Anaerobic biodegradation of biomass in sediments. Annually, plant and photosynthetic microorganisms fix 70 billion tons of carbon into biomass made up of complex biopolymers, such as cellulose, hemicellulose, lignin, proteins and lipids (Thauer et al., 2008). About 1% of this material is mineralized in various anaerobic niches of nature through a process that yields methane and carbon dioxide as end products (Figure 2). The combination of photosynthesis (GO:0015979) and macromolecule catabolism (GO:0009057) constitutes the biological component of the biogeochemical process of carbon cycling.

Cellulose is a polymer of D-glucose units connected by β (1→4) bonds. The anaerobic mineralization of cellulose (synonym of “cellulose catabolic process, anaerobic,” GO:1990488) starts with hydrolysis of the β (1→4) bonds by cellulases (GO:0008810) produced by anaerobic cellulolytic bacteria and fungi (Adney et al., 1991; Teunissen and Op Den Camp, 1993; Leschine, 1995; Li et al., 1997; Schwarz, 2001; Ljungdahl, 2008; Ransom-Jones et al., 2012). These organisms either secrete the cellulases or carry these enzymes on their cell surfaces (Teunissen and Op Den Camp, 1993; Li et al., 1997). A recent study shows that excreted enzymes with multiple catalytic sites and multiple cellulose-binding modules provide Caldicellulosiruptor bescii, an anaerobic thermophile with a high activity of cellulose degradation (Brunecky et al., 2013). The cellulose catabolic process (GO:0030245) involves the actions of endo-β-1,4-glucanases (GO:0052859) and exo-1,4-β-glucanases or cellobiohydrolases (CBH) (reducing-end-specific, GO:0033945; non-reducing-end-specific, GO:0016162) that generate cellobiose, with intermediate formation of fragments with multiple glucose units (Akin, 1980; Beguin and Aubert, 1994; Bayer et al., 1998; Perez et al., 2002; Hilden and Johansson, 2004), and hydrolysis of cellobiose to glucose (cellobiose catabolic process, GO:2000892) by β-glucosidase (GO:0008422).

The cellulose degrading anaerobic microorganisms and other non-cellulolytic anaerobes with access to the products generated by cellulolytic microbes take up and ferment D-glucose to acetate, alcohols, lactate and fatty acids (e.g., propionate, butyrate) via respective biosynthetic processes (Figure 3) (Zinder, 1993; Schink, 1997; Ahring, 2003; McInerney et al., 2009). The butyrate biosynthetic process (GO:0046358) involves an intermediate formation of acetyl-CoA (acetyl-CoA biosynthetic process, GO:0006085) whereas for propionate biosynthesis (GO:0019542) succinate generated via the tricarboxylic acid metabolic process or TCA cycle (GO:0072350) serves as the direct precursor (Zinder, 1993; Schink, 1997; Ahring, 2003; McInerney et al., 2009).

Butyrate and propionate, which are called short-chain fatty acids (SCFAs), are further fermented to acetate, hydrogen and CO₂ (fatty acid catabolic process: GO:0009062; child term, anaerobic fatty acid catabolic process, GO:1990486) via their respective catabolic processes (butyrate catabolic process, GO:0046359; propionate catabolic process, GO:0019543) (Zinder, 1993; Schink, 1997; Ahring, 2003; McInerney et al., 2009); Syntrophobacter, Syntrophomonas, Syntrophus, Smithella, and Pelotomaculum species are some of the bacteria that produce these SCFAs. Ethanol and lactate are also fermented to acetate, hydrogen and CO₂ (ethanol catabolic process, GO:0006068; anaerobic lactic acid catabolic, process GO:1990485). The hydrogen biosynthetic process (GO:1902422) is a key element of these fermentation processes and those described in the preceding paragraph for the following reason. Several steps of fermentation lead to the reduction of electron carriers such as NAD⁺ and ferredoxin, producing NADH and reduced-ferredoxin. For the fermentation process to continue, NAD⁺ and ferredoxin must be regenerated, and often the only available route to meet this requirement is the reduction of protons, yielding molecular hydrogen (H₂) (GO:1902422) (Zinder, 1993; Schink, 1997; Ahring, 2003; McInerney et al., 2009).

Degradation of hemicellulose follows a path similar to that described for cellulose (summarized in Figure 3). The term hemicellulose includes xylan (polymer of xylose), glucuronoxylan (polymer of D-glucuronate and xylose), arabinoxylan (polymer of arabinose and xylose), glucomannan (polymer of glucose and mannose), galactomannan (polymer of galactose and mannose), and xyloglucan (polymer of xylose, glucose and galactose) (Akin, 1980; Perez et al., 2002). These are degraded via specific hemicellulose catabolic processes (GO:2000895) to their respective monomers (Akin, 1980; Perez et al., 2002). The fermentation of monomers yields acetate, hydrogen and CO₂ (Wolin and Miller, 1983; Schink, 1997).

Lignin degradation in anaerobic environments (anaerobic lignin catabolic process, GO:1990487) is not well studied and is considered rare to impossible (Akin, 1980; Harwood and Gibson, 1988; Perez et al., 2002; Fuchs, 2008); the broader lignin catabolic process is generally considered an aerobic process (Perez et al., 2002). However, following the degradation of lignin by aerobic microorganisms such as fungi, a variety of aromatic compounds (catechol, benzoate, p-hydroxybenzoate, vanillate-, ferulate, syringate, p-hydroxybenzoate, p-hydroxycinnamate, and 3-methoxy-4-hydroxyphenylpyruvate) (Kaiser and Hanselmann, 1982a,b) become available in anaerobic environments. Fermentation of these aromatic compounds by anaerobic bacteria leads to acetate, CO₂ and hydrogen (Harwood and Parales, 1996; Fuchs, 2008). Anaerobic degradation of benzoate, one of the lignin monomers, has been studied in detail and this catabolic process (benzoate catabolic process via CoA ligation, GO:0010128) yields acetate and CO₂ (Figure 3). The metabolism of several other lignin monomers by anaerobes has also been investigated (Harwood and Parales, 1996; Fuchs, 2008) and some of the relevant information for vanillin, ferulate and catechol is summarized in Figure 3.

Anaerobic lipid catabolic processes also lead to acetate and hydrogen (McInerney, 1988; Zinder, 1993; Schink, 1997). The process begins with the hydrolysis of lipids (lipase activity, GO:0016298); the broader lipid catabolic process is represented by GO:0016042. The glycerol released by hydrolysis enters the glycolysis pathway generating acetate and hydrogen (Zinder, 1993) (Figure 3). The fatty acid units are degraded via the β-oxidation pathway (fatty acid beta-oxidation, GO:0006635) to acetate and the excess reducing equivalents are released as hydrogen (Figure 3). In the case of proteins, the amino acids released by the action of proteases or peptidases (peptidase activity, GO:0008233; endopeptidase activity GO:0004175, exopeptidase activity, GO:0008238) are deaminated oxidatively, releasing ammonia and hydrogen, and then the resulting ketoacids are fermented to acetate and hydrogen (McInerney, 1988; Zinder, 1993; Schink, 1997) (Figure 3).

In each of the above cases, as H₂ accumulates the oxidation of reduced electron carriers becomes thermodynamically unfavorable and consequently the fermentation process slows down or even halts (McInerney, 1988; Zinder, 1993; Schink, 1997). Methanogens consume hydrogen and reduce CO₂ to methane, thus relieving the block on fermentation (McInerney, 1988; Zinder, 1993; Schink, 1997). These archaea also convert acetate to methane and CO₂ and this action also improves the thermodynamics of biodegradation (Zinder, 1993). As CH₄ moves to aerobic zones, such as the surface of water overlaying sediments, methanotrophic bacteria oxidize this hydrocarbon to CO₂ (methane catabolic process, GO:0046188) (Kiene, 1991; Zinder, 1993; Conrad, 2007, 2009). More recent work shows that significant amount of methane is oxidized anaerobically and the microbial basis and mechanistic details of this process are beginning to emerge (Conrad, 2009; Thauer, 2011; Milucka et al., 2012; Shima et al., 2012; Haroon et al., 2013; Offre et al., 2013). Hence, by removing the hydrogen-induced thermodynamic block and converting acetate to methane, methanogens facilitate the complete degradation of the biopolymers discussed above.

In marine anaerobic sediments rich in sulfates some of the products of biomass degradation also lead to methane production. In general however, in this environment hydrogen and acetate are not available to the methanogens as, in the presence of sulfate, sulfate-reducing bacteria readily use these materials to reduce sulfate to hydrogen sulfide (dissimilatory sulfate reduction, GO:0019420), and the growth rates and affinities for H₂ of the sulfate-reducing bacteria are much higher than those of the methanogens (Widdel, 1988). However, several hydrogen-consuming methanogens belonging to the class of Methanococci have been isolated from marine environments (Whitman et al., 1986). It has been speculated that these organisms may depend primarily on formate which could arise from the catabolism of oxalate (GO:0033611) derived from plant materials (Allison et al., 1985); most methanococci are capable of consuming both hydrogen and formate (Boone et al., 1993).

A significant amount of methane is also produced from methylamines, methylsulfides and carbon monoxide (Zinder, 1993; Thauer, 1998; Deppenmeier et al., 1999; Ferry, 1999, 2011; Liu and Whitman, 2008; Thauer et al., 2008). The sources of methylamines are betaine and choline, (GO:0006579 and GO:0042426, respectively) while methylsulfides are generated from sulfur-containing compounds such as methionine and dimethylpropiothetin (GO:0009087; and GO:0047869, respectively; Figure 3) (Boone et al., 1993). In certain marine environments, carbon monoxide provided by kelp algae provides both reductant and carbon for methanogenesis (methane biosynthetic process from carbon monoxide, GO:2001134) (Rother and Metcalf, 2004; Lessner et al., 2006).

Anaerobic biodegradation of biomass in animal intestines. Foregut fermenting animals such as the ruminants (cattle, sheep, goats) as well as hindgut fermenters such as human, termites, and horse, employ variations of the overall process shown in Figure 2 for deriving nutrients from feed or food (Wolin, 1981; Wolin and Miller, 1983; Zinder, 1993; Miller and Wolin, 1996; Weimer, 1998; Hook et al., 2010; Sahakian et al., 2010). In cattle and many other foregut fermenters, the rumen serves as the first site for the degradation of forage (Wolin, 1981; Weimer, 1998; Hook et al., 2010). The residence time for the feed in rumen is rather short (5.6 h for the fluid and 35 h from particulates in rumen; compared to about 4.5 months even for nitrate, a soluble compound, in freshwater sediment) (Hristov et al., 2003), which is not conducible for significant growth and activity of slow-growing fatty acid-fermenting bacteria and acetoclastic methanogens (Zinder, 1993). Thus, in this digestive chamber the fatty acids and acetate are not converted to methane, rather are absorbed by the animal for nutrition (Zinder, 1993). The hydrogen and formate produced during the fermentation are converted to methane by methanogenic archaea. All plant material contains pectin, a methylated carbohydrate, and leaves, shoots and fruit are particularly rich in it. Anaerobic degradation of pectin (anaerobic pectin catabolic process, GO:1990489) serves as an important source of methanol in anaerobic environments (Schink et al., 1981). Thus, ruminants could carry methanogens in their rumens capable of utilizing methanol for methanogenesis and in some cases this has been shown to be true (Mukhopadhyay et al., 1992; Zinder, 1993).

In the hindgut of humans, i.e., the large intestine, the undigested material delivered from the small intestine is fermented, generating fatty acids, some hydrogen, and formate, and the latter two are converted to methane (Wolin, 1981; Zinder, 1993; Miller and Wolin, 1996; Sahakian et al., 2010; Flint, 2011). The process is beneficial to the host as it provides the fatty acids as additional nutrients. However, an uncontrolled production of fatty acids in this hindgut activity has been identified as one of the possible causes of obesity (Schmitz and Langmann, 2006; Nakamura et al., 2010; Sahakian et al., 2010; Flint, 2011).

In certain foregut fermenters such as kangaroos and wallabies and hindgut fermenters such as termites, the removal of hydrogen during biodegradation of complex polymers occurs through acetate formation and not methanogenesis (Brune and Friedrich, 2000; Gagen et al., 2010; Klieve et al., 2012).

Aerobic treatment of municipal and industrial wastes via methods such as activated sludge requires energy input for supplying oxygen (Switzenbaum, 1983; Zinder, 1993). The process also generates a significant amount of microbial biomass (Zinder, 1993), which cannot be discharged to waterways (Zinder, 1993; Paul and Debellefontaine, 2007). In contrast, anaerobic methods not only require much less energy input and produce very little microbial biomass, but also conserve most of the energy present in the waste materials in the form of methane, which can be recovered as fuel (Zinder, 1993; Gao et al., 2014). Anaerobic waste treatment and the production of methane as biofuel from renewable resources follow the basic biological process (macromolecules catabolic process, GO:0009057) that has been described above for methanogenic biodegradation of biomass in sediments (Zinder, 1993). The mixture of methane and carbon dioxide that is produced in all these cases is known as biogas (Ducom et al., 2009). The biogas obtained from waste treatment facilities as well as from bio-digesters processing high sulfur feedstock contains a substantial amount of hydrogen sulfide and nitrogen oxides (Zinder, 1993; Fdz-Polanco et al., 2001; Janssen et al., 2001; Ducom et al., 2009; Diaz and Fdz-Polanco, 2012). Several methods for the removal of these unwanted compounds have been developed and research for developing even better separation methods continues (Fdz-Polanco et al., 2001; Janssen et al., 2001; Ducom et al., 2009; Diaz and Fdz-Polanco, 2012).

This process (GO:0019386) utilizes hydrogen as the primary source of electron or reductant (Ferry, 1999; Deppenmeier and Muller, 2008; Thauer et al., 2008). It can operate with or without the involvement of cytochromes (Ferry, 1999; Deppenmeier and Muller, 2008; Thauer et al., 2008). The latter is utilized by methanogens that lack cytochromes (Ferry, 1999; Deppenmeier and Muller, 2008; Thauer et al., 2008) and is considered one of the most ancient respiratory metabolisms on earth (Leigh, 2002). We describe the process starting with the carbon transfer and reduction steps, followed by the energy production avenues.

Carbon transfer and reduction. It is believed that carbon dioxide (CO₂) is captured by methanofuran (MF) to form an unstable compound called carboxy-MF (Thauer et al., 1993) which is reduced by formyl-MF dehydrogenase (GO:0018493) in an energy-dependent (endergonic) manner to formyl-MF with a low-potential ferredoxin (Fd) serving as electron carrier (Thauer et al., 2008). Formyl-MF dehydrogenase exists in two forms, one of which contains molybdenum (Fmd) and the other tungsten (Fwd) (Thauer et al., 1993); molybdenum and tungsten are found to be bound to a molybdopterin and growth conditions dictate which metal will be incorporated (Hochheimer et al., 1995). At the next step the formyl group is transferred to H₄MPT by a transferase enzyme (Ftr, GO:0030270) to form formyl-H₄MPT (Donnelly and Wolfe, 1986; Breitung and Thauer, 1990; Thauer et al., 1993). From this stage H₄MPT carries four forms of the fixed carbon representing three oxidation states (Wolfe, 1991, 1993; Thauer et al., 1993, 2008; Ferry, 1999; Deppenmeier and Muller, 2008). First formyl-H₄MPT is dehydrated by methenyl-H₄MPT cyclohydrolase (Mch, GO:0018759) to form methenyl-H₄MPT (Donnelly et al., 1985; Dimarco et al., 1986; Mukhopadhyay and Daniels, 1989; Klein et al., 1993), which in turn is reduced to methylene-H₄MPT by the action of one of the two enzymes, F₄₂₀-dependent methylene-H₄MPT dehydrogenase (Mtd, GO:0030268) and a Fe-containing hydrogenase (Hmd, GO:0047068) (Hartzell et al., 1985; Mukhopadhyay and Daniels, 1989; Von Bunau et al., 1991; Schworer et al., 1993; Thauer et al., 1993; Mukhopadhyay et al., 1995). Mtd utilizes reduced F₄₂₀ (F₄₂₀H₂) as reductant whereas Hmd retrieves electrons from molecular hydrogen (H₂) (Hartzell et al., 1985; Mukhopadhyay and Daniels, 1989; Von Bunau et al., 1991; Schworer et al., 1993; Thauer et al., 1993; Mukhopadhyay et al., 1995). Methanogens with Hmd also carry paralogs of this protein (HmdII and HmdIII), but these proteins do not reduce methylene-H₄MPT (Lie et al., 2013). Two roles of HmdII and HmdIII have been proposed: a. guiding the maturation of Hmd and b. linking energy production and protein synthesis (Oza et al., 2012; Lie et al., 2013). Methylene-H₄MPT is reduced with F₄₂₀H₂ and by the action of F₄₂₀-dependent methylene-H₄MPT reductase (Mer, GO:0018537), providing the last H₄MPT derivative on the pathway, methyl-H₄MPT (Ma and Thauer, 1990; Te Brommelstroet et al., 1990; Ma et al., 1991; Thauer et al., 2008). The transfer of the methyl group from methyl-H₄MPT to coenzyme M is catalyzed by a membrane-bound sodium ion (Na⁺)-pumping enzyme complex called methyl-H₄MPT:coenzyme M methyl transferase (Mtr, GO:0044677) (Becher et al., 1992; Kengen et al., 1992; Gartner et al., 1993). This complex not only yields methyl-coenzyme M (CH₃-CoM), but also generates a Na⁺-gradient that is used for energy production (see below) (Ferry, 1999; Deppenmeier and Muller, 2008; Thauer et al., 2008). The next step in the sequence yields methane. This last carbon-reduction reaction is catalyzed by CH₃-CoM reductase (GO:0044674) with coenzyme B (HS-CoB or HS-HTP) serving as an electron source, resulting in a heterodisulfide, CoM-S-S-CoB, as product in addition to methane (Wolfe, 1991, 1992; Ferry, 1999; Deppenmeier and Muller, 2008; Thauer et al., 2008). The heterodisulfide is reduced by a reductase (Hdr, GO:0051912) to regenerate HS-CoM and HS-CoB (Ferry, 1999; Deppenmeier and Muller, 2008; Thauer et al., 2008). Hydrogen-oxidizing methanogens often carry two CH₃-CoM reductase isozymes (McrI and McrII) (Rospert et al., 1990), one of which is effective under high hydrogen availability and the other under low hydrogen conditions (Rospert et al., 1990).

Energy conservation. First, we describe the details for methanogens lacking cytochromes. The first site of energy conservation is the Mtr reaction (Ferry, 1999; Deppenmeier and Muller, 2008; Thauer et al., 2008). The Na⁺-gradient generated at this step is directly used for the production of ATP by a membrane-bound AoA1-ATP synthase (GO:1990490) (Deppenmeier and Muller, 2008). Under certain conditions this gradient assists two membrane-associated and energy-converting hydrogenase complexes, EhaA-T and EhbA-Q, to generate reduced Fd with the ability to deliver low redox potential electrons (Thauer et al., 2008; Costa et al., 2010; Lie et al., 2012). The reduced Fd molecules generated by EhaA-T are used for the endergonic formyl-MF dehydrogenase reaction that yields formyl-MF, and those provided by EhbA-Q are used for cellular biosynthesis (Porat et al., 2006; Thauer et al., 2008; Costa et al., 2010; Major et al., 2010; Kaster et al., 2011; Lie et al., 2012). The next energy yielding step is the reduction of CH₃-CoM and it is not clear whether the methanogens conserve this energy or the energy is released to strongly favor the forward reaction toward methane formation (Thauer et al., 2008). The reduction of CoM-S-S-CoB involves rather complex electron transfer mechanisms and also is a major site for energy conservation (Thauer et al., 2008; Costa and Leigh, 2014).

In certain methanogens without cytochromes, the reduction of CoM-S-S-CoB and formyl-MF generation is coupled via a novel mechanism called bifurcation (Thauer et al., 2008; Costa et al., 2010; Kaster et al., 2011; Lie et al., 2012) (Figure 5). Here, the Vhu hydrogenase retrieves electrons from hydrogen and transfers those to soluble heterodisulfide reductase (Hdr). Hdr utilizes these electrons for two purposes (Figure 5): (i) converting CoM-S-S-CoB to HS-CoM and HS-CoB, which requires a relatively lower investment of energy; and (ii) reducing a low potential ferredoxin, which is energetically suitable for the highly energy intensive reduction of CO₂ and generation of formyl-MF (Thauer et al., 2008; Costa et al., 2010; Kaster et al., 2011; Lie et al., 2012). This novel mechanism, where a single input (electrons of moderately low potential) is used to generate two outputs (two pools of electrons, with potentials that are higher and much lower than the input) is called electron bifurcation (GO:MENGO-UR). It is a major factor in energy conservation in methanogens as it helps to perform a highly endergonic reaction, such as the generation of formyl-MF, without an investment of ATP or an ion gradient (Thauer et al., 2008; Costa et al., 2010; Kaster et al., 2011; Lie et al., 2012). It seems to be an important tool for energy poor anaerobes (Thauer et al., 2008; Kaster et al., 2011). When withdrawal of intermediates from the methanogenesis pathway for biosynthesis causes a drop in CoM-S-S-CoB levels, the bifurcation process is rendered less efficient (Lie et al., 2012); in that case, as described in the preceding paragraph, an ion-driven hydrogenase system (EhaA-T) is employed for the generation of formyl-MF and this could be considered to be a type of anaplerosis (GO:MENGO-UR) (Lie et al., 2012).

Methanogens with cytochromes do not employ the bifurcation mechanism. Instead, a membrane-bound complex composed of a cytochrome-containing heterodisulfide reductase (HdrDE) (GO:0044678) and a hydrogenase (VhoECG) where VhoC is a b-type cytochrome is utilized (Thauer et al., 2008). The electrons derived from hydrogen by VhoECG are utilized by HdrDE for reduction of CoM-S-S-CoB (Thauer et al., 2008). The overall process is exergonic and thus, in addition to reducing CoM-S-S-CoB, the VhoECG-Hdr complex utilizes excess energy to extrude protons out of the cell. The low potential reduced ferredoxin, which is needed for the generation of formyl-MF, is provided by a proton-gradient-driven membrane-bound energy-converting hydrogenase complex (EchA-F).

As mentioned above, several methanogenesis enzymes form large protein complexes (GO:0043234) and some these are membrane bound (GO:0019898) and include specialized non-enzyme units such as ion pumps and lipid soluble small compounds. One example is the soluble heterodisulfide reductase complexes of methanogens that lack cytochromes, and can be described using a molecular function term, GO:0051912 (“CoB-CoM heterodisulfide reductase activity”) and two cellular component terms GO:0044678 (“CoB-CoM heterodisulfide reductase complex”) and GO:0043234 (“protein complex”). In the case of cytochrome-containing methanogens, one further additional cellular component GO term is available for a full description, namely GO:0019898 (“extrinsic component of membrane”).

The carbon transfer and reduction steps in this process (GO:2001127) are similar to those described above for methanogensis from H₂+CO₂ (GO: 0015948). Both the CO₂ and reducing power are derived from formate by the action of an F₄₂₀-dependent formate dehydrogenase (FdhABC) (GO:0043794) (Schauer and Ferry, 1982; Lie et al., 2012); FdhAB subunits form the enzyme that produces CO₂ and reduced F₄₂₀ or F₄₂₀H₂ (HCOO⁻ + H⁺ + F₄₂₀ → CO₂ + F₄₂₀H₂) and FdhC is thought to import formate into the cell (Wood et al., 2003). CO₂ is converted to methane using the CO₂-reduction pathway described in Figure 4. Some of the reduced F₄₂₀ (F₄₂₀H₂) participates directly in the Mtd and Mer reactions and a part of it is used by a bifurcating complex that provides electrons of appropriate redox potentials to heterodisulfide reductase and formyl-MF dehydrogenase. In the composition and some of the properties this bifurcating complex differ from the one employed for methanogenesis from H₂+CO₂ (see above). When a methanogen grows on formate, a part of the Fdh pool associates with the Hdr and Vhu/Vhc hydrogenases, and together with a formyl-MF dehydrogenase they form a bifurcating complex (Lie et al., 2012). This Fdh-containing bifurcating complex utilizes electrons from F₄₂₀H₂ (produced by Fdh) and generates high and low potential electrons, either directly or via production of hydrogen as intermediate, that are consumed in the reduction of CoM-S-S-CoB and the generation of formyl-MF, respectively (Lie et al., 2012).

Only a few methanogens can perform methanogenesis with secondary alcohol as electron source (GO:MENGO-UR; also secondary alcohol catabolic process, GO:MENGO-UR) (Widdel, 1986; Bleicher et al., 1989; Widdel and Wolfe, 1989; Schirmack et al., 2014). These substrates are oxidized to their respective ketones to provide reducing equivalents for the reduction of carbon dioxide to methane via the pathway shown in Figure 3 (Boone et al., 1993; Zinder, 1993). Ethanol, when used, is converted to acetaldehyde (methanogenesis with ethanol as electron source, GO:MENGO-UR; ethanol catabolic process, GO:0006068). These conversions are consistent with the general observation that methanogens cannot break carbon-carbon bonds in energy substrates other than that found in acetate (see below). Two types of alcohol dehydrogenase have been found in these organisms: one reduces nicotinamides (NAD⁺ or NADP⁺) (GO:0004022 and GO:0008106), and the other transfers electrons to coenzyme F₄₂₀ during alcohol oxidation (GO:0052753) (Widdel and Wolfe, 1989). Most of these enzymes have broad specificities allowing the organisms to use ethanol, 2-propanol, 2-butanol, 2-pentanol, cyclopentanol, cyclohexanol, and 2,3-butanediol (Bleicher et al., 1989).

Many methanogens can utilize carbon monoxide (CO) although higher levels of this gas inhibit growth of these archaea (Daniels et al., 1977; O'brien et al., 1984; Rother and Metcalf, 2004; Lessner et al., 2006). Three routes of CO utilization have been found in these organisms. In one, called methanogenesis from carbon monoxide (GO:2001134), CO is simply oxidized to CO₂ by carbon monoxide dehydrogenase (CODH) (GO:0008805), and the resulting two electrons are used for either hydrogen production (GO:1902422) or ferredoxin reduction (Daniels et al., 1977; Ferry, 1999; Vepachedu and Ferry, 2012). Then the hydrogen and/or reduced ferredoxin are used for methanogenesis from CO₂ (GO:0019386). Overall, for every four moles of CO oxidized, one mole of methane and 3 moles of CO₂ are produced. The second mode of CO utilization has been found in Methanosarcina acetivorans where methanogenesis (GO: 0015948) is inhibited by CO but growth is not (Rother and Metcalf, 2004). This organism uses two non-methanogenic routes for energy production (Rother and Metcalf, 2004), the primary one being acetogenic (acetate biosynthetic process, GO:0019413) and the secondary one being formate-forming (formate biosynthetic process, GO:0015943). Even under these conditions methanogenesis operates at low rates, primarily to provide cellular biosynthetic precursors (Rother and Metcalf, 2004). Here methanogenesis from CO₂ involves novel enzymes that transfer the methyl group of CH₃-H₄MPT to CH₃-CoM and serve in the accompanying energy conservation (Lessner et al., 2006); the methyl transfer step could involve a cytoplasmic methyltransferase (CmtA) in addition to a membrane-bound methyl-H₄MPT:coenzyme M methyl transferase (Mtr) (Vepachedu and Ferry, 2012). In the third route, CO promotes the production of dimethyl sulfide and methanethiol (3CO + H₂S + H₂O → CH₃SH + 2CO₂) and energy is conserved via a yet to be identified system (Moran et al., 2008).

Methanogenesis from all of these substrates involves the formation of methyl-CoM as an intermediate (Ferry, 1999). When methanol serves as the sole substrate for methanogenesis (GO:0019387), it provides both carbon and reductant for methanogenesis and this process consumes four moles of methanol for every three moles methane generated. Of these, one mole of methanol is oxidized to CO₂, generating six-electron equivalents of reductant, which are then used to convert three moles of methanol to three moles of methane (Keltjens and Vogels, 1993). The oxidation of methanol to CO₂ involves a part of the CO₂ reduction pathway, but in the reverse direction (Figure 3) (Wassenaar et al., 1998). The methyl groups enter this oxidation process at the methyl-coenzyme M stage by the action of two methyl transferases, MT1 and MT2 or MT2-M (Van Der Meijden et al., 1984a,b; Keltjens and Vogels, 1993; Wassenaar et al., 1998; Ferry, 1999). MT1 is a two-subunit enzyme (MtaBC) and MT2-M has one subunit (MtaA). The first reaction involves transfer of the methyl group of methanol by MT1 to the corrinoid co-factor of its MtaC subunit; this is an automethylation process (Van Der Meijden et al., 1984a,b; Wassenaar et al., 1998). Then MT2-M or MtaA transfers the methyl group from MtaC to HS-CoM, generating methyl-coenzyme M(Van Der Meijden et al., 1984a,b; Wassenaar et al., 1998). Existence of isozymes of MT1 catering to the growth on methanol under various conditions has also been reported (Bose et al., 2006). The methyl groups destined for oxidation are transferred from CH₃-CoM to H₄MPT by the membrane-bound methyl-H₄MPT:coenzyme M methyl transferase (Mtr) (Fischer et al., 1992; Sauer et al., 1997; Ferry, 1999). This endergonic reaction is assisted by a Na⁺-gradient and generates CH₃-H₄MPT (Deppenmeier et al., 1999; Ferry, 1999; Deppenmeier and Muller, 2008). The steps from CH₃-H₄MPT to CO₂ are a reversal of those used for CO₂-reduction, except the organisms performing this process lack Hmd and F₄₂₀-dependent Mtd performs the oxidation of methylene-H₄MPT to methenyl-H₄MPT (Thauer et al., 1993; Deppenmeier et al., 1999; Ferry, 1999; Deppenmeier and Muller, 2008).

Utilization of mono-, di- and tri-methylamines (MMA, DMA, and TMA) for methanogenesis (GO:2001128, GO:2001129, GO:2001130 respectively) follows the general process that is described above for methanol except that substrate-specific methyl transferases are involved in the transfer of methyl groups to coenzyme M. Using MT1 and MT2 of the methanol systems as the reference the methylamine-specific methyl transferases have been named as follows (Wassenaar et al., 1996, 1998; Ferguson and Krzycki, 1997; Burke et al., 1998; Ferry, 1999; Ferguson et al., 2000; Paul et al., 2000; Bose et al., 2008): for MMA, MMAMT+MMCP (MT1) and MT2-A (MT2); for DMA, DMA-MT (MT1) and MT2-A (MT2); for TMA, TMA-52+TCP (MT1) and MT2-A (MT2). For methanogenesis from TMA, MT2-A could be substituted by MT2-M (Ferry, 1999). Methanogenesis from methylated thiols (methanethiol, dimethylsulfide, or methylmercaptopropionate; GO:2001133, GO:2001131, and GO:2001132) also involves special methyl transferase proteins (Tallant et al., 2001; Bose et al., 2009). For example, dimethylsulfide is converted to methyl-CoM by the actions of MtsB (MT2) and MtsA (MT2) (Tallant et al., 2001).

Energy conservation during methanogenesis from methylated compounds occurs in at least two ways. The F₄₂₀H₂ generated during the oxidation of the methyl group of CH₃-H₄MPT to CO₂ is oxidized via the membrane-bound F₄₂₀H₂-dehydrogenase complex (reduced coenzyme F₄₂₀ dehydrogenase activity, GO:0043738), and in the process a lipid soluble membrane-resident cofactor called methanophenazine is reduced (Deppenmeier and Muller, 2008). These events lead to the extrusion of two protons per F₄₂₀H₂ oxidized (Deppenmeier and Muller, 2008). There is another avenue that produces the same outcome and it begins with the release of molecular hydrogen through the oxidation of F₄₂₀H₂ by a soluble F₄₂₀-dependent hydrogenase (Frh, GO:0050454) (Kulkarni et al., 2009). This hydrogen upon its release from the cell is captured by a membrane-bound hydrogenase complex (Vht/Vtx) (GO:MENGO-UR, GO:MENGO-UR), which transfers electrons generated from the oxidation of hydrogen to methanophenazine and releases two protons outside the cell (Kulkarni et al., 2009). In certain methanogens the latter process is the major route of F₄₂₀H₂ oxidation (Kulkarni et al., 2009). The reduced methanophenazine produced by these reactions is utilized by the membrane-bound heterodisulfide reductase (Hdr)-cytochrome b2 complex (GO:MENGO-UR) for the reduction of CoM-S-S-HTP, and this process provides two more protons outside the cell (Deppenmeier and Muller, 2008; Kulkarni et al., 2009; Welte and Deppenmeier, 2014). All these extruded protons generate proton-motive force, which drives ATP synthesis (GO:0015986) via an ATP synthase (GO:0045259) (Deppenmeier and Muller, 2008; Kulkarni et al., 2009; Welte and Deppenmeier, 2014).

A GO term for this process has recently been proposed by us (methanogenesis from H₂ and methanol, GO:1990491). Here, the methyl group of methanol is transferred to coenzyme M by two methyl transferases, MT1 and MT2, producing methyl-CoM (Keltjens and Vogels, 1993). The rest of the process, the reduction of methyl-CoM by HS-CoB, the reduction of CoM-S-S-CoB by electrons derived from hydrogen, and the energy conservation, likely follows the system described in the section on methanogenesis from H₂ plus CO₂; an exception is Methanosphaera stadtmanae, which grows only on H₂ plus methanol with a supplement of acetate (Miller and Wolin, 1985), as it would employ the cytochrome-independent system (Fricke et al., 2006).

About 70% of the biologically produced methane originates from acetate (GO:0019385) (Ferry, 1992, 1993, 1999). The methyl group of acetate is reduced to methane and the carboxyl group is oxidized to CO₂ providing the reductant for methyl reduction (Ferry, 1992, 1993, 1999, 2011; Thauer et al., 2008). The process begins with the activation of acetate by the action of one of two systems, one involving acetate kinase and phosphotransacetylase (GO:0008776 and GO:0008959) and the other catalyzed by acetyl-CoA synthase (synonym of acetate-CoA ligase activity, GO:0003987), both generating acetyl-CoA (Aceti and Ferry, 1988; Jetten et al., 1989; Lundie and Ferry, 1989; Ferry, 1992, 1993, 1999, 2011; Thauer et al., 2008). The first route generates ADP that is converted back to ATP via electron transport phosphorylation at an ATPase (Ferry, 1992, 1993, 1999). In contrast, the second route generates AMP and pyrophosphate, and AMP has to be converted to ADP by adenylate kinase (GO:0004017) through the consumption of one ATP (AMP + ATP → 2ADP) before it can used for the regeneration of ATP (ADP + P_i + energy → ATP) (Jetten et al., 1989; Zinder, 1993; Berger et al., 2012). Thus, organisms utilizing the acetyl-CoA synthase reaction are placed in an energetically unfavorable situation and exhibit slow growth rates (Zinder, 1993). However, by virtue of this investment they are able to utilize acetate even at very low concentrations and consequently are the predominant acetotrophic methanogens in many anaerobic niches of nature (Zinder, 1993). It is not known whether the energy present in pyrophosphate is conserved or is released via hydrolysis for the purpose of making the acetate activation process thermodynamically more favorable (Welte and Deppenmeier, 2014). The methanogens employing acetyl-CoA synthase carry pyrophosphatase (GO:0016462) and whether the enzyme is positioned to harvest or release energy is not known (Berger et al., 2012; Welte and Deppenmeier, 2014). The next step, the breakage of the carbon-carbon bond of the acetate moiety in acetyl-CoA, is catalyzed by an acetyl-CoA decarbonylase/synthase-carbon monoxide dehydrogenase complex (GO:0044672) (Ferry, 1993, 1999; Lu et al., 1994; Grahame, 2003; Li et al., 2006; Wang et al., 2011). The carbonyl group of acetyl-CoA is oxidized to CO₂ by the CODH component (GO:0043885) and the reducing equivalents (two-electrons) generated by this process help to reduce ferredoxin (Ferry, 1993, 1999, 2011; Lu et al., 1994; Grahame, 2003; Li et al., 2006; Wang et al., 2011). The methyl group of the acetyl group is transferred to H₄MPT via a corrinoid cofactor of the CODH/ACDS complex, producing CH₃-H₄MPT (Ferry, 1999; Grahame, 2003). The methyl group of CH₃-H₄MPT leads to methane via the actions of methyl-H₄MPT:coenzyme M methyl transferase (Mtr) and methyl-CoM reductase (Figure 4). The CO₂ produced from acetate is hydrated to bicarbonate by a membrane-bound gamma-type carbonic anhydrase (GO:0004089) and is efficiently exported out of the cell (Ferry, 2011). This process is thought to improve the thermodynamic efficiency of methanogenesis from acetate (Ferry, 2011).

There are two avenues for energy conservation in methanogenesis from acetate (Deppenmeier and Muller, 2008). One is via the use of the sodium potential generated by Mtr and has been described above. The other is through the oxidation of reduced ferredoxin through one of two complex processes. Certain acetotrophic methanogens oxidize reduced ferredoxin by use of Ech hydrogenase, generating molecular hydrogen and proton potential (Meuer et al., 1999, 2002; Kulkarni et al., 2009). The molecular hydrogen is utilized for the extrusion of additional protons and for heterodisulfide reduction via the Vho hydrogenase, methanophenazine and heterodisulfide reductase, as during methylotrophic methanogenesis (Figure 4; see above) (Kulkarni et al., 2009). In methanogens lacking Ech hydrogenase, a complex called Rnf utilizes reduced ferredoxin, producing a sodium gradient and transferring electrons to heterodisulfide reductase via methanophenazine. Thus, Rnf is considered a replacement of the Ech and Vho hydrogenases (Li et al., 2006; Wang et al., 2011). Both the H⁺ and Na⁺ potentials are utilized by an A₁A_O ATP synthase (GO:1990490) for ATP production (Deppenmeier and Muller, 2008); in some cases a Na⁺/H⁺ antiporter (GO:0015385) called Mrp adjusts the ratio of the two gradients for optimizing the thermodynamic efficiency of the ATP synthase (Li et al., 2006; Wang et al., 2011; Jasso-Chavez et al., 2013).

Many of the coenzymes involved in methanogenesis, namely methanofuran, tetrahydromethanopterin, tetrahydrosarcinapterin, coenzyme M, coenzyme F₄₂₀, coenzyme B, methanophenazine, and coenzyme F₄₃₀, have unusual properties. As a result, the respective biosynthesis pathways have attracted attention (Graham and White, 2002). This interest has increased further as some of these coenzymes have been found to perform critical functions in other organisms, such as in actinobacteria (includes mycobacteria and streptomyces groups), methanotrophic and methylotrophic bacteria, cyanobacteria, and plants (Takao et al., 1989; Batschauer, 1993; Purwantini et al., 1997; Chistoserdova et al., 2004; Krishnakumar et al., 2008). Some of the existing knowledge has been summarized at the MENGO website.