Many microbes grow by producing methane gas from carbon dioxide and hydrogen gas, and enzymes known as hydrogenases play important roles in this metabolic process. The production of methane in these microbes depends on a nickel–iron hydrogenase called Frh adding electrons to a coenzyme called F₄₂₀. This hydrogenase cleaves a hydrogen molecule into two electrons, which are transferred to the F₄₂₀ coenzyme, and two protons. The reduced form of F₄₂₀ is then used for several reactions in the methane production process. This process, which is known as methanogenesis, provides the microbes with energy.

Nickel–iron hydrogenases can be divided into five different groups, but researchers have been able to determine the detailed structures of the enzymes in just one of these groups. All nickel–iron hydrogenases contain at least two subunits: a large subunit with a catalytic center composed of both nickel and iron ions and a small subunit that contains three iron–sulfur clusters. Frh—which is short for F₄₂₀-reducing nickel–iron hydrogenase—is known to have a third subunit comprising an extra iron–sulfur cluster and a coenzyme called FAD that allows it to interact with the F₄₂₀ coenzyme. However, until now, little was known about the detailed structure of the Frh enzyme.

Mills et al. have used electron cryo-microscopy (cryo-EM) to determine the structure of Frh when it is on its own, and also when it is bound to F₄₂₀. This technique involves freezing a solution of the enzyme in a thin layer of ice and recording an image of this layer in an electron microscope. By combining a large number of images, each of which contains many identical enzymes in different orientations, it is possible to determine the 3-dimensional structure of the enzyme.

Mills et al. found that Frh forms a very large tetrahedral complex that contains six Frh dimers. And by comparing the structure with and without F₄₂₀, they identify a pocket near the FAD coenzyme that the F₄₂₀ coenzyme binds to. They also identify a fold in the third subunit that allows proteins to bind both FAD and F₄₂₀. The work demonstrates the potential of cryo-EM to elucidate structures that cannot be determined by other approaches.

[NiFe]-hydrogenases are microbial enzymes that heterolytically cleave H₂, after which the electrons from a hydride ion are reversibly transferred to electron carriers. These enzymes are involved in many metabolic pathways in microbial ecosystems, notably in methanogenesis. The members of the [NiFe]-hydrogenase family are composed of at least two subunits, large (∼60 kDa) and small (∼30 kDa), which contain a [NiFe] dinuclear metal center and three iron–sulfur (Fe-S) clusters, respectively. The family is classified into five distantly related groups based on a phylogenetic analysis of their primary structures (Vignais and Billoud, 2007; Constant et al., 2011). The cytoplasmic F₄₂₀-reducing [NiFe]-hydrogenase (Frh) belongs to the group 3 [NiFe]-hydrogenases, which catalyze the reversible reduction of the soluble hydride carrier, NAD(P) or F₄₂₀. F₄₂₀ is a deazaflavin derivative that acts as an important hydride acceptor/donor in the central methanogenic pathway. The group 3 hydrogenases contain an additional subunit that interacts with the soluble coenzymes, for example, the iron–sulfur flavoprotein FrhB. To date, the crystal structures of [NiFe]-hydrogenases only from group 1 have been determined: six derived from sulfate-reducing bacteria, including two orthologs harboring a [NiFeSe]-center as active site (Volbeda et al., 1995; Higuchi et al., 1997; Montet et al., 1997; Garcin et al., 1999; Matias et al., 2001; Marques et al., 2010), one from a photosynthetic bacterium (Ogata et al., 2010), and three from hydrogen-oxidizing bacteria and Escherichia coli, which in contrast to the other species are oxygen-tolerant and have an usual [4Fe-3S] proximal cluster (Fritsch et al., 2011; Shomura et al., 2011; Volbeda et al., 2012).

Frh is a key enzyme in the metabolism of methanogenic archaea (Thauer et al., 2010). The reduction of carbon dioxide to methane is an overall eight-electron reduction process involving the oxidation of four molecules of H₂ by several hydrogenases. Four electrons are provided through the reduced form of the coenzyme F₄₂₀, which is regenerated by Frh. Frh is a heterotrimeric enzyme composed of the large subunit FrhA (45 kDa) with a binuclear [NiFe]-center, the small subunit FrhG (26 kDa) with three [4Fe4S] clusters and the iron–sulfur flavoprotein FrhB (31 kDa) with a [4Fe4S] cluster and FAD, and the F₄₂₀-binding site. Frh accounts for about 1% of the cytoplasmic protein and has been shown by electron microscopy to form a huge complex of similar appearance in all species investigated: Methanococcus voltae (Muth et al., 1987), Methanospirillum hungatei (Sprott et al., 1987), Methanobacterium thermoautotrophicum ΔH (Wackett et al., 1987), and Methanobacterium thermoautotrophicum Marburg (Braks et al., 1994). From negative stain images, the complex was interpreted as a flat cylinder consisting of an octamer (Wackett et al., 1987) or hexamer (Braks et al., 1994) of the heterotrimer.

Recently, developments in instrumentation and image processing software have made it possible to determine structures of large macromolecular complexes to near-atomic resolution by cryo-electron microscopy. Most successful have been studies of large icosahedral viruses (Yu et al., 2008, 2011; Liu et al., 2010; Wolf et al., 2010; Zhang et al., 2010a; Chen et al., 2011; Settembre et al., 2011), but also a few smaller complexes in the 1 MDa size range of lower symmetry have yielded high-resolution structures, notably GroEL (Ludtke et al., 2008) and archaeal chaperonins (Cong et al., 2010, 2012; Zhang et al., 2010b). We present here the structure of the Frh complex from the hydrogenotrophic methanogenic archaeon, Methanothermobacter marburgensis, by cryo-electron microscopy. The Frh complex is a 1.2-MDa dodecamer with tetrahedral symmetry. The location of all cofactors were identified, and the backbone of the three proteins was traced, including FrhB that has a novel fold.

The Frh complex was highly purified from M. marburgensis under strict anaerobic conditions in the presence of FAD. Its homogeneity was confirmed by electron microscopy with negative staining. We found that all projections of the Frh complex are ring-shaped with a diameter of ∼16 nm (Figure 1A). Class averages showed twofold and threefold symmetry (Figure 1B), which is consistent with tetrahedral symmetry. We conclude that the complexes consist of six dimers of FrhABG arranged into a shell around a hollow core, not in a cylindrical arrangement as in previous EM models (Wackett et al., 1987; Braks et al., 1994). The dodecamer has a total MW of 1.2 MDa. Iterative structural refinement of a 84,000-particle cryo-EM dataset, applying tetrahedral symmetry, resulted in a 3D reconstruction (Figure 2) at a resolution where the pitch of α-helices and many side chains are recognizable in the map, and β-strands are clearly separated (Figure 2E–G, 5B). The globular protein complex showed no preferred orientations in the ice, resulting in a homogeneous coverage of angles (Figure 1C). The map shows six distinct dimers, all clearly separated from one another (Figure 2C), sitting on the faces of a cube. Each monomer contains a series of five high-density features separated by ∼10 Å, forming an arc in the interior of the protein (Figure 2D and Video 1). These features are easily assigned to the four Fe-S clusters and the [NiFe] center; the latter is recognized by its smaller size. Although at first glance the position of the FrhABG heterotrimers in the dimer is not obvious, the path of the electron transfer chains provides definitive clues to the location of an FrhABG heterotrimer. FrhABG contains one approximately 35-Å-long helix on the outer surface and a bundle of four up to 40-Å-long helices at the interface of different dimers, running from the outside to the interior of the complex. Other helices are considerably shorter.

Figure 1

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The large subunit FrhA has considerable homology to the known large subunit structures (Figure 3), and is characterized by four long α-helices. Docking the large subunit of the group 1 enzyme into the map provides a good fit for the α-helix bundle, with the [NiFe] center correctly situated in its identified location and many other structural elements of FrhA matching closely, including two three-stranded antiparallel β-sheets. FrhA is about 15 kDa smaller than the large subunits of known structure; all missing elements are located on the periphery. A structural model for FrhA (Figure 4A) was created based on this fit and a sequence alignment with the protein from Desulfovibrio vulgaris Hildenborough (Figure 3). All but the first three amino acids in the sequence could be traced in our maps. Densities for large side chains confirm the structural assignment (Figures 2, 5 and Video 2). The [NiFe] center is located near four cysteine residues, Cys 63, 66, 380, and 383, which are fully conserved in all [NiFe]-hydrogenases. The group 1 [NiFe]-hydrogenases contain another ion, typically either Mg²⁺ or Fe^2+/3+ (Higuchi et al., 1997; Garcin et al., 1999). In our map, there is strong density at this position (Figure 5D), which is part of the structurally conserved region of FrhA, and its surrounding residues, Glu44, Glu229, and His386, are fully conserved in FrhA (Figure 3A), so we interpret this density as an ion. One of the helices of the four-helix bundle has a pronounced kink. This helix was interpreted as residues 166–193, and a proline residue (Pro180) not present in the group I proteins was found exactly at the position of the kink, supporting the assignment. No density is present beyond His386, and there is no room in the structure for the final 19 amino acids of the protein. The C-terminal residues after the equivalent histidine in [NiFe]-hydrogenases are enzymatically removed during maturation of the enzymes (Menon et al., 1993; Theodoratou et al., 2005). We can conclude that FrhA is posttranslationally modified in the same way, likely by FrhD, a homologous endopeptidase contained in the Frh operon FrhADGB (Liesegang et al., 2010; Thauer et al., 2010).

Figure 3

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The fit of the small subunit of the group 1 enzyme reveals structural homology to the N-terminal domain of FrhG including the proximal Fe-S cluster. Some peripheral helices present in the group 1 protein are missing in the FrhG structure, and the similarity is generally lower than for the large subunit (Figure 6C). The N-terminal domain is characterized by a conserved parallel β-sheet flanked by short α-helices (Figure 4B). In the group 1 [NiFe]-hydrogenases, the proximal Fe-S cluster is located at the C-terminal end of the β-sheet and is coordinated by four cysteines. In the primary structure of FrhG, one of the conserved cysteine residues for the coordination of this cluster is replaced by Asp60 (Figure 6A). The bacterial hydrogenases as well as most of the archaeal species have a cysteine at this position, but aspartates have been identified before as ligands of [4Fe4S] clusters (Calzolai et al., 1995; Muraki et al., 2010; Gruner et al., 2011). Accordingly, we positioned this aspartate near the proximal Fe-S cluster.

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The C-terminal domain of FrhG has no homology to the group 1 [NiFe]-hydrogenases. Instead, the sequence indicates a ferredoxin domain, with two CxxCxxCxxxC sequences coordinating two [4Fe4S] clusters (Figure 6A) (Alex et al., 1990). Accordingly, we modeled the C-terminal domain of FrhG on ferredoxin (Figures 4B, 5A and Video 3). In our model, the residues 235–244 form a β-hairpin with Gly240 at the turn. The two β-strands were predicted and the hairpin is clearly visible in the density, confirming the assignment. Unlike FrhA, which could be traced over its full length, density of FrhG for a 12-residue stretch (188–199) is missing (Figure 6A). This region is located at the surface and may be disordered. FrhG as deduced from the M. marburgensis genome sequence is 275 residues long (Liesegang et al., 2010), but no density is present for the 45 N-terminal amino acids. The frhG gene most probably starts with the initiation codon GTG annotated as Val39 in the genome sequence (Fox et al., 1987; Alex et al., 1990), which means that density is missing in the map for only the first seven amino acids of FrhG.

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FrhB contains one Fe-S cluster, an FAD, and the F₄₂₀-binding site. There is no homolog of known structure. FrhB was therefore traced ab initio, aided by secondary structure predictions (Figure 7) and recognizable features in the map that included 10 α-helices, 11 β-strands, and the Fe-S cluster. The most prominent feature of FrhB is a ∼35 Å surface helix that corresponds to the predicted 24-residue C-terminal helix.

Figure 7

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We examined the F₄₂₀-binding proteins of known structure in the protein databank, and found no structural homology with FrhB. All these proteins have cofactors different from FAD, for example, methylene-H₄MPT or NADP (Ceh et al., 2009). Also no homologous FAD-binding proteins were found. We conclude that FrhB has a novel fold (Figure 4C and Video 4). The Fe-S cluster is most probably a 4Fe-4S cluster because it is surrounded by four cysteines (Cys 104, 134, 192, and 195), which are all fully conserved (Figure 7). The core of FrhB is a mixed six-stranded β-sheet. A density located ∼8 Å from the Fe-S cluster could be interpreted as the isoalloxazine ring of the FAD cofactor. The pyrophosphate moiety is located at the N-terminal end of an α-helix, which is a common FAD-binding motif where the dipole of the helix compensates the negative charge (Dym and Eisenberg, 2001) (Figure 8), and nearby density for the adenine moiety is visible. The FAD has a bent conformation, with the adenine and isoalloxazine ring only 4–5 Å apart (Figure 8) near the highly conserved loop Lys75-Tyr76. The FAD is completely surrounded by highly conserved protein regions (Figure 7 and Video 5), further verifying the backbone trace.

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To identify the substrate-binding site, we imaged the FrhABG complex in the presence of a large excess of F₄₂₀, and calculated maps, which were refined to a similar resolution. At all refinement stages, the map showed an additional density in a region that was empty in maps of Frh without substrate (Figure 8A,B). This density runs approximately parallel to the FAD isoalloxazine ring, at a distance of ∼4 Å. We conclude that it represents the isoalloxazine ring of the F₄₂₀ substrate. It is seated in a large pocket surrounded by highly conserved residues, with easy access to the surface of the complex. The F₄₂₀ density is close to the small residues S209 and V210 in a loop between two β-strands, which are part of the longest conserved region of FrhB (Figure 7). A predicted three-stranded β-sheet (148–172) flanked by two short helices (Figure 7) fits nicely in density. The only conserved region of this sheet, 163IGKGK, forms a turn close to the position of the isoalloxazine ring of F₄₂₀. The location between this ring and the surface of the complex suggests that one or both of the conserved lysines may interact with the phosphate group of F₄₂₀. The long α-helix on the surface that was interpreted as the C-terminal helix of FrhB is not very well resolved. The residues in the helix are mostly not conserved in FrhB, except at the N-terminal end (Figure 7). In our model, the conserved residues are located near the loop 164GKGK mentioned above. This region also contains several other conserved residues (Video 5) and likely constitutes the access channel for the substrate F₄₂₀. The map of Frh in the presence of substrate is similar to the substrate-free map, indicating that F₄₂₀ binding does not involve major conformational changes.

The residues coordinating the Fe-S cluster, FAD, and F₄₂₀ are conserved in the FrhB family, which includes subunits of F₄₂₀-dependent sulfite reductase (Fsr), F₄₂₀H₂:quinone oxidoreductase (FqoF), F₄₂₀H₂:phenazine oxidoreductase (FpoF), F₄₂₀-dependent glutamate synthase and formate dehydrogenase (FdhB) (Johnson and Mukhopadhyay, 2005) (Figure 9). Some of these proteins contain additional domains, like an N-terminal ferredoxin domain (which in Frh is part of FrhG), or a C-terminal domain containing a binding site for the substrate of the electron transfer from F₄₂₀. Clearly, these enzymes share a common fold with FrhB.

Figure 9

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Recent advances in instrumentation and image processing procedures have made cryo-electron microscopy of isolated macromolecular complexes a powerful technique in structural biology. After the first virus structure at subnanometer resolution (Böttcher et al., 1997) showed the feasibility of the method, icosahedral viruses were reconstructed in recent years to better than 4 Å resolution (Yu et al., 2008, 2011, Zhang et al., 2008, 2010a; Liu et al., 2010; Wolf et al., 2010; Chen et al., 2011; Settembre et al., 2011; ), taking advantage of the 60-fold symmetry and huge size (∼20 to 150 MDa), which together contribute to the relatively high signal-to-noise ratio for these particles that makes the determination of the orientation unambiguous. For smaller complexes, the techniques have been pioneered with the 800-kDa GroEL complex with D7 symmetry, reaching subnanometer resolution for the first time in 2004 (Ludtke et al., 2004) and ∼4 Å in 2008 (Ludtke et al., 2008). Subsequent reconstructions to this resolution were achieved for archaeal (Zhang et al., 2010b) and mammalian (Cong et al., 2010) chaperonins. The chaperonins have different conformational states, and usually only one of these gave a high-resolution map, whereas for the others the resolution was limited due to conformational flexibility. The appearance of our reconstructions is similar to the chaperonins; in agreement with this, Fourier shell correlation indicates a resolution of 3.9 and 4.0 Å for the two maps, respectively (Figure 10A). Recently it has been noted that the resolution of such reconstructions are usually overestimated due to correlation of noise at high resolution (Scheres and Chen, 2012). Using a ‘gold standard' procedure to estimate resolution, where two half-data sets are independently refined to a reference not containing high-resolution information (see ‘Materials and methods'), yielded 5.5 Å (Figure 10B). However, an objective indication of (local) resolution is provided by the appearance of secondary structure: the α-helical pitch of 5.4 Å is very well defined in many regions (Figure 2F,G), as is the separation of β-strands (axial distance ∼4.8 Å) (Figure 5B). Comparison of these areas with maps calculated from the model low-pass filtered to different resolutions suggest a resolution ∼4.5 Å. The overall resolution estimate is an average of these well-resolved protein regions and other regions that are less well resolved, especially those at the periphery of the complex. An FSC curve between the experimental map and a map calculated from the model indicated a resolution of 5.8 Å (Figure 10C); this figure also constitutes an average over the whole structure, with the model probably best for internal α-helices and worst for loops at the periphery. Since the model was derived from the map, and not from an independent experiment, this measure does not properly reflect the resolution of the map, but rather describes how well our admittedly incomplete model explains the map densities. It should be noted that we used tight protein structure restraints during modeling, in order to prevent overfitting to noise while creating an unrealistic model that would have yielded a higher model-to-map correlation.

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Several factors contributed to the successful structure determination of Frh. The tetrahedral symmetry means that there are 12 asymmetric units in each particle, less than the 14 and 16 in the D7 and D8 symmetric chaperonins, but the resulting ball-like shape ensures that the complex has no preferential orientation in the ice (Figure 1D), unlike the cylinder-shaped chaperonins that tend to be oriented with their symmetry axis parallel or perpendicular to the ice (Ludtke et al., 2001; Zhang et al., 2011). From the quality of the reconstructions, we conclude that the complex is very rigid. The maps of Frh with and without substrate are of similar resolution and quality, implying that any protein conformational changes due to F₄₂₀ binding are too small to detect at the present resolution. The density for F₄₂₀ is weak, but its position was inferred at an early stage of the analysis from the fact that maps of Frh without the substrate were consistently empty and maps with F₄₂₀ always showed some density here. The later tracing of the amino acid chain of FrhB and the FAD confirmed the assignment: F₄₂₀ is in van der Waals distance from its hydride donor, the FAD isoalloxazine ring, and the residues in the vicinity are highly conserved in FrhB and in the FrhB family of F₄₂₀-binding proteins (Figures 7, 9 and Video 5).

While the chain trace for FrhA and FrhG was relatively straightforward using the group I hydrogenase homolog, FrhB was built ab initio. The visibility of the FeS cluster density (Figure 2D and Video 1) made it possible to use the four Fe-S ligand cysteines together with a secondary structure prediction as an initial guide; large side chain densities helped to establish the correct sequence assignment. The secondary structure prediction was fully confirmed, and weak or discontinuous densities in the map were almost invariably found to represent glycine residues, further indicating a correct assignment. Density for the FAD cofactor only became clear when most of the protein chain was placed; density for the ribose and ribitol are missing (Figure 8). The pyrophosphate group is closely associated with the N-terminal α-helix 26–64 of FrhB. The N-terminus of this helix is one of the most conserved regions in the FrhB family (Figures 7, 9), and its role in compensating the negative charge of the phosphates is a common motif in FAD-binding proteins (Dym and Eisenberg, 2001). Conserved regions of FrhB were found near the FAD and surrounding the F₄₂₀-binding site and its putative entrance channel (Video 5). Of the three proteins, only FrhG is missing density for an internal stretch, residues 188–199 (Figure 6A). The flanking residues of this stretch are located less than 10 Å apart on the surface of the complex and they probably form a flexible loop. FrhA and FrhB were traced over their full length. All three proteins follow the rule that hydrophobic residues form the core of the protein and charged residues face the outside. We found several pockets lined with hydrophobic side chains, not just in FrhA and FrhG, where these features are shared with the bacterial protein used as a template for modeling, but also in FrhB. FrhB also contains a highly hydrophobic helix (27GIVTGLLAYAL), which is buried completely inside the subunit. These features all indicate a correct chain trace.

Our structure shows that each heterotrimer encompasses a complete electron transfer chain that runs from the [NiFe] cluster in FrhA via the three Fe-S clusters in FrhG and one Fe-S cluster and FAD in FrhB to F₄₂₀ (Figures 4D, 11 and Video 6). The cofactors describe a curvilinear path in the protein interior, almost parallel to the surface of the complex, with distances of ∼10 Å between neighbors, except FAD and F₄₂₀, which are in van der Waals contact to enable hydride transfer (Figure 8C). The predicted F₄₂₀-binding site is located on the outside of the complex, which facilitates access of the cytosolic F₄₂₀. The two electron transfer chains in a dimer are separated by at least 17 Å and are thus independent of each other, indicating that the functional unit is one FrhABG heterotrimer and there is no cooperativity in the complex. The three proteins in the Frh heterotrimer form a tight complex with extensive subunit interfaces and a large number of subunit interactions, including two salt bridges between loops of the two FrhA molecules in a dimer, from Asp312 of one FrhA and Arg314 of the other (Figure 5C). The heterotrimer is quite slender compared to the two-subunit [NiFe] hydrogenases of known structure, with a small contact surface especially between FrhB and the other two subunits (Figure 12), and complex formation may be necessary for stability of the complex and keeping the metal clusters at optimal distances. The same quaternary structure of Frh has been observed not only in the closely related thermophile M. thermautotrophicus (Wackett et al., 1987) but also in the phylogenetically distant mesophilic M. voltae (Muth et al., 1987) and M. hungatei (Sprott et al., 1987), so it is clear that the formation of a large complex plays an essential role for the function of Frh.

Figure 11

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Author details

1. Deryck J Mills

   Department of Structural Biology, Max Planck Institute of Biophysics, Frankfurt, Germany

   Contribution

   DJM, Prepared the cryo-EM samples, collected the EM data and processed data

   Contributed equally with

   Stella Vitt

   Competing interests

   The authors declare that no competing interests exist

2. Stella Vitt

   Max Planck Institute for Terrestrial Microbiology, Marburg, Germany

   Contribution

   SV, Purified the protein, interpreted the structure and wrote the paper with contributions from all authors

   Contributed equally with

   Deryck J Mills

   Competing interests

   The authors declare that no competing interests exist

3. Mike Strauss

   Department of Structural Biology, Max Planck Institute of Biophysics, Frankfurt, Germany

   Present address

   Department of Biological Chemistry and Molecular Pharmacology, Harvard Medical School, Boston, MA, United States

   Contribution

   MS, Processed data

   Competing interests

   The authors declare that no competing interests exist

4. Seigo Shima

   1. Max Planck Institute for Terrestrial Microbiology, Marburg, Germany
   2. PRESTO, Japan Science and Technology Agency, Kawaguchi, Japan

   Contribution

   SS, Initiated the project, interpreted the structure and wrote the paper with contributions from all authors

   Competing interests

   The authors declare that no competing interests exist

5. Janet Vonck

   Department of Structural Biology, Max Planck Institute of Biophysics, Frankfurt, Germany

   Contribution

   JV, Processed the data, built the model, interpreted the structure and wrote the paper with contributions from all authors

   For correspondence

   janet.vonck@biophys.mpg.de

   Competing interests

   The authors declare that no competing interests exist

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