The Methanogenic Bacteria

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The methanogenic bacteria are a large and diverse group that is united by three features: 1) They form large quantities of methane as the major product of their energy metabolism. 2) They are strict anaerobes. 3) They are members of the domain Archaea, or archaebacteria, (see Chapter 1) and only distantly related to the more familiar classical bacteria or eubacteria. Like the photosynthetic eubacteria, the methanogenic bacteria are related to each other primarily by their mode of energy metabolism but are very diverse with respect to their other properties.

Methanogenic bacteria obtain their energy for growth from the conversion of a limited number of substrates to methane gas. The major substrates are H₂ + CO₂, formate, and acetate. In addition, some other C-1 compounds such as methanol, trimethylamine, and dimethylsulfide and some alcohols such as isopropanol, isobutanol, cyclopentanol and ethanol are substrates for some methanogens. All of these substrates are converted stoichiometrically to methane. In this regard, the metabolism of the methanogens is strikingly different from that of the so-called “minimethane” producers, which are other anaerobic microorganisms that produce very small amounts of methane as a consequence of side reactions of their normal metabolism (for an overview of the minimethane producers, see Rimbault et al., 1988). Another feature that distinguishes the methanogens from the minimethane producers is that the methanogens are obligate methane-producers, that is, they only grow under conditions where methane is formed.

The list of substrates for growth of methanogens may be divided into three groups (Table 1). In the first group, the energy substrate (electron donor) is H₂, formate, or certain alcohols and the electron acceptor is CO₂, which is reduced to methane. The ability to utilize H₂ as an electron donor for CO₂ reduction is almost universal among methanogens. Likewise, many methanogens also utilize formate, but the ability to utilize alcohols is less common (Bleicher et al., 1989; Zellner and Winter, 1987a). Some methanogens also utilize carbon monoxide as an electron donor, but growth is very slow (Daniels et al., 1977). CO₂ reduction is the major source of methane in certain habitats such as the rumen. In other environments, such as the sediments of freshwater lakes and certain bioreactors, only about one-third of the methane is formed from CO₂ reduction. However, this reaction is still very important for maintaining the very low concentrations of H₂ and formate typical of these anaerobic habitats and facilitating the process of interspecies electron transfer.

Table 1. Reactions and standard changes in free energies for methanogenesis.^a

In the second group, the energy substrate is one of a variety of methyl-containing C-1 compounds, which can serve as substrates for a few taxa of methanogens. Usually these compounds are disproportionated. Some molecules of the substrate are oxidized to CO₂. The electron acceptors are the remaining methyl groups, which are reduced directly to methane (Table 1). Although dimethylselenide and methane thiol also serve as substrates for methanogenesis, these substrates do not support growth (Kiene et al., 1986). Methanogenesis from C-1 compounds is common where methyl-containing C-1 compounds are abundant. In marine sediments, trimethylamine may be formed from choline, glycine betaine, or trimethylamine oxide. In the large intestine of mammals, methanol may be formed from the anaerobic transformation of the methoxy groups of pectin. Dimethylsulfide is also common in anaerobic environments where it is formed from both methionine and the osmoregulant dimethylsulfoniopropionate.

In the third group, acetate is the major source of methane, but the ability to catabolize this substrate is limited to species of Methanosarcina and Methanosaeta (“Methanothrix”). Acetate is present in many environments, and methane synthesis proceeds by an aceticlastic reaction, in which the methyl carbon of acetate is reduced to methane and the carboxyl carbon is oxidized to CO₂. Methanogenesis from acetate is common in anoxic freshwater sediments where the catabolism of acetate by other anaerobes is limited by the availability of alternate electron acceptors such as sulfate or nitrate.

Methane synthesis is the major source of energy for growth of methanogens. Thus, methanogenesis may be viewed as a form of anaerobic respiration where CO₂, the methyl groups of C-1 compounds, or the methyl carbon of acetate is the electron acceptor. However, the standard change in free energy (ΔG°′) during most methanogenic reactions is very small, and the amount of ATP produced per mole of methane is probably close to or less than one (Table 1). In most natural habitats where the concentrations of H₂ and formate are very low, the change in free energy may even be much smaller.

The second distinctive feature of methanogens is their extreme sensitivity to oxygen. Thus, the methanogens are very strict anaerobes, and they are generally present in nature only in anoxic environments. For instance, the half-time for survival of one species of Methanosarcina is reported to be only 4 minutes in air-equilibrated medium (Zhilina, 1972). In part, some of the sensitivity of methanogens is probably due to the oxygen lability of many of the enzymes involved in methanogenesis. Other mechanisms are probably important as well. For instance, the methanococcal enzyme acetohydroxy acid synthase—an enzyme in the pathway of branched-chain amino acid biosynthesis—is unusually sensitive to oxygen (Xing and Whitman, 1987). Likewise, many methanogens are unable to utilize oxidized sulfur compounds. Because the most widely used sulfur source, sulfide, reacts chemically with oxygen, anaerobiosis is also required to protect the sulfur source in the medium. Paradoxically, when cultures of some species are washed and resuspended in unreduced medium in the absence of substrates, their viability in unaffected by atmospheric levels of oxygen for at least 30 hours (Kiener and Leisinger, 1983).

The third distinctive feature of the methanogens is that they are archaebacteria (Jones et al., 1987; Woese, 1987). Other archaebacteria include the extreme halophiles and the sulfur-dependent extreme thermophiles. Some properties the methanogens share with other archaebacteria are listed in Table 2 and are described in more detail in subsequent sections. However, the methanogens are different from other archaebacteria because they are abundant in environments of moderate temperature, pH, and salinity. Although some extremely halophilic or extremely thermophilic methanogens have been described, most species are found in moderate environments where they are often closely associated with anaerobic eubacteria and eukaryotes. Likewise, the methanogens are unlike other archaebacteria in that they contain large amounts of coenzymes essential for methane synthesis (see below). Some of these, such as coenzyme F₄₂₀ and coenzyme M, may serve as distinctive biomarkers.

Table 2. Distinctive features found in various members of the domain Archaea (archaebacteria).

Habitats of Methanogenic Bacteria

Methanogenic bacteria are abundant in habitats where electron acceptors such as O₂, NO₃ ⁻, Fe³⁺ and SO₄ ²⁻ are limiting. Common habitats for methanogens are anaerobic digestors, anoxic sediments, flooded soils, and gastrointestinal tracts. Methanogens are generally absent from the water column of unstratified lakes and rivers because convection currents rapidly aerate the deep waters. However, the diffusion of O₂ between the layers of stratified lakes is often too slow to maintain oxic conditions in the lower layers. Similarly, the physical structure of sediments limits dispersive mechanisms, so deeper sediments are often anoxic and harbor methanogens. Soil environments may be divided into the vadose zone, which is not saturated with water, and the zone below the water table, which is saturated. In the vadose zone, O₂ diffusion is rapid and anoxic vadose zones are extremely rare, although anoxic microenvironments may occur. Thus, the activity of methanogens in the vadose zone is limited. O₂ diffusion through water is slower than through soil, so the water-saturated zone is often anoxic. Because air-saturated water at 20°C contains less than 0.3 mM O₂, even small amounts of organic pollution cause groundwater to become anoxic and suitable for development of methanogenic populations.

In axenic culture, methanogenic bacteria are extremely sensitive to small amounts of O₂. However, in natural habitats, the activities of other organisms protect methanogens in apparently oxic environments. For instance, methanogens have been isolated from large dental caries and subgingival plaque in the human mouth (Belay et al., 1988; Brusa et al., 1987). Methanogens may survive in such environments because O₂-uptake by aerobic and euryoxic bacteria creates anoxic microenvironments.

In anoxic environments, the presence of NO₃ ⁻, Fe³⁺, and SO₄ ²⁻ inhibits methanogenesis by allowing other organisms to outcompete methanogens for reduced substrates. For instance, in the presence of sulfate, sulfate-reducing bacteria utilize H₂ at concentrations lower than the minimum concentration which can be utilized by methanogens (Kristjansson et al., 1982; Lovley, 1985). Presumably, the ability of the sulfate-reducing bacteria to outcompete the methanogens is a direct consequence of the more-positive reduction potential of SO₄ ²⁻ compared to that of CO₂.

In environments with sufficient quantities of sulfate, hydrogen sulfide is the predominant reduced product, and the major fate of biodegradable organic carbon is oxidation to CO₂. If sulfate becomes limiting, methane replaces hydrogen sulfide as the reduced product, and the organic carbon is disproportionated to CO₂ and methane. The fate of the methyl group of acetate is an indicator of whether methanogenesis is a dominant catabolic pathway in an ecosystem. If acetate is catabolized by the sulfate-reducing bacteria, CO₂ is formed from the methyl group of acetate. In a methanogenic system, acetate is catabolized by the aceticlastic reaction, which forms methane from the methyl group. Thus, methanogenic degradation of [2-¹⁴C]-acetate leads to ¹⁴CH₄, whereas sulfidogenic degradation leads to ¹⁴CO₂, and the ratio of these labeled gaseous products indicates whether methanogenesis predominates (Winfrey and Zeikus, 1979). However, this test fails in some thermophilic ecosystems where acetate is oxidized to CO₂ prior to formation of methane (see below).

Sulfidogenesis normally dominates in estuarine, marine, and hypersaline sediments, where sulfate diffuses from overlying water. Beneath areas of high productivity, such as kelp forests, sulfate may be limiting in the deep organic sediments. Under these conditions, aceticlastic and H₂- or formate-using methanogens develop. Even in surface layers where sulfate is in excess, some methanogenesis may occur from methylated compounds such as trimethylamine and dimethylsulfide. Trimethylamine is often found in marine sediments, where it is formed from betaine glycine or other related osmoprotectants which are produced by marine plants and bacteria to balance the osmolarity of their cytoplasm with that of the seawater. Trimethylamine is not easily utilized by sulfidogenic bacteria, but it is rapidly fermented by methanogens to methane, CO₂, and ammonia. Thus, trimethylamine has been termed a “noncompetitive” substrate for methanogens. Trimethylamine-degrading methanogens from marine environments are all in the family Methanosarcinaceae, and all methanogens that have been isolated to date from hypersaline environments use trimethylamine as catabolic substrate.

Interspecies Electron Transfer and Obligate Syntrophy

Because of their limited substrate range, methanogens depend on fermentative bacteria to convert a wide range of organic compounds into methanogenic substrates. In environments where organic matter is completely degraded to methane and CO₂, the methanogenic precursors are predominantly acetate, formate, and H₂ + CO₂. The organic matter is initially fermented mainly to volatile organic acids, H₂, and CO₂. Methanogens can directly catabolize H₂ + CO₂, formate, and acetate, but longer-chain volatile organic acids (with three or more carbon atoms) such as propionate and butyrate must be metabolized to one or more of these methanogenic precursors by a specialized group of microbes called syntrophs. These syntrophs form methanogenic substrates only in the presence of vanishingly low concentrations of H₂ or formate (for reviews see Boone and Mah, 1988; Wolin and Miller, 1987).

During the catabolism of sugars and amino acids, acetate production is more favorable for fermentative bacteria because an extra ATP is produced from acetyl CoA, which is derived from pyruvate oxidation. However, if acetate is formed, the cells require an alternative mechanism to reoxidize the NADH generated during the fermentation. Many fermentative bacteria have NADH-linked hydrogenases or formate dehydrogenases, but the thermodynamics for NADH oxidation are unfavorable except at very low concentrations of H₂ or formate. For instance, the H₂ concentration must be less than about 1 µM and the formate concentration less than 100 µM for these substances to be produced from NADH. Thus, in pure cultures, H₂ or formate production from NADH is inhibited by end-product accumulation. However, when these bacteria are grown in coculture with H₂ and formate-utilizing methanogens, the concentrations of H₂ and formate remain low, and they become important products for fermentative bacteria. In such methanogenic cocultures, the fermentative bacteria produce more acetate and less reduced products such as propionate, butyrate, lactate, and ethanol. Although the fermentative bacteria can generally grow without methanogens, additional energy is obtained from phosphoroclastic acetate production when methanogens are present. Because the activities of the methanogens are not required by the fermentative bacteria, this type of interspecies electron transfer is called nonobligate interspecies electron transfer.

A second type of interspecies electron transfer cannot proceed without the activities of H₂ or formate-utilizing species. A specialized group of bacteria called obligate syntrophs oxidize compounds such as propionate, longer-chain volatile organic acids, and aromatic compounds. The obligate syntrophs must dispose of the electrons by the reduction of protons to H₂ or of CO₂ to formate. They lack alternative fermentative reactions and cannot produce other reduced organic compounds. When the concentrations of H₂ and formate are high, end-product inhibition prevents the oxidation of the syntrophic substrates. Thus, propionate and butyrate oxidation are accomplished by obligate syntrophy of fatty acid-oxidizing bacteria and methanogens. The activities of CO₂-reducing methanogens keep the concentrations of H₂ and formate low, allowing the exergonic oxidation of fatty acids by the syntrophs.

Until recently it was assumed that H₂ is the major precursor for CO₂-reduction to methane, although many of the CO₂-reducing methanogens can use H₂ or formate equally well. The butyrate-oxidizing bacterium Syntrophomonas wolfei can produce either H₂ or formate, so the relative importance of these substances as interspecies electron carriers may depend on the enzyme kinetics of their production and degradation or the rate of diffusion between cells. Although the diffusion coefficient of formate is only one-fifth as large as that of H₂, the formate concentration may be one hundred times greater than that of H₂. Under this condition, formate may be responsible for the bulk of interspecies electron transfer in some environments (Boone et al., 1989; Thiele and Zeikus, 1988).

Habitats of Special Interest.

When organic matter is completely catabolized to methane and CO₂, the major substrates of methanogens are usually acetate, formate, and H₂ + CO₂. However, in some environments the growth of aceticlastic methanogens and obligate syntrophs is too slow to maintain a large population in the system. For instance, in the rumen and colon, acetate accumulates to concentrations of 50 to 100 mM. Although this is well above the concentration required for aceticlastic growth of methanogens such as Methanosarcina, these organisms do not catabolize significant quantities of acetate because their growth rate on this substrate is too slow to maintain the population in a rapid-turnover ecosystem. However, when methylamine or methanol is present, the cell numbers of Methanosarcina in the rumen may reach 10⁵ to 10⁶ per milliliter because these substrates support a faster growth rate. Propionate and butyrate are also present in the rumen at significant concentrations, but the slowly growing propionate- and butyrate-degrading organisms are not found in abundance. In the rumen, a wide range of CO₂-reducing methanogens may be found, including Methanobrevibacter ruminantium, Methanobacterium formicicum, and Methanomicrobium mobile. Methanobrevibacter species are the most commonly found CO₂-reducing methanogen in nonruminant intestinal tracts. Methanosphaera species have also been isolated from colonic environments; they only grow by using H₂ to reduce methanol to methane.

Methanogenic bacteria form mutualistic associations with rumen, freshwater and marine ciliates and amoebae, growing as ecto- or endosymbionts. The physical associations of these microbes with protozoa may improve the efficiency of interspecies H₂- or formate-transfer. The methanogenic partners in the endosymbioses have been classified as Methanoplanus endosymbiosus, Methanobrevibacter species, and Methanobacterium formicicum. It is quite possible that in some habitats such as sapropel (aquatic sludge rich in organic matter), the majority of the methanogens are involved in these symbioses (Lee et al., 1987; van Bruggen et al., 1985).

In some environments, such as hot springs and solfataras (volcanic fissures that emit steam and other gases), the predominant substrate is geothermal H₂ rather than decaying organic matter. In such environments, methanogens do not rely on the activities of other organisms for provision of their substrates. The waters near these sources are often thermal as well, and thermophilic methanogens have been isolated from hot springs in Yellowstone National Park (Zeikus et al., 1980) and in Iceland (Huber et al., 1982; Lauerer et al., 1986), and from submarine thermal vents (Jones et al., 1983a).

Atmospheric Methane

At its current atmospheric concentration of 1.7 ppm, methane is the second most abundant carbon-containing gas, and its atmospheric concentration is increasing at a rate of about 1% per year (for a review, see Cicerone and Oremland, 1988). Because methane is a major greenhouse gas, its sources and atmospheric chemistry are of considerable interest. Each year, about 400–640 × 10¹² g of methane are released into the atmosphere. Estimates based upon the isotopic composition of the atmospheric methane suggest that about 74% of this methane is derived from recent microbiological activity. The total amount of methane that is produced may be far greater, and in some important habitats, microbial methane oxidation is known to be very significant. Thus, the methane-producing bacteria are an important component of the earth’s carbon cycle.

The major sources of atmospheric methane include: enteric fermentations in animals; wetlands such as bogs, swamps, and rice paddies; landfills; and termites. Little methane is obtained from marine systems where competition with the sulfate-reducing bacteria and anaerobic methane-oxidizing organisms limits its production and release, respectively.

Isolation

Methanogenic bacteria are extremely sensitive to oxygen, and strict anaerobiosis and prereduced media are required for their isolation. Two general strategies are employed: 1) Enrichment techniques are generally very successful because few other microorganisms are capable of utilizing the major methanogenic substrates under anaerobic conditions. In addition, the sulfate- and sulfur-reducing bacteria, the denitrifying bacteria, and the photosynthetic bacteria may be further excluded by omitting sulfate, sulfur, and nitrate from the enrichment medium and performing the incubations in the dark. While the chemolithotrophic acetogenic bacteria may also be enriched under these conditions, they may be eliminated in subsequent steps. 2) Antibiotics can be employed as selective inhibitors of the eubacteria. Since the methanogens are archaebacteria, many of their enzyme systems are unaffected by a wide variety of common antibiotics.

Enrichments are performed in media that simulate the source of the sample and mimic the environmental pH, salt concentration, and temperature. The presence of methanogens in the enrichment is determined by gas chromatographic analysis of the headspace for methane. Consumption of H₂ + CO₂ or gas production from acetate or methylamines is a less reliable indicator of methanogenic activity. For example, acetogenic bacteria will also consume H₂ + CO₂ gas under the same conditions as the methanogens (see Chapter 21). Likewise, in enrichments containing sediments or large amounts of other organic material, gas may be produced by fermentative bacteria. If the samples are believed to contain high numbers of methanogens, they are first serially diluted in medium, and after incubation, the highest dilution with methanogenic activity is processed further. Mineral medium is used to obtain autotrophic methanogens. Although most of the heterotrophic contaminants will be eliminated upon successive transfers, low levels of some contaminants may persist due to growth on exudates of the methanogen or on lysed cells. Because many methanogens have nutritional requirements for low levels of acetate, amino acids, volatile fatty acids, or vitamins, a complex enrichment medium is often more suitable. For very fastidious methanogens, the enrichment medium may also be amended with 30% rumen fluid or sludge extract (Mah and Smith, 1981). In this case, the heterotrophic contaminants must be eliminated by colony isolation on solid medium. Antibiotics may also reduce the numbers of contaminants. However, because of the rapid selection of antibiotic resistant contaminants, they are seldom sufficient alone for the isolation of pure cultures.

Antibiotics are especially useful when used in conjunction with enrichment techniques. Because some antibiotics are toxic to certain methanogens (Böck and Kandler, 1985), it is also useful to test their effects on the enrichment culture. Success has been obtained with a variety of antibiotics, used either singly or in combination (Table 3). However, to avoid the selection of resistant contaminants, the antibiotics are only utilized for one or two transfers. In addition, the antibiotics must be omitted from the medium when determining the purity of the isolated methanogen.

Table 3. The use of antibiotics for isolation of methanogens.

When a successful enrichment has been obtained, the methanogen may be observed by fluorescence microscopy (Doddema and Vogels, 1978). For optimal fluorescence, the excitation wavelength should be between 350 and 420 nm. Under these conditions, most species autofluoresce a blue-green color due to an abundance of coenzyme F₄₂₀. However, the intensity of the fluorescence varies greatly, and it may not be observed following growth of Methanosaeta on acetate (Zehnder et al., 1980). The fluorescence may also rapidly fade within a few seconds or be absent in older and inactive cells. In spite of these limitations, fluorescence microscopy is useful for determining if more than one type of methanogen is present in an enrichment or for checking the relative purity of the culture.

The isolation of single colonies on solid medium is usually necessary for obtaining axenic cultures of methanogens. Solid medium is prepared by the roll-tube technique or in petri plates in an anaerobic glove box. One advantage of roll tubes is that colonies of methanogens may be frequently visualized through the wall of the tube by their fluorescence. In any case, the anaerobic techniques must be stringent enough to ensure that the plating efficiency is high, since, otherwise, most of the colonies may contain contaminants that are less oxygen-sensitive than the methanogen. In a minimal medium, it may not be possible to obtain isolated colonies because the methanogen may require a nutrient produced by a contaminating organism. For instance, in media lacking acetate, this nutrient may be produced by the fermentation of amino acids by a contaminating bacterium. Under such conditions, if acetate is required for growth of the methanogen, colonies will contain a mixture of both organisms.

In addition to the normal microbiological tests for axenic cultures, certain growth tests are useful to establish culture purity. Growth should not be observed in complex medium prepared without the substrates for methanogenesis. The medium is prepared with the same salt and trace nutrient composition as the enrichment medium, but 0.2–0.4% trypticase peptones or casamino acids and 0.2% glucose are added. Yeast extract, a source of B-vitamins, frequently contains small amounts of formate and acetate and should be avoided. Growth in a complex medium without a methanogenic substrate indicates the presence of a heterotrophic contaminant. The plating efficiency of the methanogenic culture in the enrichment medium should also be greater than 50%. If it is much lower, it is possible that growth is dependent upon crossfeeding between the methanogen and another organism.

Techniques for Culturing Methanogenic Bacteria

The methodology for culturing methanogenic bacteria is based upon the pioneering work of Hungate on anaerobic rumen bacteria (1969). Anaerobiosis is achieved by the replacement of air with oxygen-free gases and the addition of a reducing agent. The culture vessels are made of glass and sealed with butyl rubber stoppers. Oxygen-permeable rubber and plastic are avoided whenever possible. Media are dispensed in an anaerobic glove box, under a stream of oxygen-free gas, or with a glass syringe. Commercial gases are scrubbed to remove traces of contaminating oxygen. Details of the procedures are described in the primary literature (Balch and Wolfe, 1976; Macy et al., 1972) and in several recent reviews (Ljungdahl and Wiegel, 1986; Mah and Smith, 1981).

Enrichment Media for Growth of Methanogens

A large number of different types of media have been formulated for the growth of methanogens. Many are described in the original species descriptions cited elsewhere in this chapter. Therefore, only a few general media will be described. Although most methanogens will grow in one of these media, optimal growth may require modification of the concentrations of some of the components or special additions.

Medium 1 for Freshwater, Sewage, and Intestinal Species

Its composition is similar to media described earlier (Balch et al., 1979; Mah and Smith, 1981). Its composition is (per liter of medium):

Yeast extract	2.0 g
Trypticase peptones	2.0 g
Salt solution A (see below)	10 ml
Phosphate solution (200 g/l of K2HPO4· 3H2O)	2 ml
Resazurin solution (0.5 g/l in water)	2 ml
Sodium acetate solution (136 g/l of Na acetate · 3H2O)	10 ml
Trace element solution (see below)	10 ml
Vitamin solution (see below)	10 ml
NaHCO3	5.0 g
Cysteine/hydrochloride	0.5 g
Sulfide solution (see below)	20 ml

Salt solution A is composed of (per liter):

NH4Cl	100 g
MgCl2 · 6H2O	100 g
CaCl2 · 2H2O	40 g

After dissolving the salts, the pH is adjusted to 4 with HCl.

The trace element solution is modified from Wolin et al. 1963 and is composed of (per liter):

Nitrilotriacetic acid	1.5 g
Fe(NH4)2(SO4)2 · 6H2O	0.2 g
Na2SeO3	0.2 g
CoCl2 · 6H2O	0.1 g
MnSO4 · 2H2O	0.1 g
Na2MoO4 · 2H2O	0.1 g
Na2WO4 · 2H2O	0.1 g
ZnSO4 · 7H2O	0.1 g
AlCl3 · 6H2O	0.04 g
NiCl2 · 6H2O	0.025 g
H3BO3	0.01 g
CuSO4 · 5H2O	0.01 g

To prepare the trace element solution, dissolve the nitrilotriacetic acid in 800 ml of water and adjust the pH to 6.5 with KOH. Then dissolve the minerals in order, adjust the pH to 7.0, and bring the volume to 1 liter.

The vitamin solution is modified from Bryant et al. 1971 and is composed of (per liter):

p-Aminobenzoic acid	10 mg
Nicotinic acid	10 mg
Calcium pantothenate	10 mg
Pyridoxine hydrochloride	10 mg
Riboflavin	10 mg
Thiamine hydrochloride	10 mg
Biotin	5 mg
Folic acid	5 mg
α-Lipoic acid	5 mg
Vitamin B12 (stored in dark at 5°C)	5 mg

The sodium sulfide solution is prepared in anoxic water under N₂ gas. A 10 mM solution of NaOH, 110 ml, is boiled under a stream of N₂ gas until the volume is reduced to 100 ml. It is then allowed to cool under a stream of N₂ gas in a fume hood. Because sodium sulfide is very toxic, precautions must be taken to avoid contact with the solid and the solution, as well as inhalation of the vapor. To remove sulfur oxides from the surface, a large crystal of Na₂S·9H₂O (about 3 g) is washed in 50 ml of water for a few seconds. The crystal is blotted dry on a paper towel and weighed. About 2.5 g should remain and is immediately added to the anaerobic solution of NaOH. The sulfide solution is then dispensed anaerobically into tubes or bottles, pressurized with 100 kPa of N₂, and autoclaved. If an anaerobic glove box is available, the tubes or bottles and their stoppers should be placed in the glove box 24 hours before use to remove adsorbed O₂. The stoppers must fit tightly and should not be greased. They may be lubricated with anaerobic water to facilitate insertion and twisting into place. After autoclaving, the sulfide solution may be stored for up to two weeks or until the solution becomes cloudy. For organisms like Methanococcus, best results are obtained if the sulfide solution is stored in the anaerobic chamber immediately after autoclaving.

Medium 2 for Methanogens Isolated from Marine Environments

It is modified from Romesser et al. 1979, and it is composed of (per liter):

Salt solution B (see below)	500 ml
Phosphate solution (14 g/l of K2HPO4· 3H2O)	10 ml
Trace element solution	10 ml
Vitamin solution	10 ml
Iron stock solution (see below)	5 ml
Sodium acetate solution (136 g/l of sodium acetate · 3H2O)	10 ml
Resazurin solution (0.5 g/l in water)	2 ml
NaCl solution (293 g/l)	75 ml
Yeast extract	2 g
NaHCO3	5 g
Cysteine hydrochloride	0.5 g
Sulfide solution	20 ml

Salt solution B is composed of (per liter):

CaCl2 · 2H2O	0.28 g
KCl	0.67 g
NH4Cl	1.00 g
MgCl2 · 6H2O	5.50 g
MgSO4 · 7H2O	6.90 g

To prepare the iron stock solution, add 0.2 g of Fe(NH₄)₂(SO₄)₂ · 6H₂O to a screw-top bottle. Then add 0.1 ml of concentrated HCl followed by 100 ml of H₂O. The solution should be replaced every four weeks.

Medium 3 for Moderately Halophilic Methanogens

It contains the same components as Medium 1 except that 88 g/l of NaCl is added.

As described above, the media do not contain a substrate for methanogenesis. For growth on H₂ + CO₂ (80:20 vol/vol), the gas mixture is added after dispensing and sterilizing the medium, and the culture tubes are pressurized to 100 kPa above atmospheric pressure. After inoculation, the culture tubes are pressurized to 240 kPa. For growth on other substrates, the gas atmosphere is 100 kPa of N₂ + CO₂ (80:20 vol/vol), and the NaHCO₃ concentration is reduced to 2 g/l. For growth on acetate, 50 ml of the sodium acetate solution (1 M) is added. For growth on trimethylamine/hydrochloride, 12 ml of a 1 M solution (95.5 g/l) is added. For growth on methanol, 4 ml of methanol is added per liter. For growth on 2-propanol, 1-propanol, ethanol, 2-butanol, and 1-butanol, the alcohols are added at a final concentration of 20–30 mM (Widdel, 1986). For growth on formate, 4 g/l of sodium formate is added. To control the pH in batch cultures, a formic-acid reservoir may also be added to the culture tube (Schauer and Whitman, 1989). Regardless of the substrate, care must be taken to avoid exceeding the pressure limit of the culture vessels. For instance, 10 ml of medium containing 50 mM acetate will generate nearly 15 ml of gas if the acetate is completely converted to CO₂ and CH₄. Therefore, the headspace of the culture vessel must be large enough to prevent an excessive increase in pressure, which could lead to an explosion. To minimize this danger, the culture vessels should be handled in wire baskets or metal cans. During growth on H₂ + CO₂, the opposite problem occurs. Because the methanogens consume 5 moles of gas for every mole of CH₄ produced, the vessel will quickly develop a negative pressure. Also, as the partial pressure of CO₂ decreases, the medium will become strongly alkaline, which may inhibit growth and cause cell lysis. To minimize these problems, the volume of the headspace should be at least five times the volume of the medium, and the culture vessel should be pressurized periodically throughout the growth period. For better control of the pH, the headspace may also be repressurized with H₂ + CO₂ (75:25 vol/vol) to fully replenish the CO₂ consumed. For the thermophilic species, attention must also be given to the expansion of gas at higher temperatures.

All three media may be prepared by the same method. All the components are combined except the gaseous substrates and the reducing agents, cysteine hydrochloride and the sulfide solution. The medium is then brought to a boil under a stream of N₂ + CO₂ gas (80:20 vol/vol) and immediately removed from the heat. Solid cysteine hydrochloride is added, and the medium is allowed to cool under a stream of N₂ + CO₂. During this time, the oxygen indicator resazurin changes from blue to pink to colorless. The change from blue to pink occurs upon the pH-dependent formation of resorufin. The change from pink to clear occurs upon the reduction of resorufin to dihydroresorufin. When the medium has cooled to about 50°C, it can be dispensed anaerobically to the culture tubes or bottles under a stream of O₂-free N₂ + CO₂ gas or in an anaerobic glove box (Ljungdahl and Wiegel, 1986; Mah and Smith, 1981). For growth on H₂ + CO₂, the gas atmosphere of the culture tubes is exchanged before autoclaving. After autoclaving, the pH of the bicarbonate-CO₂ buffered medium is 6.8–7.0.

The media may be stored for several months in the anaerobic glove box or for about one week on the laboratory bench prior to the addition of the sulfide solution. One hour before inoculation, one part of the sterile sulfide solution is added anaerobically to 50 parts of sterile medium. Some species appear to be especially sensitive to the products of sulfide oxidation. When small inocula are used, more reproductible results are obtained by adding the sulfide solution 24 hours before the inoculation.

For growth of fastidious methanogens, the media may be amended (per liter) with 100–300 ml of clarified rumen fluid, 10 ml of a volatile fatty acid mixture, or 2 ml of boiled cell extract (BCE) of a methanogen (Mah and Smith, 1981; Tanner and Wolfe, 1988). To prepare rumen fluid, samples of the rumen contents are obtained from a fistulated animal, a mouth tube, or a slaughtered animal. The fluid is separated from the large solids by filtration through 8 layers of cheese cloth into an Erlenmeyer flask. The fluid is then centrifuged under anoxic conditions at 10,000 × g for 20 minutes to remove microbial cells and small particulate matter. The supernatant is decanted and sparged slowly with CO₂ overnight. Then it is dispensed into serum bottles, flushed with CO₂, and autoclaved. The volatile fatty acid mixture is modified from Bryant et al. 1971 and is composed as follows: 46 ml of butyric, 46 ml of isobutyric, 55 ml of isovaleric, 55 ml of DL-2-methylbutyric, 37 ml of propionic, and 54 ml of valeric acids are added to 500 ml of water in a fume hood, neutralized (by litmus) with 2 M NaOH, and brought to a final volume of 1 liter.

Some methanogens require growth factors that are only produced by other methanogenic bacteria. These factors may be obtained from rumen fluid or BCE (Tanner and Wolfe, 1988). To prepare BCE, 10 g wet weight of methanogen cells are resuspended in 20 ml of anaerobic 20 mM potassium phosphate buffer, pH 7.0, and incubated under a stream of N₂ gas in a boiling water bath for one hour. After cooling, the suspension is centrifuged anaerobically at 20,000× g for 30 min at 4°C. The resulting supernatant, BCE, is stored anaerobically at −20°C.

Mineral media may be prepared by omitting the organic supplements: yeast extract, trypticase, acetate, and cysteine. However, cysteine also functions as a reducing agent. Therefore, the medium will remain oxidized until the sulfide solution is added. In this case, the sulfide solution should be added 24 h before inoculation.

For growth of the alkalophilic methanogens, the medium is prepared under a N₂ atmosphere. The NaHCO₃ concentration is increased to 10 g/l (pH 8.5) or replaced with 3 g/l of NaHCO₃ plus 2 g/l of Na₂CO₃ (pH 9.3; Mathrani et al., 1988; Worakit et al., 1986). If H₂ is the substrate, the culture is initially pressurized with 50 kPa H₂. CO₂ is not added initially to avoid acidification of the medium. The gas is then replenished with H₂ + CO₂ (75:25 vol/vol) during growth, stoichiometrically replacing the CO₂ reduced to methane. For mass culture, gas additions are controlled by a pH auxostat (Boone et al., 1987).

Solid medium is prepared by the addition of agar or Gelrite during the medium preparation. For roll tubes or bottles, the medium plus sulfide with 2% agar or 1% Gelrite is allowed to coat the walls to provide a surface for inoculation (Harris, 1985; Hermann et al., 1986; Hungate, 1969). The agar surface may be streaked or the cells may be added as a soft agar overlay. Alternatively, molten agar may be cooled to 45°C and inoculated prior to solidification. Petri plates containing 0.8–2% agar or Gelrite (Kelco Div., Merck & Co., Inc., San Diego, Calif.) may be prepared in an anaerobic glove box (Balch et al., 1979). In this case, the plates are prepared without sulfide (Jones et al., 1983c). After inoculation, the plates are pressurized with H₂ + CO₂ or N₂ + CO₂ in a cannister and removed from the chamber. An open tube containing 0.75 ml of 20% Na₂S·9H₂O per liter of cannister volume is added just prior to pressurization. Because the CO₂ neutralizes the sulfide solution, volatile H₂S (about 1.5%) is generated inside the cannister. Because H₂S is toxic, the cannister must either be opened in a fume hood or be flushed with N₂ + CO₂ gas before opening in the anaerobic glove box.

Measurement of Growth

In many cases, the growth of methanogens can be measured by normal microbiological techniques. However, some organisms grow as aggregates or have very low cell yields, and it is difficult to measure growth turbidimetrically or by other common techniques. An alternative is to measure CH₄ accumulation in the headspace by gas chromatography. However, whenever product formation is used to estimate growth rate, it is important to insure that cell growth is balanced, that is, product formation is proportional to the increase in cell mass. Thus, control experiments should demonstrate a correlation between CH₄ formation and cell mass. This method is additionally complicated because upon inoculation, only cells and not the initial product are transferred (Powell, 1983). Therefore, plots of the logarithm of CH₄ accumulated versus time are nonlinear for about four generations, and the growth curves appear to be convex. This artifact results in an overestimate of the specific growth rate. To correct for this effect, the logarithm of the rate of methane formation versus time should be plotted (Powell, 1983). This replot is linear, and the specific growth rate is equal to the slope.

Maintenance of Stock Cultures

Many of the same techniques used for storage of stock cultures of other anaerobes are suitable for at least some of the methanogens (Hippe, 1984). While freezing in liquid nitrogen is a very reliable method, less-expensive alternatives may be employed for laboratory collections. Lyophilization in horse serum plus 7.5% glucose and 3 mg/ml of ferrous sulfide followed by storage at either 8° or −70°C is suitable for many species of Methanobacteriaceae and Methanosarcina (Hippe, 1984). However, this method is less effective for Methanococcus and Methanospirillum. Many methanogens can also be stored on agar slants at 4°C for one year if oxygen is excluded (Winter, 1983). Likewise, storage at −18°C in medium containing 50% glycerol in sealed glass ampules is effective for 20 months. Methanococcus cultures can be preserved in medium containing 25% glycerol in screw-top vials at −70°C for 30 months without special anaerobic precautions (Whitman et al., 1986). Methanobrevibacter species and some other methanogens can be stored in biphasic cultures at −76°C for 6–12 months (Miller, 1989). Culture tubes are prepared with a slant of double-strength medium containing 3% agar and twice the usual concentration of reducing agents. One-third volume of reduced, single-strength broth is added. The tube is inoculated and allowed to grow to a heavy density. The culture is then cooled for 1 hour at 4°C prior to freezing at −76°C.

Cultures of the type strains of most species of methanogens are available through the Deutsche Sammlung von Mikroorganismen (Braunschweig, FRG) and the OGC Collection of Methanogenic Archaeobacteria (Oregon Graduate Institute, Beaverton, Oregon, USA).

Identification

Three major difficulties are encountered in the identification of isolates of methanogenic bacteria: 1) Most of the species descriptions are based upon the examination of only a few strains. Therefore, it is frequently not known if the phenotypic characterization is representative of the species. 2) Probably only a fraction of the methanogens in nature have been described. Therefore, a new isolate may represent an undescribed species or subspecies. 3) Because of their restricted catabolism coupled with extreme genetic diversity, phenotypic characters alone are often insufficient to identify methanogens. For these reasons, phylogenetic methods such as DNA-DNA hybridization and ribosomal RNA sequence analysis are frequently necessary for a definitive assignment (Boone and Whitman, 1988). Other techniques that have proven useful in the identification of methanogens include analysis of their antigenic relationships, polyamine content, molecular weights of the methylreductase subunits, and molecular weights of the polar lipids. Cross-reactivity of specific immunoglobulins has proven useful for identifying methanogens at the species and genus level (Conway de Macario et al., 1981). For distinguishing among members of different families, data on the relative distribution of putrescine, spermidine, spermine, and homospermidine, as well as the molecular weight of the small subunit of component C of the methylreductase, are useful (Kneifel et al., 1986; Rouvière and Wolfe, 1987). Likewise, the distribution of polar lipids distinguishes the families of methanogens (Koga et al., 1987; Morii et al., 1988).

Nineteen genera and more than 50 species of methanogenic bacteria have been described. The current taxonomy reflects the phylogeny described by the oligonucleotide catalogs of the 16S rRNAs (Balch et al., 1979). More recent information, which includes nearly complete sequences of the 16S rRNA and many new species, has confirmed many of these earlier conclusions. The current phylogenetic tree supports the classification of the methanogens into three major groups that correspond to the orders proposed by Balch et al., 1979 (Fig. 1). The orders have been further divided into six families, and the creation of additional families may be warranted to include other deep branches of the major lineages. However, the proposal to place Methanoplanus in a separate family (Wildgruber et al., 1982) is not supported by the most recent information.

Fig. 1.

Phylogenetic tree for the methanogenic bacteria and other archaebacteria based upon 16S rRNA sequences. Mc. Methanococcus; Mt., Methanothermus; M., Methanobacterium; Mbb., Methanobrevibacter; Ms., Methanospirillum; Mp., Methanoplanus; Msa., Methanosarcina; Msae., Methanosaeta; H., Halobacterium; T., Thermoplasma; A., Archaeoglobus; Tc., Thermococcus; Tp., Thermoproteus; Tf., Thermofilum; P., Pyrodictium; S., Sulfolobus; and D., Desulfurococcus. (P. E. Rouvière and C. R. Woese, personal communication.)

At present, three orders and six families are recognized within the methanogens (Tables 4 and 5). As indicated in the footnotes to Table 4, most genera have not been formally assigned to an order and family. While the present scheme is consistent with the current phylogenetic and phenotypic data, future changes are to be expected.

Table 4. Major taxonomic groups of methanogenic bacteria.

Table 5. Some characteristics of the methanogen families.

Two families, the Methanobacteriaceae and the Methanothermaceae, are closely related. These methanogens possess cell walls composed in part of pseudomurein (Kandler and König, 1985). The cell walls of the Methanothermaceae also contain an additional surface layer composed of protein. The family Methanothermaceae contains one genus, Methanothermus, both species of which are extremely thermophilic bacilli with temperature optima of 83–88°C. Like in many of the Methanobacteriaceae, the only substrate for methanogenesis is H₂ + CO₂. The family Methanobacteriaceae contains two genera, composed of thermophilic as well as mesophilic species. These genera, Methanobacterium and Methanobrevibacter, are bacilli that utilize either H₂ + CO₂ alone or H₂ + CO₂ and formate as substrates for methanogenesis. An additional genus, Methanosphaera, is closely related to the Methanobacteriaceae, although it has not yet been formally placed within this family. It includes two coccoid species that utilize H₂ + methanol as substrates. These organisms are unique in that they are incapable of both the reduction of CO₂ to methane and the oxidation of methanol to CO₂ (Miller and Wolin, 1983).

The family Methanococcaceae contains one genus of irregular cocci, Methanococcus, most species of which utilize both H₂ + CO₂ and formate as substrates. The six known species of Methanococcus contain protein cell walls and are either mesophilic or extremely thermophilic. Most species are also autotrophic. By the criterion of the 16S rRNA sequences, this genus is very diverse (Fig. 1). To maintain genera of equal phylogenetic depth, the two thermophilic species should be placed in two new genera.

The family Methanosarcinaceae contains all the methanogens capable of utilizing acetate as a substrate for methanogenesis or oxidizing the methyl groups of C-1 compounds. Formate is not utilized by any species in this family and most species are also unable to grow on H₂ + CO₂. Seven genera have been described. The genus Methanosarcina contains six species of cocci, which may occur singly, in packets, or in large pseudoparenchyma (Zhilina, 1976). The aggregates are held together by methanochondroitin, a polymer of N-acetyl-D-galactosamine and D-glucuronic acid similar to animal chondroitin (Kreisl and Kandler, 1986). Pseudomurein is absent. Pseudoparenchyma formation is species and strain specific and varies even in strains that do form aggregates, depending upon the culture conditions and growth stage. Therefore, the absence of aggregation is not sufficient evidence to exclude an isolate from this genus. Another typical property of this genus is the ability to utilize both acetate and C-1 compounds as substrates for methano-genesis. In addition, some species also utilize H₂ + CO₂. Other genera in this family that contain cocci are distinctive in their ability to utilize only C-1 compounds as substrates for methanogenesis. These genera include Methanolobus, Methanococcoides, Methanohalobium, Halomethanococcus, and Methanohalophilus. The last three genera contain the moderately and extremely halophilic species of methanogens, but have not been formally placed within this family. An additional species, Methanococcus halophilus, should probably also be classified with these organisms on the basis of its 5S rRNA sequence (Lysenko and Zhilina, 1985). The last genus, Methanosaeta (“Methanothrix”), includes large sheathed rods that can only utilize acetate as a substrate for methanogenesis.

The family Methanomicrobiaceae contains six genera of different morphological types with similar physiology. With one exception, all species utilize both H₂ + CO₂ and formate as substrates. Two genera, Methanogenium and Methanoculleus, are cocci that can be distinguished from the methanococci by their slower growth rates under optimal conditions, more complex nutritional requirements, higher GC content, and lower NaCl requirements. The genus Methanospirillum has a distinctive spiral morphology, and the genus Methanoplanus contains plate-like or irregular disk-shaped species. Lastly, the genera Methanomicrobium and Methanolacinia contain rod-shaped species. These species may be distinguished from Methanobrevibacter because they stain Gram-negative due to the absence of pseudomurein and they have a higher GC content.

The family Methanocorpusculaceae is related to the family Methanomicrobiaceae. It contains one genus, Methanocorpusculum, of very small cocci. The complex nutritional requirements of the species of this family distinguishes them from species of Methanococcus. The absence of a requirement for NaCl for good growth distinguishes them from species of Methanogenium and Methanoculleus.

Characteristics of Methanogenic Bacteria

Some of the most important and most distinctive features of methanogenic species will be described in this section. More detailed descriptions of the various taxa of bacteria may be found in the original citations, which are included in the footnotes to Tables 6 7 8 9 10 11 12 13 14 15 16. Whenever possible, the descriptions summarize the properties of all the described strains as well as the type strain. Because many of the growth descriptions depend greatly upon the experimental conditions, some caution must be exercised in the evaluation of results from different laboratories. In particular, variation in the growth optima may occur depending on whether growth is measured by turbidity, methane formation, growth rate, or growth yield.

Table 6. Some characteristic properties of species of the genus.

Table 7. Some characteristic properties of species of the genus Methanobrevibacter.

Table 8. Some characteristic properties of species of the genus Methanosphaera.

Table 9. Some characteristic properties of species of the genus Methanothermus.

Table 10. Some characteristic properties of species of the genus Methanococcus.

Table 11. Some characteristic properties of species of the genera Methanogenium and Methanoculleus.

Table 12. Some characteristic properties of species of the genera Methanolacinia, Methanomicrobium, Methanoplanus, and Methanospirillum.

Table 13. Some characteristic properties of species of the genus Methanocorpusculum.

Table 14. Some characteristic properties of species of the genus Methanosarcina.

Table 15. Some characteristic properties of the obligately methylotrophic cocci.

Table 16. Some characteristic properties of species of the genus Methanosaeta (“Methanothrix”).

The Family Methanobacteriaceae

The presence of a peptidoglycan chemically different from murein (called pseudomurein) is ubiquitous among species of the family Methanobacteriaceae. This peptidoglycan, like its counterpart in eubacteria, confers shape to these rods and coccobacilli, which have a wall structure typical of Gram-positive bacteria (Kandler and König, 1985). While the cells frequently stain Gram-positive, they are not formally considered Gram-positive because the cell wall is not composed of true murein. Many of the rod-shaped methanogens are in this family, and the morphologies of representative species are shown in Fig. 2. The exceptions are a few other species that have protein cell walls and are sensitive to lysis by detergents (Methanomicrobium) or are enclosed within a sheath that appears to confer the shape (Methanosaeta). The family Methanobacteriaceae contains three genera.

Fig. 2.

Phase contrast photomicrographs of some rod-shaped methanogenic bacteria of the genera Methanobrevibacter, Methanobacterium, and Methanomicrobium. (A) Methanobrevibacter ruminantium; (B) Methanomicrobium mobile (stained preparation); (C) Methanobacterium formicicum; (D) Methanobacterium bryantii; (E) Methanobrevibacter arboriphilicus; (F) Methanobacterium thermoautotrophicum. (From Mah and Smith, 1981.)

Methanobacterium.

The species of Methanobacterium vary widely in length, and filaments are common (Table 6). One species, M. thermoaggregans, forms large multicellular aggregates. The rod-shaped cells are often irregularly crooked. All species grow with H₂ + CO₂ as a substrate for methanogenesis. In addition, formate is used by M. formicicum, “M. palustre,” and M. thermoformicicum. The secondary alcohols 2-propanol and 2-butanol are also utilized by M. bryantii, M. formicicum, and ”M. palustre” (Widdel et al., 1988; Zellner and Winter, 1987a). In addition, 2-propanol and 2-butanol support low levels of methane synthesis but not growth of the moderately acidophilic species M. espanolae (Patel et al., 1990). Most species of Methanobacterium are capable of autotrophic growth with CO₂ as a sole carbon source. However, acetate, cysteine, and yeast extract are frequently stimulatory. M. bryantii is also stimulated by a mixture of B-vitamins. The alkaliphilic species, M. alcaliphilum and M. thermoalcaliphilum, both require yeast extract for growth, and their major carbon sources are not known.

While the nitrogen and sulfur sources have not been investigated systematically, all species of Methanobacterium that have been tested use ammonium, sulfide, and elemental sulfur. In addition, M. ivanovii uses glutamine as a sole nitrogen source (Bhatnagar et al., 1984) and M. thermoautotrophicum uses glutamine and urea. Note that at the high temperature at which the thermophile is grown glutamine decomposes rapidly in the medium (Friedmann and Thauer, 1987). M. ivanovii, M. bryantii, and M. thermoautotrophicum use cysteine as a sole sulfur source and M. ivanovii also uses methionine. M. thermoautotrophicum also uses sulfite and thiosulfate as sole sulfur sources (Daniels et al., 1986).

The species of Methanobacterium have been isolated from anaerobic digestors, sewage sludge, manure, groundwater, and formation water of oil-bearing rocks. In general, these habitats contain low NaCl concentrations, which are optimal for growth. Concentrations above 0.2 M are frequently inhibitory. M. wolfei also has a growth requirement for 8 µM tungstate, which is an unusually high concentration.

Methanobrevibacter.

Species of the genus Methanobrevibacter are oblong cocci or very short rods (Table 7). Flagella or fimbriae are not observed. All species utilize H₂ + CO₂ as substrates for methanogenesis. Formate is utilized by some species. Secondary alcohols were not utilized by the species tested, M. arboriphilicus and M. smithii (Widdel et al., 1988; Zellner and Winter, 1987a). The optimum pH for growth is near 7, except for M. arboriphilicus, which has an optimum near 8.0. M. ruminantium was isolated from the rumen and has very complex nutritional requirements. These requirements may be met by rumen fluid, acetate, 2-methylbutyrate, coenzyme M, or a mixture of amino acids. In addition, several similar strains have been isolated from the rumen that do not require coenzyme M (Lovley et al., 1984; Miller et al., 1986). At present, it is not known whether these strains represent new species or atypical strains of M. ruminantium. M. smithii appears to be a common methanogen in the human gastrointestinal tract (Miller and Wolin, 1982; Weaver et al., 1986). Acetate, trypticase, yeast extract, and B-vitamins are required or stimulatory for growth. In contrast to M. ruminantium and M. arboriphilicus, growth of M. smithii is not inhibited by bile salts. Methanobrevibacter species have also been isolated from the feces of the horse, pig, goose, and rat (Miller and Wolin, 1986). The strain from the rat may be a new and undescribed species. On the basis of serological evidence, it has also been proposed that an additional species of Methanobrevibacter may predominate in some human feces (Misawa et al., 1986).

Methanosphaera.

Methanosphaera is distinct from other Methanobacteriaceae because it is coccoid and cannot reduce CO₂ to CH₄. Cells are capable of growth only by using H₂ to reduce methanol. However, the cell walls contain pseudomurein, and sequence analysis of the 16S rRNA suggests that it is closely related to the Methanobacteriaceae. Cells are spherical and occur singly or in pairs, tetrads, or small clusters (Table 8). So far, they are found only in the mammalian colon, and they are resistant to bile salts. Cells are chemoorganotrophic, and they require CO₂, acetate, some vitamins, and amino acids for growth. In addition to the amino acids, M. stadtmaniae requires NH₃ as a nitrogen source. Although the two species are very similar phenotypically, they can be easily distinguished by DNA-DNA hybridization or serology.

The Family Methanothermaceae

This family contains one genus, Methanothermus, which is extremely thermophilic and rod-shaped (Table 9). The temperature optimum is between 80° and 90°C and no growth occurs below 60°C or above 97°C. Its habitat is geothermal (85° to 101°C) waters and muds in Iceland, and it has not been found in similar habitats in other geographical locations. H₂ + CO₂ is the only substrate for methanogenesis. Colonies do not grow on agar plates; therefore, polysilicate plates have been used. Unlike those of the Methanobacteriaceae, the cell wall of Methanothermus has an S-layer external to the pseudomurein layer. Two species of Methanothermus have been described: M. fervidus grows as dispersed cells and never forms filaments whereas M. sociabilis forms aggregates as large as 3 mm in diameter and contains pili-like appendages. These species also differ in their lipid and pseudomurein composition (Lauerer et al., 1986). Growth of each species is inhibited in vessels made of borosilicate glass, so soda-lime glass vessels are used.

The Family Methanococcaceae

Methanococcaceae contains one genus, Methanococcus, which is composed of six species of mesophilic and thermophilic organisms (Table 10). An additional species has been proposed, M. deltae, but it is probably a subspecies of M. maripaludis (Corder et al., 1983; Whitman, 1989). The methanococci are all irregular cocci, 1–2 µm in diameter (Fig. 3). Thus, they are not easily distinguished morphologically from each other or other coccal methanogens. The cell wall is composed of an S-layer containing hexagonally arranged protein subunits (Koval and Jarrell, 1987; Nusser and König, 1987). The molecular weights of all the major wall proteins are less than 90,000, which distinguishes them from the S-layer proteins of the members of the family Methanomicrobiaceae. In addition, the S-layer proteins of the methanococci are not glycosylated. Cells lyse rapidly in 0.01% sodium dodecyl sulfate and in distilled water and they disintegrate upon heat fixation to microscope slides. The methanococci are motile, with one or two polar tufts of flagella. Isolated flagella of M. voltae are composed of two flagellins with molecular weights of 31,000 and 33,000 (Kalmokoff et al., 1988). The basal body is knob-like and lacks the complex ultrastructure typical of the eubacterial basal body.

Fig. 3.

Phase contrast photomicrograph of Methanococcus voltae. (From Mah and Smith, 1981.)

The methanococci utilize H₂ + CO₂ as a substrate for methanogenesis. Formate is also an electron donor for all the methanococci except for two of the three described strains of M. jannaschii. None of the methanococci utilize acetate, methyl compounds, or alcohols as substrates for methanogenesis. Except for M. voltae, the methanococci are capable of rapid autotrophic growth with CO₂ as the sole carbon source. M. voltae requires acetate, isoleucine (or propionate or 2-methylbutyrate), and leucine (or isovalerate) as carbon sources (for a review see Jarrell and Koval, 1989). M. voltae grows mixotrophically, and its cellular carbon is derived from both the assimilation of exogenous acetate and autotrophic acetyl-CoA biosynthesis (Shieh et al., 1988). M. maripaludis also assimilates acetate, glycine, and a variety of nonpolar and basic amino acids (Whitman et al., 1987). In contrast, M. vannielii and “M. aeolicus” do not assimilate amino acids when they are provided at low concentrations, and acetate is not stimulatory to growth. Pantoyl lactone stimulates the growth of M. voltae, but water-soluble vitamins do not affect the growth rates of the other mesophilic methanococci.

The methanococci utilize ammonia as a sole nitrogen source. M. maripaludis and M. thermolithotrophicus can also use N₂; M. maripaludis can use alanine; and M. vannielii can use purines as sole nitrogen sources (DeMoll and Tsai, 1986). The methanococci utilize H₂S and elemental sulfur as sole sulfur sources. In addition to these, M. thermolithotrophicus utilizes thiosulfate, sulfite, and sulfate as sole sulfur sources (Daniels et al., 1986). The methanococci are not known to use organic sulfur sources, although M. voltae readily assimilates coenzyme M (Santoro and Konisky, 1987).

The methanococci appear to be restricted to marine environments. M. vannielii was isolated from a formate enrichment of black mud from San Francisco Bay, CA, USA. Only a single strain has been described. A number of strains of M. voltae and M. maripaludis were isolated from estuarine sediments and Spartina alterniflora marshes. The source of “M. aeolicus” has not been reported. The single described strain of M. thermolithotrophicus was isolated from geothermally heated sediments at Stufe di Nerone near Naples, Italy. Three strains of M. jannaschii were isolated from hydrothermal vents in the East Pacific Rise and the Guaymas Basin.

The Family Methanomicrobiaceae

The family Methanomicrobiaceae contains H₂-utilizing cocci (Methanogenium and Methanoculleus), as well as disc-shaped (Methanoplanus), rod-shaped (Methanomicrobium and Methanolacinia), and spiral-shaped (Methanospirillum) methanogens. The phylogentic tree is shown in Fig. 4. The coccoid Methanomicrobiaceae have protein S-layers, which cause them to stain Gram-negative and to be osmotically fragile. Cells lyse when exposed to dilute detergents or hypotonic shock. They all grow by reducing CO₂, using H₂ and formate as electron donors. Acetate is generally required as a growth factor and peptones are often required or stimulatory as well; many strains require tungstate and nickel. Motility is rare, although electron micrographs indicate that several strains have flagella.

Fig. 4.

Phylogenetic tree for the families Methanomicrobiaceae (spirillum group) and Methanosarcinaceae (sarcina group) based upon 16S rRNA sequences. Msar., Methanosarcina; Mlo., Methanolobus; Mco., Methanococcoides; SF1 is a strain closely related to Methanohalophilus mahii; Mha., Methanohalophilus; Msae., Methanosaeta; Mg., Methanogenium; Mpl., Methanoplanus; Mm., Methanomicrobium; Mcul., Methanoculleus; Mcr., Methanocorpusculum; Ms., Methanospirillum; and Mc., Methanococcus. (P. E. Rouvière and C. R. Woese, personal communication.)

Methanogenium and Methanoculleus.

Nine species have been described within the genera Methanogenium and Methanoculleus (Table 11). In addition, Methanogenium aggregans, which is not listed, has recently been reclassified as Methanocorpusculum (Xun et al., 1989). Since the time when the genus Methanogenium was originally defined (Romesser et al., 1979), the circumscription of the genus has gradually and informally been broadened to include most coccoid, H₂- and formate-utilizing methanogens that did not fit into the genus Methanococcus. Recently, these organisms were shown to fall into three phylogenetic groups (Xun et al., 1989): The first group, comprised of Methanogenium cariaci (the type species) (Fig. 5) and M. organophilum, includes the two species with a requirement for concentrations of NaCl close to that found in seawater. The second group, which is the most diverse physiologically, has been reclassified as a new genus called Methanoculleus (Maestrojuán et al., 1990). This group includes both mesophiles and thermophiles that require low concentrations of NaCl for optimal growth but may also be halotolerant. The species in this group are Methanoculleus marisnigri, M. bourgense, M. olentangyi, and M. thermophilicum. Methanogenium tationis constitutes the third group, and, on the basis of DNA-DNA hybridization, it is not closely related to other species of Methanogenium or Methanoculleus. However, at this time, it has not been reclassified. Likewise, the phylogenetic positions of “M. frittonii” and “M. liminatans” have not been examined in detail at this time.

Fig. 5.

Methanogenium cariaci. Bar = 10 µm. (Courtesy of D. Boone.)

The substrates for methanogenesis for all nine species in this group include H₂ + CO₂ and formate. Four species, M. marisnigri, M. thermophilicum (one out of three strains), M. organophilum, and “M. liminatans,” also utilize secondary alcohols. In addition, M. organophilum utilizes ethanol and 1-propanol. With the exception of “M. frittonii,” all species require organic carbon supplements. Growth of M. marisnigri requires trypticase, and other peptones and yeast extract do not substitute. Peptones or yeast extract also stimulate the growth of M. organophilum, M. thermophilicum, M. bourgense, M. tationis, and “M. frittonii.” For M. tationis, peptones can be partially replaced by a heavy metal solution. Certain strains of M. thermophilicum also have growth requirements for 4-aminobenzoate, biotin, nickel, molybdate, or tungstate.

Methanogenium and Methanoculleus species have been isolated from a variety of habitats. M. cariaci, M. marisnigri, M. organophilum, and M. thermophilicum were isolated from marine sediments. Additional strains of M. thermophilicum were also isolated from apple, potato, and kelp fed bioreactors. M. bourgense was isolated from a bioreactor fed tannery byproducts. M. olentangyi and “M. frittonii” were isolated from freshwater sediments. M. tationis was isolated from a moderately thermophilic solfataric pool in Chile. “M. liminatans” was isolated from treated industrial wastewater. Because the Methanogenium species grow more slowly than many other H₂- and formate-utilizing methanogens, they may be missed during viable-cell enumerations. In broth culture, they may be overgrown by Methanococcus species or other organisms. In roll tubes, colonies of Methanogenium may take several weeks to become visible. The addition of 0.05–0.20 M NaCl may improve their recovery, however.

Methanoplanus.

The cellular morphology of Methanoplanus is very distinctive (Table 12): by phase contrast microscopy, M. limicola forms rectangular plates, and M. endosymbiosus is disc shaped. As the cells rotate in solution, they can be viewed on edge and appear as very thin rods. The dimensions of M. limicola are 0.07–0.30 µm thick, 1.6–2.8 µm long, and 1.5 µm wide. M. endosymbiosus is more disc shaped, with a diameter of 1.6–3.4 µm. Both species utilize H₂ and formate as substrates for methanogenesis. Best growth is obtained at neutral pH, mesophilic temperatures, and low NaCl (0.20–0.25 M). Growth of M. limicola is stimulated by yeast extract or peptones plus vitamins. The nutritional requirements of M. endosymbiosus have not been fully investigated. The cell envelope of each species contains an S-layer composed of hexagonally arranged subunits. The major proteins stain with the periodate-Schiff reagent and have apparent molecular weights of 143,000 and 110,000 for M. limicola and M. endosymbiosus, respectively. M. limicola was isolated from a small swamp formed from drilling wastes. M. endosymbiosis was isolated from a marine sapropelic ciliate.

Methanomicrobium and Methanolacinia.

Methanomicrobium cells are short rods, but unlike the Methanobacteriaceae, they contain no pseudomurein (Table 12). The physical structure of the cell is apparently conferred by a protein cell wall. Exposure to low concentrations of sodium dodecyl sulfate and other detergents causes rapid cell lysis. Cells stain Gram-negative (Fig. 2). H₂ and formate are substrates for methanogenesis. Growth is most rapid at temperatures near 40°C and between pH 6.1 to 6.9. M. mobile has exceptionally complex nutritional requirements and was isolated from the rumen of a fistulated heifer fed alfalfa hay, where it was very abundant.

Methanolacinia cells are pleomorphic, and they have been described both as short rods and as highly irregular and lobed cocci. Although flagella have been observed, motility has not. Until recently, the single species, M. paynteri, was classified in the genus Methanomicrobium. The single isolate utilizes H₂ and secondary alcohols as substrates for methanogenesis (Zellner and Winter, 1987a). Growth is stimulated by yeast extract, trypticase, and 0.15 M NaCl. The temperature and pH optima are 40°C and pH 7, respectively. M. paynteri may readily be distinguished from Methanomicrobium mobile by differences in its polyamine and lipid content (Zellner et al., 1989b). Methanolacinia paynteri was isolated from marine sediment.

Methanospirillum

Methanospirillum is a spiral-shaped cell (0.4 µm wide by 7.4–10 µm long), twisted into a gentle helix (Table 12). The helical shape may be conferred by a sheath that surrounds the cell (Patel et al., 1986). Cells may form either long chains, or filaments, encased by a continuous sheath or be single (Fig. 6; Patel et al., 1979). M. hungatei also lyses slowly in 1% sodium dodecyl sulfate. Colonies in solidified medium have a wavy appearance, probably conferred by the helical shape of the cells. M. hungatei is the only species in this genus that has been described. H₂+CO₂ and formate are substrates for methanogenesis, and some strains use 2-propanol and 2-butanol as well. The temperature optimum for growth is 35–40°C. Acetate is required or stimulatory. In addition, one strain is stimulated by peptones and B-vitamins. M. hungatei has been isolated from sewage sludge and a pear-waste digestor.

Fig. 6.

Methanospirillum (A) Long, wavy filaments. (B) Individual cells. (From Mah and Smith, 1981.)

The Family Methanocorpusculaceae

This family contains only one genus, Methanocorpusculum, and is more closely related to the Methanomicrobiaceae than to any other families of methanogens. The five species of the genus are very small, irregular cocci that utilize H₂ and formate as substrates for methanogenesis (Table 13). M. parvum and M. bavaricum also utilize secondary alcohols (Zellner and Winter, 1987a; Zellner et al., 1989c). The diameter of the cells varies somewhat with the growth conditions. For M. parvum, the cell diameter is less than or equal to 1 µm. The cell diameters of M. labreanum and M. aggregans vary from about 0.5–2.0 µm. M. aggregans also forms large, multicellular clumps. All species have complex nutritional requirements for peptones, rumen fluid, or yeast extract. In addition, M. labreanum is stimulated by acetate, and M. parvum requires 1 µM tungstate. M. parvum was isolated from a whey digestor and is unusually halotolerant. M. aggregans was isolated from a sewage digestor. M. labreanum was isolated from lake sediment near the La Brea Tar Pits, California. M. sinense was isolated from a biogas plant in Chengdu, China. M. bavaricum was isolated from a wastewater treatment pond in Germany.

The Family Methanosarcinaceae

All of the aceticlastic methanogens belong to this family, as well as all methanogens that disproportionate methanol, methylamines, or other methyl-containing compounds. The only other species that catabolize methanol are Methanosphaera species, which require H₂ to reduce it to methane. The family Methanosarcinaceae can be divided into three physiological and morphological types: 1) the genus Methanosarcina, which contains coccoid and pseudosarcinal cells and can disproportionate methanol and catabolize acetate and H₂ + CO₂; 2) the genus Methanosaeta, which grows only by the aceticlastic reaction; 3) the halophilic, methylotrophic organisms of the genera Methanolobus, Methanococcoides, and Methanohalophilus. Phylogenetically, only the first of these groups is composed of closely related species (Fig. 4).

Methanosarcina.

Methanosarcina species disproportionate methanol and methylamines, forming CH₄ and CO₂ in a ratio of approximately 3:1 (Table 14). Most strains also use acetate, and most also use H₂ to reduce either CO₂ or methanol to CH₄. Formate is never used. Cells appear to have a protein cell wall adjacent to the cytoplasmic membrane. External to the protein wall, many strains have a layer of methanochondroitin, referred to as a matrix. Cells without the matrix are individual, irregular cocci, but when the matrix is present, packets or pseudoparenchyma are formed (Fig. 7). Within the packets, the cells are irregular in shape, the division planes are not always perpendicular, and the volume of the daughter cells is not always equal. Three morphotypes of Methanosarcina have been described. Morphotype 1 includes some strains of M. barkeri that form aggregates of packets of cocci termed pseudoparenchyma, which may be visible to the unaided eye (Zhilina, 1976). Morphotype 2 includes M. vacuolata, which forms small packets and tends to remain dispersed and settles slowly in liquid medium. Morphotype 3 includes M. mazei, which forms single cocci, packets of pseudococci, and large macrocysts (Zhilina and Zavarzin, 1979b; Robinson, 1986).

Fig. 7.

Methanosarcina barkeri strain MS. Bar = 10 µm. (Courtesy of D. Boone.)

While morphology and substrate range are important characteristics of Methanosarcina, they are not always reliable indicators of species differences. These characteristics frequently vary when closely related strains are tested, or the same strain may give varied results under different conditions. For example, a single enzyme may be responsible for changing large pseudoparenchymal aggregates into individual coccoid cells, and this enzyme may be produced only under certain growth conditions (Harris, 1987; Sowers and Gunsalus, 1988; Xun et al., 1988). The ability to use substrates often depends on the conditions of the inoculum (Boone et al., 1987), and the ability to form gas vacuoles is not a constant characteristic of strains (R. A. Mah, personal communication). Thus, variations in these characteristics within species may be commonplace, and they may not always be reliable for the placement of strains into species.

The diameter of the cocci or cells within the aggregates is generally 1–2 µm, although both smaller and larger cells are observed. The cells of Methanosarcina are nonmotile, and in those species that stain Gram negative, the cells lyse in the presence of sodium dodecyl sulfate. Cells of M. vacuolata contain numerous gas vacuoles tightly clustered in the cytoplasm. The Methanosarcina do not require additional carbon sources, and species that use H₂ + CO₂ as a substrate for methanogenesis can grow autotrophically. However, peptones or yeast extract are stimulatory for M. mazei, M. vacuolata, and M. frisia, and riboflavin is stimulatory or required for at least one strain of M. barkeri (Scherer and Sahm, 1981a). The chemicals 4-aminobenzoate and calcium chloride (0.7 mM) are required for growth of M. thermophila. M. barkeri grows with dinitrogen as its sole nitrogen source (Bomar et al., 1985; Murray and Zinder, 1984). Methionine, cysteine, thiosulfate, and elemental sulfur are also sulfur sources for some strains of M. barkeri (Mazumder et al., 1986; Scherer and Sahm, 1981b).

M. barkeri, M. mazei, M. thermophila, and M. vacuolata were isolated from anaerobic digestors. M. barkeri is also found in freshwater and marine sediments, rumens of ungulates, and animal-waste lagoons. M. mazei has been found in garden soil, sewage sludge, and various other sources. M. vacuolata is common in freshwater sediments, marshes and wetlands. M. acetivorans and M. frisia were isolated from marine sediments and for optimal growth require 0.2 and 0.3–0.4 M NaCl, respectively. M. acetivorans also requires 0.05–0.10 M magnesium for optimal growth. In addition to the type strain of M. thermophila, two strains of thermophilic Methanosarcina have been described (Ollivier et al., 1984; Touzel et al., 1985), whose nutritional characteristics resemble those of M. thermophila. However, both the pseudoparenchymal and coccoid morphologies are present, and the taxonomic status of these latter strains is uncertain.

Obligately Methylotrophic Cocci.

Eight species and five genera of obligately methylotrophic, irregular cocci have been described (Table 15). In addition, two species of Methanolobus, M. siciliae and M. vulcani, have been proposed, but a complete description is unavailable (Stetter, 1989). At present, the taxonomy of these organisms is uncertain. Therefore, they have been arranged in the table in the order of increasing optimal salinity for growth, which varies from marine to extremely halophilic. Phylogenetic studies by RNA sequencing, RNA-DNA hybridization and DNA-DNA hybridization suggest that Methanococcoides methylutens, Methanolobus tindarius, Methanohalophilus zhilinae, M. mahii, and Methanohalobium evestigatum probably represent five different genera rather than only four (Sowers et al., 1984b; Chumakov et al., 1987; also see Fig. 4). The placement of the three remaining species listed in Table 15 is not known.

These species grow by disproportionation of methanol, methylamines, and other C-1 compounds. H₂, formate, and acetate are not substrates. Cells are small irregular cocci, and some species form small aggregates. Cells of M. evestigatum are flat and polygonal, and they often occur in sheets (Fig. 8). Methanococcus halophilus produces slime. All the species are nonmotile, except for Methanolobus tindarius, which has monotrichous flagellation. With exception of Methanohalobium evestigatum, they stain Gram negative and lyse rapidly in dilute sodium dodecyl sulfate (or Sarkosyl for Methanococcus halophilus). These species are mesophilic, and their temperature optima are 30–35°C with three exceptions: the temperature optimum for Methanolobus tindarius is 25°C; the temperature optimum for Methanohalophilus zhilinae is 45°C; the temperature optimum for Methanohalobium evestigatum is 50°C. Methanohalophilus zhilinae and M. oregonense are also alkalophilic.

Fig. 8.

Methanohalobium evestigatum. (A) Cells and aggregates, anoptral contrast Reichert. Bar = 5 µm. (B) Ultrathin section of late exponential phase cells. Layers of the envelope slip from the cell surface. Nuclear region and star-shaped granules (possibly glycogen) are present. Bar = 0.5 µm. (Courtesy of T. Zhilina.)

In addition to NaCl, M. euhalobius and M. mahii require high concentrations of magnesium and calcium for good growth. M. methylutens and M. oregonense require high concentrations of magnesium. Although the organic growth factors of the type strain of M. mahii are not known, strain SF-1, which appears to be closely related, requires yeast extract. Vitamins stimulate the growth of M. tindarius and peptones that of M. zhilinae. In contrast, greater than 0.05% yeast extract or peptones inhibit the growth of M. halophilus and M. evestigatum.

M. tindarius and M. methylutens were isolated from marine sediments. M. mahii, Halomethanococcus doii, M. zhilinae, and M. oregonense were isolated from sediments of saline or saline and alkaline lakes. M. halophilus was isolated from a sample containing the cyanobacterial mat and associated mud of a stromatolite in Shark Bay, Australia. M. evestigatum was isolated from a cyanobacterial mat in a hypersaline lagoon close to Arabat, Sivash. M. euhalobius was isolated from the stratal liquid of an exploratory oil well.

Methanosaeta.

In contrast to the other genera of the family Methanosarcinaceae, which are coccoid, the species of Methanosaeta (formerly called Methanothrix) are sheathed rods. These organisms grow only by the aceticlastic reaction (Table 16). Acetate is the only substrate for methanogenesis, and H₂, formate, and methyl compounds are not utilized. Because of their very long generation time, 1 to 3 days under optimal conditions, pure cultures have been difficult to obtain. For instance, the original culture of “M. soehngenii” is not pure, and vancomycin is required to suppress the growth of contaminants (Patel and Sprott, 1990; Touzel et al., 1988). Likewise, the description of M. thermoacetophila is based upon a monoculture, although a pure culture has recently been obtained. Two other pure cultures have also been described. M. concilii appears to be closely related to the methanogen in “M. soehngenii” cultures, and strain CALS-1 appears to be similar to M. thermoacetophila, but insufficient cell mass has been obtained to complete the species description.

Methanosaeta is a short, fat, nonmotile rod with flat ends. Cells stain Gram negative and are sheathed. Except for CALS-1, stationary cultures may form long filaments with a contiguous sheath (Fig. 9). The filaments may form large bundles, mats, or flocs. In shaken cultures, the filaments tend to fragment into short filaments or individual cells. M. thermoacetophila and CALS-1 contain gas vacuoles and M. concilii produces a capsule.

Fig. 9.

Methanosaeta concilii. Bar = 10 µm. (Courtesy of D. Boone.)

Marine or halophilic species have not been described, and NaCl does not need to be added to the media above trace amounts. The pH optima of “M. soehngenii,” M. concilii, and CALS-1 are 7.4–7.8, 7.1–7.5, and 6.5, respectively. Yeast extract inhibits the growth of M. concilii, and penicillin inhibits the growth of “M. soehngenii” and M. thermoacetophila. CALS-1 is very sensitive to the reducing agents in the medium, and 1 mM sulfide and 1 mM 2-mercaptoethanesulfonate appear to be optimal.

Methanosaeta is widely distributed in nature. Its distinctive morphology has allowed it to be identified in sewage sludge, digestors, animal wastes, and sanitary landfills. Monocultures of the thermophiles have been obtained from different types of thermophilic digestors and thermal sediments. “M. soehngenii” and CALS-1 were obtained from sludge digestors. M. concilii was obtained from a pear waste digestor. M. thermoacetophila was obtained from the sediments of a thermal lake.

Methanogens of Uncertain Affiliation

A thermophile, TAM, that morphologically resembles Methanosaeta has been isolated from a sewage sludge digestor (Ahring and Westermann, 1984, 1985). Cells are 0.8–1 µm wide and 4–5 µm in length, and they normally form short filaments. A sheath is present. Cells stain Gram positive to variable. In contrast to Methanosaeta, H₂ + CO₂ and formate are substrates for methanogenesis in addition to acetate. Growth factors are not required during growth on acetate. The pH optimum is 7.3–7.5, and the temperature optimum is 60°C. Penicillin and other typical eubacterial antibiotics are inhibitory at high concentrations.

The isolation of a wall-less methanogen, “Methanoplasma elizabethii,” from an anaerobic chemostat fed glucose was reported but not confirmed (Rose and Pirt, 1981). “M. elizabethii” uses H₂ + CO₂ and formate as substrates for methanogenesis.

Biochemical and Physiological Properties

Coenzymes of Methanogenesis

Methanogens contain many novel coenzymes that are associated with the biochemistry of methane synthesis. Thus, what should be a chemically simple reduction of C-1 compounds to methane is biochemically complex. Because the structure and synthesis of many of the coenzymes have recently been reviewed (Jones et al., 1987), only the most recent citations are included below.

Methanofuran (MFR) (Fig. 10A) functions as a formyl carrier in methanogenesis. It is required for the initial activation of CO₂ to formyl-MFR, the first stable product of CO₂ reduction. MFR has been found in all methanogens examined at levels ranging from 0.5 to 2.5 mg per kg dry weight of cells. At least five different forms of methanofuran exist among the methanogens (White, 1988). This coenzyme has not been detected in either nonmethanogenic archaebacteria, with the exception of Archaeoglobus fulgidus, or eubacteria. The 2,4-disubstituted furan moiety and the 4,5-dicarboxy octanoate moiety are unique structures in nature.

Fig. 10.

The seven coenzymes of methanogenesis.

Tetrahydromethanopterin (H₄MPT) is a one-carbon carrier similar to folate in structure and function (Fig. 10B). Likewise, biosynthesis of H₄MPT is also similar to tetrahydrofolate biosynthesis in eubacteria. The coenzyme was first noticed as a yellow fluorescent compound (YFC) that was rapidly labeled by ¹⁴CO₂ in cell extracts (Daniels and Zeikus, 1978). H₄MPT is also required for several biosynthetic reactions that are folate-dependent in eubacteria, such as serine hydroxymethyltransferase.

A third one-carbon carrier unique to methanogens is coenzyme M (HS-CoM) (Fig. 10C), which chemically is 2-mercaptoethanesulfonic acid. HS-CoM serves as the terminal carbon carrier in methanogenesis where methyl coenzyme M (CH₃-S-CoM) (Fig. 10D) is reduced to CH₄. Since HS-CoM has been found in all methanogens examined, it may serve as a sensitive biomarker for the qualitative identification of methanogens in various ecological niches.

7-Mercaptoheptanoylthreonine phosphate (HS-HTP) (Fig. 10E) is a colorless coenzyme that is required for the final step of methanogenesis. This coenzyme is the electron donor for CH₃-S-CoM reduction, and it is proposed to participate in the energy-conserving step of methanogenesis (Ellermann et al., 1988). It is also required for the activation of CO₂ reduction (Bobik and Wolfe, 1988). HS-HTP is probably biosynthesized from α-ketoglutarate by repeated α-keto-acid chain elongation in a process similar to the aminoadipate pathway of lysine biosynthesis in eubacteria (White, 1989).

Coenzyme F₄₃₀ (Fig. 10F) is a nickel-containing coenzyme required for the final step of methanogenesis, and it is tightly associated with component C of the methylreductase system. The coenzyme was first noticed in cell extracts by J. LeGall and reported by Gunsalus and Wolfe (1978). It is interesting to note that biosynthesis of coenzyme F₄₃₀ proceeds from 5-aminolevulenate via uroporphyrinogen III, which indicates that nickel tetrapyrroles share a common biosynthetic pathway with all other porphinoid compounds (Pfaltz et al, 1987).

Autofluorescence of methanogenic bacteria is largely due to the presence of high levels of coenzyme F₄₂₀ (Fig. 10G). Coenzyme F₄₂₀ is ubiquitous among methanogens, and it is found at low levels in some other organisms. It is a deazaflavin and participates in two-electron transfer reactions. In this regard, it functions in a manner analogous to NADH, but its redox potential of −340 to −350 mV is lower. The coenzyme is an electron donor in methanogenesis. In addition, coenzyme F₄₂₀ is coupled to hydrogenase, formate dehydrogenase, carbon monoxide dehydrogenase, NADP+ reductase, pyruvate synthase, and α-ketoglutarate synthase (Keltjens and van der Drift, 1986).

In addition to the novel coenzymes described above, methanogens contain a number of common vitamins (Noll and Barber, 1988). These include thiamine, riboflavin, pyridoxine, corrinoids, biotin, niacin, and pantothenate. Flavins are known to function as electron carriers for hydrogenase, NADH reductase, formate dehydrogenase, and the methylreductase system in Methanobacterium. Many methanogens contain abundant amounts of unusual corrinoids, with factor III or pseudovitamin B₁₂ being the most predominant (Stupperich and Kräutler, 1988). Corrinoids have also been implicated in methane synthesis, where they function as methyl carriers. In contrast, folates are absent or present in very small amounts in methanogens (Worrell et al., 1988). Presumably, methanopterin substitutes for folate as a methyl carrier. Methanogens have also been found to contain ferredoxin, thioredoxin, and cytochromes b and c. Ferredoxin is abundant in the aceticlastic methanogens, where it is coupled with the catabolic carbon monoxide dehydrogenase, and has been isolated from Methanococcus thermolithotrophicus (Hatchikian et al., 1989; Terlesky and Ferry, 1988a). Thioredoxin was isolated from Methanobacterium thermoautotrophicum, but its function is unknown (Schlicht et al., 1985). Cytochromes function as electron carriers during methanogenesis from methanol, methylamines, and acetate. They are implicated in the oxidation of these substrates during their disproportionation (Terlesky and Ferry, 1988b). In support of this hypothesis, cytochromes are absent from Methanosphaera, which must use H₂ as a reductant during growth on methanol.

The Pathway of Methanogenesis

The pathway of methanogenesis has been extensively reviewed (Jones et al., 1987; Rouvière et al., 1988; Rouvière and Wolfe, 1988). The reduction of CO₂ to CH₄ involves seven steps. The source of electrons may be either H₂ via hydrogenase or formate via formate dehydrogenase. Formate itself does not appear to be an intermediate. Initially, CO₂ is activated to form formyl-methanofuran. Next, the formyl group is transferred to H₄MPT, where it is reduced to the methylene and methyl levels. Last, the methyl group is transferred to coenzyme M and reduced to methane by the methylreductase system.

The first step of CO₂ activation is not clearly understood (Fig. 11, step 1). It requires ATP, but not in stoichiometric amounts. The energy for the reaction is derived from the final step of methane formation (Fig. 11, step 7). This conclusion is supported by the observation that CO₂ activation in vitro requires small amounts of CH₃-S-CoM or a strong reductant such as titanium citrate (Bobik and Wolfe, 1989). The formyl group is transferred from MFR to H₄MPT by formyl methanofuran:tetra-hydromethanopterin formyltransferase (Fig. 11, step 2). The formyl group then undergoes sequential reduction beginning with its conversion to 5,10-methenyl-H₄MPT by the enzyme 5,10-methenyltetrahydromethanopterin cyclohydrolase (Fig. 11, step 3). The methenyl group is reduced by coenzyme F₄₂₀ in a reaction catalyzed by methylenetetrahydromethanopterin:coenzyme F₄₂₀ oxidoreductase to yield 5,10-methylene-H₄MPT (Fig. 11, step 4). The enzymes for steps 5 and 6 have not been purified to homogeneity. Step 5 is presumably catalyzed by methylene-H₄MPT reductase to yield methyl-H₄MPT. Conversion of methyl-H₄MPT to CH₃-S-CoM involves a corrinoid-containing methyltransferase (Kengen et al., 1988; Fig. 11, step 6).

Fig. 11.

The pathway of methane formation from acetate, methanol, and CO₂. Methyl-CoM is the key intermediate for the reduction of CO₂, acetate, and methanol to methane. The numbers refer to the seven steps of the cycle. (Adapted from Rouvière et al., 1988.)

The terminal reaction, catalyzed by the methylreductase system, represents the completion of the cycle with the release of CH₄ and the activation of CO₂ (Fig. 11, step 7). The methylreductase system catalyzes the conversion of CH₃-S-CoM and HS-HTP to CH₄ and CoM-S-S-HTP. Although the exact process is not known, methane formation is also coupled to ATP synthesis via a chemiosmotic mechanism (Blaut and Gottschalk, 1985). However, this coupling may occur during the reduction of the disulfide bond of CoM-S-S-HTP rather than during the reduction of CH₃-S-CoM (Ellermann et al., 1988). CoM-S-S-HTP is also required for the coupling of CH₃-S-CoM reduction to CO₂ activation (Bobik and Wolfe, 1988).

The process of CH₃-S-CoM reduction is complex and requires four protein components and at least three unique coenzymes. Components A1, A2, A3, and C are proteinaceous. A1 and A3 are oxygen-labile and have not been highly purified. A1 has coenzyme F₄₂₀-dependent hydrogenase activity and is probably required for the reduction of CoM-S-S-HTP (Rouvière et al., 1988). A3 has been resolved into two fractions: A3a and A3b. A3a is a large iron-sulfur protein, which may be involved in providing electrons for component C. A3b has methyl viologen-dependent hydrogenase activity (Rouvière and Wolfe, 1989). A2 is an oxygen-stable component and has been purified to homogeneity. A2 and A3 are probably required for the ATP-dependent reductive activation of component C (Rouvière et al., 1988). Component C is an oxygen-stable protein, which is the methyl-S-CoM methylreductase proper. It contains 2 moles of F₄₃₀ and 2 moles of HS-CoM per mole of enzyme, and it has three types of subunits with a stoichiometry of α₂β₂γ₂. The enzyme represents about 10% of the total cellular protein. The genes for component C are organized into a five-gene cluster, mcrBDCGA. The primary sequences of these genes are highly conserved in all methanogens examined. Subunits α, β, and γ are coded for by mcrA, mcrB, and mcrG, respectively. The functions of mcrC and mcrD are unknown (Allmansberger et al., 1989; Weil et al., 1989).

The methylreductase system is also required for methanogenesis from substrates other than CO₂ (Fig. 11). During the catabolism of acetate, the methyl and carboxyl groups are converted to CH₄ and CO₂, respectively, via an aceticlastic reaction. This reaction involves the initial activation of acetate as acetyl-CoA (Terlesky et al., 1987). The carbon-carbon bond is then cleaved by the carbon monoxide dehydrogenase system, producing HS-CoA, an enzyme-bound CO, and a methyl group. The methyl group is transferred to HS-CoM via H₄MPT with the aid of an unidentified corrinoid enzyme (Fischer and Thauer, 1989; van de Wijngaard et al., 1988). Oxidation of the enzyme-bound CO to H₂ and CO₂ provides electrons for the reduction of CH₃-S-CoM to CH₄ (Terlesky and Ferry, 1988b; Bott and Thauer, 1989).

Methane is also produced from the catabolism of methanol and methylamines. Methanogenesis from methanol can occur in two different ways depending on the electron donor: 1) if it is H₂, the methyl group of methanol is transferred to HS-CoM via two methyltransferases (MT1 and MT2). The former contains an oxygen-sensitive cobamide moiety as a methyl carrier. CH₃-S-CoM is then reduced to CH₄ by H₂. 2) When H₂ is absent, some of the methanol is oxidized in a disproportionation reaction to serve as reductant for the methylreductase system. Likewise, for methylamines the methyl group is either reduced to CH₄ or oxidized to CO₂. After growth in the presence of trimethylamine, Methanosarcina barkeri contains a trimethylamine:HS-coenzyme M methyltransferase similar to the methanol-induced methyltransferases.

Carbon Metabolism

Methanogens use only simple organic or inorganic carbon compounds as energy sources. About one-half of methanogens are also capable of autotrophic growth, and they obtain all their organic carbon from the assimilation of CO₂. The proposed methanogenic autotrophic pathway, the Ljungdahl-Wood pathway, is unlike classic autrotrophic CO₂ fixation (the ribulose bisphosphate pathway) since the primary intermediate is acetyl-CoA. The Ljungdahl-Wood pathway was discovered in the acetigenic clostridia and has been extensively studied in Clostridium thermoaceticum (Fig. 12A; Ljungdahl, 1986; Wood et al., 1986). The key enzyme is a nickel-containing carbon monoxide dehydrogenase system (CODH). In methanogens, the one-carbon carrier of the clostridial pathway, folate, is replaced by tetrahydromethanopterin (H₄MPT) (Fig. 12B; Fuchs, 1986). The pathways of autotrophy and methanogenesis are closely linked, and they share common intermediates. Thus, many of the methanogenic coenzymes serve both anabolic and catabolic functions. CO₂ is first reduced to formyl-MFR. The formyl group is then transferred to H₄MPT and reduced to a methyl group. At this point the anabolic and catabolic pathways diverge. The subsequent transfer of this methyl group to HS-CoA is believed to involve a cobamide-containing protein. The carboxyl moiety is derived from CO₂ via CODH and a carbonylation reaction. At this step the carboxyl group freely exchanges with CO in the environment. Although acetogenesis in methanogens appears similar to the pathway in eubacteria, it is distinctive due to the absence of folates and the lack of an ATP requirement for the activation of CO₂.

Fig. 12.

The two acetogenic pathways for the autotrophic fixation of CO₂ to acetyl-CoA in (A) Clostridium thermoaceticum and (B) Methanobacterium thermoautrotrophicum. THF, tetrahydrofolate; E1, carbon monoxide dehydrogenase; E2, the corrinoid enzyme involved in methyl transfer. (Adapted from Jones et al., 1987.)

Methanogens utilize the anabolic tricarboxylic acid (TCA) cycle to synthesize cell carbon from acetyl-CoA (Jones et al., 1987; Fig. 13). However, the exact nature of the cycle varies among different groups of methanogens. In Methanosarcina barkeri, α-ketoglutarate is formed by an oxidative branch. Methanospirillum, Methanococcus, and Methanobacterium utilize the reductive or reverse pathway (Fuchs and Stupperich, 1986). Hexoses are derived from acetate by gluconeogenesis. The major storage product for carbohydrates is glycogen. Very little information is available on how this polysaccharide reserve is used, since no known methanogens can catabolize exogenous sugars. Methanosarcina thermophila is capable of mobilizing its reserves and actually producing small amounts of acetate when starved for acetate (Murray and Zinder, 1987). Likewise, Methanolobus produces small amounts of methane upon mobilization of its glycogen reserves (König et al., 1985).

Fig. 13.

The pathways of carbon assimilation in methanogens. Solid lines represent the incomplete reductive TCA cycle found in Methanobacterium thermoautotrophicum and Methanococcus maripaludis. Broken arrows represent the incomplete oxidative TCA cycle found in Methanosarcina barkeri. (Shieh and Whitman, 1987.)

Heterotrophic methanogens rely on exogenous acetate for growth. An incomplete, reductive TCA cycle has been found in Methanococcus voltae, indicating the presence of a mechanism of anabolic carbon metabolism that is similar to the one found in autotrophic methanogens (Shieh and Whitman, 1987).

Nitrogen and Sulfur Metabolism

Ammonia serves as the sole nitrogen source during autotrophic growth, although some methanogens are capable of utilizing a variety of other nitrogenous compounds. The presence of glutamate synthase in Methanobacterium thermoautotrophicum enables it to use glutamate as a sole nitrogen source (Kenealy et al., 1982). Some methanogens have also been shown to utilize urea, purines, and dinitrogen (Bhatnagar et al., 1984; DeMoll and Tsai, 1986; Lobo and Zinder, 1988). Nitrogen fixation (diazotrophy) in Methanosarcina barkeri requires molybdenum and is inhibited by tungsten, like in the eubacteria (Lobo and Zinder, 1988). For at least one strain, vanadium will substitute for molybdenum (Scherer, 1989). The rate of biosynthesis of the nitrogenase enzyme decreases in the presence of ammonia. In addition, the decrease in growth rate suggests diazotrophy is an energetically expensive process, as it is in eubacteria. Although the enzyme has not been purified, genetic evidence suggests sequence homology between methanogen and eubacterial nitrogenase genes (Sibold et al., 1985).

Sulfide is commonly included as a reductant and a sulfur source in the cultivation of methanogens. However, several species have the ability to use other inorganic sulfur sources, including elemental sulfur, sulfate, sulfite, and thiosulfate (Daniels et al., 1986). The observed reduction of sulfate and sulfite is strictly assimilatory in methanogens. Therefore, it is unlikely that these compounds can serve as alternate electron acceptors. Organic sulfur sources include methionine and cysteine (Mazumder et al., 1986; Scherer and Sahm, 1981b).

Cell Envelope Structure

The diversity of the methanogenic bacteria is demonstrated by the large differences in cell envelope structure. These topics are reviewed in detail by König (1988). The three known types of cell walls are composed of pseudomurein, protein, and heteropolysaccharide.

The cell wall of the Methanobacteriaceae (Fig. 14A and B) is composed of pseudomurein, a peptidoglycan analogous to eubacterial murein (Fig. 15). Pseudomurein contains L-talosaminuronic acid instead of muramic acid, β(1–3) linkages instead of β(1–4) linkages in the polysaccharide backbone, and different sequences of crosslinking amino acids. When these differences are considered, it is obvious why Methanobacterium is resistant to the β-lactam antibiotics that inhibit eubacterial cell wall biosynthesis. Variation of pseudomurein structure within the Methanobacteriaceae occurs by alteration of the amino acid residues.

Fig. 14.

(A) Methanobacterium thermoautotrophicum prepared by freeze-substitution in osmium-acetone. The wall (W) averages 15–16 nm in thinkness and the membrane (PM) 8 nm. (B) M. thermoautotrophicum prepared by chemical fixation: glutaraldehyde-formaldehyde mixture followed by osmium tetroxide. Appearance is very similar to freeze-substituted cell (A and B from Aldrich et al., 1987). (C) Chemically fixed cells of Methanosarcina mazei. Wall (CW) in this preparation appears expanded and fluffy. Wall thickness is 40–50 nm. Small arrows indicate plasma membrane. (D) Freeze-fractured cell of Methanosarcina barkeri, showing large expanse of plasma membrane. Arrows delimit areas in which 12–13-nm-diameter intramembrane particles are arranged in regular arrays. (E) Cell of M. barkeri freeze-substituted in osmium-acetone. Wall (CW) appears dense and compact and measures about 20 nm in thickness. Trilaminar plasma membrane (PM) measures 7 nm. (F) Freeze-substituted cell of M. mazei. Wall and membrane are similar in size and appearance to those shown in (E). Bars = 100 nm. (Figs. 10C–F are courtesy of H.C. Aldrich.)

Fig. 15.

Comparison of murein and pseudomurein structure. D-GlcNAc, D-N-acetylglucosamine; D-MurNAc, D-N-acetylmuramic acid; L-NAcTalNOA, L-N-acetyltalosaminuronic acid; L-DA, L-diamino acid; (I), interpeptide bridge. (Adapted from Kandler and König, 1985.)

In contrast is the simple protein cell wall of the Methanococcaceae and most of the Methanomicrobiaceae. It consists of a crystalline arrangement of proteins or glycoproteins called an S-layer (Fig. 16). Unlike the rigid pseudomurein, the S-layer provides only limited support; therefore, the cells are osmotically fragile. The composition of the S-layer varies between genera based on the presence of glycosylation and between species by molecular weight of the subunits and antigenicity.

Fig. 16.

(A) Thin section of Methanococcus voltae showing the S-layer (S) and plasma membrane (PM). (From Jarrell and Koval, 1989.) (B) Freeze-etch replica of Methanococcus voltae showing the hexagonal arrangement of the S-layer protein (arrows). (From Koval and Jarrell, 1987.) Bars = 100 nm.

Members of the Methanosarcinaceae have a third type of cell wall, referred to as a heteropolysaccharide. The cell wall of Methanosarcina is composed of a polysaccharide matrix (Fig. 14 C–F), which is similar in structure to eukaryotic chondroitin found in connective tissue. This polysaccharide stabilizes the cell aggregates formed by Methanosarcina species. The work of Aldrich et al. (1986) and Kreisl and Kandler (1986) has elucidated the chemical structure and arrangement of these polysaccharides on the cell surface as a loose matrix.

Methanosaeta (“Methanothrix”) and Methanospirillum, which are characteristically filamentous, have an outer protein sheath (Figs. 17 and 18). The cells are surrounded by an electron-dense inner wall of unknown composition and are separated by septa or plugs. The crystalline structure of the sheath has been studied in considerable detail by Beveridge et al. (1986) and Patel et al. (1986).

Fig. 17.

(A) Thin section of a Methanosaeta consilii filament showing spacer plugs (arrow heads) separating the cells and the amorphous matrix (M) surrounding each cell. Bar = 1 µm. (B) Thin section showing the spacer plug (P) and the matrix (M) between two cells. Bar = 100 nm. (C) Thin section showing the sheath (S), the matrix (M), and the plasma membrane (PM). Bar = 100 nm. (From Beveridge et al., 1986.)

Fig. 18.

(A) Thin section of a Methanospirillum hungatei filament showing a spacer plug (arrow head) separating two cells. Bar = 500 nm. (B) Thin section showing spacer plug (P) and sheath (S). Bar = 100 nm. (C) Thin section showing sheath (S), cell wall (W), and plasma membrane (PM). Bar = 100 nm. (From Beveridge et al., 1987.)

The cell membranes of methanogens are characterized by several unusual lipids (see reviews by De Rosa and Gambacorta, 1988; Langworthy, 1985; and Jones et al., 1987). In the majority of methanogens, the lipids are polar, either diphytanyl-glycerol-diethers or dibiphytanyl-diglycerol-tetraethers (Fig. 19). Thus, the membrane structure in many methanogens differs from the lipid bilayer typical of eubacteria. The tetraethers appear to span the membrane, with the polar head groups at opposite sides. Therefore, the membranes are probably arranged in a monolayer with bilayer regions resulting from interspersed diethers. As in eubacteria and eukaryotes, these lipids are often present as phospho- or glyco-derivatives. The major nonpolar lipids consist of isoprenoid and hydroisoprenoid hydrocarbons.

Fig. 19.

Structures of a characteristic diether lipid and tetraether lipid found in the membranes of methanogenic bacteria. (Adapted from Jones et al., 1987.)

Molecular Biology of the Methanogens

Because the methanogens and other archaebacteria represent a very ancient line of descent, there has been a great deal of interest in their molecular biology. Thus, the molecular features that the methanogens share with the eubacteria and eukaryotes may have been inherited from the common ancestor of all three kingdoms. Only a brief overview of this topic is presented here, but a number of detailed reviews on different aspects are available (Brown et al., 1989; Dennis, 1986; Fox, 1985; Jones et al., 1987; Kandler and Zillig, 1986; Matheson, 1985; Thomm et al., 1986; Woese and Wolfe, 1985; Zillig et al., 1988). Furthermore, an issue of the Canadian Journal of Microbiology (1989; vol. 35, no. 1) is devoted to this topic.

Methanogens have some genetic characteristics of both eubacteria and eukaryotes. Sequence analysis of methanogen genes suggests that they are arranged in operons similar to those in eubacteria. However, the process of transcription initiation and the termination sequences of certain genes are more similar to those found in eukaryotes. Moreover, the methanogen DNA polymerases that have been purified resemble both eubacterial and eukaryotic enzymes.

The DNA-dependent RNA polymerase of methanogens is genetically more closely related to the enzyme of eukaryotes than of eubacteria (Bernghofer et al., 1988). As a consequence, it might be expected that the methanogenic promoter sequences should also resemble eukaryotic sequences. Analysis of these sequences in numerous methanogen operons has revealed two conserved sequences. One has the consensus sequence of AAANNTTTATATA, where N represents any of the four nucleotides (Allmansberger et al., 1988). This sequence is found 25 bp upstream of the start site of transcription and resembles the “TATA” box found in eukaryotes. Binding of purified RNA polymerase to this sequence has been demonstrated in the methylreductase operon, hisA, and in the ribosomal RNA genes of Methanococcus vannielii (Thomm et al., 1989). The second conserved region, TGCAAGT, has been found at the start site of transcription of the ribosomal RNA genes. Termination of transcription has not been studied in detail. However, the gene mcrA, which codes for the α subunit of methylreductase, has a sequence analogous to the rho-independent terminator of eubacteria. In contrast, analysis of rRNA genes suggests a termination mechanism similar to that of eukaryotes, which use AT-rich sequences.

DNA polymerases in methanogens are heterogeneous. The enzyme purified from Methanococcus vannielii resembles the type A enzyme of eukaryotes in its sensitivity to the inhibitor aphidicolin (Zabel et al., 1987). In contrast, the purified enzyme of Methanobacterium thermoautotrophicum is resistant to aphidicolin and is similar to DNA polymerase I of Escherichia coli (Klimczak et al., 1986). The enzymes of M. vannielii and M. thermoautotrophicum also differ from each other in molecular weight.

The methanogen ribosome has a “typical” prokaryotic composition. The ribosome is 70S in size and dissociates into 30S and 50S subunits. The 30S subunit contains 16S rRNA, and the 50S subunit contains 5S and 23S rRNAs. A 5.8S rRNA, as found in eukaryotes, is not present. However, the structure of the methanogen ribosome differs from the eubacterial ribosome in that the 30S subunit has the characteristic archaebacterial “bill,” a small protrusion for which the function is unknown. A similar “bill” is also present on the eukaryotic 40S subunit (Lake et al., 1982). Ribosomal proteins of methanogens and other archaebacteria are unusually acidic, in contrast to the basic ribosomal proteins of eubacteria and eukaryotes. However, the number of acidic ribosomal proteins correlates well with the internal K⁺ concentration in the strains studied. The ribosomal proteins of M. vannielii are encoded by a transcriptional unit homologous in organization to the “spectinomycin operon” of E. coli. However, the deduced amino acid sequences of the methanogen proteins more closely resemble those of their eukaryotic homologs (Auer et al., 1989; Köpke and Wittmann-Liebold, 1989).

Sequencing of methanogen rRNAs has been instrumental in establishing their phylogenetic relationship with eubacteria and eukaryotes. The secondary structure of the methanogen 5S rRNA has characteristics of both the eubacterial and eukaryotic molecule. However, the methanogen rRNA molecule has characteristics that are unique to archaebacteria. Sequence data for the 16S rRNA support the hypothesis of the extreme diversity of the archaebacteria. The secondary structure of the methanogen 16S rRNA is more similar to that of the eubacterial structure than the eukaryotic secondary structure. Less information is available concerning the secondary structure of the 23S rRNA molecule. However, it does have characteristics of both the eubacterial and eukaryotic molecules. The genes for rRNA are arranged in a typical operon similar to that of eubacteria. The 16S, 23S, and 5S genes are closely linked and interspersed with several tRNA genes. Methanococcus vannielii and M. voltae also have unlinked 5S rRNA genes that are associated with tRNA-encoding genes.

Antibiotic Sensitivity and Resistance

The lack of sensitivity of methanogens and other archaebacteria to most antibiotics is a reflection of their unique biochemical nature and has been reviewed previously (Böck and Kandler, 1985). Methanogens are resistant to many commonly used antibiotics simply because the specific target is not present. Therefore, comparison of the antibiotic sensitivity spectrum of methanogens with those of eubacteria and eukaryotes can provide insight into their biochemical similarities and differences.

The differences in chemical structure of methanogen cell envelopes to those of eubacteria have been well established. These differences are supported by the lack of sensitivity of methanogens to many cell-wall inhibitors (Table 17). For example, Methanobacterium thermoautotrophicum is resistant to penicillin G, since the crosslinking of pseudomurein, which replaces peptidoglycan in the cell wall, is not inhibited by the antibiotic. However, antibiotics such as bacitracin, which inhibit the formation of lipid-bound precursors of murein in eubacteria, may inhibit formation of similar lipid-bound precursors involved in biosynthesis of lipids and glycoproteins in methanogens.

Table 17. Comparison of the antibiotic sensitivity of several species of methanogens, eubacteria and eukaryotes.

In addition to genetic homology, the antibiotic sensitivity of DNA-dependent RNA polymerase reflects the similarity of the methanogen enzyme to the eukaryotic enzyme (Table 17). Both enzymes are resistant to rifampicin, an antibiotic which inhibits RNA polymerase of eubacteria. The methanogen enzyme is also resistant to α-amanitin, which inhibits the type B and C polymerases of eukaryotes but not the type A enzyme.

Antibiotics that inhibit protein synthesis may block aminoacyl-tRNA synthesis, elongation, or translation. Pseudomonic acid is a potent inhibitor of Methanococcus voltae and inhibits isoleucyl-tRNA synthetase activity (Possot et al., 1988). Translation in methanogens is resistant to many of the antibiotics that inhibit translation by either the eubacterial 70S or eukaryotic 80S ribosomes (Böck and Kandler, 1985). While growth of the methanogens is sensitive to chloramphenicol, this inhibition is not due to an effect on protein synthesis. Instead, inhibition is probably due to the sensitivity of the particulate hydrogenase of Methanobacterium to this antibiotic and to other halogenated compounds. However, the elongation factor aEF-2 is sensitive to diphtheria toxin, which is also characteristic of the eukaryotic protein (Lechner et al., 1988). Sequence analysis of the genes encoding aEF-2 indicates that the methanogen enzyme is more closely related to its eukaryotic than its eubacterial homolog. In addition, all archaebacterial aEF-2 genes examined possess a histidine residue that is posttranslationally modified to diphthamide, a phenomenon analogous to that of the eukaryotic EF-2. Translation in the methanogens is also resistant to most antibiotics in the streptomycin class of aminoglycosides, as well as to tetracycline and the macrolides. However, methanogens are sensitive to deoxystreptamine aminoglycosides, such as gentamycin and neomycin, as well as to virginiamycin. Therefore, the translation process in methanogens is different from that of the eubacterial process with respect to antibiotic sensitivity.

Antibiotics that act on membrane integrity also have varying effects on methanogens, as they do with eubacteria. Ionophores such as monensin, lasalocid, and gramicidin S are very effective inhibitors of methanogens. This inhibition is presumably the result of interference with Na⁺ transport and thus with methanogenesis (Perski et al., 1982). Monensin and lasalocid inhibit both growth and methanogenesis (Dellinger and Ferry, 1984). However, this inhibition depends on the culture conditions, since they both stimulate methanogenesis at low Na⁺ concentrations in vivo (Schönheit and Beimborn, 1986).

Motility

Motility in methanogens has been associated with the presence of flagella. However, little is known about their structure, attachment to the cell envelope, and the energetics of motility. Kalmokoff et al. (1988) found that the flagella of Methanococcus voltae, consisting of two protein subunits with molecular weights of 33,000 and 31,000, have a filament diameter of 13 nm, unlike those of eubacteria which generally consist of only one monomer. The basal structure is knob-like in appearance, as opposed to the multiple ring structure found in eubacteria. This may be due to the simpler structure of the methanococcal S-layer. It will be interesting to compare the basal structures of methanogens that have more complex cell envelopes. The arrangement of flagella varies greatly among methanogen genera, as it does in eubacteria. Methanococcus jannaschii has lophotrichous flagella with two bundles of flagella at one pole arranged in a “corkscrew” fashion (Jones et al., 1983a). In contrast, Methanogenium species have peritrichous flagella (Romesser et al., 1979).

Chemotaxis in methanogens has been described in Methanospirillum hungatei, for which acetate is a chemoattractant (Migas et al., 1989). In addition, Methanococcus voltae responds to acetate, isoleucine, and leucine as attractants but not to histidine (Sment and Konisky, 1989). M. voltae has an absolute requirement for all these nutrients except histidine. Therefore, it is evident that some methanogens respond positively to higher nutrient concentrations. The advantage of this in certain habitats, such as marine marshes where substrate concentrations may be low, is self evident.

Applications

Methanogenic bacteria have numerous applications, especially in the degradation of wastes rich in organic matter to methane and CO₂. Methane is the major component of natural gas, a clean fuel which is already in wide use. This topic has been reviewed extensively (Daniels, 1984; Hobson et al., 1981; van den Berg, 1984; Wodzinski et al., 1987), and only a few major points will be discussed here.

Because of the narrow substrate specificity of methanogens, pure cultures have limited usefulness in waste treatment. For most applications, consortia containing both methanogenic bacteria and heterotrophic bacteria capable of converting the complex organic materials common in waste materials to methanogenic substrates are used. The consortia are usually obtained from an inoculum of manure, sewage sludge, freshwater sediment, or some other natural source, and they are formed upon enrichment with the waste material of interest under methanogenic conditions. In this manner, consortia have been obtained for the degradation of a wide variety of wastes and other materials. Moreover, thermophilic consortia have been obtained by performing the enrichments at high temperatures.

For example, methanogenic consortia that degrade whey, a major by-product in milk and cheese production, have been described (Chartrain and Zeikus, 1986a, 1986b; Wildenauer and Winter, 1985; Zellner et al., 1987b; Zellner and Winter, 1987b). These consortia were composed of sugar- and protein-fermenting strains of Lactobacillus, Eubacterium, Clostridium, Klebsiella, and Leuconostoc. Their fermentation products were then metabolized by acetogenic strains of Desulfovibrio, Eubacterium, and Clostridium. The acetate, H₂, and formate formed were substrates in turn for the methanogens present, Methanobacterium, Methanobrevibacter, Methanospirillum, Methanocorpusculum, Methanosarcina, and Methanosaeta. These complex consortia were capable of the nearly complete conversion of whey to methane and CO₂ within 5 days.

Another common application of methanogens is in the treatment of domestic sewage. Whether it be a simple cesspool or an advanced anaerobic digestor, the methane produced from sewage can be collected and used to offset the cost of an otherwise obligatory process. In countries such as India and China, where the economic resources required for such digestors are not available, animal and human wastes are still used as a source of energy. Instead of advanced digestors, these countries have developed inexpensive small scale digestors for production of methane. In addition, the spent slurry can be used as a natural fertilizer. Obvious drawbacks of these digestors are their limited size and requirement for manual loading and emptying. The digestors are also limited to temperate climates since they must rely on the ambient temperature to eliminate the additional heating costs required to maintain an active biomass.

While consortia can be obtained for the conversion of many types of organic wastes to methane and CO₂, the growth of these consortia is usually slow when compared to growth of those composed of aerobic microorganisms. This can cause problems since high waste input can result in loss of cell biomass, and thus waste decomposition is limited by the growth rate of the bacteria. Therefore, to increase efficiency, specialized bioreactors have been designed to retain the microbial population in the bioreactor while allowing for a high input of waste. In this way, the rate of waste decomposition is not limited by the growth rate of the bacteria. The five types of bioreactors commonly used have been reviewed by van den Berg (1984) and are described below:

• 1. The anaerobic contact reactor uses a sludge, which settles to the bottom and contacts the raw waste. Thus, the settling ability and mixing of the sludge with the waste are important in the efficiency of bioconversion. This bioreactor type works well for particulate wastes, which settle easily and are completely biodegradable.

• 2. The anaerobic filter reactor retains suspended bacteria and waste in a packing material or solid support where degradation occurs. It is advantageous for treatment of dilute soluble wastes. However, it is easily plugged by suspended particulate waste.

• 3. The upflow anaerobic sludge bed reactor avoids plugging by reducing the volume of the packing material and using a gas collector to encourage settling of the sludge. High concentrations of microbial biomass are needed in addition to proper development of a granular sludge to promote settling.

• 4. The anaerobic fluidized or expanded bed reactors allow growth of biomass on inert particles, which settle and are suspended by a rapid flow of waste.

• 5. The downflow stationary fixed-film reactor avoids plugging of the anaerobic filter by forming an active biomass film. This reactor can accommodate a wide variety of wastes.

Another area of potential applications of methanogenic consortia is in degradation of xenobiotics (for reviews see Berry et al., 1987; Young, 1984). Especially important is the degradation of halogenated aromatic and aliphatic compounds, which are common pollutants. For instance, tetrachloroethylene is a common pollutant in ground waters due to improper disposal in landfills (Fatherpure and Boyd, 1988). In the presence of sewage sludge or pure cultures of methanogens, it is reductively dehalogenated to trichloroethylene. In contrast, tetrachloroethylene is not rapidly degraded under aerobic conditions. Likewise, low concentrations of chloroform, carbon tetrachloride, and 1,2-dichloroethane are almost completely oxidized to CO₂ by methanogenic consortia (Bouwer and McCarty, 1983). Haloaromatic compounds are also reductively dehalogenated by methanogenic consortia (Suflita et al., 1983). Thus, chlorinated phenols and chlorinated benzoates are converted to phenol and benzoate, respectively. Phenol may then be carboxylated to form benzoate (Knoll and Winter, 1989). Benzoate is then converted to methane and CO₂, presumably by a reductive route via cyclohexane carboxylate.

Methanogens have also been implicated in anaerobic biocorrosion, which is a significant economic problem. By the process of cathodic depolarization, elemental iron is oxidized and protons in water are reduced to H₂ (Daniels et al., 1987). Methanogens accelerate this process by maintaining a very low partial pressure of H₂. The mechanism is similar to that proposed for sulfate-reducing bacteria, which also consume H₂. Protons from water combine with electrons from iron to form hydrogen, which is utilized by the hydrogenase enzyme.