters, Biosynthesis of complex iron– sulfur enzymes, Curr. Opin. Chem. Biol. 15 (2011) 319–327. [62] M. Fontecave, S. Ollagnier-de-Choudens, Iron–sulfur cluster biosynthesis in bacteria: mechanisms of cluster assembly and transfer, Arch. Biochem. Biophys. 474 (2008) 226–237. [63] J. Fritsch, O. Lenz, B. Friedrich, The maturation factors HoxR and HoxT contribute to oxygen tolerance of membrane-bound [NiFe] hydrogenase in Ralstonia eutropha H16, J. Bacteriol. 193 (2011) 2487–2497.

1575

[64] B.M. Wolfe, S.M. Lui, H. Manyani, L. Rey, J.M. Palacios, J. Imperial, T. Ruiz-Argueso, Gene products of the hupGHIJ Operon are involved in maturation of the iron– sulfur subunit of the [NiFe] hydrogenase from Rhizobium leguminosarum bv. viciae, J. Bacteriol. 187 (2005) 7018–7026. [65] L.A. Sazanov, P. Hinchliffe, Structure of the hydrophilic domain of respiratory complex I from Thermus thermophilus, Science 311 (2006) 1430–1436. [66] S. Sham, L. Calzolai, P.L. Wang, K. Bren, H. Haarklau, P.S. Brereton, M.W.W. Adams, G.N. La Mar, A solution NMR molecular model for the aspartate-ligated, cubane cluster containing ferredoxin from the hyperthermophilic archeaon Pyrococcus furiosus, Biochemistry 41 (2002) 12498–12508. [67] I. Gruner, C. Fradrich, L.H. Bottger, A.X. Trautwein, D. Jahn, E. Hartig, Aspartate 141 is the fourth ligand of the oxygen-sensing [4Fe–4S](2 +) cluster of Bacillus subtilis transcriptional regulator Fnr, J. Biol. Chem. 286 (2011) 2017–2021. [68] O. Schröder, B. Bleijlevens, T.E. de Jongh, Z.J. Chen, T.S. Li, J. Fischer, J. Forster, C.G. Friedrich, K.A. Bagley, S.P.J. Albracht, W. Lubitz, Characterization of a cyanobacteriallike uptake [NiFe] hydrogenase: EPR and FTIR spectroscopic studies of the enzyme from Acidithiobacillus ferrooxidans, J. Biol. Inorg. Chem. 12 (2007) 212–233.