Organohalide respiratory chains: composition, topology and key enzymes

Organohalide respiratory chains: composition, topology and key enzymes Abstract The utilization of halogenated organic compounds as terminal electron acceptors separates the phylogenetically diverse organohalide-respiring bacteria from other respiratory anaerobes that predominantly use nitrate, fumarate, sulfate or oxidized metals. Organohalide respiration is unique in recruiting a cobamide-containing iron–sulfur protein, the extracellular membrane-bound reductive dehalogenase, as terminal reductase in the electron transfer chain. In recent years substantial contributions have been made to the understanding of how electron transfer paths couple mechanistically to chemiosmosis in the organohalide-respiring bacteria. The structural analysis of a respiratory and a non-respiratory reductive dehalogenase revealed the intramolecular electron transfer via two cubane iron–sulfur clusters to the cobamide at the active site. Based on whether quinones are involved, two types of intermolecular electron transfer chains have been identified, which differ in their composition and mode of proton translocation. Indeed, various respiratory chain architectures have been unraveled and evidence for different putative coupling mechanisms presented. The identification of a multienzyme respiratory complex that combines uptake hydrogenase, a complex iron–sulfur molybdoenzyme and a reductive dehalogenase in Dehalococcoides mccartyi strain CBDB1 has raised new questions regarding the mode of energy conservation in these enigmatic microbes. In this mini-review, we highlight these findings and provide an outlook on potential future developments. anaerobic respiration, reductive dehalogenation, chemiosmosis, organohalides, reductive dehalogenase, cobamides INTRODUCTION Halogenated organic compounds (organohalides) are produced by geogenic, biological or synthetic processes (Gribble 2003; IARC 2014, 2016, 2017) and are present in various anoxic habitats as energy and carbon sources for the growth of anaerobic microbes. Several biological transformations that lead to the degradation of organohalides under anoxic conditions have been identified (Janssen, Pries and van der Ploeg 1994; Fetzner 1998), among them microbial reductive dehalogenation (Bouwer and McCarty 1983a,b; Horowitz, Suflita and Tiedje 1983). The latter process can precede the complete biological mineralization of organohalides in oxygen-depleted zones under methanogenic, iron-, sulfate-, or nitrate-reducing conditions (Monserrate and Häggblom 1997; Lohner and Spormann 2013; Tiedt et al. 2016) or at oxic–anoxic interfaces (Kurt, Mack and Spain 2014; Atashgahi et al.2017; Weatherill et al. 2018), where it can be coupled to oxic degradation processes. Taking into account the ubiquity and diversity of naturally occurring or man-made organohalides, the study of biological reductive dehalogenation is of ecological and environmental relevance. The exploration of how organohalides are used as energy or carbon source is important in terms of ecosystem remediation but is also essential for a complete understanding of microbial metabolic interactions in the environment. Particular anaerobic bacteria that carry out reductive dehalogenation have attracted attention because of their enigmatic mode of energy conservation, which often relies on respiration with hazardous halogenated chemicals (Holliger and Schumacher 1994; Holliger, Wohlfarth and Diekert 1998). This mini-review summarizes the advances made in recent years to unravel the molecular basis of microbial reductive dehalogenation. It also describes how this process is coupled to energy conservation via chemiosmosis. Desulfomonile tiedjei, a sulfate-reducer, was the first isolate observed to grow by coupling the oxidation of formate or hydrogen to the reduction of 3-chlorobenzoate (Shelton and Tiedje 1984; Dolfing and Tiedje 1987; DeWeerd et al. 1990; DeWeerd and Suflita 1990; Mohn and Tiedje 1990). This finding revealed the existence of a so-far-unknown respiratory process whereby reductive dehalogenation of organohalides is coupled to a chemiosmotic mechanism (organohalide respiration) (Louie and Mohn 1999). Meanwhile, various phylogenetically diverse bacteria (including Chloroflexi, Firmicutes, Beta-, Delta- and Epsilonproteobacteria; summarized in Maphosa, de Vos and Smidt 2010) have been identified that conserve energy using organohalides as terminal electron acceptors of membrane-associated electron transfer chains. These microorganisms are termed organohalide-respiring bacteria (OHRB) (Maphosa, de Vos and Smidt 2010; Hug et al. 2013; Adrian and Löffler 2016; Jugder et al. 2016; Atashgahi, Häggblom and Smidt 2017). OHRB can be divided into those that rely exclusively on organohalide respiration for energy conservation (obligate OHRB) and those that can alternatively use non-halogenated electron acceptors (versatile OHRB) including nitrate, fumarate, sulfate and oxidized metal ions (Maphosa, de Vos and Smidt 2010; Hug et al. 2013; Jugder et al. 2016). Selected versatile OHRB may even use oxygen as acceptor under microoxic conditions (Sanford, Cole and Tiedje 2002; Goris et al. 2014; Gadkari et al. 2018). The unifying feature of the diverse OHRB is the presence of reductive dehalogenases (RDases), which are located at the outer face of the cytoplasmic membrane and function as terminal reductases in membrane-associated electron transfer chains (Hug et al. 2013; Jugder et al. 2016; Fincker and Spormann 2017). However, the presence of such an exoplasmic and membrane-bound RDase does not necessarily indicate that the respective organism is able to perform organohalide respiration, especially when complex electron donors are utilized (e.g. pyruvate or lactate). In such cases, a fermentative lifestyle based on energy conservation via substrate level phosphorylation, with the organohalides serving exclusively as an electron sink (facilitated fermentation), is feasible (van de Pas et al. 2001) and needs to be ruled out before a respiratory lifestyle is ascertained. Furthermore, non-respiratory, cytoplasmically located RDases have been described (Chen et al. 2013; Payne et al. 2015). These enzymes ‘prime’ organohalides by the removal of halogen substituents for the subsequent degradation of the carbon backbone. As the name suggests, non-respiratory RDases are not involved in energy conservation via electron-transport phosphorylation. Hence, biological reductive dehalogenation by microbes is not necessarily linked to chemiosmosis. During respiratory RDase-mediated catalysis the net transfer of two electrons onto a (poly-)halogenated substrate results in the release of a chloride, a bromide or an iodide ion (hydrogenolysis). Alternatively, two halide ions can be released in a β-elimination reaction (dihaloelimination/vicinal reduction). Dehalogenation of fluorinated organohalides by a RDase has not been observed to date. The extraordinary stability of the carbon–fluorine bond represents a substantial obstacle to reductive cleavage. The standard redox potentials (E0′) of organohalides range between +240 and +580 mV (Dolfing and Janssen 1994; Dolfing and Novak 2015), which indicates that they are highly suitable electron acceptors for anaerobic respiration, and in terms of thermodynamics are comparable to nitrate with E0′ (NO3−/NO2−) = +433 mV (Thauer, Jungermann and Decker 1977). However, the low H+/e− ratio of about 1 measured for H2-dependent reduction (E0′ (H+/H2) = −414 mV) of tetrachloroethene (PCE) by Dehalobacter restrictus (Schumacher and Holliger 1996) and of 3-chlorobenzoate by D. tiedjei (Louie and Mohn 1999) suggests that either mechanistic or thermodynamic restrictions hinder a complete exploitation of the free energy available in the reaction: $$\rm{C_{2}Cl_{4} + H_{2}\rightarrow C_{2}HC_{3}+H^{+}+Cl^{-} \quad \Delta {\rm G}^{0'}=-189 \, kJ \, mol^{-1}H_{2}}$$ (Holliger, Wohlfarth and Diekert 1998) $$\rm{3-Cl-benzoate+H_{2}\rightarrow benzoate+H^{+}+Cl^{-} \quad \Delta {\rm G}^{0'}=-125 \, kJ \, mol^{-} \, H_{2}}$$ (Dolfing and Tiedje 1987) Taking into consideration a potential difference of approximately 100–200 mV, which is required for the translocation of 1 mol H+ across the cell membrane (Thauer, Jungermann and Decker 1977; Simon, van Spanning and Richardson 2008), the low H+/e−ratio falls substantially below the maximal theoretical yield. The formation of ATP from ADP plus Pi consumes 3–4 protons, depending on the architecture of the ATP synthase (Weber and Senior 2003; Mayer and Müller 2014; Silverstein 2014). Hence, between one-third and two-thirds of an ATP is formed per halide ion released in organohalide respiration. This is also supported by the low growth yields of S. multivorans (1.4 g cell protein per mole chloride released) when growing on hydrogen plus tetrachloroethene (Scholz-Muramatsu et al. 1995) and of an enrichment culture of D. restrictus (2.3 g cell protein per mole chloride released) (Holliger et al. 1993). Both the molecular mechanism and the structural basis for the proton translocation in organohalide respiratory chains remain to be elucidated. However, based on recent genomic and proteomic studies (summarized in Türkowsky et al. 2018 in this Special Issue), it is becoming increasingly clear that a uniform organization of the electron transfer chains in the diverse OHRB does not exist. Rather, diverse modes of coupling reductive dehalogenation to proton translocation seem to have evolved. In addition, based on the 3D-structure of a respiratory RDase (Bommer et al. 2014), as well as that of a non-respiratory RDase (Payne et al. 2015), novel insights have emerged regarding both intramolecular electron transfer and the catalytic mechanism of RDases. REDUCTIVE DEHALOGENASES During the last two decades several hundred entries of RDase protein sequences have been deposited in databases (Hug et al. 2013). Although overlapping substrate ranges have been identified, the sequence variability among RDases is high. For example, the respiratory tetrachloroethene RDase (PceASmul) of the Gram-negative epsilonproteobacterium S. multivorans (Neumann et al. 1996) and PceAY51 of the Firmicute Desulfitobacterium hafniense strain Y51 (Suyama et al. 2002) share only 27% amino acid sequence identity, yet both enzymes efficiently convert PCE. Orthologous groups of RDases with elevated sequence similarity can be identified (Hug et al. 2013), but due to the lack of their detailed biochemical characterization, a correlation of these groups with substrate classes or catalytic mechanisms is difficult to define (for examples see Bommer et al. 2014; Parthasarathy et al. 2015; Payne et al. 2015; Alfán-Guzmán et al. 2017; Kunze et al. 2017; Kunze, Diekert and Schubert 2017). RDases are metalloproteins and all representatives analyzed so far, with one exception, contain a cobamide. The exception is the 3-chlorobenzoate-converting RDase from D. tiedjei, which was reported to contain a heme cofactor (Ni, Fredrickson and Xun 1995). Cobalt-, iron-, and nickel-containing porphyrinoids share the ability to catalyze abiotic reductive dehalogenation (Schrauzer, Deutsch and Windgassen 1968; Wood, Kennedy and Wolfe 1968; Krone et al. 1989; Krone, Thauer and Hogenkamp 1989; Gantzer and Wackett 1991), but the identification of a heme rather than a cobamide as cofactor in a RDase remains unique. Further biochemical analyses are needed to define the role of alternative transition metal-containing porphyrinoids in enzymatic reductive dehalogenation. It is also currently unknown whether RDase enzymes have evolved from a single progenitor. The similarity of structural features within the nitroreductase fold that binds the cobamide cofactor at the core of RDases (Bommer et al. 2014; Payne et al. 2015) indicates an evolutionary conservation of common traits. However, a conserved motif for the binding of the cobamide by RDases has not been identified. Besides the well-characterized adenosyl-cobamide-dependent enzymes (mutases and eliminases) or the methyl-cobamide-dependent methyltransferases, RDases form a novel class of cobamide-dependent enzymes together with the queuosine biosynthetic enzyme QueG (Bridwell-Rabb and Drennan 2017). RDase sequences have been identified in archaea belonging to the Asgard superphylum (Zaremba-Niedzwiedzka et al. 2017), which suggests that RDase structural elements might have evolved prior to the division of bacteria and archaea. Substrate range Information on the mode of catalysis and the substrate range of RDases has been obtained either from the characterization of purified proteins or via enzyme activity measurements performed with RDases enriched on native polyacrylamide gels (Adrian et al. 2007b; Tang et al. 2013) or produced heterologously (Mac Nelly et al. 2014; Parthasarathy et al. 2015; Payne et al. 2015; Kunze, Diekert and Schubert 2017; Jugder et al. 2018). Before conclusions on the mechanism or substrate range of a given RDase can be drawn, cobamide-mediated abiotic conversion of organohalides (Schrauzer, Deutsch and Windgassen 1968; Wood, Kennedy and Wolfe 1968; Krone, Thauer and Hogenkamp 1989; Gantzer and Wackett 1991) has to be excluded. Whether the protein environment of the substrate-binding site has a direct impact on the conversion of a given substrate must be tested with heat-inactivated samples, as has been demonstrated for the dihaloelimination of 1,2-dibromoethane or 1,1,2,2-tetrachloroethane (Kunze, Diekert and Schubert 2017). Two general modes of RDase function have been documented: (i) hydrogenolysis/halogen substitution and (ii) vicinal reduction/dihaloelimination (Fig. 1A). While hydrogenolysis involves a proton transfer in the catalytic cycle, recent findings indicate that a proton might be dispensable for the dihaloelimination reactions catalyzed by RDases (Franke et al. 2017; Kunze, Diekert and Schubert 2017). Information is available about the involvement of RDases specialized for halogen substitution in organohalide respiratory chains (see below), but less is known about the coupling of dihaloeliminating enzymes to chemiosmosis in organohalide-respiring bacteria (De Wildeman et al. 2003; Moe et al. 2009; Bowman et al. 2013; Ding, Zhao and He 2014; Tang et al. 2016; Wong et al. 2016; Key et al. 2017). This area requires intensive future investigation. Figure 1. View largeDownload slide RDase catalysis and substrate range. (A) RDases that are known to catalyze both halogen substitution and dihaloelimination. (B) Selected substrates converted by PceASmul from S. multivorans. PCE, tetrachloroethene; TCE, trichloroethene; cDCE, cis-dichloroethene; VC, vinyl chloride; 1,2-DCA, 1,2-dichloroethane; 1,1,2-TCA, 1,1,2-trichloroethane; TBE, tribromoethene; 1,1-DBE, 1,1-dibromoethene; tDBE, trans-dibromoethene; cDBE, cis-1,2-dibromoethene; t-1,3-DCP, trans-1,3-dichloropropene; t-1-CP, trans-1-chloropropene; c-1-CP, cis-1-chloropropene; 3-CP, 3-chloropropene; 2,4,6-TBP, 2,4,6-tribromophenol; 2,4-DBP, 2,4-dibromophenol. Figure 1. View largeDownload slide RDase catalysis and substrate range. (A) RDases that are known to catalyze both halogen substitution and dihaloelimination. (B) Selected substrates converted by PceASmul from S. multivorans. PCE, tetrachloroethene; TCE, trichloroethene; cDCE, cis-dichloroethene; VC, vinyl chloride; 1,2-DCA, 1,2-dichloroethane; 1,1,2-TCA, 1,1,2-trichloroethane; TBE, tribromoethene; 1,1-DBE, 1,1-dibromoethene; tDBE, trans-dibromoethene; cDBE, cis-1,2-dibromoethene; t-1,3-DCP, trans-1,3-dichloropropene; t-1-CP, trans-1-chloropropene; c-1-CP, cis-1-chloropropene; 3-CP, 3-chloropropene; 2,4,6-TBP, 2,4,6-tribromophenol; 2,4-DBP, 2,4-dibromophenol. Although halogen substitution and dihaloelimination were both presumed to be catalyzed by distinct types of RDases, actual case reports show that, depending on the substrate, a single enzyme can perform both reactions (Fig. 1A). For example, the vinyl chloride (VC) RDase (VcrA) from D. mccartyi strain VS has been found to mediate the reductive halogen substitution at VC and forms ethene as well as catalyzing the dihaloelimination of 1,2-dichloroethane (1,2-DCA, synonym: ethylene dichloride, EDC) to ethene (Parthasarathy et al. 2015). The conversion rate with 1,2-DCA was only two-fold lower compared with the rate measured with VC as an electron acceptor. Moreover, the heterologously produced DcaA of Desulfitobacterium dichloroeliminans catalyzed the dihaloelimination of 1,2-DCA but also mediated PCE conversion to trichloroethene (TCE) and finally to cis-1,2-dichloroethene (cDCE) via hydrogenolysis (Kunze, Diekert and Schubert 2017). Here, the activity was 10-fold lower with PCE compared with 1,2-DCA. More recently it was shown that 1,1,2-trichloroethane (1,1,2-TCA) was converted to a mixture of vinyl chloride and 1,2-DCA by the trichloromethane RDase (TmrA) of Dehalobacter sp. strain UNSWDHB (Jugder et al.2017). In this case both mechanisms, halogen substitution and dihaloelimination, appeared to be applied by a single enzyme to the same substrate. Nevertheless, it still must be determined in every case whether all enzyme substrates identified for a single RDase allow coupling of the reductive dehalogenation to chemiosmosis in the respective organohalide-respiring bacterium. Besides numerous tests on substrate promiscuity of RDases, systematic analyses of the ability of RDases to convert halogenated aliphatic and aromatic substrates are needed to understand fully the flexibility of these enzymes. Furthermore, the influence of heteroatoms in aromatic substrates and the impact of flanking substituents on the reductive dehalogenation have to be tested more intensively (Cooper et al. 2015; Zhang et al. 2017). An example of an OHRB that shows enzymatic conversion of a structurally diverse set of organohalides is D. mccartyi strain CBDB1. The substrate range of this organism encompasses different congeners of polychlorinated biphenyls (Adrian et al. 2009), chlorinated dioxins and other aromatic compounds such as halogenated benzenes, phenols, anilines, or benzonitriles (Adrian et al. 2007a; Wagner et al. 2012; Cooper et al. 2015). Aliphatic organohalides, e.g. chlorinated ethenes, are also converted by D. mccartyi strain CBDB1 (Fung et al. 2007; Marco-Urrea, Nijenhuis and Adrian 2011). Whether a single enzyme of the 32 RDase complement encoded in the genome of D. mccartyi strain CBDB1 (Kube et al. 2005) is responsible for all the conversions listed above is not known. However, it has to be assumed that a single enzyme converts structurally different substrates, since the organism's substrate range is independent of the growth substrate that might induce the production of different RDases. Furthermore, it appears feasible that the conversion of aliphatic substrates by a given RDase does not preclude the dehalogenation of aromatic compounds and vice versa. Detailed information on the substrate range of PceASmul from S. multivorans is available (Neumann et al. 2002; Schmitz et al. 2007). This enzyme was initially thought to convert only aliphatic substrates such as chlorinated ethenes or propenes (Fig. 1B), but subsequent studies showed the efficient dehalogenation of bromoethenes and bromophenols (Ye et al. 2010; Kunze et al. 2017). Another example is RdhA3 of D. hafniense DCB-2, a RDase that displays a preference for the conversion of chlorophenols; however, it also dehalogenated PCE at lower rates (Mac Nelly et al. 2014). Such observations suggest a hidden potential of RDases to dehalogenate structurally diverse substrates and these results should be taken into account when the substrate range of newly discovered RDase is investigated. The RDase active-site cavity is connected via a hydrophobic substrate channel to the solvent (Bommer et al. 2014; Payne et al. 2015). Most of the amino acids involved in forming the channel are part of the enzyme's N-terminus, which in general shows little sequence conservation. This diversity might reflect an adaptation of the various RDases to the huge variety of halogenated substrates and increased amino acid sequence identity in this region might indicate an overlapping substrate range. However, a high overall sequence similarity does not necessarily mean that two RDases share the same substrate range or form the same products (Buttet, Holliger and Maillard 2013; Alfan-Guzman et al. 2017; Kunze, Diekert and Schubert 2017). Predicting the substrate preferences of a given RDase based on sequence comparison or structural modeling is hampered by the fact that even minor changes in the RDase architecture modulate substrate preference (Kunze, Diekert and Schubert 2017). For example, PceASmul of S. multivorans reduces PCE via TCE to cDCE (Neumann et al. 1996) and it shares this property with the highly similar PceAShal from Sulfurospirillum halorespirans (92% amino acid sequence identity for the processed enzyme without the twin arginine translocation signal peptide) (Luijten et al. 2003; Goris et al. 2017). In the enriched Sulfurospirillum sp. SL-2, a PceA variant, termed PceA (TCE), was identified that also shares 92% amino acid sequence identity with PceASmul but converts PCE only to TCE (Buttet, Holliger and Maillard 2013). A structural alignment of the PceASmul crystal structure and an in silico structural model of PceA (TCE) identified slight modifications in the architecture of the active site cavity that might be responsible for the difference in substrate range (Fig. 2). Figure 2. View largeDownload slide Comparison of PceA enzymes from Sulfurospirillum spp. (A) Multiple sequence alignment of PceASmul of S. multivorans (accession no. AHJ12791.1), PceAShal of S. halorespirans (acc. no. AOO65270.1) and PceA (TCE) of Sulfurospirillum sp. SL-2 (acc. no. ARU48750.1). Twin arginine translocation (Tat) signal peptides were not included in the alignment. Amino acid residues involved in the formation of the active site cavity are labeled in grey, while amino acid exchanges are highlighted in red. The cysteines binding the two cubane iron–sulfur clusters are marked in yellow. (B) In silico structural model of the active site cavity of PceA (TCE) from Sulfurospirillum sp. SL2 (petrol) overlaid with the structure of PceASmul (gray) (Bommer et al. 2014) depicted in two orientations. The structural model (C-score: 0.97) was generated with the I-TASSER server for protein structure and function prediction (Zhang 2008; Yang et al. 2015). The residues that differ between the enzymes are shown in red frames. The two orientations identified for TCE at the active site of PceASmul are depicted in an overlay. The red sphere between the cobamide cofactor and the substrate represents a water or hydroxyl group. Figure 2. View largeDownload slide Comparison of PceA enzymes from Sulfurospirillum spp. (A) Multiple sequence alignment of PceASmul of S. multivorans (accession no. AHJ12791.1), PceAShal of S. halorespirans (acc. no. AOO65270.1) and PceA (TCE) of Sulfurospirillum sp. SL-2 (acc. no. ARU48750.1). Twin arginine translocation (Tat) signal peptides were not included in the alignment. Amino acid residues involved in the formation of the active site cavity are labeled in grey, while amino acid exchanges are highlighted in red. The cysteines binding the two cubane iron–sulfur clusters are marked in yellow. (B) In silico structural model of the active site cavity of PceA (TCE) from Sulfurospirillum sp. SL2 (petrol) overlaid with the structure of PceASmul (gray) (Bommer et al. 2014) depicted in two orientations. The structural model (C-score: 0.97) was generated with the I-TASSER server for protein structure and function prediction (Zhang 2008; Yang et al. 2015). The residues that differ between the enzymes are shown in red frames. The two orientations identified for TCE at the active site of PceASmul are depicted in an overlay. The red sphere between the cobamide cofactor and the substrate represents a water or hydroxyl group. Heterologous production The low growth yields of OHRB and the toxicity of their halogenated substrates limit the amount of biomass available to deliver sufficient amounts of enzyme for biochemical studies. Thus, only a few RDases have been purified and biochemically characterized to date (summarized in Hug et al. 2013; Lu et al. 2015; Jugder et al. 2016; Schubert and Diekert 2016; Fincker and Spormann 2017). Given the fact that RDases display high amino acid sequence variability and diverse substrate ranges, the results obtained from this group of selected enzymes might only provide limited insight into RDase substrate specificity. Functional heterologous production of catalytically active RDases is complicated because of the need for cobamides and Fe–S clusters, and their synthesis and insertion often hamper such approaches. Nevertheless, the PCE-converting PceAY51 of D. hafniense Y51, the chlorophenol-dehalogenating RdhA3 of D. hafniense DCB-2, the 1,2-DCA-dihaloeliminating DcaA of D. dichloroeliminans, the chloroform-dehalogenating TmrA of Dehalobacter sp. strain UNSWDHB, and the non-respiratory NpRdhA of N. pacificus were functionally produced in a heterologous host (Mac Nelly et al. 2014; Payne et al. 2015; Kunze, Diekert and Schubert 2017; Jugder et al. 2018). Successful reconstitution of a catalytically active RDase from the apoprotein has been described only once for the vinyl chloride RDase (VcrA) from Dehalococcoides mccartyi strain VS (Parthasarathy et al. 2015). Cobamide cofactor The demand for cobamides (i.e. corrinoids with a complete nucleotide loop) as essential cofactors of the key enzyme of organohalide respiration is satisfied differently by the various OHRB. Cobamide producers such as S. multivorans (Goris et al. 2014; Keller et al. 2014), D. hafniense Y51 (Nonaka et al. 2006; Reinhold et al. 2012), or Dehalobacter sp. strains TCA1, CF, DCA (Sun et al. 2002; Tang et al. 2016; Wang et al. 2017) and UNSWDHB (Desphande et al. 2013; Wong et al. 2016) have been identified. In addition, cobamide auxotrophs have been isolated, e.g. Dehalobacter restrictus (Rupakula et al. 2013) and Dehalococcoides mccartyi (Löffler et al. 2013), that have to take up cobamides from the environment. The cobamides produced by the OHRB, which are able to perform de novo biosynthesis, differ in the structure of their nucleotide loop (Kräutler et al. 2003; Yan et al. 2018). The nucleotide loop of the norpseudo-B12 produced by S. multivorans lacks a methyl group at position 176 and harbors an adeninyl moiety as the lower base (Fig. 3A) (Kräutler et al. 2003). Whether the lack of the methyl group represents a specific adaptation to the cofactor requirements of PceASmul is not definitively resolved (Bommer et al. 2014; Keller et al. 2016; Schubert 2017). Recently, the structure of the cobamide produced by D. hafniense strains was elucidated and shown to differ from vitamin B12 (Fig. 3B) by the presence of unsubstituted purine as lower base (Yan et al. 2018). Figure 3. View largeDownload slide Cobamide structures. (A) The norpseudo-B12 is depicted in the base-off conformation with a hydroxyl group as upper ligand. (B) Vitamin B12 is shown in base-on conformation with the artificial cyano-group as upper ligand (alternative name cyanocobalamin). DMB = 5,6-dimethylbenzimidazole (C) Alternative norcobamides produced by S. multivorans in the presence of exogenous benzimidazoles. The functional utilization of these norcobamides as cofactors of PceASmul is indicated by the colored circles (green, functional; orange, reduced functionality; red, substantial impairment) (data from Keller et al. 2018). (D) Alternative cobamides amended to cultures of D. mccartyi strain 195 cultivated with TCE. The functional utilization of these cobamides as cofactors in organohalide respiration is indicated by colored circles (green, functional; red, substantial impairment) (data from Yi et al. 2012). In addition, 2-methylmercaptoadeninyl-cobamide (not shown) was tested with D. mccartyi strain 195, but did not functionally replace vitamin B12. Figure 3. View largeDownload slide Cobamide structures. (A) The norpseudo-B12 is depicted in the base-off conformation with a hydroxyl group as upper ligand. (B) Vitamin B12 is shown in base-on conformation with the artificial cyano-group as upper ligand (alternative name cyanocobalamin). DMB = 5,6-dimethylbenzimidazole (C) Alternative norcobamides produced by S. multivorans in the presence of exogenous benzimidazoles. The functional utilization of these norcobamides as cofactors of PceASmul is indicated by the colored circles (green, functional; orange, reduced functionality; red, substantial impairment) (data from Keller et al. 2018). (D) Alternative cobamides amended to cultures of D. mccartyi strain 195 cultivated with TCE. The functional utilization of these cobamides as cofactors in organohalide respiration is indicated by colored circles (green, functional; red, substantial impairment) (data from Yi et al. 2012). In addition, 2-methylmercaptoadeninyl-cobamide (not shown) was tested with D. mccartyi strain 195, but did not functionally replace vitamin B12. Preferences of some RDases for specific lower bases within cobamide cofactors have been observed and alterations in cobamide structure, i.e. exchange of the lower ligand base, interferes with RDase function (Fig. 3C and D) (Yan et al. 2012; Yi et al. 2012; Yan et al. 2013; Keller et al. 2014; Yan et al. 2016; Keller et al. 2018). The structural or mechanistic reasons for these cofactor incompatibilities have not been unraveled yet. Solving the three-dimensional structure of PceASmul of S. multivorans and the non-respiratory bromophenol RDase (NpRdhA) of the marine Nitratireductor pacificus pht-3B, uncovered the mode of cobamide binding (Bommer et al. 2014; Payne et al. 2015). In both RDases the cobamide is bound deep within the protein, with the nucleotide loop oriented away from the corrin ring. Several hydrogen bonds between the nucleotide loop and the protein environment indicate a role for the cobamide's tail in cofactor positioning. Hence, negative effects on RDase activity of structural variations in the nucleotide loop of the cobamide cofactor might be based on a selectivity in cofactor incorporation rather than resulting from direct interference of the nucleotide loop structure with the catalytic cycle or substrate range. So far, no evidence for an involvement of the cobamide cofactor's nucleotide loop in the catalytic cycle of RDases has been found (Bommer et al. 2014; Payne et al. 2015). Based on the static PceASmul structure it was not possible to identify a barrier that prevents the incorporation of distinct cobamides into the enzyme. Consequently, an incompatibility of the cobamide structure and the enzyme's folding process has been suggested (Keller et al. 2018). ELECTRON TRANSFER WITHIN REDUCTIVE DEHALOGENASE ENZYMES Evidence has been presented that the super-reduced state [CoI] of the cobalt ion in the cobamide is the reactive form of the cofactor that attacks the organohalide (Fig. 4) (Miller, Wohlfarth and Diekert 1996; Schumacher et al. 1997; van de Pas et al. 1999; Parthasarathy et al. 2015; Kunze et al. 2017). Several purified RDases have been studied by electron paramagnetic resonance (EPR) spectroscopy and shown to harbor a cobamide cofactor in the [CoII] oxidation state (see below). This ‘as isolated’ state appears not to be reactive. Low potential electron donors, e.g. reduced methyl viologen (E0′ = −446 mV), are required for the conversion of the substrate in the enzymatic assay (Miller, Wohlfarth and Diekert 1996). This necessity is indicative of the need for the [CoI] oxidation state to attack the organohalide. The electrons provided by artificial electron donors are probably transferred via the two Fe–S clusters to the cobamide cofactor. The efficient inhibition of PceASmul by 1-iodopropane (propyl iodide) supports the assumption that [CoI] functions as the reactive oxidation state (Neumann, Wohlfarth and Diekert 1995). Based on the active site architecture of PceASmul and NpRdhA (Bommer et al. 2014; Payne et al. 2015) and the positioning of the substrate (Kunze et al. 2017), a nucleophilic attack of the [CoI] on the electron-deficient carbon backbone of the substrate is unlikely. However, this mechanism was reported earlier for the abiotic reductive dehalogenation mediated by protein-free cobamides (Fig. 4A) (Schrauzer, Deutsch and Windgassen 1968; Wood, Kennedy and Wolfe 1968; Krone, Thauer and Hogenkamp 1989). Proposals for the initial attack on the enzyme-bound substrate include either a long-range electron transfer from the cobalt to the substrate (Fig. 4B) (Schmitz et al. 2007; Bommer et al. 2014; Kunze et al. 2017), or a direct reaction of [CoI] with a halogen substituent (Fig. 4C) (Payneet al. 2015; Cooperet al. 2015; Johannissen, Leys and Hay 2017). The mechanistic diversity of cobamide-mediated enzymatic and abiotic reductive dehalogenation have been subject to stable isotope fractionation analyses, which were reviewed previously and are not discussed here (Elsner 2010; Nijenhuis et al. 2016). Figure 4. View largeDownload slide Proposed modes for the initial attack of the super-reduced [CoI] onto the halogenated RDase substrate. (A) Nucleophilic attack of [CoI] at the electron-deficient carbon backbone of the organohalide results in the alkylation of the cobalt. (B) Substrate radical formation via long-range electron transfer. The proposed subsequent formation of a carbanion after the elimination of the halogen substituent is not depicted. (C) [CoI] attacks directly at the halogen substituent and the carbon-halogen bond is either cleaved heterolytically (left) or homolytically (right). [CoI-II], oxidation states of the cobalt ion in the cobamide cofactor; R, hydrocarbon backbone; X, halogen substituent (adapted from Kunze et al. 2017). Figure 4. View largeDownload slide Proposed modes for the initial attack of the super-reduced [CoI] onto the halogenated RDase substrate. (A) Nucleophilic attack of [CoI] at the electron-deficient carbon backbone of the organohalide results in the alkylation of the cobalt. (B) Substrate radical formation via long-range electron transfer. The proposed subsequent formation of a carbanion after the elimination of the halogen substituent is not depicted. (C) [CoI] attacks directly at the halogen substituent and the carbon-halogen bond is either cleaved heterolytically (left) or homolytically (right). [CoI-II], oxidation states of the cobalt ion in the cobamide cofactor; R, hydrocarbon backbone; X, halogen substituent (adapted from Kunze et al. 2017). RDases in the ‘as isolated’ form studied by EPR spectroscopy showed the cobalt ion to be in the [CoII] oxidation state (Schumacher et al. 1997; van de Pas et al. 1999; Kräutler et al. 2003; Parthasarathy et al. 2015; Payne et al. 2015; Kunze et al. 2017). The midpoint redox potential of the couple [CoII]/[CoI] in the enzyme-bound cofactor has been determined for the PCE-RDase of D. restrictus (PceADres; −350 ± 20 mV, pH 8.0; Schumacher et al. 1997), PceASmul (−380 mV, pH 7.5; Kräutler et al. 2003)and the chlorophenol RDase of Desulfitobacterium dehalogenans (−370 mV, pH 7.8; van de Pas et al. 1999). The two [4Fe–4S] clusters of PceADres were found to have more negative midpoint redox potentials of about −480 mV (Schumacher et al. 1997), which is commensurate with an unimpeded electron transfer to the cobamide at the active site. A similar value (approximately −440 mV) was obtained for the Fe–S clusters in PceASmul, when an EPR-coupled redox titration of the two cubane iron–sulfur clusters was carried out (C. Kunze and F. Hagen, unpublished results); purified and catalytically active PceASmul enzyme was used in this experiment. The enzyme fraction contained mainly the mature form of PceASmul. The result of the redox titration of PceASmul was in accord with the onset potential (−450 mV versus standard hydrogen electrode at pH 7) needed to initiate reductive dehalogenation of PCE and TCE by PceASmul bound to a redox electrode (Siritanaratkul et al. 2016). Based on the topology of the redox-active metal centers inside the RDase, it was hypothesized that the electrons delivered by an external donor are conducted via a short ‘electronic wire’ comprising both [4Fe–4S] clusters and the cobamide (Bommer et al. 2014; Payne et al. 2015). The presence of two [4Fe–4S] clusters has also been confirmed for VcrA of D. mccartyi VS and NpRdhA, but their respective redox potentials have not been determined (Parthasarathy et al. 2015; Payne et al. 2015). Although the Fe–S cluster binding motifs in CprA of D. dehalogenans indicate the presence of two cubane clusters, a [4Fe–4S] cluster with a midpoint redox potential of −440 mV and one with a [3Fe–4S] cluster with a positive redox potential of +70 mV were identified (van de Pas et al. 1999). The role of the latter cluster in the reduction reaction remains to be determined. The corrin ring of the cobamide cofactor defines the base of the active site cavity, which is mainly lined by hydrophobic amino acids (Fig. 2B). A network of hydrogen bonds positions the cofactor in RDases. The cobamide cofactor is bound in the base-off conformation in all RDases analyzed so far by EPR spectroscopy or X-ray crystallography (Schumacher et al. 1997; van de Pas et al. 1999; Bommer et al. 2014; Payne et al. 2015; Keller et al. 2018). The term ‘base-off’ describes the absence of a coordinative bond between the lower base and the central cobalt ion (Fig. 3A). The reduction of the cobalt ion in protein-free cobamides from [CoII] to the [CoI] ion is accompanied by a structural conversion from the base-on to the base-off conformation. The protein environment in PceASmul and NpRdhA fixes the cobamide cofactor in the base-off conformation with no alternative ligand bound to the cobalt and it prevents a switch from the base-off to the base-on state. This feature lends support to the hypothesis that the protein environment influences the reduction of the cobalt ion in the RDase-bound cofactor and this is reflected in the elevated midpoint redox potential of the RDase-bound cofactor compared with the protein-free form (e.g. −490 mV vs. standard hydrogen electrode at pH 7 for the protein-free norpseudovitamin B12 compared with −380 mV for the enzyme-bound norpseudo-B12 at pH 7.5; Kräutler et al. 2003). An exception to the rule of base-off binding might have been identified for VcrA of D. mccartyi VS, which was analyzed by EPR and found to incorporate the cobamide cofactor in the base-on conformation with the lower base of the cofactor coordinating the central metal ion directly (Parthasarathy et al. 2015). Redox potentials of the different oxidation states of the cobamide cofactor have not yet been determined in this case. Given the findings that RDases are highly diverse and cobamides display a multifaceted chemistry, alternative modes of cobamide cofactor binding and utilization cannot be excluded. In analogy to the cobamide-binding motif present in other B12-dependent enzymes (Dowling, Croft and Drennan 2012), putative cobamide-binding sequences (DHXG-X39-S-X32-G) were identified in dehalogenating Chloroflexi (Hölscher et al. 2004; Padilla-Crespo et al. 2014). However, as visualized by structure prediction based on the PceASmul and NpRdhA 3D-structures, the histidine residue does not come close to the cobalt to serve as an alternative lower ligand. Most primary amino acid sequences of Chloroflexi RDases form a distinct cluster in phylogenetic trees (Hug et al. 2013; Fincker and Spormann et al. 2017). This distinctiveness might reflect differences in the catalytic mechanism of these enzymes, in their mode of cobamide cofactor binding, or in the composition of the organohalide respiratory chain of these organisms. SUBCELLULAR LOCALIZATION OF RESPIRATORY RDases and the RdhB PROTEINS The localization of the membrane-associated primary electron-donating enzyme and the terminal reductase of a respiratory chain is critical in determining the H+/e− ratio (Simon, van Spanning and Richardson 2008). In general, RDases are synthesized and equipped with cofactors in the cytoplasm (John et al. 2006; Reinhold et al. 2012). Since a characteristic property of the cytoplasmic RDase precursors is the presence of an N-terminal twin-arginine translocation (Tat) (Palmer and Berks 2012) signal peptide, it is assumed that the Tat machinery translocates the enzymes across the cytoplasmic membrane. The findings of early studies, however, proposed a binding of the enzyme to the inner face of the cytoplasmic membrane, mainly based on the results obtained from enzyme activity measurements with intact or permeabilized cells conducted with the membrane-impermeant methyl viologen as an artificial electron donor (Holliger and Schumacher 1994; Miller, Wohlfarth and Diekert 1996, 1997). Later, the same experimental approach applied to D. mccartyi strain 195 led to the conclusion that the RDase in this organism has an exoplasmic localization (Nijenhuis and Zinder 2005). In a subsequent study, the subcellular localization of PceASmul was re-examined via freeze-fracture replica immunogold labeling, which allowed direct detection of the membrane-bound state of the enzyme in cells of S. multivorans (John et al. 2006). This study showed unequivocally that the PceASmul molecules were indeed located on the outer face of the cytoplasmic membrane in PCE-grown cells. A cytoplasmic localization of PceASmul, observed in earlier studies, has been attributed to a reduced transport of the enzyme's precursor in fumarate-grown cells. An exoplasmic localization of PceA was also demonstrated for D. hafniense Y51 (Reinhold et al. 2012), while treatment of D. restrictus with proteinase K resulted in the loss of PceADres from these cells, which also indicated an exoplasmic localization of the RDase in this organism (C. Holliger; personal communication). Hence, it is currently assumed that the physiologically active RDase is located at the outer rather than the inner face of the lipid bilayer. In most cases, a small open reading frame encoding a hydrophobic protein (rdhB) accompanies the structural gene for the respiratory RDase enzyme (rdhA). It was proposed that RdhB functions as a membrane anchor for RdhA (Neumann, Wohlfarth and Diekert 1998). However, the experimental proof for this assumption is still lacking and the formation of the rdhB gene products could not be verified for a long time. Recently, however, the synthesis of the B protein in S. multivorans has been demonstrated by proteomic analysis of PCE-grown cells (Goris et al. 2015). Furthermore, the involvement of the B protein in the formation of a multienzyme organohalide respiratory complex in D. mccartyi was confirmed via peptide mass fingerprinting (Kublik et al. 2016). Although a substantial number of rdhA genes are indeed accompanied by a dedicated rdhB gene, also solitary rdhA genes have been identified. In Dehalogenimonas species, the number of rdhB genes is much lower than the number of rdhA genes, which might indicate that the RdhB protein is dispensable for some RDases in this organism, that a single RdhB can anchor different RdhA proteins, or that other proteins might be involved in RDase membrane binding (Siddaramappa et al. 2012; Molenda, Quaile and Edwards 2015; Key et al. 2016; Yang et al. 2017). Most of the RDases encoded in Dehalogenimonas species have a Tat signal peptide for membrane export, implying they have an exoplasmic localization. Whether all of the rdhA gene products in this organism are functional in energy conservation or have another function remains to be elucidated. Topology calculations of RdhB proteins showed in most cases three transmembrane helices in addition to an exoplasmic N-terminus and a short exoplasmic loop between helices 2 and 3 (Fig. 5A). The corresponding B protein of the PceASmul enzyme represents an exception in having only two transmembrane helices and most probably both termini are exposed to the cytoplasm (Fig. 5B). Since respiratory RDase does not appear to have membrane integral domains that insert into the lipid bilayer, the enzymes must interact with the exoplasmic loops or domains of other integral membrane proteins or membrane-associated proteins. Although the membrane-spanning helices of B proteins share little sequence identity, two conserved glutamic acid residues can be identified in the exoplasmic loop of most RdhBs and these might play a role in the RdhA–RdhB interaction. No typical cofactor-binding site is present in RdhB proteins. Consequently, their direct involvement in the electron transfer to the RDase is unlikely. Whether anchoring the RDase to the membrane is the sole function of the B proteins or whether they also serve as a facilitator between the RDase and other proteins involved in electron transfer is unclear. Figure 5. View largeDownload slide The PceB protein of D. hafniense Y51 and of S. multivorans. (A and B) Predicted topology of PceB in the cytoplasmic membrane (TMHMM Server v. 2.0; Sonnhammer, von Heijne and Krogh 1998) is shown. Figure 5. View largeDownload slide The PceB protein of D. hafniense Y51 and of S. multivorans. (A and B) Predicted topology of PceB in the cytoplasmic membrane (TMHMM Server v. 2.0; Sonnhammer, von Heijne and Krogh 1998) is shown. QUINONE-DEPENDENT ORGANOHALIDE RESPIRATION The electron transfer path in the organohalide respiratory chain from the oxidation of the electron donor to the terminal RDase differs in the various OHRB. The paths can be classified into those using quinones for electron and proton shuttling within the membrane and those possessing a quinone-independent electron transfer chain. In theory, the involvement of quinones in respiratory chains allows a proton transfer via a redox loop mechanism with interaction sites for quinone reduction and quinol oxidation being at opposite sides of the cytoplasmic membrane (Jormakka et al. 2002; Simon, van Spanning and Richardson 2008). In the absence of quinones a conformational proton pump might be responsible for generating the proton gradient (Simon, van Spanning and Richardson 2008; Efremov, Baradaran and Sazanov 2010). The obligate organohalide-respiring D. restrictus was shown to contain different menaquinones (MK; mainly MK-7 and MK-8) (Holliger et al. 1998). The organism harbors a complete set of genes encoding the enzymes of menaquinone biosynthesis (Dairi et al. 2011; Kruse et al. 2013). Comparative spectral analysis of D. restrictus membranes reduced with H2 and oxidized with PCE displayed a clear change in the absorbance spectrum of MK revealing its potential participation in the PCE respiratory chain (Schumacher and Holliger 1996). The reductive dehalogenation of PCE in whole cells of D. restrictus was driven by the addition of reduced 2,3-dimethyl-1,4-naphthoquinone (DMNH2), a water-soluble MK analog, and was blocked in the presence of the quinone antagonist 2-n-heptyl-4-hydroxyquinoline N-oxide (HQNO; 10 nmol mg−1 protein). An inhibitory effect of HQNO at an elevated concentration (150 nmol mg−1 protein) was also found for D. tiedjei (Louie and Mohn 1999) and for S. multivorans (>80 nmol mg−1 protein; Gadkari 2017). Menaquinones have been extracted from S. multivorans (Scholz-Muramatsu et al. 1995) and the genetic information for menaquinone biosynthesis via the alternative futalosine pathway was identified (Goris et al. 2014). The presence of (mena-)quinones has also been confirmed for D. tiedjei (Louie and Mohn 1999) and D. dehalogenans (Kruse et al. 2015). In the latter organism, oxidation of menaquinol (MKH2) by 3-chloro-4-hydroxy-phenylacetate (ClOHPA) was observed. All of these results indicate the involvement of quinones in the intermolecular electron transfer of the listed OHRB. Due to the lack of functional motifs, an interaction of the B proteins with the MK pool in quinone-dependent OHRB is unlikely, which leaves the question of the mode of electron transfer from the MKH2 to the RDase unanswered. Proteome analysis of S. multivorans identified a putative quinol dehydrogenase found exclusively in PCE-grown cells, which might represent the ‘missing link’ in the electron transfer chain (Fig. 6; Goris et al. 2015). The genes for the heterodimeric protein (locus tags: SMUL_1541 and SMUL_1542) are located in proximity to the gene encoding PceASmul (Goris et al. 2014). The two gene products show sequence similarity to the NapGH quinol dehydrogenase involved in electron transfer to the periplasmic nitrate reductase (NapA) (Kern and Simon 2008; Simon and Klotz 2013), but are part of a separate clade based on a phylogenetic analysis of the NapGH-like quinol dehydrogenase protein family (Goris et al. 2014). The localization of RDase genes in proximity to genes encoding a NapGH-like quinol dehydrogenase can also be found in the genome sequence of other dehalogenating Sulfurospirillum species, such as S. halorespirans (Goris et al. 2017) and Sulfurospirillum sp. SL2–1 (RefSeq. NZ_CP021416.1), as well as in the sulfate-reducers D. tiedjei (RefSeq. NC_018025.1), Pseudodesulfovibrio indicus (RefSeq. NZ_CP014206; Cao et al.2016), Desulfovibrio bizertensis (RefSeq. NZ_FUYA01000002), and Desulforhopalus singaporensis (RefSeq. FNJI01000016). However, the analysis of co-expression of these genes with the RDase structural genes still needs to be performed. Figure 6. View largeDownload slide Simplified scheme for the composition of the quinone-dependent organohalide respiratory chain in Sulfurospirillum multivorans. HQNO, 2-n-heptyl-4-hydroxyquinoline N-oxide; DMNH2, 2,3-dimethyl-1,4-naphthoquinone; MNH2, 2-methyl-1,4-naphthoquinone; MKH2, menaquinol; Q-DH, quinol dehydrogenase. The inhibitory effect of HQNO and the electron-donating function of MNH2 are indicated in red and green, respectively. The question mark designates the quinol-oxidizing component, which has not been identified so far. The putative membrane anchor of the RDase (PceB, locus tag: SMUL_1532) is depicted in white, since its interaction with the RdhA protein lacks experimental verification. Figure 6. View largeDownload slide Simplified scheme for the composition of the quinone-dependent organohalide respiratory chain in Sulfurospirillum multivorans. HQNO, 2-n-heptyl-4-hydroxyquinoline N-oxide; DMNH2, 2,3-dimethyl-1,4-naphthoquinone; MNH2, 2-methyl-1,4-naphthoquinone; MKH2, menaquinol; Q-DH, quinol dehydrogenase. The inhibitory effect of HQNO and the electron-donating function of MNH2 are indicated in red and green, respectively. The question mark designates the quinol-oxidizing component, which has not been identified so far. The putative membrane anchor of the RDase (PceB, locus tag: SMUL_1532) is depicted in white, since its interaction with the RdhA protein lacks experimental verification. The quinol-oxidizing reactivity of NapGH homologs has not been verified experimentally and very recent data indicate the involvement of other components in quinol-oxidation (i.e. Rieske/cytochrome bc complex) in the nitrate respiratory chain of the non-dehalogenating Wolinella succinogenes (Hein, Witt and Simon 2017). NapG homologs are characterized by the presence of 16 cysteine residues that are proposed to bind four [4Fe–4S] clusters. The NapG precursor bears an N-terminal Tat signal peptide that identifies the protein as a substrate of the Tat translocase. The mature NapG protein of W. succinogenes was detected in the periplasm (Kern and Simon 2008), where it is expected to interact with the periplasmic nitrate reductase for electron transfer. Assuming that the gene product of SMUL_1541, which shows homology to NapG, interacts with the membrane-attached PceASmul, then the final step in the electron transfer to the RDase proceeds outside the membrane. The iron–sulfur clusters within the membrane-associated quinol dehydrogenase must come into proximity with the Fe–S clusters in PceASmul to allow efficient electron transfer. This assumption has to be taken into consideration when suggestions for the positioning and orientation of PceASmul at the membrane surface are made (Bommer et al. 2014). It is assumed that the SMUL_1541 protein is bound to the outer face of the cytoplasmic membrane by the membrane-integral NapH-like SMUL_1542 gene product. NapH homologues are predicted to contain four transmembrane helices. Eight cysteine residues are conserved in the C-terminal part of the protein (polyferredoxin domain), which is located in the cytoplasm (Brondijk et al. 2004), and these cysteine residues are proposed to bind two [4Fe–4S] clusters. The function of the metal centers in the electron transfer pathway to the periplasmic NapG at the opposite face of the membrane is unknown. Quinone-dependent OHRBs are posed with a thermodynamic problem. The midpoint redox potential of MK (E0′ (MK/MKH2) = −74 mV) (Thauer, Jungermann and Decker 1977) is considerably more positive than the [CoII]/[CoI] redox couple (about −370 mV) of the enzyme-bound cobamide cofactor and the low-potential cubane Fe–S clusters in the RDase. Hence, a driving force is required to convert the thermodynamically unfavorable redox-reaction into a favorable electron transfer. Based on the sensitivity of PCE respiration in S. multivorans to the presence of uncoupling agents such as carbonyl cyanide-p-(trifluoromethoxy)-phenylhydrazone (15 nmol mg−1 protein) a reverse electron flow driven by the proton gradient was concluded as being necessary to overcome the thermodynamic barrier (Miller, Wohlfarth and Diekert 1996). A similar effect resulted when the membrane integrity was perturbed by mild treatment with detergent. An interruption in the electron transfer from hydrogen oxidation to PCE reductive dechlorination by protonophors was not detected in D. restrictus (Schumacher and Holliger 1996) and D. hafniense Y51 (Gadkari 2017). The insensitivity towards uncoupling agents found in the latter two OHRB indicates a different mechanistic solution for facilitating the electron transfer from MKH2 to the low-potential metal centers of the RDase. The distant phylogenetic relationship between the pceC gene product, encoded in the pceABCT operons of D. restrictus (Kruse et al. 2013) and D. hafniense Y51 (Nonaka et al. 2006) and the membrane-integral NapH might indicate a role for PceC in electron transfer comparable to the role proposed for the NapGH-like quinol dehydrogenase in S. multivorans. Low similarity to NosR-like proteins involved in N2O-respiration (Wunsch and Zumft 2005) is also in accord with a role for PceC in electron transfer. PceC is a putative membrane-protein that lacks the polyferredoxin domain present in the C-terminal part of NapH. In contrast, a putative N-terminally bound flavin predicted to be exposed to the exoplasm might form part of the electron transfer chain from MKH2 to the RDase. Soluble, periplasmic flavin-containing proteins in D. dehalogenans might fulfill a similar function (Kruse et al. 2015). This is an interesting proposal, because reduced flavins are able to transfer two electrons at different redox potentials (electron bifurcation; Buckel and Thauer 2013; Peters et al. 2016) and could in theory provide a single low-potential electron for the RDase reaction. So far, there is no concrete evidence to support such a mechanism, but the involvement of flavin radicals or even quinone radicals in electron transfer might be an alternative possibility to circumvent the thermodynamically unfavorable difference in the redox potentials between the quinone pool and the RDase. Since such mechanisms are independent of the proton gradient across the cytoplasmic membrane, uncoupling agents should not have a direct effect. It was shown earlier that the proton gradient appeared to be dispensable for the reductive dehalogenation of ClOHPA coupled to hydrogen oxidation in D. dehalogenans, since the treatment of cells with mild detergents had no effect (van de Pas et al. 2001). Along with the PceC-ortholog CprC in D. dehalogenans, other flavin-containing proteins were also more abundant in cells cultivated with ClOHPA and were discussed as being involved in electron transfer in the ClOHPA respiratory chain (Kruse et al. 2015). Further studies are clearly necessary to unravel the exact role of flavin-containing electron-transferring proteins in cells actively respiring organohalides. Different paths are feasible for the reduction of the MK pool in the quinone-dependent OHRB: (i) the electrons could be transferred directly from a quinone-reactive, membrane-bound dehydrogenase (e.g. membrane-bound uptake hydrogenase or formate dehydrogenase) to the MK pool, or (ii) the electrons could be provided by a soluble dehydrogenase via reduced pyridine nucleotides or soluble electron-transferring proteins, such as ferredoxin/flavodoxin, to a membrane-bound NADH-quinone oxidoreductase or ferredoxin/flavodoxin-quinone oxidoreductase, respectively, which in turn reduces MK. Both options have been discussed to play a role in the PCE respiration of S. multivorans when the organism is cultivated with different electron donors. Based on both transcriptional and biochemical studies, a trimeric membrane-bound [ Ni–Fe] hydrogenase is proposed to be involved in hydrogen oxidation and MK reduction (Kruse et al. 2017). The enriched trimeric membrane-bound [Ni–Fe] hydrogenase, including the quinone-reactive, membrane-integral cytochrome b subunit, was effective in reducing the soluble quinone-analog 2,3-dimethyl-1,4-naphthoquinone (DMN), underpinning the assumption that it reduces the MK pool in H2-grown cells of the organism. When formate serves as electron donor, a membrane-bound formate dehydrogenase might be responsible for MK reduction in S. multivorans (Schmitz and Diekert 2003). During growth on pyruvate and PCE, the electrons derived from pyruvate oxidation by the pyruvate-ferredoxin oxidoreductase were proposed to be transferred via ferredoxin and a ferredoxin-reactive complex I to the MK pool (Goris et al. 2015). QUINONE-INDEPENDENT ORGANOHALIDE RESPIRATION An early study with D. mccartyi strains BAV1 and FL2 reported the presence of quinones in these isolates (White et al. 2005), which suggested a quinone-dependent organohalide respiration in these obligate OHRB. However, D. mccartyi genomes lack the genes for complete biosynthesis of quinones (Kube et al. 2005; Seshadri et al. 2005) questioning the involvement of quinones in organohalide respiration in D. mccartyi strains. Since cultivation of D. mccartyi in synthetic mineral medium without the addition of quinones as vitamins has been reported (Schipp et al. 2013), the only alternative possibility to synthesize quinones would be via a novel biosynthetic pathway. The lack of standard quinone-reactive cytochromes also negates a requirement for quinones. Moreover, a recent study failed to detect any quinoid compound in extracts of D. mccartyi strain CBDB1 using sensitive mass spectrometric analyses (Kublik et al. 2016) and these results clearly distinguish D. mccartyi from other OHRB. Organohalide respiration in D. mccartyi strain CBDB1 was neither inhibited by HQNO (Jayachandran, Görisch and Adrian 2004) nor driven by reduced quinones such as DMNH2, menadiol, menaquinol-4, or diverse ubiquinols (Q-0, Q-4, or Q-10) (Jayachandran, Görisch and Adrian 2004; Kublik et al. 2016). Finally, the addition of protonophors that dissipate the proton gradient, such as carbonylcyanide m-chlorophenyl m-hydrazone and 3,3′,4′,5-tetrachlorosalicylanilide, did not inhibit dehalogenation by whole cells of D. mccartyi strain CBDB1 demonstrating that a reverse electron flow, as proposed for S. multivorans, is not necessary (Jayachandran, Görisch and Adrian 2004). Together, all these findings strongly support the notion of a quinone-independent respiratory chain in D. mccartyi and indicate that it is fundamentally different from that described for other OHRB. The obligate organohalide-respiring Dehalogenimonas species resemble D. mccartyi in lacking quinone biosynthetic genes, which indicates a similar mode of electron transfer (Siddaramappa et al. 2012; Molenda, Quaile and Edwards 2015; Key et al., 2016, 2017). While Dehalogenimonas spp. oxidize hydrogen and formate (Key et al. 2017), D. mccartyi strains are limited to using hydrogen as sole electron donor for organohalide respiration (Löffler et al. 2013). In the absence of quinones, the alternative is protein-mediated electron transfer from a hydrogenase to the RDase. Four different [Ni–Fe] hydrogenases are encoded in the genome of D. mccartyi strain CBDB1, but only one has been proposed to play a role in energy conservation (Kube et al. 2005). The abundance of the hydrogen-uptake (Hup) hydrogenase in D. mccartyi cells (Morris et al. 2007) and its dependence on Tat (Hartwig et al. 2015) predict that it is localized at the exoplasmic face of the cytoplasmic membrane, making its involvement in the organohalide respiratory chain highly likely (Kube et al. 2005; Mansfeldt et al. 2014). However, although the hup gene cluster encodes the catalytic large subunit (HupL) and the Fe–S-harboring small subunit (HupS), it lacks a hupC gene encoding a membrane-anchoring cytochrome b subunit. Moreover, no cytochromes have been detected in D. mccartyi (Löffler et al. 2013; Schipp et al. 2013). Typically, in other H2-oxidizing organisms, the membrane integral subunit facilitates electron transfer to the quinone pool via b-type cytochromes. The presence of a ferredoxin-like subunit (named HupX; locus tag: cbdbA131 in D. mccartyi strain CBDB1), which is encoded in the hup operon and which is also found in the hydrogenase 2 enzyme of E. coli (Pinske et al. 2015), might indicate alternative means of electron conduction from HupSL to the RDase. Although the hupX gene does not encode its own N-terminal Tat signal peptide, its co-translocation together with the HupSL heterodimer is feasible. This would need HupSLX complex formation in the cytoplasm prior to membrane translocation. A protein-mediated electron transfer path would require a transient or continuous interaction between the hydrogenase, the HupX protein and the RDase. In 2016, (Kublik et al. 2016) were able to show complex formation between HupSLX, a RDase, and a complex iron–sulfur molybdoenzyme in D. mccartyi strain CBDB1 by extracting membrane proteins under native conditions, separating protein complexes by native polyacrylamide gel electrophoresis, and subsequently identifying the polypeptides via mass peptide fingerprinting (Fig. 7). More recently, this complex has been isolated from the membrane of D. mccartyi strain CBDB1, enriched and been shown to catalyze hydrogen-dependent reduction of 1,2,3-trichlorobenzene (Hartwig et al. 2017), providing further support for quinone-independent electron transfer within the complex. Figure 7. View largeDownload slide Tentative scheme for the composition of the quinone-independent organohalide respiratory chain of D. mccartyi strain CBDB1 (Kublik et al. 2016). MGD, molybdopterin guanine dinucleotide; OmeAB, organohalide-respiration involved molybdoenzyme subunits A and B. Figure 7. View largeDownload slide Tentative scheme for the composition of the quinone-independent organohalide respiratory chain of D. mccartyi strain CBDB1 (Kublik et al. 2016). MGD, molybdopterin guanine dinucleotide; OmeAB, organohalide-respiration involved molybdoenzyme subunits A and B. While no direct interaction between HupSL and the RDase was observed, both enzymes were found to interact with HupX and the heterodimeric complex iron–sulfur molybdoenzyme complex. This complex consists of two proteins encoded by the genes cbdbA195 and cbdbA193 (Kublik et al. 2016). Here we propose to name the two genes omeA and omeB, for organohalide-respiration involved molybdoenzyme subunits A and B (see Fig. 7). OmeA has a Tat signal peptide at its N-terminus and is transported to the exoplasm like HupSL and the RDases. OmeB contains 10 transmembrane helices and notably is similar to the membrane anchor of hydrogenase 2-like enzymes (Pinske et al. 2015; Hartwig et al. 2017), suggesting that OmeB represents the docking site for HupSL. The transcription of the ome and the hup operons is tightly linked to the presence of halogenated compounds such as trichlorobenzenes, which strongly supports their functional linkage to reductive dehalogenation (Hartwig et al. 2017). Although OmeA shows amino acid sequence similarity to respiratory formate dehydrogenases, it lacks the key active site (seleno)cysteine and histidine residues (Hartmann, Schwanhold and Leimkühler 2015) required to catalyze formate oxidation. Moreover, formate oxidation has never been observed in D. mccartyi cultures and thus OmeA clearly has a different function (Hartwig et al. 2017). The polytopic membrane-integral OmeB could play a role in coupling electron transfer through the respiratory complex to proton translocation across the membrane (Fig. 7). Based on amino acid sequence similarity, its role could resemble that of the membrane anchor of H2-oxidizing hydrogenase 2. Sequence analysis of OmeB revealed the presence of a conserved glutamate in the membrane-spanning part of the protein, which is consistent with an involvement in proton translocation coupled to a conformational change (Zinder 2016). CONCLUSIONS AND OUTLOOK The exploration of the mechanisms coupling reductive dehalogenation to ATP synthesis via electron transport phosphorylation in OHRB has brought a diverse and complex picture to light. Breakthroughs in the analysis of structure–function relationships in cobamide-containing reductive dehalogenases, the description of novel respiratory chain components, and the identification of the potential mode of proton transfer have substantially improved our understanding of the physiology of OHRB. However, although structural information on reductive dehalogenases is available, neither the catalytic mechanism(s) nor the broad substrate spectrum of these enzymes is understood in detail. The molecular basis of the differentiation into RDases catalyzing hydrogenolysis/halogen substitution or vicinal reduction/dihaloelimination or both needs to be elucidated. Furthermore, the definition of the coupling sites that link the membrane-bound electron transfer chain to the translocation of protons across the cytoplasmic membrane is unclear. Thus, there remain many aspects of this exciting field of research that require further study. ACKNOWLEDGEMENTS This work was supported by the DFG Research Unit FOR1530. The authors would like to thank Cindy Kunze for her assistance in generating the overlay of the RDase structures. Tobias Goris is gratefully acknowledged for helpful discussions. Conflict of interest. None declared. REFERENCES Adrian L, Dudkova V, Demnerova K et al.   “Dehalococcoides” sp. strain CBDB1 extensively dechlorinates the commercial polychlorinated biphenyl mixture Aroclor 1260. Appl Environ Microbiol . 2009; 75: 4516– 24. Google Scholar CrossRef Search ADS PubMed  Adrian L, Hansen SK, Fung JM et al.   Growth of Dehalococcoides strains with chlorophenols as electron acceptors. Environ Sci Technol . 2007a; 41: 2318– 23. Google Scholar CrossRef Search ADS   Adrian L, Löffler FE. Organohalide-Respiring Bacteria . Berlin, Heidelberg: Springer, 2016. 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Environ Microbiol . 2016; 18: 2773– 5. Google Scholar CrossRef Search ADS PubMed  © FEMS 2018. All rights reserved. For permissions, please e-mail: journals.permissions@oup.com http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png FEMS Microbiology Ecology Oxford University Press

Organohalide respiratory chains: composition, topology and key enzymes

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Abstract

Abstract The utilization of halogenated organic compounds as terminal electron acceptors separates the phylogenetically diverse organohalide-respiring bacteria from other respiratory anaerobes that predominantly use nitrate, fumarate, sulfate or oxidized metals. Organohalide respiration is unique in recruiting a cobamide-containing iron–sulfur protein, the extracellular membrane-bound reductive dehalogenase, as terminal reductase in the electron transfer chain. In recent years substantial contributions have been made to the understanding of how electron transfer paths couple mechanistically to chemiosmosis in the organohalide-respiring bacteria. The structural analysis of a respiratory and a non-respiratory reductive dehalogenase revealed the intramolecular electron transfer via two cubane iron–sulfur clusters to the cobamide at the active site. Based on whether quinones are involved, two types of intermolecular electron transfer chains have been identified, which differ in their composition and mode of proton translocation. Indeed, various respiratory chain architectures have been unraveled and evidence for different putative coupling mechanisms presented. The identification of a multienzyme respiratory complex that combines uptake hydrogenase, a complex iron–sulfur molybdoenzyme and a reductive dehalogenase in Dehalococcoides mccartyi strain CBDB1 has raised new questions regarding the mode of energy conservation in these enigmatic microbes. In this mini-review, we highlight these findings and provide an outlook on potential future developments. anaerobic respiration, reductive dehalogenation, chemiosmosis, organohalides, reductive dehalogenase, cobamides INTRODUCTION Halogenated organic compounds (organohalides) are produced by geogenic, biological or synthetic processes (Gribble 2003; IARC 2014, 2016, 2017) and are present in various anoxic habitats as energy and carbon sources for the growth of anaerobic microbes. Several biological transformations that lead to the degradation of organohalides under anoxic conditions have been identified (Janssen, Pries and van der Ploeg 1994; Fetzner 1998), among them microbial reductive dehalogenation (Bouwer and McCarty 1983a,b; Horowitz, Suflita and Tiedje 1983). The latter process can precede the complete biological mineralization of organohalides in oxygen-depleted zones under methanogenic, iron-, sulfate-, or nitrate-reducing conditions (Monserrate and Häggblom 1997; Lohner and Spormann 2013; Tiedt et al. 2016) or at oxic–anoxic interfaces (Kurt, Mack and Spain 2014; Atashgahi et al.2017; Weatherill et al. 2018), where it can be coupled to oxic degradation processes. Taking into account the ubiquity and diversity of naturally occurring or man-made organohalides, the study of biological reductive dehalogenation is of ecological and environmental relevance. The exploration of how organohalides are used as energy or carbon source is important in terms of ecosystem remediation but is also essential for a complete understanding of microbial metabolic interactions in the environment. Particular anaerobic bacteria that carry out reductive dehalogenation have attracted attention because of their enigmatic mode of energy conservation, which often relies on respiration with hazardous halogenated chemicals (Holliger and Schumacher 1994; Holliger, Wohlfarth and Diekert 1998). This mini-review summarizes the advances made in recent years to unravel the molecular basis of microbial reductive dehalogenation. It also describes how this process is coupled to energy conservation via chemiosmosis. Desulfomonile tiedjei, a sulfate-reducer, was the first isolate observed to grow by coupling the oxidation of formate or hydrogen to the reduction of 3-chlorobenzoate (Shelton and Tiedje 1984; Dolfing and Tiedje 1987; DeWeerd et al. 1990; DeWeerd and Suflita 1990; Mohn and Tiedje 1990). This finding revealed the existence of a so-far-unknown respiratory process whereby reductive dehalogenation of organohalides is coupled to a chemiosmotic mechanism (organohalide respiration) (Louie and Mohn 1999). Meanwhile, various phylogenetically diverse bacteria (including Chloroflexi, Firmicutes, Beta-, Delta- and Epsilonproteobacteria; summarized in Maphosa, de Vos and Smidt 2010) have been identified that conserve energy using organohalides as terminal electron acceptors of membrane-associated electron transfer chains. These microorganisms are termed organohalide-respiring bacteria (OHRB) (Maphosa, de Vos and Smidt 2010; Hug et al. 2013; Adrian and Löffler 2016; Jugder et al. 2016; Atashgahi, Häggblom and Smidt 2017). OHRB can be divided into those that rely exclusively on organohalide respiration for energy conservation (obligate OHRB) and those that can alternatively use non-halogenated electron acceptors (versatile OHRB) including nitrate, fumarate, sulfate and oxidized metal ions (Maphosa, de Vos and Smidt 2010; Hug et al. 2013; Jugder et al. 2016). Selected versatile OHRB may even use oxygen as acceptor under microoxic conditions (Sanford, Cole and Tiedje 2002; Goris et al. 2014; Gadkari et al. 2018). The unifying feature of the diverse OHRB is the presence of reductive dehalogenases (RDases), which are located at the outer face of the cytoplasmic membrane and function as terminal reductases in membrane-associated electron transfer chains (Hug et al. 2013; Jugder et al. 2016; Fincker and Spormann 2017). However, the presence of such an exoplasmic and membrane-bound RDase does not necessarily indicate that the respective organism is able to perform organohalide respiration, especially when complex electron donors are utilized (e.g. pyruvate or lactate). In such cases, a fermentative lifestyle based on energy conservation via substrate level phosphorylation, with the organohalides serving exclusively as an electron sink (facilitated fermentation), is feasible (van de Pas et al. 2001) and needs to be ruled out before a respiratory lifestyle is ascertained. Furthermore, non-respiratory, cytoplasmically located RDases have been described (Chen et al. 2013; Payne et al. 2015). These enzymes ‘prime’ organohalides by the removal of halogen substituents for the subsequent degradation of the carbon backbone. As the name suggests, non-respiratory RDases are not involved in energy conservation via electron-transport phosphorylation. Hence, biological reductive dehalogenation by microbes is not necessarily linked to chemiosmosis. During respiratory RDase-mediated catalysis the net transfer of two electrons onto a (poly-)halogenated substrate results in the release of a chloride, a bromide or an iodide ion (hydrogenolysis). Alternatively, two halide ions can be released in a β-elimination reaction (dihaloelimination/vicinal reduction). Dehalogenation of fluorinated organohalides by a RDase has not been observed to date. The extraordinary stability of the carbon–fluorine bond represents a substantial obstacle to reductive cleavage. The standard redox potentials (E0′) of organohalides range between +240 and +580 mV (Dolfing and Janssen 1994; Dolfing and Novak 2015), which indicates that they are highly suitable electron acceptors for anaerobic respiration, and in terms of thermodynamics are comparable to nitrate with E0′ (NO3−/NO2−) = +433 mV (Thauer, Jungermann and Decker 1977). However, the low H+/e− ratio of about 1 measured for H2-dependent reduction (E0′ (H+/H2) = −414 mV) of tetrachloroethene (PCE) by Dehalobacter restrictus (Schumacher and Holliger 1996) and of 3-chlorobenzoate by D. tiedjei (Louie and Mohn 1999) suggests that either mechanistic or thermodynamic restrictions hinder a complete exploitation of the free energy available in the reaction: $$\rm{C_{2}Cl_{4} + H_{2}\rightarrow C_{2}HC_{3}+H^{+}+Cl^{-} \quad \Delta {\rm G}^{0'}=-189 \, kJ \, mol^{-1}H_{2}}$$ (Holliger, Wohlfarth and Diekert 1998) $$\rm{3-Cl-benzoate+H_{2}\rightarrow benzoate+H^{+}+Cl^{-} \quad \Delta {\rm G}^{0'}=-125 \, kJ \, mol^{-} \, H_{2}}$$ (Dolfing and Tiedje 1987) Taking into consideration a potential difference of approximately 100–200 mV, which is required for the translocation of 1 mol H+ across the cell membrane (Thauer, Jungermann and Decker 1977; Simon, van Spanning and Richardson 2008), the low H+/e−ratio falls substantially below the maximal theoretical yield. The formation of ATP from ADP plus Pi consumes 3–4 protons, depending on the architecture of the ATP synthase (Weber and Senior 2003; Mayer and Müller 2014; Silverstein 2014). Hence, between one-third and two-thirds of an ATP is formed per halide ion released in organohalide respiration. This is also supported by the low growth yields of S. multivorans (1.4 g cell protein per mole chloride released) when growing on hydrogen plus tetrachloroethene (Scholz-Muramatsu et al. 1995) and of an enrichment culture of D. restrictus (2.3 g cell protein per mole chloride released) (Holliger et al. 1993). Both the molecular mechanism and the structural basis for the proton translocation in organohalide respiratory chains remain to be elucidated. However, based on recent genomic and proteomic studies (summarized in Türkowsky et al. 2018 in this Special Issue), it is becoming increasingly clear that a uniform organization of the electron transfer chains in the diverse OHRB does not exist. Rather, diverse modes of coupling reductive dehalogenation to proton translocation seem to have evolved. In addition, based on the 3D-structure of a respiratory RDase (Bommer et al. 2014), as well as that of a non-respiratory RDase (Payne et al. 2015), novel insights have emerged regarding both intramolecular electron transfer and the catalytic mechanism of RDases. REDUCTIVE DEHALOGENASES During the last two decades several hundred entries of RDase protein sequences have been deposited in databases (Hug et al. 2013). Although overlapping substrate ranges have been identified, the sequence variability among RDases is high. For example, the respiratory tetrachloroethene RDase (PceASmul) of the Gram-negative epsilonproteobacterium S. multivorans (Neumann et al. 1996) and PceAY51 of the Firmicute Desulfitobacterium hafniense strain Y51 (Suyama et al. 2002) share only 27% amino acid sequence identity, yet both enzymes efficiently convert PCE. Orthologous groups of RDases with elevated sequence similarity can be identified (Hug et al. 2013), but due to the lack of their detailed biochemical characterization, a correlation of these groups with substrate classes or catalytic mechanisms is difficult to define (for examples see Bommer et al. 2014; Parthasarathy et al. 2015; Payne et al. 2015; Alfán-Guzmán et al. 2017; Kunze et al. 2017; Kunze, Diekert and Schubert 2017). RDases are metalloproteins and all representatives analyzed so far, with one exception, contain a cobamide. The exception is the 3-chlorobenzoate-converting RDase from D. tiedjei, which was reported to contain a heme cofactor (Ni, Fredrickson and Xun 1995). Cobalt-, iron-, and nickel-containing porphyrinoids share the ability to catalyze abiotic reductive dehalogenation (Schrauzer, Deutsch and Windgassen 1968; Wood, Kennedy and Wolfe 1968; Krone et al. 1989; Krone, Thauer and Hogenkamp 1989; Gantzer and Wackett 1991), but the identification of a heme rather than a cobamide as cofactor in a RDase remains unique. Further biochemical analyses are needed to define the role of alternative transition metal-containing porphyrinoids in enzymatic reductive dehalogenation. It is also currently unknown whether RDase enzymes have evolved from a single progenitor. The similarity of structural features within the nitroreductase fold that binds the cobamide cofactor at the core of RDases (Bommer et al. 2014; Payne et al. 2015) indicates an evolutionary conservation of common traits. However, a conserved motif for the binding of the cobamide by RDases has not been identified. Besides the well-characterized adenosyl-cobamide-dependent enzymes (mutases and eliminases) or the methyl-cobamide-dependent methyltransferases, RDases form a novel class of cobamide-dependent enzymes together with the queuosine biosynthetic enzyme QueG (Bridwell-Rabb and Drennan 2017). RDase sequences have been identified in archaea belonging to the Asgard superphylum (Zaremba-Niedzwiedzka et al. 2017), which suggests that RDase structural elements might have evolved prior to the division of bacteria and archaea. Substrate range Information on the mode of catalysis and the substrate range of RDases has been obtained either from the characterization of purified proteins or via enzyme activity measurements performed with RDases enriched on native polyacrylamide gels (Adrian et al. 2007b; Tang et al. 2013) or produced heterologously (Mac Nelly et al. 2014; Parthasarathy et al. 2015; Payne et al. 2015; Kunze, Diekert and Schubert 2017; Jugder et al. 2018). Before conclusions on the mechanism or substrate range of a given RDase can be drawn, cobamide-mediated abiotic conversion of organohalides (Schrauzer, Deutsch and Windgassen 1968; Wood, Kennedy and Wolfe 1968; Krone, Thauer and Hogenkamp 1989; Gantzer and Wackett 1991) has to be excluded. Whether the protein environment of the substrate-binding site has a direct impact on the conversion of a given substrate must be tested with heat-inactivated samples, as has been demonstrated for the dihaloelimination of 1,2-dibromoethane or 1,1,2,2-tetrachloroethane (Kunze, Diekert and Schubert 2017). Two general modes of RDase function have been documented: (i) hydrogenolysis/halogen substitution and (ii) vicinal reduction/dihaloelimination (Fig. 1A). While hydrogenolysis involves a proton transfer in the catalytic cycle, recent findings indicate that a proton might be dispensable for the dihaloelimination reactions catalyzed by RDases (Franke et al. 2017; Kunze, Diekert and Schubert 2017). Information is available about the involvement of RDases specialized for halogen substitution in organohalide respiratory chains (see below), but less is known about the coupling of dihaloeliminating enzymes to chemiosmosis in organohalide-respiring bacteria (De Wildeman et al. 2003; Moe et al. 2009; Bowman et al. 2013; Ding, Zhao and He 2014; Tang et al. 2016; Wong et al. 2016; Key et al. 2017). This area requires intensive future investigation. Figure 1. View largeDownload slide RDase catalysis and substrate range. (A) RDases that are known to catalyze both halogen substitution and dihaloelimination. (B) Selected substrates converted by PceASmul from S. multivorans. PCE, tetrachloroethene; TCE, trichloroethene; cDCE, cis-dichloroethene; VC, vinyl chloride; 1,2-DCA, 1,2-dichloroethane; 1,1,2-TCA, 1,1,2-trichloroethane; TBE, tribromoethene; 1,1-DBE, 1,1-dibromoethene; tDBE, trans-dibromoethene; cDBE, cis-1,2-dibromoethene; t-1,3-DCP, trans-1,3-dichloropropene; t-1-CP, trans-1-chloropropene; c-1-CP, cis-1-chloropropene; 3-CP, 3-chloropropene; 2,4,6-TBP, 2,4,6-tribromophenol; 2,4-DBP, 2,4-dibromophenol. Figure 1. View largeDownload slide RDase catalysis and substrate range. (A) RDases that are known to catalyze both halogen substitution and dihaloelimination. (B) Selected substrates converted by PceASmul from S. multivorans. PCE, tetrachloroethene; TCE, trichloroethene; cDCE, cis-dichloroethene; VC, vinyl chloride; 1,2-DCA, 1,2-dichloroethane; 1,1,2-TCA, 1,1,2-trichloroethane; TBE, tribromoethene; 1,1-DBE, 1,1-dibromoethene; tDBE, trans-dibromoethene; cDBE, cis-1,2-dibromoethene; t-1,3-DCP, trans-1,3-dichloropropene; t-1-CP, trans-1-chloropropene; c-1-CP, cis-1-chloropropene; 3-CP, 3-chloropropene; 2,4,6-TBP, 2,4,6-tribromophenol; 2,4-DBP, 2,4-dibromophenol. Although halogen substitution and dihaloelimination were both presumed to be catalyzed by distinct types of RDases, actual case reports show that, depending on the substrate, a single enzyme can perform both reactions (Fig. 1A). For example, the vinyl chloride (VC) RDase (VcrA) from D. mccartyi strain VS has been found to mediate the reductive halogen substitution at VC and forms ethene as well as catalyzing the dihaloelimination of 1,2-dichloroethane (1,2-DCA, synonym: ethylene dichloride, EDC) to ethene (Parthasarathy et al. 2015). The conversion rate with 1,2-DCA was only two-fold lower compared with the rate measured with VC as an electron acceptor. Moreover, the heterologously produced DcaA of Desulfitobacterium dichloroeliminans catalyzed the dihaloelimination of 1,2-DCA but also mediated PCE conversion to trichloroethene (TCE) and finally to cis-1,2-dichloroethene (cDCE) via hydrogenolysis (Kunze, Diekert and Schubert 2017). Here, the activity was 10-fold lower with PCE compared with 1,2-DCA. More recently it was shown that 1,1,2-trichloroethane (1,1,2-TCA) was converted to a mixture of vinyl chloride and 1,2-DCA by the trichloromethane RDase (TmrA) of Dehalobacter sp. strain UNSWDHB (Jugder et al.2017). In this case both mechanisms, halogen substitution and dihaloelimination, appeared to be applied by a single enzyme to the same substrate. Nevertheless, it still must be determined in every case whether all enzyme substrates identified for a single RDase allow coupling of the reductive dehalogenation to chemiosmosis in the respective organohalide-respiring bacterium. Besides numerous tests on substrate promiscuity of RDases, systematic analyses of the ability of RDases to convert halogenated aliphatic and aromatic substrates are needed to understand fully the flexibility of these enzymes. Furthermore, the influence of heteroatoms in aromatic substrates and the impact of flanking substituents on the reductive dehalogenation have to be tested more intensively (Cooper et al. 2015; Zhang et al. 2017). An example of an OHRB that shows enzymatic conversion of a structurally diverse set of organohalides is D. mccartyi strain CBDB1. The substrate range of this organism encompasses different congeners of polychlorinated biphenyls (Adrian et al. 2009), chlorinated dioxins and other aromatic compounds such as halogenated benzenes, phenols, anilines, or benzonitriles (Adrian et al. 2007a; Wagner et al. 2012; Cooper et al. 2015). Aliphatic organohalides, e.g. chlorinated ethenes, are also converted by D. mccartyi strain CBDB1 (Fung et al. 2007; Marco-Urrea, Nijenhuis and Adrian 2011). Whether a single enzyme of the 32 RDase complement encoded in the genome of D. mccartyi strain CBDB1 (Kube et al. 2005) is responsible for all the conversions listed above is not known. However, it has to be assumed that a single enzyme converts structurally different substrates, since the organism's substrate range is independent of the growth substrate that might induce the production of different RDases. Furthermore, it appears feasible that the conversion of aliphatic substrates by a given RDase does not preclude the dehalogenation of aromatic compounds and vice versa. Detailed information on the substrate range of PceASmul from S. multivorans is available (Neumann et al. 2002; Schmitz et al. 2007). This enzyme was initially thought to convert only aliphatic substrates such as chlorinated ethenes or propenes (Fig. 1B), but subsequent studies showed the efficient dehalogenation of bromoethenes and bromophenols (Ye et al. 2010; Kunze et al. 2017). Another example is RdhA3 of D. hafniense DCB-2, a RDase that displays a preference for the conversion of chlorophenols; however, it also dehalogenated PCE at lower rates (Mac Nelly et al. 2014). Such observations suggest a hidden potential of RDases to dehalogenate structurally diverse substrates and these results should be taken into account when the substrate range of newly discovered RDase is investigated. The RDase active-site cavity is connected via a hydrophobic substrate channel to the solvent (Bommer et al. 2014; Payne et al. 2015). Most of the amino acids involved in forming the channel are part of the enzyme's N-terminus, which in general shows little sequence conservation. This diversity might reflect an adaptation of the various RDases to the huge variety of halogenated substrates and increased amino acid sequence identity in this region might indicate an overlapping substrate range. However, a high overall sequence similarity does not necessarily mean that two RDases share the same substrate range or form the same products (Buttet, Holliger and Maillard 2013; Alfan-Guzman et al. 2017; Kunze, Diekert and Schubert 2017). Predicting the substrate preferences of a given RDase based on sequence comparison or structural modeling is hampered by the fact that even minor changes in the RDase architecture modulate substrate preference (Kunze, Diekert and Schubert 2017). For example, PceASmul of S. multivorans reduces PCE via TCE to cDCE (Neumann et al. 1996) and it shares this property with the highly similar PceAShal from Sulfurospirillum halorespirans (92% amino acid sequence identity for the processed enzyme without the twin arginine translocation signal peptide) (Luijten et al. 2003; Goris et al. 2017). In the enriched Sulfurospirillum sp. SL-2, a PceA variant, termed PceA (TCE), was identified that also shares 92% amino acid sequence identity with PceASmul but converts PCE only to TCE (Buttet, Holliger and Maillard 2013). A structural alignment of the PceASmul crystal structure and an in silico structural model of PceA (TCE) identified slight modifications in the architecture of the active site cavity that might be responsible for the difference in substrate range (Fig. 2). Figure 2. View largeDownload slide Comparison of PceA enzymes from Sulfurospirillum spp. (A) Multiple sequence alignment of PceASmul of S. multivorans (accession no. AHJ12791.1), PceAShal of S. halorespirans (acc. no. AOO65270.1) and PceA (TCE) of Sulfurospirillum sp. SL-2 (acc. no. ARU48750.1). Twin arginine translocation (Tat) signal peptides were not included in the alignment. Amino acid residues involved in the formation of the active site cavity are labeled in grey, while amino acid exchanges are highlighted in red. The cysteines binding the two cubane iron–sulfur clusters are marked in yellow. (B) In silico structural model of the active site cavity of PceA (TCE) from Sulfurospirillum sp. SL2 (petrol) overlaid with the structure of PceASmul (gray) (Bommer et al. 2014) depicted in two orientations. The structural model (C-score: 0.97) was generated with the I-TASSER server for protein structure and function prediction (Zhang 2008; Yang et al. 2015). The residues that differ between the enzymes are shown in red frames. The two orientations identified for TCE at the active site of PceASmul are depicted in an overlay. The red sphere between the cobamide cofactor and the substrate represents a water or hydroxyl group. Figure 2. View largeDownload slide Comparison of PceA enzymes from Sulfurospirillum spp. (A) Multiple sequence alignment of PceASmul of S. multivorans (accession no. AHJ12791.1), PceAShal of S. halorespirans (acc. no. AOO65270.1) and PceA (TCE) of Sulfurospirillum sp. SL-2 (acc. no. ARU48750.1). Twin arginine translocation (Tat) signal peptides were not included in the alignment. Amino acid residues involved in the formation of the active site cavity are labeled in grey, while amino acid exchanges are highlighted in red. The cysteines binding the two cubane iron–sulfur clusters are marked in yellow. (B) In silico structural model of the active site cavity of PceA (TCE) from Sulfurospirillum sp. SL2 (petrol) overlaid with the structure of PceASmul (gray) (Bommer et al. 2014) depicted in two orientations. The structural model (C-score: 0.97) was generated with the I-TASSER server for protein structure and function prediction (Zhang 2008; Yang et al. 2015). The residues that differ between the enzymes are shown in red frames. The two orientations identified for TCE at the active site of PceASmul are depicted in an overlay. The red sphere between the cobamide cofactor and the substrate represents a water or hydroxyl group. Heterologous production The low growth yields of OHRB and the toxicity of their halogenated substrates limit the amount of biomass available to deliver sufficient amounts of enzyme for biochemical studies. Thus, only a few RDases have been purified and biochemically characterized to date (summarized in Hug et al. 2013; Lu et al. 2015; Jugder et al. 2016; Schubert and Diekert 2016; Fincker and Spormann 2017). Given the fact that RDases display high amino acid sequence variability and diverse substrate ranges, the results obtained from this group of selected enzymes might only provide limited insight into RDase substrate specificity. Functional heterologous production of catalytically active RDases is complicated because of the need for cobamides and Fe–S clusters, and their synthesis and insertion often hamper such approaches. Nevertheless, the PCE-converting PceAY51 of D. hafniense Y51, the chlorophenol-dehalogenating RdhA3 of D. hafniense DCB-2, the 1,2-DCA-dihaloeliminating DcaA of D. dichloroeliminans, the chloroform-dehalogenating TmrA of Dehalobacter sp. strain UNSWDHB, and the non-respiratory NpRdhA of N. pacificus were functionally produced in a heterologous host (Mac Nelly et al. 2014; Payne et al. 2015; Kunze, Diekert and Schubert 2017; Jugder et al. 2018). Successful reconstitution of a catalytically active RDase from the apoprotein has been described only once for the vinyl chloride RDase (VcrA) from Dehalococcoides mccartyi strain VS (Parthasarathy et al. 2015). Cobamide cofactor The demand for cobamides (i.e. corrinoids with a complete nucleotide loop) as essential cofactors of the key enzyme of organohalide respiration is satisfied differently by the various OHRB. Cobamide producers such as S. multivorans (Goris et al. 2014; Keller et al. 2014), D. hafniense Y51 (Nonaka et al. 2006; Reinhold et al. 2012), or Dehalobacter sp. strains TCA1, CF, DCA (Sun et al. 2002; Tang et al. 2016; Wang et al. 2017) and UNSWDHB (Desphande et al. 2013; Wong et al. 2016) have been identified. In addition, cobamide auxotrophs have been isolated, e.g. Dehalobacter restrictus (Rupakula et al. 2013) and Dehalococcoides mccartyi (Löffler et al. 2013), that have to take up cobamides from the environment. The cobamides produced by the OHRB, which are able to perform de novo biosynthesis, differ in the structure of their nucleotide loop (Kräutler et al. 2003; Yan et al. 2018). The nucleotide loop of the norpseudo-B12 produced by S. multivorans lacks a methyl group at position 176 and harbors an adeninyl moiety as the lower base (Fig. 3A) (Kräutler et al. 2003). Whether the lack of the methyl group represents a specific adaptation to the cofactor requirements of PceASmul is not definitively resolved (Bommer et al. 2014; Keller et al. 2016; Schubert 2017). Recently, the structure of the cobamide produced by D. hafniense strains was elucidated and shown to differ from vitamin B12 (Fig. 3B) by the presence of unsubstituted purine as lower base (Yan et al. 2018). Figure 3. View largeDownload slide Cobamide structures. (A) The norpseudo-B12 is depicted in the base-off conformation with a hydroxyl group as upper ligand. (B) Vitamin B12 is shown in base-on conformation with the artificial cyano-group as upper ligand (alternative name cyanocobalamin). DMB = 5,6-dimethylbenzimidazole (C) Alternative norcobamides produced by S. multivorans in the presence of exogenous benzimidazoles. The functional utilization of these norcobamides as cofactors of PceASmul is indicated by the colored circles (green, functional; orange, reduced functionality; red, substantial impairment) (data from Keller et al. 2018). (D) Alternative cobamides amended to cultures of D. mccartyi strain 195 cultivated with TCE. The functional utilization of these cobamides as cofactors in organohalide respiration is indicated by colored circles (green, functional; red, substantial impairment) (data from Yi et al. 2012). In addition, 2-methylmercaptoadeninyl-cobamide (not shown) was tested with D. mccartyi strain 195, but did not functionally replace vitamin B12. Figure 3. View largeDownload slide Cobamide structures. (A) The norpseudo-B12 is depicted in the base-off conformation with a hydroxyl group as upper ligand. (B) Vitamin B12 is shown in base-on conformation with the artificial cyano-group as upper ligand (alternative name cyanocobalamin). DMB = 5,6-dimethylbenzimidazole (C) Alternative norcobamides produced by S. multivorans in the presence of exogenous benzimidazoles. The functional utilization of these norcobamides as cofactors of PceASmul is indicated by the colored circles (green, functional; orange, reduced functionality; red, substantial impairment) (data from Keller et al. 2018). (D) Alternative cobamides amended to cultures of D. mccartyi strain 195 cultivated with TCE. The functional utilization of these cobamides as cofactors in organohalide respiration is indicated by colored circles (green, functional; red, substantial impairment) (data from Yi et al. 2012). In addition, 2-methylmercaptoadeninyl-cobamide (not shown) was tested with D. mccartyi strain 195, but did not functionally replace vitamin B12. Preferences of some RDases for specific lower bases within cobamide cofactors have been observed and alterations in cobamide structure, i.e. exchange of the lower ligand base, interferes with RDase function (Fig. 3C and D) (Yan et al. 2012; Yi et al. 2012; Yan et al. 2013; Keller et al. 2014; Yan et al. 2016; Keller et al. 2018). The structural or mechanistic reasons for these cofactor incompatibilities have not been unraveled yet. Solving the three-dimensional structure of PceASmul of S. multivorans and the non-respiratory bromophenol RDase (NpRdhA) of the marine Nitratireductor pacificus pht-3B, uncovered the mode of cobamide binding (Bommer et al. 2014; Payne et al. 2015). In both RDases the cobamide is bound deep within the protein, with the nucleotide loop oriented away from the corrin ring. Several hydrogen bonds between the nucleotide loop and the protein environment indicate a role for the cobamide's tail in cofactor positioning. Hence, negative effects on RDase activity of structural variations in the nucleotide loop of the cobamide cofactor might be based on a selectivity in cofactor incorporation rather than resulting from direct interference of the nucleotide loop structure with the catalytic cycle or substrate range. So far, no evidence for an involvement of the cobamide cofactor's nucleotide loop in the catalytic cycle of RDases has been found (Bommer et al. 2014; Payne et al. 2015). Based on the static PceASmul structure it was not possible to identify a barrier that prevents the incorporation of distinct cobamides into the enzyme. Consequently, an incompatibility of the cobamide structure and the enzyme's folding process has been suggested (Keller et al. 2018). ELECTRON TRANSFER WITHIN REDUCTIVE DEHALOGENASE ENZYMES Evidence has been presented that the super-reduced state [CoI] of the cobalt ion in the cobamide is the reactive form of the cofactor that attacks the organohalide (Fig. 4) (Miller, Wohlfarth and Diekert 1996; Schumacher et al. 1997; van de Pas et al. 1999; Parthasarathy et al. 2015; Kunze et al. 2017). Several purified RDases have been studied by electron paramagnetic resonance (EPR) spectroscopy and shown to harbor a cobamide cofactor in the [CoII] oxidation state (see below). This ‘as isolated’ state appears not to be reactive. Low potential electron donors, e.g. reduced methyl viologen (E0′ = −446 mV), are required for the conversion of the substrate in the enzymatic assay (Miller, Wohlfarth and Diekert 1996). This necessity is indicative of the need for the [CoI] oxidation state to attack the organohalide. The electrons provided by artificial electron donors are probably transferred via the two Fe–S clusters to the cobamide cofactor. The efficient inhibition of PceASmul by 1-iodopropane (propyl iodide) supports the assumption that [CoI] functions as the reactive oxidation state (Neumann, Wohlfarth and Diekert 1995). Based on the active site architecture of PceASmul and NpRdhA (Bommer et al. 2014; Payne et al. 2015) and the positioning of the substrate (Kunze et al. 2017), a nucleophilic attack of the [CoI] on the electron-deficient carbon backbone of the substrate is unlikely. However, this mechanism was reported earlier for the abiotic reductive dehalogenation mediated by protein-free cobamides (Fig. 4A) (Schrauzer, Deutsch and Windgassen 1968; Wood, Kennedy and Wolfe 1968; Krone, Thauer and Hogenkamp 1989). Proposals for the initial attack on the enzyme-bound substrate include either a long-range electron transfer from the cobalt to the substrate (Fig. 4B) (Schmitz et al. 2007; Bommer et al. 2014; Kunze et al. 2017), or a direct reaction of [CoI] with a halogen substituent (Fig. 4C) (Payneet al. 2015; Cooperet al. 2015; Johannissen, Leys and Hay 2017). The mechanistic diversity of cobamide-mediated enzymatic and abiotic reductive dehalogenation have been subject to stable isotope fractionation analyses, which were reviewed previously and are not discussed here (Elsner 2010; Nijenhuis et al. 2016). Figure 4. View largeDownload slide Proposed modes for the initial attack of the super-reduced [CoI] onto the halogenated RDase substrate. (A) Nucleophilic attack of [CoI] at the electron-deficient carbon backbone of the organohalide results in the alkylation of the cobalt. (B) Substrate radical formation via long-range electron transfer. The proposed subsequent formation of a carbanion after the elimination of the halogen substituent is not depicted. (C) [CoI] attacks directly at the halogen substituent and the carbon-halogen bond is either cleaved heterolytically (left) or homolytically (right). [CoI-II], oxidation states of the cobalt ion in the cobamide cofactor; R, hydrocarbon backbone; X, halogen substituent (adapted from Kunze et al. 2017). Figure 4. View largeDownload slide Proposed modes for the initial attack of the super-reduced [CoI] onto the halogenated RDase substrate. (A) Nucleophilic attack of [CoI] at the electron-deficient carbon backbone of the organohalide results in the alkylation of the cobalt. (B) Substrate radical formation via long-range electron transfer. The proposed subsequent formation of a carbanion after the elimination of the halogen substituent is not depicted. (C) [CoI] attacks directly at the halogen substituent and the carbon-halogen bond is either cleaved heterolytically (left) or homolytically (right). [CoI-II], oxidation states of the cobalt ion in the cobamide cofactor; R, hydrocarbon backbone; X, halogen substituent (adapted from Kunze et al. 2017). RDases in the ‘as isolated’ form studied by EPR spectroscopy showed the cobalt ion to be in the [CoII] oxidation state (Schumacher et al. 1997; van de Pas et al. 1999; Kräutler et al. 2003; Parthasarathy et al. 2015; Payne et al. 2015; Kunze et al. 2017). The midpoint redox potential of the couple [CoII]/[CoI] in the enzyme-bound cofactor has been determined for the PCE-RDase of D. restrictus (PceADres; −350 ± 20 mV, pH 8.0; Schumacher et al. 1997), PceASmul (−380 mV, pH 7.5; Kräutler et al. 2003)and the chlorophenol RDase of Desulfitobacterium dehalogenans (−370 mV, pH 7.8; van de Pas et al. 1999). The two [4Fe–4S] clusters of PceADres were found to have more negative midpoint redox potentials of about −480 mV (Schumacher et al. 1997), which is commensurate with an unimpeded electron transfer to the cobamide at the active site. A similar value (approximately −440 mV) was obtained for the Fe–S clusters in PceASmul, when an EPR-coupled redox titration of the two cubane iron–sulfur clusters was carried out (C. Kunze and F. Hagen, unpublished results); purified and catalytically active PceASmul enzyme was used in this experiment. The enzyme fraction contained mainly the mature form of PceASmul. The result of the redox titration of PceASmul was in accord with the onset potential (−450 mV versus standard hydrogen electrode at pH 7) needed to initiate reductive dehalogenation of PCE and TCE by PceASmul bound to a redox electrode (Siritanaratkul et al. 2016). Based on the topology of the redox-active metal centers inside the RDase, it was hypothesized that the electrons delivered by an external donor are conducted via a short ‘electronic wire’ comprising both [4Fe–4S] clusters and the cobamide (Bommer et al. 2014; Payne et al. 2015). The presence of two [4Fe–4S] clusters has also been confirmed for VcrA of D. mccartyi VS and NpRdhA, but their respective redox potentials have not been determined (Parthasarathy et al. 2015; Payne et al. 2015). Although the Fe–S cluster binding motifs in CprA of D. dehalogenans indicate the presence of two cubane clusters, a [4Fe–4S] cluster with a midpoint redox potential of −440 mV and one with a [3Fe–4S] cluster with a positive redox potential of +70 mV were identified (van de Pas et al. 1999). The role of the latter cluster in the reduction reaction remains to be determined. The corrin ring of the cobamide cofactor defines the base of the active site cavity, which is mainly lined by hydrophobic amino acids (Fig. 2B). A network of hydrogen bonds positions the cofactor in RDases. The cobamide cofactor is bound in the base-off conformation in all RDases analyzed so far by EPR spectroscopy or X-ray crystallography (Schumacher et al. 1997; van de Pas et al. 1999; Bommer et al. 2014; Payne et al. 2015; Keller et al. 2018). The term ‘base-off’ describes the absence of a coordinative bond between the lower base and the central cobalt ion (Fig. 3A). The reduction of the cobalt ion in protein-free cobamides from [CoII] to the [CoI] ion is accompanied by a structural conversion from the base-on to the base-off conformation. The protein environment in PceASmul and NpRdhA fixes the cobamide cofactor in the base-off conformation with no alternative ligand bound to the cobalt and it prevents a switch from the base-off to the base-on state. This feature lends support to the hypothesis that the protein environment influences the reduction of the cobalt ion in the RDase-bound cofactor and this is reflected in the elevated midpoint redox potential of the RDase-bound cofactor compared with the protein-free form (e.g. −490 mV vs. standard hydrogen electrode at pH 7 for the protein-free norpseudovitamin B12 compared with −380 mV for the enzyme-bound norpseudo-B12 at pH 7.5; Kräutler et al. 2003). An exception to the rule of base-off binding might have been identified for VcrA of D. mccartyi VS, which was analyzed by EPR and found to incorporate the cobamide cofactor in the base-on conformation with the lower base of the cofactor coordinating the central metal ion directly (Parthasarathy et al. 2015). Redox potentials of the different oxidation states of the cobamide cofactor have not yet been determined in this case. Given the findings that RDases are highly diverse and cobamides display a multifaceted chemistry, alternative modes of cobamide cofactor binding and utilization cannot be excluded. In analogy to the cobamide-binding motif present in other B12-dependent enzymes (Dowling, Croft and Drennan 2012), putative cobamide-binding sequences (DHXG-X39-S-X32-G) were identified in dehalogenating Chloroflexi (Hölscher et al. 2004; Padilla-Crespo et al. 2014). However, as visualized by structure prediction based on the PceASmul and NpRdhA 3D-structures, the histidine residue does not come close to the cobalt to serve as an alternative lower ligand. Most primary amino acid sequences of Chloroflexi RDases form a distinct cluster in phylogenetic trees (Hug et al. 2013; Fincker and Spormann et al. 2017). This distinctiveness might reflect differences in the catalytic mechanism of these enzymes, in their mode of cobamide cofactor binding, or in the composition of the organohalide respiratory chain of these organisms. SUBCELLULAR LOCALIZATION OF RESPIRATORY RDases and the RdhB PROTEINS The localization of the membrane-associated primary electron-donating enzyme and the terminal reductase of a respiratory chain is critical in determining the H+/e− ratio (Simon, van Spanning and Richardson 2008). In general, RDases are synthesized and equipped with cofactors in the cytoplasm (John et al. 2006; Reinhold et al. 2012). Since a characteristic property of the cytoplasmic RDase precursors is the presence of an N-terminal twin-arginine translocation (Tat) (Palmer and Berks 2012) signal peptide, it is assumed that the Tat machinery translocates the enzymes across the cytoplasmic membrane. The findings of early studies, however, proposed a binding of the enzyme to the inner face of the cytoplasmic membrane, mainly based on the results obtained from enzyme activity measurements with intact or permeabilized cells conducted with the membrane-impermeant methyl viologen as an artificial electron donor (Holliger and Schumacher 1994; Miller, Wohlfarth and Diekert 1996, 1997). Later, the same experimental approach applied to D. mccartyi strain 195 led to the conclusion that the RDase in this organism has an exoplasmic localization (Nijenhuis and Zinder 2005). In a subsequent study, the subcellular localization of PceASmul was re-examined via freeze-fracture replica immunogold labeling, which allowed direct detection of the membrane-bound state of the enzyme in cells of S. multivorans (John et al. 2006). This study showed unequivocally that the PceASmul molecules were indeed located on the outer face of the cytoplasmic membrane in PCE-grown cells. A cytoplasmic localization of PceASmul, observed in earlier studies, has been attributed to a reduced transport of the enzyme's precursor in fumarate-grown cells. An exoplasmic localization of PceA was also demonstrated for D. hafniense Y51 (Reinhold et al. 2012), while treatment of D. restrictus with proteinase K resulted in the loss of PceADres from these cells, which also indicated an exoplasmic localization of the RDase in this organism (C. Holliger; personal communication). Hence, it is currently assumed that the physiologically active RDase is located at the outer rather than the inner face of the lipid bilayer. In most cases, a small open reading frame encoding a hydrophobic protein (rdhB) accompanies the structural gene for the respiratory RDase enzyme (rdhA). It was proposed that RdhB functions as a membrane anchor for RdhA (Neumann, Wohlfarth and Diekert 1998). However, the experimental proof for this assumption is still lacking and the formation of the rdhB gene products could not be verified for a long time. Recently, however, the synthesis of the B protein in S. multivorans has been demonstrated by proteomic analysis of PCE-grown cells (Goris et al. 2015). Furthermore, the involvement of the B protein in the formation of a multienzyme organohalide respiratory complex in D. mccartyi was confirmed via peptide mass fingerprinting (Kublik et al. 2016). Although a substantial number of rdhA genes are indeed accompanied by a dedicated rdhB gene, also solitary rdhA genes have been identified. In Dehalogenimonas species, the number of rdhB genes is much lower than the number of rdhA genes, which might indicate that the RdhB protein is dispensable for some RDases in this organism, that a single RdhB can anchor different RdhA proteins, or that other proteins might be involved in RDase membrane binding (Siddaramappa et al. 2012; Molenda, Quaile and Edwards 2015; Key et al. 2016; Yang et al. 2017). Most of the RDases encoded in Dehalogenimonas species have a Tat signal peptide for membrane export, implying they have an exoplasmic localization. Whether all of the rdhA gene products in this organism are functional in energy conservation or have another function remains to be elucidated. Topology calculations of RdhB proteins showed in most cases three transmembrane helices in addition to an exoplasmic N-terminus and a short exoplasmic loop between helices 2 and 3 (Fig. 5A). The corresponding B protein of the PceASmul enzyme represents an exception in having only two transmembrane helices and most probably both termini are exposed to the cytoplasm (Fig. 5B). Since respiratory RDase does not appear to have membrane integral domains that insert into the lipid bilayer, the enzymes must interact with the exoplasmic loops or domains of other integral membrane proteins or membrane-associated proteins. Although the membrane-spanning helices of B proteins share little sequence identity, two conserved glutamic acid residues can be identified in the exoplasmic loop of most RdhBs and these might play a role in the RdhA–RdhB interaction. No typical cofactor-binding site is present in RdhB proteins. Consequently, their direct involvement in the electron transfer to the RDase is unlikely. Whether anchoring the RDase to the membrane is the sole function of the B proteins or whether they also serve as a facilitator between the RDase and other proteins involved in electron transfer is unclear. Figure 5. View largeDownload slide The PceB protein of D. hafniense Y51 and of S. multivorans. (A and B) Predicted topology of PceB in the cytoplasmic membrane (TMHMM Server v. 2.0; Sonnhammer, von Heijne and Krogh 1998) is shown. Figure 5. View largeDownload slide The PceB protein of D. hafniense Y51 and of S. multivorans. (A and B) Predicted topology of PceB in the cytoplasmic membrane (TMHMM Server v. 2.0; Sonnhammer, von Heijne and Krogh 1998) is shown. QUINONE-DEPENDENT ORGANOHALIDE RESPIRATION The electron transfer path in the organohalide respiratory chain from the oxidation of the electron donor to the terminal RDase differs in the various OHRB. The paths can be classified into those using quinones for electron and proton shuttling within the membrane and those possessing a quinone-independent electron transfer chain. In theory, the involvement of quinones in respiratory chains allows a proton transfer via a redox loop mechanism with interaction sites for quinone reduction and quinol oxidation being at opposite sides of the cytoplasmic membrane (Jormakka et al. 2002; Simon, van Spanning and Richardson 2008). In the absence of quinones a conformational proton pump might be responsible for generating the proton gradient (Simon, van Spanning and Richardson 2008; Efremov, Baradaran and Sazanov 2010). The obligate organohalide-respiring D. restrictus was shown to contain different menaquinones (MK; mainly MK-7 and MK-8) (Holliger et al. 1998). The organism harbors a complete set of genes encoding the enzymes of menaquinone biosynthesis (Dairi et al. 2011; Kruse et al. 2013). Comparative spectral analysis of D. restrictus membranes reduced with H2 and oxidized with PCE displayed a clear change in the absorbance spectrum of MK revealing its potential participation in the PCE respiratory chain (Schumacher and Holliger 1996). The reductive dehalogenation of PCE in whole cells of D. restrictus was driven by the addition of reduced 2,3-dimethyl-1,4-naphthoquinone (DMNH2), a water-soluble MK analog, and was blocked in the presence of the quinone antagonist 2-n-heptyl-4-hydroxyquinoline N-oxide (HQNO; 10 nmol mg−1 protein). An inhibitory effect of HQNO at an elevated concentration (150 nmol mg−1 protein) was also found for D. tiedjei (Louie and Mohn 1999) and for S. multivorans (>80 nmol mg−1 protein; Gadkari 2017). Menaquinones have been extracted from S. multivorans (Scholz-Muramatsu et al. 1995) and the genetic information for menaquinone biosynthesis via the alternative futalosine pathway was identified (Goris et al. 2014). The presence of (mena-)quinones has also been confirmed for D. tiedjei (Louie and Mohn 1999) and D. dehalogenans (Kruse et al. 2015). In the latter organism, oxidation of menaquinol (MKH2) by 3-chloro-4-hydroxy-phenylacetate (ClOHPA) was observed. All of these results indicate the involvement of quinones in the intermolecular electron transfer of the listed OHRB. Due to the lack of functional motifs, an interaction of the B proteins with the MK pool in quinone-dependent OHRB is unlikely, which leaves the question of the mode of electron transfer from the MKH2 to the RDase unanswered. Proteome analysis of S. multivorans identified a putative quinol dehydrogenase found exclusively in PCE-grown cells, which might represent the ‘missing link’ in the electron transfer chain (Fig. 6; Goris et al. 2015). The genes for the heterodimeric protein (locus tags: SMUL_1541 and SMUL_1542) are located in proximity to the gene encoding PceASmul (Goris et al. 2014). The two gene products show sequence similarity to the NapGH quinol dehydrogenase involved in electron transfer to the periplasmic nitrate reductase (NapA) (Kern and Simon 2008; Simon and Klotz 2013), but are part of a separate clade based on a phylogenetic analysis of the NapGH-like quinol dehydrogenase protein family (Goris et al. 2014). The localization of RDase genes in proximity to genes encoding a NapGH-like quinol dehydrogenase can also be found in the genome sequence of other dehalogenating Sulfurospirillum species, such as S. halorespirans (Goris et al. 2017) and Sulfurospirillum sp. SL2–1 (RefSeq. NZ_CP021416.1), as well as in the sulfate-reducers D. tiedjei (RefSeq. NC_018025.1), Pseudodesulfovibrio indicus (RefSeq. NZ_CP014206; Cao et al.2016), Desulfovibrio bizertensis (RefSeq. NZ_FUYA01000002), and Desulforhopalus singaporensis (RefSeq. FNJI01000016). However, the analysis of co-expression of these genes with the RDase structural genes still needs to be performed. Figure 6. View largeDownload slide Simplified scheme for the composition of the quinone-dependent organohalide respiratory chain in Sulfurospirillum multivorans. HQNO, 2-n-heptyl-4-hydroxyquinoline N-oxide; DMNH2, 2,3-dimethyl-1,4-naphthoquinone; MNH2, 2-methyl-1,4-naphthoquinone; MKH2, menaquinol; Q-DH, quinol dehydrogenase. The inhibitory effect of HQNO and the electron-donating function of MNH2 are indicated in red and green, respectively. The question mark designates the quinol-oxidizing component, which has not been identified so far. The putative membrane anchor of the RDase (PceB, locus tag: SMUL_1532) is depicted in white, since its interaction with the RdhA protein lacks experimental verification. Figure 6. View largeDownload slide Simplified scheme for the composition of the quinone-dependent organohalide respiratory chain in Sulfurospirillum multivorans. HQNO, 2-n-heptyl-4-hydroxyquinoline N-oxide; DMNH2, 2,3-dimethyl-1,4-naphthoquinone; MNH2, 2-methyl-1,4-naphthoquinone; MKH2, menaquinol; Q-DH, quinol dehydrogenase. The inhibitory effect of HQNO and the electron-donating function of MNH2 are indicated in red and green, respectively. The question mark designates the quinol-oxidizing component, which has not been identified so far. The putative membrane anchor of the RDase (PceB, locus tag: SMUL_1532) is depicted in white, since its interaction with the RdhA protein lacks experimental verification. The quinol-oxidizing reactivity of NapGH homologs has not been verified experimentally and very recent data indicate the involvement of other components in quinol-oxidation (i.e. Rieske/cytochrome bc complex) in the nitrate respiratory chain of the non-dehalogenating Wolinella succinogenes (Hein, Witt and Simon 2017). NapG homologs are characterized by the presence of 16 cysteine residues that are proposed to bind four [4Fe–4S] clusters. The NapG precursor bears an N-terminal Tat signal peptide that identifies the protein as a substrate of the Tat translocase. The mature NapG protein of W. succinogenes was detected in the periplasm (Kern and Simon 2008), where it is expected to interact with the periplasmic nitrate reductase for electron transfer. Assuming that the gene product of SMUL_1541, which shows homology to NapG, interacts with the membrane-attached PceASmul, then the final step in the electron transfer to the RDase proceeds outside the membrane. The iron–sulfur clusters within the membrane-associated quinol dehydrogenase must come into proximity with the Fe–S clusters in PceASmul to allow efficient electron transfer. This assumption has to be taken into consideration when suggestions for the positioning and orientation of PceASmul at the membrane surface are made (Bommer et al. 2014). It is assumed that the SMUL_1541 protein is bound to the outer face of the cytoplasmic membrane by the membrane-integral NapH-like SMUL_1542 gene product. NapH homologues are predicted to contain four transmembrane helices. Eight cysteine residues are conserved in the C-terminal part of the protein (polyferredoxin domain), which is located in the cytoplasm (Brondijk et al. 2004), and these cysteine residues are proposed to bind two [4Fe–4S] clusters. The function of the metal centers in the electron transfer pathway to the periplasmic NapG at the opposite face of the membrane is unknown. Quinone-dependent OHRBs are posed with a thermodynamic problem. The midpoint redox potential of MK (E0′ (MK/MKH2) = −74 mV) (Thauer, Jungermann and Decker 1977) is considerably more positive than the [CoII]/[CoI] redox couple (about −370 mV) of the enzyme-bound cobamide cofactor and the low-potential cubane Fe–S clusters in the RDase. Hence, a driving force is required to convert the thermodynamically unfavorable redox-reaction into a favorable electron transfer. Based on the sensitivity of PCE respiration in S. multivorans to the presence of uncoupling agents such as carbonyl cyanide-p-(trifluoromethoxy)-phenylhydrazone (15 nmol mg−1 protein) a reverse electron flow driven by the proton gradient was concluded as being necessary to overcome the thermodynamic barrier (Miller, Wohlfarth and Diekert 1996). A similar effect resulted when the membrane integrity was perturbed by mild treatment with detergent. An interruption in the electron transfer from hydrogen oxidation to PCE reductive dechlorination by protonophors was not detected in D. restrictus (Schumacher and Holliger 1996) and D. hafniense Y51 (Gadkari 2017). The insensitivity towards uncoupling agents found in the latter two OHRB indicates a different mechanistic solution for facilitating the electron transfer from MKH2 to the low-potential metal centers of the RDase. The distant phylogenetic relationship between the pceC gene product, encoded in the pceABCT operons of D. restrictus (Kruse et al. 2013) and D. hafniense Y51 (Nonaka et al. 2006) and the membrane-integral NapH might indicate a role for PceC in electron transfer comparable to the role proposed for the NapGH-like quinol dehydrogenase in S. multivorans. Low similarity to NosR-like proteins involved in N2O-respiration (Wunsch and Zumft 2005) is also in accord with a role for PceC in electron transfer. PceC is a putative membrane-protein that lacks the polyferredoxin domain present in the C-terminal part of NapH. In contrast, a putative N-terminally bound flavin predicted to be exposed to the exoplasm might form part of the electron transfer chain from MKH2 to the RDase. Soluble, periplasmic flavin-containing proteins in D. dehalogenans might fulfill a similar function (Kruse et al. 2015). This is an interesting proposal, because reduced flavins are able to transfer two electrons at different redox potentials (electron bifurcation; Buckel and Thauer 2013; Peters et al. 2016) and could in theory provide a single low-potential electron for the RDase reaction. So far, there is no concrete evidence to support such a mechanism, but the involvement of flavin radicals or even quinone radicals in electron transfer might be an alternative possibility to circumvent the thermodynamically unfavorable difference in the redox potentials between the quinone pool and the RDase. Since such mechanisms are independent of the proton gradient across the cytoplasmic membrane, uncoupling agents should not have a direct effect. It was shown earlier that the proton gradient appeared to be dispensable for the reductive dehalogenation of ClOHPA coupled to hydrogen oxidation in D. dehalogenans, since the treatment of cells with mild detergents had no effect (van de Pas et al. 2001). Along with the PceC-ortholog CprC in D. dehalogenans, other flavin-containing proteins were also more abundant in cells cultivated with ClOHPA and were discussed as being involved in electron transfer in the ClOHPA respiratory chain (Kruse et al. 2015). Further studies are clearly necessary to unravel the exact role of flavin-containing electron-transferring proteins in cells actively respiring organohalides. Different paths are feasible for the reduction of the MK pool in the quinone-dependent OHRB: (i) the electrons could be transferred directly from a quinone-reactive, membrane-bound dehydrogenase (e.g. membrane-bound uptake hydrogenase or formate dehydrogenase) to the MK pool, or (ii) the electrons could be provided by a soluble dehydrogenase via reduced pyridine nucleotides or soluble electron-transferring proteins, such as ferredoxin/flavodoxin, to a membrane-bound NADH-quinone oxidoreductase or ferredoxin/flavodoxin-quinone oxidoreductase, respectively, which in turn reduces MK. Both options have been discussed to play a role in the PCE respiration of S. multivorans when the organism is cultivated with different electron donors. Based on both transcriptional and biochemical studies, a trimeric membrane-bound [ Ni–Fe] hydrogenase is proposed to be involved in hydrogen oxidation and MK reduction (Kruse et al. 2017). The enriched trimeric membrane-bound [Ni–Fe] hydrogenase, including the quinone-reactive, membrane-integral cytochrome b subunit, was effective in reducing the soluble quinone-analog 2,3-dimethyl-1,4-naphthoquinone (DMN), underpinning the assumption that it reduces the MK pool in H2-grown cells of the organism. When formate serves as electron donor, a membrane-bound formate dehydrogenase might be responsible for MK reduction in S. multivorans (Schmitz and Diekert 2003). During growth on pyruvate and PCE, the electrons derived from pyruvate oxidation by the pyruvate-ferredoxin oxidoreductase were proposed to be transferred via ferredoxin and a ferredoxin-reactive complex I to the MK pool (Goris et al. 2015). QUINONE-INDEPENDENT ORGANOHALIDE RESPIRATION An early study with D. mccartyi strains BAV1 and FL2 reported the presence of quinones in these isolates (White et al. 2005), which suggested a quinone-dependent organohalide respiration in these obligate OHRB. However, D. mccartyi genomes lack the genes for complete biosynthesis of quinones (Kube et al. 2005; Seshadri et al. 2005) questioning the involvement of quinones in organohalide respiration in D. mccartyi strains. Since cultivation of D. mccartyi in synthetic mineral medium without the addition of quinones as vitamins has been reported (Schipp et al. 2013), the only alternative possibility to synthesize quinones would be via a novel biosynthetic pathway. The lack of standard quinone-reactive cytochromes also negates a requirement for quinones. Moreover, a recent study failed to detect any quinoid compound in extracts of D. mccartyi strain CBDB1 using sensitive mass spectrometric analyses (Kublik et al. 2016) and these results clearly distinguish D. mccartyi from other OHRB. Organohalide respiration in D. mccartyi strain CBDB1 was neither inhibited by HQNO (Jayachandran, Görisch and Adrian 2004) nor driven by reduced quinones such as DMNH2, menadiol, menaquinol-4, or diverse ubiquinols (Q-0, Q-4, or Q-10) (Jayachandran, Görisch and Adrian 2004; Kublik et al. 2016). Finally, the addition of protonophors that dissipate the proton gradient, such as carbonylcyanide m-chlorophenyl m-hydrazone and 3,3′,4′,5-tetrachlorosalicylanilide, did not inhibit dehalogenation by whole cells of D. mccartyi strain CBDB1 demonstrating that a reverse electron flow, as proposed for S. multivorans, is not necessary (Jayachandran, Görisch and Adrian 2004). Together, all these findings strongly support the notion of a quinone-independent respiratory chain in D. mccartyi and indicate that it is fundamentally different from that described for other OHRB. The obligate organohalide-respiring Dehalogenimonas species resemble D. mccartyi in lacking quinone biosynthetic genes, which indicates a similar mode of electron transfer (Siddaramappa et al. 2012; Molenda, Quaile and Edwards 2015; Key et al., 2016, 2017). While Dehalogenimonas spp. oxidize hydrogen and formate (Key et al. 2017), D. mccartyi strains are limited to using hydrogen as sole electron donor for organohalide respiration (Löffler et al. 2013). In the absence of quinones, the alternative is protein-mediated electron transfer from a hydrogenase to the RDase. Four different [Ni–Fe] hydrogenases are encoded in the genome of D. mccartyi strain CBDB1, but only one has been proposed to play a role in energy conservation (Kube et al. 2005). The abundance of the hydrogen-uptake (Hup) hydrogenase in D. mccartyi cells (Morris et al. 2007) and its dependence on Tat (Hartwig et al. 2015) predict that it is localized at the exoplasmic face of the cytoplasmic membrane, making its involvement in the organohalide respiratory chain highly likely (Kube et al. 2005; Mansfeldt et al. 2014). However, although the hup gene cluster encodes the catalytic large subunit (HupL) and the Fe–S-harboring small subunit (HupS), it lacks a hupC gene encoding a membrane-anchoring cytochrome b subunit. Moreover, no cytochromes have been detected in D. mccartyi (Löffler et al. 2013; Schipp et al. 2013). Typically, in other H2-oxidizing organisms, the membrane integral subunit facilitates electron transfer to the quinone pool via b-type cytochromes. The presence of a ferredoxin-like subunit (named HupX; locus tag: cbdbA131 in D. mccartyi strain CBDB1), which is encoded in the hup operon and which is also found in the hydrogenase 2 enzyme of E. coli (Pinske et al. 2015), might indicate alternative means of electron conduction from HupSL to the RDase. Although the hupX gene does not encode its own N-terminal Tat signal peptide, its co-translocation together with the HupSL heterodimer is feasible. This would need HupSLX complex formation in the cytoplasm prior to membrane translocation. A protein-mediated electron transfer path would require a transient or continuous interaction between the hydrogenase, the HupX protein and the RDase. In 2016, (Kublik et al. 2016) were able to show complex formation between HupSLX, a RDase, and a complex iron–sulfur molybdoenzyme in D. mccartyi strain CBDB1 by extracting membrane proteins under native conditions, separating protein complexes by native polyacrylamide gel electrophoresis, and subsequently identifying the polypeptides via mass peptide fingerprinting (Fig. 7). More recently, this complex has been isolated from the membrane of D. mccartyi strain CBDB1, enriched and been shown to catalyze hydrogen-dependent reduction of 1,2,3-trichlorobenzene (Hartwig et al. 2017), providing further support for quinone-independent electron transfer within the complex. Figure 7. View largeDownload slide Tentative scheme for the composition of the quinone-independent organohalide respiratory chain of D. mccartyi strain CBDB1 (Kublik et al. 2016). MGD, molybdopterin guanine dinucleotide; OmeAB, organohalide-respiration involved molybdoenzyme subunits A and B. Figure 7. View largeDownload slide Tentative scheme for the composition of the quinone-independent organohalide respiratory chain of D. mccartyi strain CBDB1 (Kublik et al. 2016). MGD, molybdopterin guanine dinucleotide; OmeAB, organohalide-respiration involved molybdoenzyme subunits A and B. While no direct interaction between HupSL and the RDase was observed, both enzymes were found to interact with HupX and the heterodimeric complex iron–sulfur molybdoenzyme complex. This complex consists of two proteins encoded by the genes cbdbA195 and cbdbA193 (Kublik et al. 2016). Here we propose to name the two genes omeA and omeB, for organohalide-respiration involved molybdoenzyme subunits A and B (see Fig. 7). OmeA has a Tat signal peptide at its N-terminus and is transported to the exoplasm like HupSL and the RDases. OmeB contains 10 transmembrane helices and notably is similar to the membrane anchor of hydrogenase 2-like enzymes (Pinske et al. 2015; Hartwig et al. 2017), suggesting that OmeB represents the docking site for HupSL. The transcription of the ome and the hup operons is tightly linked to the presence of halogenated compounds such as trichlorobenzenes, which strongly supports their functional linkage to reductive dehalogenation (Hartwig et al. 2017). Although OmeA shows amino acid sequence similarity to respiratory formate dehydrogenases, it lacks the key active site (seleno)cysteine and histidine residues (Hartmann, Schwanhold and Leimkühler 2015) required to catalyze formate oxidation. Moreover, formate oxidation has never been observed in D. mccartyi cultures and thus OmeA clearly has a different function (Hartwig et al. 2017). The polytopic membrane-integral OmeB could play a role in coupling electron transfer through the respiratory complex to proton translocation across the membrane (Fig. 7). Based on amino acid sequence similarity, its role could resemble that of the membrane anchor of H2-oxidizing hydrogenase 2. Sequence analysis of OmeB revealed the presence of a conserved glutamate in the membrane-spanning part of the protein, which is consistent with an involvement in proton translocation coupled to a conformational change (Zinder 2016). CONCLUSIONS AND OUTLOOK The exploration of the mechanisms coupling reductive dehalogenation to ATP synthesis via electron transport phosphorylation in OHRB has brought a diverse and complex picture to light. Breakthroughs in the analysis of structure–function relationships in cobamide-containing reductive dehalogenases, the description of novel respiratory chain components, and the identification of the potential mode of proton transfer have substantially improved our understanding of the physiology of OHRB. However, although structural information on reductive dehalogenases is available, neither the catalytic mechanism(s) nor the broad substrate spectrum of these enzymes is understood in detail. The molecular basis of the differentiation into RDases catalyzing hydrogenolysis/halogen substitution or vicinal reduction/dihaloelimination or both needs to be elucidated. Furthermore, the definition of the coupling sites that link the membrane-bound electron transfer chain to the translocation of protons across the cytoplasmic membrane is unclear. Thus, there remain many aspects of this exciting field of research that require further study. 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FEMS Microbiology EcologyOxford University Press

Published: Apr 1, 2018

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