Genomic and metagenomic surveys of hydrogenase distribution indicate H₂ is a widely utilised energy source for microbial growth and survival

Abstract

Recent physiological and ecological studies have challenged the long-held belief that microbial metabolism of molecular hydrogen (H₂) is a niche process. To gain a broader insight into the importance of microbial H₂ metabolism, we comprehensively surveyed the genomic and metagenomic distribution of hydrogenases, the reversible enzymes that catalyse the oxidation and evolution of H₂. The protein sequences of 3286 non-redundant putative hydrogenases were curated from publicly available databases. These metalloenzymes were classified into multiple groups based on (1) amino acid sequence phylogeny, (2) metal-binding motifs, (3) predicted genetic organisation and (4) reported biochemical characteristics. Four groups (22 subgroups) of [NiFe]-hydrogenase, three groups (6 subtypes) of [FeFe]-hydrogenases and a small group of [Fe]-hydrogenases were identified. We predict that this hydrogenase diversity supports H₂-based respiration, fermentation and carbon fixation processes in both oxic and anoxic environments, in addition to various H₂-sensing, electron-bifurcation and energy-conversion mechanisms. Hydrogenase-encoding genes were identified in 51 bacterial and archaeal phyla, suggesting strong pressure for both vertical and lateral acquisition. Furthermore, hydrogenase genes could be recovered from diverse terrestrial, aquatic and host-associated metagenomes in varying proportions, indicating a broad ecological distribution and utilisation. Oxygen content (pO₂) appears to be a central factor driving the phylum- and ecosystem-level distribution of these genes. In addition to compounding evidence that H₂ was the first electron donor for life, our analysis suggests that the great diversification of hydrogenases has enabled H₂ metabolism to sustain the growth or survival of microorganisms in a wide range of ecosystems to the present day. This work also provides a comprehensive expanded system for classifying hydrogenases and identifies new prospects for investigating H₂ metabolism.

Introduction

Molecular hydrogen (H₂) has several physical properties desirable for biological systems, notably its redox potential (E°'=−0.42 V) and diffusion coefficient (4 × 10⁻⁹ m² s⁻¹). Microorganisms are able to harness these properties by consuming and producing H₂ using specialised metalloenzymes called hydrogenases (Schwartz et al., 2013). There are three phylogenetically unrelated classes of hydrogenase distinguishable based on the metal content of their H₂-binding sites: the [NiFe]-, [FeFe]- and [Fe]-hydrogenases (Volbeda et al., 1995; Peters et al., 1998; Shima et al., 2008). H₂ oxidation by such enzymes yields low-potential electrons that are transduced through respiratory chains or used to fix inorganic carbon. In contrast, H₂ evolution efficiently dissipates excess reductant as a diffusible gas during microbial fermentation and photobiological processes (Schwartz et al., 2013). Certain hydrogenases are also part of low-potential ion-translocating complexes that use protons as terminal electron acceptors (Buckel and Thauer, 2013). Since the discovery of microbial H₂ oxidation in the 1900s (Kaserer, 1906; Stephenson and Stickland, 1931), H₂ metabolism has been observed in multiple bacterial, archaeal and eukaryotic phyla. It is increasingly recognised that H₂ metabolism is important for a wide range of microorganisms: lithotrophs and phototrophs, respirers and fermenters and aerobes and anaerobes alike (Vignais and Billoud, 2007; Schwartz et al., 2013; Peters et al., 2014). Furthermore, it is widely hypothesised that H₂ was the primordial electron donor, suggesting early and sustained evolutionary importance (Lane et al., 2010).

Several recent studies demonstrated that microbial H₂ metabolism is more widespread than previously reported. It was recently shown that some aerobic soil actinobacteria and acidobacteria persist by scavenging H₂ from the lower atmosphere (Constant et al., 2010; Greening et al., 2014, 2015a, b), overturning long-held beliefs that hydrogen metabolism is restricted to low O₂, high H₂ environments and highlighting the importance of H₂ for survival in addition to growth (Greening and Cook, 2014). Biochemists have simultaneously elucidated mechanisms dependent on reversed electron flow that enable certain hydrogenases to function in the presence of the oxygen (traditionally an inhibitor of their active sites) (Fritsch et al., 2011; Shomura et al., 2011; Horch et al., 2015). In anaerobic systems, ultra-minimalistic hydrogenase-containing respiratory chains have been described that efficiently generate energy within oligotrophic environments (Kim et al., 2010; Lim et al., 2014). In parallel, the discovery of electron bifurcation has expanded our understanding of how energy is conserved in anaerobic processes such as cellulolytic fermentation, acetogenesis and methanogenesis (Schut and Adams, 2009; Kaster et al., 2011; Buckel and Thauer, 2013; Schuchmann and Muller, 2014). Other themes, including H₂ sensing within anaerobes (Zheng et al., 2014) and H₂ fermentation in aerobes (Berney et al., 2014), are emerging.

Despite this progress, there remains much to be discovered about microbial H₂ metabolism on both the microscopic and macroscopic levels. Most studies on microbial H₂ metabolism focus on only a few branches of the hydrogenase phylogenetic tree and a small subset of organisms within the universal tree of life. Physiological and biochemical characterisations have focussed on model organisms from within five phyla, Proteobacteria, Firmicutes, Cyanobacteria, Euryarchaeota and Chlorophyta (Schwartz et al., 2013; Lubitz et al., 2014). Furthermore, detailed biochemical information and atomic-resolution structures are available for only a subset of hydrogenases (Volbeda et al., 1995; Peters et al., 1998; Shima et al., 2008; Fritsch et al., 2011; Mills et al., 2013). Although the contribution of H₂ metabolism to total ecosystem processes is recognised in some environments (for example, anoxic sediments, animal guts and hydrothermal vents; Vignais and Billoud, 2007; Schwartz et al., 2013), the role of hydrogenases in general soil and aquatic ecosystems remains largely unresolved (Barz et al., 2010; Constant et al., 2011; Beimgraben et al., 2014; Greening et al., 2015b). Consequently, the influence of H₂ evolution and consumption on community structuring and global biogeochemical cycling requires further investigation (Schwartz et al., 2013; Greening et al., 2015b).

Hydrogenase gene surveys are vital for understanding microbial H₂ metabolism at the global scale. Current knowledge on the evolution and diversity of hydrogenases relies heavily on the progressive surveys conducted by Wu and Vignais (Wu and Mandrand, 1993; Vignais et al., 2001; Vignais and Billoud, 2007); these studies revealed that the primary sequences and subunit architectures of [NiFe]- and [FeFe]-hydrogenases have diversified to enable them to adopt a wide range of physiological roles (whereas the [Fe]-hydrogenase is constrained to a single function). In the eight years following these studies (Vignais and Billoud, 2007), the emergence of sequencing technologies has resulted in the rapid expansion of genome and metagenome sequence data. Genomes are now available for a far greater range of organisms, spanning model laboratory specimens, representatives of dominant environmental phyla, and poorly described ‘Microbial Dark Matter' (Wu et al., 2009; Rinke et al., 2013). Furthermore, metagenomes enable the metabolic capability of entire communities to be described in silico (Tringe et al., 2005; Morales and Holben, 2011; Wrighton et al., 2012). In this work, we used publicly available genome and metagenome resources to comprehensively analyse the distribution of hydrogenases. Our findings suggest that H₂ metabolism is more diverse and widespread on both the taxonomic and community levels than previously reported.

Materials and methods

Hydrogenase sequence retrieval

Amino acid sequences of all non-redundant putative hydrogenase catalytic subunits represented in the National Center for Biotechnology Information (NCBI) Reference Sequence (RefSeq) (Pruitt et al., 2007) and Joint Genome Institute (JGI) Microbial Dark Matter (MDM) (Rinke et al., 2013) databases were retrieved by Protein BLAST (Altschul et al., 1990) during August 2014. The retrieved sequences were verified as hydrogenase encoding through screening for the presence of conserved cysteine residues required to ligate H₂-binding metal centres (L1 and L2 motifs for [NiFe]-hydrogenases (Vignais and Billoud, 2007); P1, P2 and P3 motifs for [FeFe]-hydrogenases (Vignais and Billoud, 2007); and Cys176 in [Fe]-hydrogenases (Shima et al., 2008)). The analysis omitted protein families homologous to [NiFe]-hydrogenases (Ehr/Mbx, NuoD), [FeFe]-hydrogenases (Narf/Nar1p) and [Fe]-hydrogenases (HmdII) that appear to lack the capacity to metabolise H₂. Our analysis did not include nitrogenases, alkaline phosphatases, formate dehydrogenases and carbon monoxide dehydrogenases that have been shown to catalyse side-reactions resulting in H₂ oxidation or evolution (Schwartz et al., 2013).

Hydrogenase classification and analysis

Protein sequences encoding the catalytic subunits ([NiFe]-hydrogenases, [Fe]-hydrogenases) or catalytic domains ([FeFe]-hydrogenases) of hydrogenases were aligned using the ClustalW (Larkin et al., 2007) and MUSCLE (Edgar, 2004) algorithms. Evolutionary relationships were analysed using neighbour-joining phylogenetic trees (Saitou and Nei, 1987) constructed with MEGA6 (Tamura et al., 2013). All trees were bootstrapped using 500 replicates and were rooted with ancestral sequences where available. The robustness of the analysis was confirmed by varying the number of ingroup sequences tested and the nature of the outgroup sequence used. The Microbial Genomic Context Viewer (MGcV) was used to compare genome regions encoding homologous hydrogenases (Overmars et al., 2013). Domains were predicted by searching the Conserved Domain Database (CDD) (Marchler-Bauer et al., 2011) and using multiple sequence alignments to identify signature conserved residues. WebLogo (Crooks et al., 2004) was used to visualise conserved metal-binding motifs forming the active sites and redox centres. Using a combination of the information derived from these methods, the [NiFe]- and [FeFe]-hydrogenases were further divided into phylogenetically distinct groups and subgroups.

Metagenome analysis

Metagenome sequence libraries derived from 10 ecosystems including soil (farmland, forest, permafrost, bog), gut (termite, human) and water (fresh water, hot spring, coastal upwelling, deep ocean) environments were identified from publicly available databases. For each ecosystem, two libraries were selected. All selected libraries were sequenced with paired-end reads on an Illumina (San Diego, CA, USA) platform and represented read sizes of between 201 and 280 nucleotides. BLAST (Altschul et al., 1990) analyses were performed using a local BLAST database containing the protein sequences of the catalytic subunit ([NiFe]-hydrogenases, [Fe]-hydrogenases) or catalytic domain ([FeFe]-hydrogenases) of all sequenced hydrogenases (curated as described above). Low complexity regions for all reference sequences were masked using the SEG algorithm (Wootton and Federhen, 1996) of BLAST+ (Camacho et al., 2008) and a reference BLAST database was created. All metagenome libraries were randomly subsampled to an equal depth (1 million reads) and read length >201 nucleotides before analyses. A translated BLAST screening of all subsamples was performed using blastx (word size 3 and e-value 10). To minimise false positives, hits within the initial screen were sieved by removing any result with a minimum percentage identity 60% and minimum query coverage 40 amino acids. Identified reads for each class are recorded as relative percentage abundance.

Results

An expanded hydrogenase classification scheme predictive of biological function

The first aim of this work was to identify and classify all putative hydrogenases represented in public databases. Initially, we retrieved non-redundant sequences encoding the catalytic subunits of all hydrogenases in the NCBI database and verified that 3286 of them contained the sufficient residues required to bind their metal centres (Supplementary Tables S1 and S2). In order to develop the classification scheme (Table 1), we correlated the phylogenetic clustering of the hydrogenases with functional information and predictors. For all hydrogenases, we analysed: (1) primary phylogeny to determine their evolutionary relationships (Figure 1), (2) metal- and cofactor-binding motifs to predict redox centres (Table 2), (3) genetic organisation to identify probable partner proteins (Figure 2) and (4) previous literature reports to probe biochemical characteristics and physiological roles (Table 1). Integrating this information, we were able to classify hydrogenases into multiple groups and subgroups/subtypes likely to have distinct cellular functions.

Table 1. New scheme for the classification of hydrogenases.

Group	Proposed function	References	PDB
Group 1: membrane-bound H2-uptake [NiFe]-hydrogenases
Group 1a: ancestral	Liberates electrons for sulphate, metal, organohalide and methanogenic heterodisulphide respiration. Includes [NiFeSe] variants.	Desulfovibrio vulgaris (Marques et al., 2010) Desulfomicrobium baculatum (Garcin et al., 1999) Methanosarcina mazei (Deppenmeier and Blaut, 1995)	2WPN 1CC1
Group 1b: prototypical	Liberates electrons for sulphate, fumarate and nitrate respiration.	Desulfovibrio gigas (Volbeda et al., 1995) Wolinella succinogenes (Gross et al., 1998) Helicobacter pylori (Olson and Maier, 2002)	1FRV
Group 1c: Hyb type	Liberates electrons primarily for fumarate respiration. Possibly bidirectional.	Escherichia coli (Lukey et al., 2010) Salmonella enterica (Maier et al., 2013)
Group 1d: oxygen tolerant	Electron input for aerobic respiration and oxygen-tolerant anaerobic respiration.	Ralstonia eutropha (Fritsch et al., 2011) Escherichia coli (Volbeda et al., 2013) Aquifex aeolicus (Brugna-Guiral et al., 2003)	3RGW 4GD3
Group 1e: Isp type	Liberates electrons primarily for sulphur respiration. Possibly bidirectional.	Allochromatium vinosum (Ogata et al., 2010) Aquifex aeolicus (Brugna-Guiral et al., 2003) Thiocapsa roseopersicina (Tengölics et al., 2014)	3MYR
Group 1f: oxygen protecting	Unresolved. May liberate electrons to reduce reactive oxygen species.	Geobacter sulfurreducens (Tremblay and Lovley, 2012) Frankia sp. (Leul et al., 2007)
Group 1g: Crenarchaeota type	Unresolved. May liberate electrons primarily for sulphur respiration.	Pyrodictium brockii* (Pihl et al., 1989) Acidianus ambivalens* (Laska et al., 2003)
Group 1h/5: Actinobacteria type	Scavenges electrons from tropospheric H2 to sustain aerobic respiration during starvation.	Ralstonia eutropha (Schäfer et al., 2013) Mycobacterium smegmatis (Greening et al., 2014) Streptomyces avermitilis (Constant et al., 2010)	5AA5
Group 2: Cytosolic H2-uptake [NiFe]-hydrogenases
Group 2a: Cyanobacteria type	Electron input for aerobic respiration and recycling H2 produced by cellular processes (for example, nitrogenase, fermentation).	Anabaena sp. (Houchins and Burris, 1981) Mycobacterium smegmatis (Greening et al., 2014) Nitrospira moscoviensis* (Koch et al., 2014)
Group 2b: HK linked	Senses H2 and activates two-component cascade controlling hydrogenase expression.	Ralstonia eutropha (Lenz and Friedrich, 1998) Rhodobacter capsulatus (Vignais et al., 2005)
Group 2c: DGC linked (putative)	Unknown. Predicted to sense H2 and induce cyclic di-GMP production.	Uncharacterised.
Group 2d: Aquificae type	Unknown. May generate reductant for carbon fixation or have a regulatory role.	Aquifex aeolicus (Brugna-Guiral et al., 2003)
Group 3: Cytosolic bidirectional [NiFe]-hydrogenases
Group 3a: F420 coupled	Directly couples oxidation of H2 to reduction of F420 during methanogenesis. Reverse reaction may also occur. Includes [NiFeSe] variants.	Methanothermobacter marburgensis (Mills et al., 2013) Methanosarcina barkeri (Kulkarni et al., 2009)	4OMF
Group 3b: NADP coupled	Directly couples oxidation of NADPH to evolution of H2. May be reversible. Some complexes are proposed to have sulfhydrogenase activity.	Pyrococcus furiosus (Ma et al., 1993) Thermococcus kodakarensis (Kanai et al., 2011) Mycobacterium smegmatis (Berney et al., 2014)
Group 3c: HDR linked	Bifurcates electrons from H2 to heterodisulphide and ferredoxin in methanogens without cytochromes.	Methanothermobacter marburgensis (Kaster et al., 2011)
Group 3d: NAD coupled	Directly interconverts electrons between H2 and NAD depending on redox state.	Anabaena sp. (Houchins and Burris, 1981) Ralstonia eutropha (Burgdorf et al., 2005) Thiocapsa roseopersicina (Rákhely et al., 2004)
Group 4: Membrane-bound H2-evolving [NiFe]-hydrogenases
Group 4a: formate hydrogenlyases	Couples oxidation of formate to fermentative evolution of H2. Hyf-type complexes may translocate protons via antiporter modules.	Escherichia coli (McDowall et al., 2014; Andrews et al., 1997) Salmonella enterica (Sawers et al., 1986)
Group 4b: Mrp linked	Couples oxidation of formate or carbon monoxide to proton reduction. Generates sodium-motive force via Mrp antiporter modules.	Pyrococcus furiosus (Sapra et al., 2003) Thermococcus onnurineus (Lim et al., 2014) Thermococcus kodakarensis (Kanai et al., 2011)
Group 4c: CODH linked	Forms complex with carbon monoxide dehydrogenase to anaerobically respire CO using protons as terminal electron acceptors.	Carboxydothermus hydrogenoformans (Soboh et al., 2002) Rhodospirillum rubrum (Fox et al., 1996)
Group 4d: Eha/Ehb type	Multimeric complexes that couple H2 oxidation to ferredoxin reduction for anaplerotic (Eha) and anabolic (Ehb) purposes. H+/Na+ driven.	Methanococcus maripaludis (Lie et al., 2012; Porat et al., 2006)
Group 4e: Ech type	Couples ferredoxin oxidation to H2 evolution. This process is physiologically reversible via H+/Na+ translocation.	Methanosarcina barkeri (Meuer and Kuettner, 2002) Desulfovibrio gigas (Morais-Silva et al., 2013) Thermoanaerobacter tengcongensis (Soboh et al., 2004)
Group 4f: Ehf type (putative)	Unknown. May couple oxidation of a one-carbon compound to proton reduction concurrent with proton translocation. Related to Ehr complexes.	Uncharacterised.
[FeFe]-hydrogenases
Group A1: prototypical	Couples oxidation of ferredoxin to fermentative or photobiological evolution of H2.	Clostridium pasteurianum (Peters et al., 1998) Desulfovibrio desulfuricans (Nicolet et al., 1999) Chlamydomonas reinhardtii (Happe and Naber, 1993)	1FEH 1HFE 3LX4
Group A2: glutamate synthase linked (putative)	Unknown. Predicted to transfer electrons from H2 to NAD, generating reducing equivalents for glutamate synthase.	Uncharacterised.
Group A3: bifurcating	Reversibly bifurcates electrons from H2 to ferredoxin and NAD in anaerobic bacteria.	Thermotoga maritima (Schut and Adams, 2009) Acetobacterium woodii (Schuchmann and Müller, 2012) Moorella thermoacetica (Wang et al., 2013)
Group A4: formate dehydrogenase linked	Couples formate oxidation to evolution of H2. Some can also bifurcate electrons from H2 to ferredoxin and NADP.	Clostridium autoethanogenum (Wang et al., 2013)
Group B: ancestral (putative)	Unknown. May couple oxidation of ferredoxin to fermentative evolution of H2.	Uncharacterised.
Group C: sensory (putative)	Unknown. Predicted to sense hydrogen and induce cascades via co-transcribed regulatory elements, for example, kinases and phosphatases.	Thermoanaerobacterium saccharolyticum (Shaw et al., 2009) Ruminococcus albus (Zheng et al., 2014)
[Fe]-hydrogenases
HmdI: methenyl-H4MPT dehydrogenase	Couples oxidation of H2 to reduction of 5,10-methenyltetrahydromethanopterin in methanogens. Physiologically-reversible and important during nickel limitation.	Methanocaldococcus jannaschii (Shima et al., 2008) Methanothermobacter thermoautotrophicum (Afting et al., 1998)	3DAG

This scheme correlates the phylogenetic clustering of the hydrogenases with their probable functions. The groups A, B and C [FeFe]-hydrogenases and several subgroups of the [NiFe]-hydrogenases (2a, 2b, 3a, 3b, 3c, 3d) were previously defined (Vignais et al., 2001; Vignais and Billoud, 2007; Calusinska et al., 2010). The group 1h [NiFe]-hydrogenases have also been defined as the group 5 [NiFe]-hydrogenases; however, this work shows they are they are descended from other group 1 [NiFe]-hydrogenases. References are provided to structural, biochemical and physiological characterisations of representative hydrogenases from each subgroup/subtype. Structural characterisations are listed first and Protein Database (PDB) structures are provided (see Lubitz et al., 2014 for a full list of solved hydrogenase structures). Asterisked organisms have yet to be sequenced, hence their [NiFe]-hydrogenases are not represented on the phylogenetic trees.

Figure 1.

Classification and phylogeny of hydrogenases. These neighbour-joining skeleton trees show the phylogenetic relationships of all 3286 hydrogenases identified in this work. The trees are colour coded by [NiFe]-hydrogenase subgroup and [FeFe]-hydrogenase group. The nodes separating the major clades are encircled and coloured according to their bootstrap values, that is, black circles for well-supported nodes (bootstrap values >0.75) and red circles for unsupported nodes (bootstrap values <0.75). Group A [FeFe]-hydrogenases cannot be reliably subdivided phylogenetically and can only be classified into subtypes based on their genetic organisation. The expanded trees, including taxon names and bootstrap values, are shown in Supplementary Figures S1 to S6.

Table 2. Metal centres of hydrogenases.

[NiFe]	L1 motif	L2 motif	[FeS]proximal	[FeS]medial	[FeS]distal	Extension
Group 1a	xxRICGVCPxxH	SFDPCxxCxxH*	4Cys[4Fe4S]	4Cys[4Fe4S]	3Cys1His/Asp[4Fe4S]	Y
Group 1b	xQRxCGVCTxxH	xxDPCxACxVH	4Cys[4Fe4S]/3Cys1Asn[4Fe4S]	3Cys[3Fe4S]	3Cys1His[4Fe4S]	Y
Group 1c	xQRICGVCTTVH	SFDPCxxCAVH	4Cys[4Fe4S]	3Cys[3Fe4S]	3Cys1His[4Fe4S]	Y
Group 1d	xxRICGVCTxxH	SFDPCLACxxH	6Cys[4Fe3S]	3Cys[3Fe4S]	3Cys1His[4Fe4S]	Y
Group 1e	xxRICGVCTxVH	SFDPCxACxxH	4Cys[4Fe4S]	3Cys[3Fe4S]	3Cys1His[4Fe4S]	Y
Group 1f	xQRxCGVCTxVH	SFDPCxACxVH	4Cys[4Fe4S]/3Cys1Asp/Asn[4Fe4S]	3Cys[3Fe4S]	3Cys1His[4Fe4S]	Y
Group 1g	xSRxCGVCGxxH	SFDPCxxCxVH	3Cys1Asp/Asn[4Fe4S]	3Cys[3Fe4S]	3Cys1His/Arg[4Fe4S]	Y
Group 1h	TSRICGICGDNH	SFDPCLPCGVH	3Cys1Asp[4Fe4S]	4Cys[4Fe4S]	3Cys1His[4Fe4S]	Y
Group 2a	xxRICGICGxxH	SFDxCLVCTVH	3Cys1Asn[4Fe4S]	3Cys[3Fe4S]	3Cys1Glu[4Fe4S]	Y
Group 2b	xPRICGICSxSQ	SFDPCMVCTVH	4Cys[4Fe4S]	4Cys[4Fe4S]	3Cys1His[4Fe4S]	N
Group 2c	xxRxCGICxxxH	SxDPCxxCTVH	3Cys1Asp/Glu[4Fe4S]	3Cys[3Fe4S]	3Cys1His[4Fe4S]	Y
Group 2d	xPRxCGICGxAH	xFDxCSVCTTH	3Cys1Asn[4Fe4S]	3Cys[3Fe4S]	3Cys1His[4Fe4S]	N
Group 3a	xxRxCGxCxxxH	xYDxCxSCATH*	4Cys[4Fe4S]/3Cys1Asp[4Fe4S]	4Cys[4Fe4S]	4Cys[4Fe4S]	Y
Group 3b	xxRICxxCxxxx	xxDPCISCxxH	4Cys[4Fe4S]	4Cys[4Fe4S]	4Cys[4Fe4S]	Y
Group 3c	xxxICGxCxxxH	AYDPCxxCATH*	4Cys[4Fe4S]	4Cys[4Fe4S]	4Cys[4Fe4S]	Y
Group 3d	xxRxCGICPVSH	xxDPCLSCxTH	4Cys[4Fe4S]	Absent	Absent	Y
Group 4a	xxRVCGICGxxH	SLDPCYSCTDR	4Cys[4Fe4S]	Absent	Absent	Y
Group 4b	xERICGICxxxH	SIDPCxSCTxR	4Cys[4Fe4S]	Absent	Absent	Y
Group 4c	xExxCxLCSNxH	SIDPCISCxER	4Cys[4Fe4S]	Absent	Absent	N
Group 4d	xExxCGICSxxH	xxDPCxxCxxR	4Cys[4Fe4S]	Absent	Absent	Y
Group 4e	xxRxCGICSxxH	xIDPCIxCxER	4Cys[4Fe4S]	Absent	Absent	N
Group 4g	xERVCGVCxxSH	SFELCYxCxDR	4Cys[4Fe4S]	Absent	Absent	N
[FeFe]	P1 motif	P2 motif	P3 motif	Domains
Group A1	xTSCCPxWV	xxxPCxxKK	ExMxCxxGCxxGG	M1, M2 or M3
Group A2	FTSCCPxWx	CxMPCxAKK	ExMACPGGCxxGG	M3
Group A3	xTSCC/SPxW	xxMPCxAKK	ExMxCxGGCxxGG	M2, M3 or M4
Group A4	FTSCCPxWV	xxMPCTCKx	EVMxCPxGCxxGG	M2 or M3
Group B	xTSC[C]CPxxx	FxGPCxAKK	ExMxCxGGCxxGP	M2 or M3
Group C	IxxxCPxxx	FxxPCxxKx	ExxxCxxGCxxGP	M2P
[Fe]	Fe motif
HmdI	xTHACTIPT

Abbreviations: N, no; Y, yes.

The consensus motifs surrounding the metal-ligating cysteine residues are shown for each hydrogenase type: L1 and L2 motifs for [NiFe]-hydrogenases (Vignais and Billoud, 2007); P1, P2 and P3 motifs for [FeFe]-hydrogenases (Vignais and Billoud, 2007); and the Cys176-based binding motif in [Fe]-hydrogenases (Shima et al., 2008). For [NiFe]-hydrogenases, the number, configuration and ligands of the iron–sulphur clusters in the small subunit are listed (the consensus metal-binding motifs for these clusters are listed in Supplementary Table S5). ‘Extension' refers to whether the C-terminus of the [NiFe]-hydrogenase large subunit is predicted to be cleaved by an endopeptidase during maturation. For [FeFe]-hydrogenases, the domain structures of the catalytic subunit are shown. Building on existing schemes (Vignais and Billoud, 2007; Calusinska et al., 2010), we defined the domain organisation of the catalytic subunit as M1 (H-cluster only), M2 (H-cluster, two FeS clusters), M3 (H-cluster, four FeS clusters), M4 (H-cluster, five FeS clusters) or M2P (H-cluster, PAS domain, two FeS clusters) (see also Figure 2 and Supplementary Table S1).

Figure 2.

Genetic organisation of hydrogenases. The genes surrounding the catalytic subunit of representatives of each subtype/subclass are shown to-scale. Genes/domains are colour coded as follows: green=catalytic site; blue=small subunit; yellow=electron acceptor or donor; red=redox subunit; light orange=maturation factor; dark orange=ion-translocation module; purple=regulatory module; grey=conserved hypothetical. Redox-active centres are shown in circles, where: orange=heme; red=[4Fe4S] cluster; yellow=[2Fe2S] cluster; green=[3Fe4S] cluster; purple=[4Fe3S] cluster. Genes are named according to nomenclature if previously defined. There are often variations in the genetic organisation within subgroups, for example, cytochrome c subunits replace cytochrome b subunits in most group 1a and 1b [NiFe]-hydrogenases in δ-Proteobacteria. However, the organisations depicted reflect the most common organisation, as inferred using the Microbial Genomic Context Viewer.

All hydrogenases could be classified into eight previously described major lineages (Vignais and Billoud, 2007; Calusinska et al., 2010): groups 1 to 4 [NiFe]-hydrogenases, groups A to C [FeFe]-hydrogenases and [Fe]-hydrogenases. However, pre-existing classification schemes did not sufficiently reflect the variety in the functions of the experimentally studied hydrogenases within most of these groups. For example, existing schemes do not account for the great heterogeneity in primary sequence phylogeny, genetic organisation and physiological roles of the group 1 and group 4 [NiFe]-hydrogenases (Vignais and Billoud, 2007), as well as the group A [FeFe]-hydrogenases (Calusinska et al., 2010). In addition, the group 2 [NiFe]-hydrogenases of recently sequenced Aquificae and methanotrophs form distinct lineages from the presently recognised 2a and 2b subgroups (Vignais and Billoud, 2007). We therefore expanded the [NiFe] enzymes into 22 functionally distinct subgroups (groups 1a to 1h, 2a to 2d, 3a to 3d and 4a to 4f) and the group A [FeFe]-hydrogenases into four subtypes (groups A1 to A4). A description of the hydrogenase subgroups/subtypes defined here is provided in Table 1.

Functional diversity of hydrogenases is reflected in phylogenetic clustering, genetic organisation and metal-binding motifs

The hydrogenase classification scheme was developed primarily on the basis of amino acid sequence phylogeny (Figure 1). Our analysis shows that the [NiFe]-hydrogenases are the most diverse and widespread of the hydrogenases, and can be initially divided into H₂-uptake (groups 1 and 2), bidirectional (group 3) and H₂-evolving (group 4) clades. On the basis of phylogeny, these enzymes can be further divided into 22 subgroups (Figure 1; Supplementary Figures S1–S4) predicted to have distinct physiological roles described in Table 1. These subgroups are monophyletic, well populated (with more than 15 unique sequences) and statistically supported (with bootstrap values above 0.75), with the exceptions of certain group 4 clades (see Supplementary Figure S4 legend). On the basis of phylogeny, three [FeFe]-hydrogenase groups were defined: a main group represented by fermentative and bifurcating hydrogenases (group A), an ancestral group of unknown function (group B) and a group containing putative sensory hydrogenases (group C) (Figure 1). However, we were unable to subdivide these enzymes further by phylogeny alone, owing to poor bootstrapping of the group A subclades (Supplementary Figure S5) and lack of functional information on the group B and C enzymes.

The genetic organisation of the hydrogenases also serves as a reliable indicator of function between subgroups. As detailed in Figure 2, genes encoding hydrogenase structural components are proximal to those encoding diverse electron-transfer proteins, ion-translocating subunits, regulatory components, maturation factors, hypothetical proteins and partner enzymes. Although many of these associations have been previously described (Vignais and Billoud, 2007; Schwartz et al., 2013), novel findings included the association of surprising regulatory components (for example, diguanylate cyclases/phosphodiesterases) with putative group C [FeFe]-hydrogenases and group 2c [NiFe]-hydrogenases. Genome architecture is well conserved within subgroups of [NiFe]-hydrogenases, but varies extensively between enzymes of different functions (Figure 2). Genome architecture is also able to discriminate the sometimes poorly bootstrapped lineages of the group 4 enzymes (Supplementary Figure S4) and therefore is a valuable hydrogenase classification tool. Variations in the domain organisation and probable quaternary structure of the [FeFe]-hydrogenases were also apparent. On this basis, group A [FeFe]-hydrogenases can be subdivided into four functionally distinct subtypes: stand-alone enzymes (group A1) and those associated with putative glutamate synthases (group A2), NADH dehydrogenases (group A3) and formate dehydrogenases (group A4) (Figure 2).

As detailed in Table 2, the content of the hydrogenase metal centres also differed between subgroups, further confirming phylogenetic placements. Cysteine residues that bind the metal ions of the catalytic centres of the three types of hydrogenase are conserved, suggesting the chemical structures of the active site do not vary. However, the neighbouring residues in the motifs binding the [NiFe]-centre (L1, L2) and [FeFe]-centre (P1, P2, P3) vary between subgroups that may influence the catalytic behaviour of the enzymes. The number, configuration and ligands of the iron–sulphur clusters of the [NiFe]-hydrogenase small subunits differ between subgroups (Table 2). Nonstandard ligands for iron–sulphur clusters, that is, Asp, Glu, Asn and His, were common in the clusters proximal and distal to the active site. The number of iron–sulphur clusters associated with the [FeFe]-hydrogenase catalytic domain also varies (Table 2 and Figure 2).

The determinants of hydrogen metabolism are widely distributed in bacterial and archaeal genomes

By curating the sequences of all hydrogenases in public sequence databases, we were able to comprehensively map the distribution and diversity of hydrogenases across sequenced microorganisms. Genes encoding putative hydrogenases were detected in 1397 species (Supplementary Table S3) across 55 phyla (Supplementary Table S4) (note that not all species within these phyla contain hydrogenases). The [NiFe]-hydrogenases were the most widespread of the enzymes, occurring in 36 bacterial and 6 archaeal phyla. Consistent with our current knowledge, they were widespread in all classes of Proteobacteria, as well as Firmicutes, Cyanobacteria, Aquificae, Euryarchaeota and Crenarchaeota. [NiFe]-hydrogenases were also common in multiple phyla where hydrogenases have yet to be described, notably Bacteroidetes, Chlorobi, Chloroflexi, Planctomycetes and Verrucomicrobia. Putative group A1 [FeFe]-hydrogenases were detected in 12 phyla of anaerobic bacteria, 5 phyla of unicellular eukaryotes and, surprisingly, the single-amplified genome associated to the archaeal candidate phylum Diapherotrites (pMC2A384). Other types of [FeFe]-hydrogenases, including bifurcating, ancestral and sensory varieties, were exclusive to anaerobic bacteria such as Firmicutes, Bacteroidetes, Spirochaetes, Thermotogae and Fusobacteria. Both [NiFe]- and [FeFe]-hydrogenase genes were also identified within most genomes of newly characterised phyla, including the databases of the MDM project (Figure 3). In contrast, genes encoding the functionally restricted [Fe]-hydrogenases were exclusively found within 25 methanogen genomes (Supplementary Figure S6). Phylogenetic distribution correlates with oxygen preference: obligate anaerobes encoded O₂-sensitive [NiFe]- and [FeFe]-hydrogenases; obligate aerobes encoded O₂-tolerant [NiFe]-hydrogenases (groups 1d, 1h, 2a, 3b, 3d); and genomes of facultative anaerobes contained a diverse range (Figure 3). Many hydrogenases have a mosaic distribution that poorly reflects 16S rRNA gene sequence phylogeny, for example, the abundant group 1d and group 3b [NiFe]-hydrogenases (Supplementary Figures S1 and S3), suggesting strong pressure for lateral acquisition of these enzymes.

Figure 3.

Distribution of hydrogenases in microorganisms. (a) Distribution by hydrogenase type. (b) Distribution by phyla. The cells are shaded by the number of hydrogenases detected in each phyla (light=few hydrogenases, dark=many hydrogenases, grey=no hydrogenases). Hydrogenases were subdivided into the following seven types based on their determined or predicted functions: [NiFe] aerobic uptake (groups 1d, 1h, 2a) [NiFe] anaerobic uptake (groups 1a, 1b, 1c, 1e, 1f, 1g, 3a), [NiFe] bidirectional (groups 3b, 3c, 3d), [NiFe] evolving (groups 4a, 4b, 4c, 4d, 4e, 4f), [FeFe] evolving (groups A, B), [NiFe] regulatory (groups 2b, 2c) and [FeFe] regulatory (group C).

The determinants of microbial hydrogen metabolism are widely distributed in ecosystems

We subsequently gained insight into the distribution of hydrogenases at the ecosystem level through identifying and analysing hydrogenase sequence reads in 20 publicly available metagenomes (Supplementary Table S5). Sequence reads corresponding to the catalytic subunits/domains of hydrogenases were detected in all metagenome samples analysed, and all 22 [NiFe]-hydrogenase subgroups and 3 [FeFe]-hydrogenase groups (but not the methanogen-specific [Fe]-hydrogenases) were detected in at least two samples each. The normalised abundance of hydrogenase reads ranged approximately ∼50-fold, from <0.001% of the total sequence reads in lake and coastal waters to >0.04% reads in permafrost soils and hot springs (Supplementary Table S5). The distribution of hydrogenase-encoding genes in the sequence reads vary depending on the aeration state of the samples: oxic agricultural and forest soils were dominated by aerobically adapted uptake and bidirectional [NiFe]-hydrogenase reads (groups 1d, 1h, 3b, 3d); anoxic termite and human guts contained a high abundance of fermentative and putative sensory [FeFe]-hydrogenase reads (groups A, B, C); and the bog soils, permafrost soils and hot springs contained diverse [NiFe]- and [FeFe]-hydrogenase reads (Figure 4 and Supplementary Figure S7). Hydrogenase-encoding genes were far less abundant within the aquatic ecosystems tested compared with soil and enteric systems; nevertheless, numerous sequence reads homologous to the Robiginitalea biformata hydrogenase of the still-uncharacterised group 1g [NiFe]-hydrogenases were detected in deep ocean samples. The quantity and distribution of hydrogenase reads was consistent between paired metagenome samples, including samples from equivalent ecosystems taken at different locations (for example, Atlantic Ocean vs Indian Ocean) (Figure 4). An interesting exception was the termite gut samples, with hydrogenases predicted to be 10 times more abundant in a gut sample of an African termite (Cubitermes sp.) compared with an American termite (Nasutitermes sp.) (Supplementary Table S5).

Figure 4.

Distribution of hydrogenases in ecosystems. The distribution of different hydrogenase types was analysed in 20 metagenomes. Hydrogenases were subdivided into seven types as described in the legend of Figure 3. Metagenomes were screened using the sequences of the catalytic subunits ([NiFe]-hydrogenases, [Fe]-hydrogenases) or catalytic domains ([FeFe]-hydrogenases) listed in Supplementary Table S1. (a) Percentage of sequence reads for each hydrogenase type identified within 1 million random metagenome reads. (b) Percentage of sequence reads for each hydrogenase type compared with total hydrogenase sequence reads. Supplementary Figure S5 shows the metagenome distribution by [NiFe]-hydrogenase subgroup and [FeFe]-hydrogenase group. Note that no [Fe]-hydrogenases were detected in these metagenomes.

Discussion

Molecular hydrogen is a major electron donor for respiration in both anoxic and oxic ecosystems

Molecular hydrogen occurs ubiquitously in the environment, as a result of production from biological, geothermal and atmospheric sources (Schwartz et al., 2013). Our analysis suggests microorganisms are capable of respiring this fuel source in a wide variety of ecosystems, ranging from the hypoxic H₂-enriched environments of animal guts and bog soils to aerated soils and waters containing trace concentrations of H₂. Group 1 and 2 [NiFe]-hydrogenases that mediate respiratory H₂ uptake were encoded in 19 of the 20 ecosystems surveyed (Figure 4), and were widely distributed in the bacterial and archaeal phyla (Figure 3). As summarised in Table 1, these enzymes have differentiated into multiple subgroups that differ in their redox couplings, oxygen tolerance, affinities and cellular interactions. This enables these enzymes to support life across a wide range of ecological niches (Pandelia et al., 2012; Schwartz et al., 2013).

Integrating our analysis with the wider literature, we suggest oxygen partial pressure (pO₂) principally drove the evolution and distribution of respiratory hydrogenases. Phylogenetic analysis reveals the deepest-branching forms of these enzymes (groups 1a, 1b) are O₂ sensitive and mediate anaerobic respiration in strictly anaerobes (Supplementary Figures S1 and S2). Such enzymes are abundant in hypoxic soils (for example, bog soils, permafrost soils) (Supplementary Figure S7), and are predominantly found in the genomes of anaerobic Firmicutes and δ-Proteobacteria (Supplementary Figure S1) capable of H₂-dependent sulphate reduction, metal reduction and dehalorespiration (Schwartz et al., 2013). The mid-branching heterotetrameric hydrogenases (groups 1c, 1e, 1g) were more taxonomically dispersed (Supplementary Figure S1) and appear to support roles in fumarate and nitrate respiration, anoxygenic photosynthesis and chemolithoautotrophy across a diversity of taxa (Pandelia et al., 2012). In contrast, the more recently branching lineages (groups 1d, 1h and 2a) appear to be oxygen-tolerant enzymes that mediate respiration in aerobes and facultative anaerobes (Table 1). These enzymes were predominant in aerated samples (Figure 4), with 0.004 to 0.009 % of total metagenome sequence reads in agricultural and forest soils corresponding to the group 1h enzyme (Supplementary Table S5). Our analysis suggests that these subgroups independently developed mechanisms to tolerate O₂ following the emergence of oxygenic photosynthesis (Figure 1); consistently, the well-reported proximal 6Cys[4Fe3S] cluster (Fritsch et al., 2011) is exclusive to 1d enzymes (Table 2), indicating the 1h and 2a subgroups use alternative mechanisms (possibly using other nonstandard clusters) to prevent or reverse formation of O₂-inhibited states.

Oxygen-tolerant uptake hydrogenases are significantly more widespread than the literature currently reports. Aerobic H₂ uptake has only been reported in three dominant soil phyla to date: many α-, β- and γ-Proteobacteria (for example, Ralstonia eutropha; Schwartz et al., 2013) can grow chemolithoautotrophically using biologically evolved H₂, whereas certain model Actinobacteria (for example, Mycobacterium smegmatis; Greening et al., 2014) and Acidobacteria (that is, Pyrinomonas methylaliphatogenes; Greening et al., 2015a) enhance their persistence by scavenging atmospheric H₂. However, Figure 3 reveals that the group 1d, 1h, and 2a hydrogenases mediating such processes are also encoded in some 17 bacterial and archaeal phyla, among them representatives of all 9 of the most dominant phyla in global soils (Janssen, 2006). Most significantly, the group 1h [NiFe]-hydrogenases that mediate tropospheric H₂ oxidation are encoded in multiple representatives of undercultured, slow-growing phyla (that is, Acidobacteria, Verrucomicrobia, Chloroflexi and Planctomycetes). These findings are consistent with our recent hypothesis that H₂ serves as an energy source for the maintenance of dormant soil bacteria (Greening et al., 2015b). Hydrogenase-encoding genes were also identified in the genomes of multiple seemingly obligate methane oxidisers, ammonia oxidisers and nitrite oxidisers (Supplementary Table S1), suggesting H₂ may serve as a fuel source for growth or survival of these bacteria and archaea. In line with this, it was recently demonstrated that Nitrospira moscoviensis of the phylum Nitrospirae is capable of hydrogenotrophic growth using a group 2a [NiFe]-hydrogenase (Koch et al., 2014). Aerobic H₂ oxidation may therefore provide hitherto-unrecognised metabolic flexibility in microorganisms controlling the methane and nitrogen cycles.

Determinants of fermentative hydrogen production are universally distributed

Since its discovery in the early twentieth century (Stephenson and Stickland, 1932), it has been widely believed that fermentative H₂ evolution occurs exclusively in anaerobic microorganisms. This notion was recently challenged by the discovery that the obligately aerobic soil bacterium Mycobacterium smegmatis evolves H₂ using a tightly regulated hydrogenase to maintain redox balance under hypoxia (Berney et al., 2014). The surveys presented in this work demonstrate that homologues of the group 3b [NiFe]-hydrogenase mediating mycobacterial H₂ evolution actually have the most extensive distribution (phylum level) of all the subgroups in our database. Once thought to be confined to anaerobic archaea (Ma et al., 1993), these enzymes actually occur in at least 27 bacterial and archaeal phyla, among them multiple representatives of the ‘MDM' (Rinke et al., 2013) (Figure 3). In addition to the versatile group 3d [NiFe]-hydrogenases (Burgdorf et al., 2005), these oxygen-tolerant enzymes are proposed to serve as redox valves that interconvert electrons between NAD(P)H and H₂ depending on the availability of exogenous electron acceptors (Greening and Cook, 2014). These enzymes are also abundant at the metagenome level, constituting dominant groups in aerated soils and hot spring ecosystems (Figure 4). These findings may help to explain why the communities of such ecosystems are relatively stable despite pO₂ fluctuations. Unsurprisingly, the classical determinants of H₂ fermentation were abundant in anoxic ecosystems (Figure 4). Figure 3 shows that formate hydrogenlyases (group 4a [NiFe]-hydrogenases) are widespread in enteric bacteria that adopt a facultatively fermentative lifestyle. The group A1 [FeFe]-hydrogenases, which mediate ferredoxin-dependent H₂ production (Peters et al., 1998), are distributed in numerous obligately fermentative bacteria (for example, clostridia), eukaryotes containing hydrogenosomes (for example, Trichomonas vaginalis) and unicellular algae mediating photobiological H₂ production (for example, Chlamydomonas reinhardtii). On the basis of domain conservation and phylogenetic similarity, we predict the still-uncharacterised group B [FeFe]-hydrogenases serve a similar function.

Energy-converting and electron-bifurcating complexes enhance the efficiency and flexibility of anaerobe-type hydrogenases

Although group 4 [NiFe]-hydrogenases are traditionally known for their roles in fermentation, the majority of these enzymes have a respiratory function. They associate into complexes comprising primary dehydrogenases and terminal hydrogenases and conserve the energy liberated during electron transfer as a proton- or sodium-motive force (Buckel and Thauer, 2013). Our analysis shows these enzymes have retained roles in anaerobic microorganisms, especially Firmicutes, Proteobacteria (γ, δ and ɛ classes) and methanogens (Figure 3), and contribute to hydrogenase diversity in metagenomes (Figure 4). They appear to have diverse physiological roles, as reflected by their wide-branching phylogeny (Figure 1) and highly modular genetic organisation (Figure 2). This enables them to liberate electrons from low-potential donors, namely formate (group 4a, 4b, possibly 4f), carbon monoxide (group 4b, 4c) or ferredoxin (group 4d, 4e), whereas protons serve as the terminal electron acceptor. Though minimalistic, the respiratory chains they form are often highly efficient and may provide a primary strategy for energy generation within particularly oligotrophic environments; this was emphasised by the recent discovery of a complex in the deep-sea vent archaeon Thermococcus onnurineus that sustains growth across a narrow energy bracket by transferring electrons from formate to protons (Kim et al., 2010; Lim et al., 2014). Others are highly flexible, as demonstrated by the multifaceted roles of the physiologically reversible Ech hydrogenase (group 4e) in hydrogenotrophic vs aceticlastic methanogenesis (Meuer and Kuettner, 2002). Our phylogenetic analyses suggest that the ancestral forms of the group 4 enzymes—and likely [NiFe]-hydrogenases as a whole—may have been formate-oxidising, H₂-evolving, energy-transducing complexes. We discovered a deep-branching lineage of these enzymes in Firmicutes (candidate group 4f [NiFe]-hydrogenases) that align closely with the functionally cryptic Ehr complexes (homologues of group 4 [NiFe]-hydrogenases lacking Ni-binding cysteine residues; elaborated on in Marreiros et al., 2013) and yet possess the critical cysteine residues required for [NiFe]-centre ligation.

Many of the anaerobe-type hydrogenases we identified are predicted to mediate electron bifurcation, a recently discovered third mode of energy conservation. Electron-bifurcating hydrogenases are bidirectional enzymes that energise the endergonic reaction of the reduction of ferredoxin with H₂ by simultaneously reducing a relatively electropositive acceptor (for example, heterodisulphide, NAD, NADP) (Buckel and Thauer, 2013). The group 3c [NiFe]-hydrogenase in functional complex with heterodisulphide reductase, for example, simultaneously reduces ferredoxin and heterodisulphide during H₂ oxidation (Kaster et al., 2011); these enzymes complete the recently elucidated Wolfe cycle of methanogenesis (Thauer, 2012), and are also distributed in some bacteria (for example, δ-Proteobacteria) (Figure 4). The group A3 [FeFe]-hydrogenases reversibly bifurcate electrons from H₂ to ferredoxin and NAD using trimeric or tetrameric complexes; in the reverse reaction, energy conserved during the oxidation of ferredoxin is used to drive the thermodynamically unfavourable production of H₂ from NADH (Schut and Adams, 2009; Schuchmann and Müller, 2012). A subtype of the group A4 [FeFe]-hydrogenases can also bifurcate electrons from H₂ to NADP and ferredoxin, and act physiologically in hexameric complexes with formate dehydrogenase (Wang et al., 2013). We analysed the genetic organisation of the 705 group A [FeFe]-hydrogenases represented in our database in order to identify putative electron-bifurcating complexes (Figure 2 and Supplementary Table S1). This analysis suggested that whereas putative NADP-dependent bifurcating complexes are rare (7 sequences), putative NAD-dependent bifurcating complexes are very abundant in anaerobic bacteria (391 sequences). The group A3 [FeFe]-hydrogenases are highly flexible, capable of both dissipating excess reductant during fermentation (for example, cellulose fermentation) and generating reduced electrons for carbon fixation (for example, acetogenesis) and respiration (via the sodium-motive ferredoxin-NAD oxidoreductase complex) (Buckel and Thauer, 2013; Schuchmann and Muller, 2014). Supported by PCR amplicon sequencing (Zheng et al., 2013), metagenome analysis (Figure 4) demonstrates that group A [FeFe]-hydrogenases, including probable bifurcating hydrogenases, are abundant in termite guts.

Hydrogen sensing may be more important than previously recognised

Our exploration of hydrogenase sequences across a diversity of environments also uncovered evidence that hydrogen-based signal transduction cascades are more significant than previously anticipated. The only characterised sensory hydrogenases to date are the group 2b [NiFe]-hydrogenases (for example, Ralstonia eutropha, Rhodobacter capsulatus) (Lenz and Friedrich, 1998; Vignais et al., 2005); these enzymes have adapted the [NiFe] active site to sense high partial pressures of H₂ and in turn activate two-component regulatory cascades that control expression of respiratory hydrogenases (Greening and Cook, 2014). Our analysis shows these enzymes are restricted to Proteobacteria (α, β and γ classes) (Figure 3 and Supplementary Figure S2) and are present in soil environments characterised by logarithmic variations in pH₂ (Figure 4 and Supplementary Figure S7). We identified a sister lineage, the group 2c [NiFe]-hydrogenases, that appear to be co-transcribed with diguanylate cyclases/phosphodiesterases (Figure 2). Through modulation of cyclic di-GMP production, we hypothesise these enzymes regulate global cellular functions during adaptation to H₂-rich vs H₂-deprived environments; currently, however, H₂-dependent signal cascades have only been shown to regulate the expression of other hydrogenases. Our genome and metagenome surveys suggest these enzymes are rare (Figures 3 and 4), and are primarily found in methane-oxidising bacteria and sulphate-reducing bacteria that inhabit aquatic environments (Supplementary Figure S2). The presence of helix-turn-helix protein-encoding genes immediately downstream of group 2 [NiFe]-hydrogenase genes in some Aquificae and Crenarchaeota (Supplementary Figure S2) is also suggestive of a regulatory role and requires further study.

Looking more widely, it is probable that the group C [FeFe]-hydrogenases of anaerobic bacteria have a sensory role. Our conserved domain analysis suggests these enzymes are expressed or fused with putative regulatory components, namely serine/threonine phosphatases, histidine kinases, AAA+-type transcriptional activators, methyl-accepting chemotaxis proteins and again diguanylate cyclases/phosphdiesterases (Figure 2). As with the group 2b and 2c [NiFe]-hydrogenases, the operons encoding these hydrogenases contain predicted PAS domains (Figure 2) that likely transduce the signal of hydrogenase activity to downstream components via a redox-active heme. There is some transcriptional evidence that the putative phosphatase-linked sensory hydrogenases of Thermoanaerobacterium saccharolyticum (Shaw et al., 2009) and Ruminococcus albus (Zheng et al., 2014) regulate the transcription of group A [FeFe]-hydrogenases, but it has yet to be biochemically confirmed that these enzymes have a regulatory role. Other sensory hydrogenases may regulate wider cellular functions (for example, motility) in response to changes of pH₂ in anoxic environments. Group C [FeFe]-hydrogenases are abundant in strictly anaerobic bacteria of the phyla Firmicutes, Bacteroidetes, Spirochaetes and Thermotogae. These hydrogenases are highly abundant in termite guts and strongly associated with group A [FeFe]-hydrogenases (Supplementary Figure S7).

Conclusions

The surveys reported here suggest that hydrogenases are highly diverse, ancient and widespread. Our work collectively supports the hypothesis that H₂ serves as a widely utilised energy source for microbial growth and survival. Before this study, it was already well established that H₂ metabolism played major roles in certain specific microorganisms and ecosystems (Schwartz et al., 2013). However, by comprehensively surveying the distribution of hydrogenases, we have provided evidence that microbial H₂ metabolism is significantly more extensive and elaborate than previously anticipated. Integrating analysis of primary phylogeny, genetic organisation and metal-binding motifs, we demonstrate that hydrogenases have evolved into numerous functionally distinct subgroups/subtypes. This diversification has enabled the primordial process of H₂ metabolism to sustain roles in most major phyla and ecosystems. We showed that some 51 bacterial and archaeal phyla have the genetic capacity to oxidise or evolve H₂—vastly more than the 13 phyla experimentally shown to metabolise H₂ (Schwartz et al., 2013; Greening et al., 2015a)—and emphasised through metagenome analysis that microbial H₂ metabolism is likely to be highly important in both oxic and anoxic environments. Our analysis has emphasised that the evolution and distribution of hydrogenases is particularly influenced by pO₂; however, other factors such as pH₂, pH, temperature and metal ion availability are also likely to be profoundly significant (Schwartz et al., 2013; Greening and Cook, 2014). However, experimental studies are required to gain a deeper understanding of the ecological significance of H₂ oxidation and evolution.

In light of this work, there are now numerous new avenues to investigate microbial hydrogen metabolism at the microscopic and macroscopic levels: What are the functions of multiple newly defined types of [NiFe]-hydrogenase (groups 1e, 1g, 2c, 2d, 4f) and [FeFe]-hydrogenases (groups A2, B, C)? Why are hydrogenases found in the genomes of microorganisms as diverse as Acidobacteria, Chlorobi, Crenarchaeota and Bacteroidetes? What environmental and physiological signals lead to the regulation of the genetic determinants of hydrogen metabolism? How does microbial H₂ metabolism influence anthropogenic ecosystems (for example, wastewater treatment) and how can the reported diversity of hydrogenases be exploited for bioremediation, biofuel production and fuel cell development? How does microbial H₂ metabolism influence community structuring and biogeochemical cycling in soil and aquatic environments? Some of these research questions will be addressed as we further investigate the physiological roles of the hydrogenases described here and the influence of H₂ metabolism in different ecosystems.

The authors declare no conflict of interest.