Methane-Oxidizing Microorganisms › content › mmbr › 45 › 4 › 556.full.pdf · METHANE-OXIDIZING MICROORGANISMS 557 rized in Table 1, whichwascompiled fromthe data of Ehhalt

MICROBIOLOGICAL REVIEWS, Dec. 1981, p. 556-590 Vol. 45, No. 40146-0749/81/040556-35$02.00/0

Methane-Oxidizing MicroorganismsI. J. HIGGINS,'` D. J. BEST,' R. C. HAMMOND,2 AND D. SCOTT'

Biotechnology Unit, Cranfield Institute of Technology, Cranfield, Bedford, MK43 OAL,' and UnileverResearch, Colworth Laboratory, Shamrbrook, Bedford, MK44 1L9,2 United Kingdom

INTRODUCTION............5....5............5.....6BIOLOGY ..................................................................7...57Taxonomy .......................................5...........................57Ecology ............................................. .............. 658Morphology and Physiology ................................................ 558

Fine structure of type I methane-utilizing bacteria.559Fine structure of type II methane-utilizing bacteria.559Effect of growth conditions on intracytoplasmic membranes.560Phospholipid and fatty acid compositions ofmethanotrophs560Exospore and cyst formation by methanotrophs.562Ultrastructure of methane-utilizing yeasts.563

CARBON METABOLISM.563The Serine Pathway.563The Ribulose Monophosphate (Quayle) Cycle and Other Aspects of Carbon

Assimilation by Type I Methanotrophs.567The Dissimilatory Ribulose Monophosphate Pathway.570Intracellular and Extracellular Polymers Produced by Methane-Utilizing

Bacteria ................................................................ 570Intermediary Metabolism, the Tricarboxylic Acid Cycle, and Utilization of

Ancillary Carbon Sources . 571Methane-Utilizing Yeasts 5.73

ENERGY METABOLISM.573Methane Monooxygenase.573Comparison of methane monooxygenases from different species.575Mechanisms of oxidation by methane monooxygenase.576

Methanol Dehydrogenase.577Formaldehyde Dehydrogenase.577Formate Dehydrogenase.579Secondary Alcohol Dehydrogenase.579Oxidation of Multicarbon Compounds.579Energy Transduction and Growth Yields.579Methanotrophic Yeasts and Anaerobic Methane Oxida.tion .580

NITROGEN METABOLISM 680GENETCS OF METANOTROPHS .581EVOLUTION OF METIANOTROPHS .581BIOTECHNOLOGICAL APPLICATIONS OF METHANOTROPHS.583ITMRATURE CITED.583

INTRODUCTION

Despite studies of methane-oxidizing micro-organisms (methanotrophs) spanning some 75years, their importance in global elemental cy-cles is neither widely acknowledged nor fullyunderstood. Nevertheless, particularly in thelast 15 years, many groups have accepted thechallenge of methanotrophic microbiology, insome cases spurred on by the prospects of com-mercial exploitation of these microorganisms,initially for single-cell protein production andmore recently as biocatalysts or sources of usefulpolymers. There is now a substantial body ofknowledge of methanotrophy, but it is not ourintention here to offer an extensive historicalreview; rather, we wish to highlight recent dis-

coveries and to assess the environmental impor-tance and possibilities for commercial exploita-tion of this fascinating group. For complemen-tary information on methanotrophs, the recentreviews by Colby et al. (20), Hanson (50), Wolfeand Higgins (195), and Higgins (59) should beconsulted. The microbial transformation of C1compounds in general is discussed by Higginsand Hammond (63), and two excellent articleson the microbial assimilation of C, compoundswere recently published by Quayle (131, 132).The magnitude of turnover in the biosphere

of the major carbon source for methanotrophs,methane, is not widely recognized. Approxi-mately 50% of total organic carbon degraded byanaerobic microflora is converted to methane.Sources of atmospheric methane are summa-

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METHANE-OXIDIZING MICROORGANISMS 557

rized in Table 1, which was compiled from thedata of Ehhalt (41). Biogenic methane is themajor source of atmospheric methane and isquantitatively similar to the output from naturalgas wells. It is equivalent to about 0.5% of theannual production of dry organic matter, butrecent environmental studies (discussed in Bi-ology) indicate this to be only a small fraction oftotal biogenic methane, most ofwhich is oxidizedmicrobiologically to C02 before the carbonreaches the atmosphere. Methane is, therefore,one of the most abundant organic compoundson this planet, and this is reflected by the abun-dance of methanotrophs in the environment.They can represent up to 8% of the total "het-erotrophic" population, and in some sedimentsmethanotroph counts of 5 x 107/ml have beenreported (184). Thus, methanotrophs constituteone of the major groups of free-living microor-ganisns, and their metabolic activities play a

major role in determining the nature of theenvironment. Although it is self-evident thatmethanotrophs may play a role in maintaining

low levels of methane in the atmosphere (1.4ppm, vol/vol), other consequences of their activ-ities are less well understood or quantifiable atthe time of writing. There is, nevertheless, ac-

cumulating evidence that this group of micro-organisms- plays important roles in the nitrogencycle and in the degradation of complex, partic-ularly hydrophobic, organic compounds (60).

In the last few years, several concepts aboutthe nature of methanotrophy have been ques-tioned by a number of studies. The most inter-esting aspects concern the prevalence and natureof obligate methanotrophy versus facultativemethanotrophy, anaerobic methane oxidation,ancillary metabolism of complex compounds,the multiplicity of C, incorporation pathways insome species, the nature of methane monooxy-

genases (MMOs), and the role of intracyto-plasmic membranes. These aspects are exam-

ined in this review, and an attempt is made togive a contemporary view of the environmentalsignificance of these microorgasms and to as-

sess the potential for their industrial exploita-tion.

BIOLOGYTaxonomy

Most methanotrophic bacteria that have beenisolated are obligate for reduced C, substratesand gram-negative, obligately aerobic rods,vibrios, or cocci. The most extensive amount ofpublished data on isolation of these organis isdue to Whittenbury and his colleagues (188),who classified the isolates into type I (Methylo-coccus, Methylomonas, and Methylobacter) and

TABLE 1. Sources of atmospheric methaneGlobal methane

Source production(Mt/yr)

BiogenicaEnteric fermentation in animals 101-220Paddy fields .................... 280Swamps, marshes ........ ....... 130-260Freshwater lakes ......... ....... 1.3-25Upland fields ................... 10Forests ......................... 0.4Tundra ........................ 0.8-8OceansOpen ........................ 4-6.7Continental shelf ........ ...... 0.07-1.4

Total biogenic ........... ....... 528-812

Other sourcesCoal mining .................... 6.3-22ignite mining ........... ....... 1.6-5.7Industrial losses .......... ....... 7-21Automobile exhaust ....... ...... 0.5Volcanic emissions ........ ...... 0.2

Total other sources ........ ...... 15.6-49.4

TOTAL ALL SOURCES .......... 544-862

Other relevant figures for comparisonOutput of natural gas wells for con-

sumption (1965) ....... ...... 520Dry organic matter ........ ...... 165,000Atmospheric methane (about 1.4

ppm, vol/vol) ........ ....... 4,000a These figures indicate only methane released into

the atmosphere and do not include biogenic methaneoxidized by methanotrophs.

type II (Methylosinus and Methylocystis). Themajor division between types I and II was basedupon differences in intracytoplasmic membranearrangement and carbon incorporation path-ways. Most workers accept this as a basis fordevelopment of a formal classification, and mostorganisms isolated since this work fall more orless into one of these groups. As more detailedknowledge has accumulated, the need for twosubgroups (A and B) above the genus level hasarisen for classification oftype I organisms (188),and the new genus offacultative methanotrophs,Methylobacterium (117, 122), has been accom-modated as a subgroup within type II. It shareswith type II species, which use only reduced C1substrates, the characteristics of paired periph-eral membranes when growing on methane andC, incorporation via the serine pathway. Recentdeoxyribonucleic acid (DNA) homology studies(50) suggest, however, that this classification isinappropriate, for there is little homology be-tween the DNAs of Methylobacterium organo-philum XX and a representative type II species,

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Methylosinus trichosporium OB3b. Type I spe-cies have membranes that are bundles of vesic-ular disks, and they use the ribulose monophos-phate (RMP) cycle. Subgroup B species alsopossess the ribulose diphosphate pathway ofC02 fixation. Attempts are now being made todevise a formal taxonomy and nomenclature ofmethanotrophs (140, 142), although there is acontinuing need to accommodate the unex-pected, e.g., Methylococcus capsulatus (Bath),which has three pathways for incorporation ofC, units: ribulose diphosphate, RMP, and serinepathways (166). This may represent a third ma-jor group, type X (185).

Recently, yeasts have been isolated which arefacultative methanotrophs (192) and have beenidentified as strains of Sporobolomyces roseus,Sporobolomyces gracilis, Rhodotorula glutinis,and Rhodotorula rubra (193).Some bacterial species can oxidize methane

anaerobically by coupling the oxidation to sul-fate reduction (109), but these remain to beidentified. The recent finding that well-charac-terized methanogenic bacteria can also oxidizemethane, albeit slowly, is fascinating (198). Al-though the rates involved are only 0.001 to 0.3%ofthe rate ofmethane formation, the mechanismappears to be distinct and not a simple reversalof the methanogenic system. There is inconclu-sive evidence that some algae and fungi canoxidize methane (42, 197).

EcologyThe vast majority ofmethane-utilizing species

isolated from a wide range of natural environ-ments are aerobic, obligate, methanotrophicbacteria. Whether this is a true reflection ofrelative abundance or an artifact due to isolationprocedures remains to be established.Methane is continually generated in anaerobic

environments, and there is good evidence for itsanaerobic oxidation linked to sulfate reductionby uncharacterized microorganisms present insediments (50, 109). In most situations wheremethane diffuses into aerobic environments,large populations of aerobic methanotrophs canbe found, e.g., soils, surface layers of sediments,and natural waters. As mentioned in Introduc-tion, their numbers can be extremely high insediments, and counts of 2.5 x 105/ml have beenmeasured in lake water (77). The greatest num-bers in water columns are found in the thermo-cline zone of meromictic lakes. The indigenouspopulation is often highly effective in oxidizingmethane. For example, in experiments with col-umns of sediments, in some cases none of themethane generated in the lower, anaerobic re-gions is released into the atmosphere above the

sediment (184). Similarly, gas bubbles releasedfrom a lake bottom showed a 75% decrease inmethane content over a vertical distance of 10m (41). It is clear, even from the relatively fewdeterminations of numbers of methanotrophs inthe environment, that they are ubiquitous andrepresent a significant proportion of the naturalmicroflora. There is no indication that particularspecies are habitat specific; rather, a variety ofspecies are found in all habitats (184). Thermo-philic and thermotolerant strains have been iso-lated (96, 184, 188).With a fluorescent antibody staining tech-

nique, Reed and Dugan (134) surveyed the dis-tribution of Methylomonas methanica and M.trichosporium in Cleveland Harbor (Cleveland,Ohio). Concentrations ofthe two methanotrophswere inversely proportional to sampling depth,suggesting that they grew primarily in or about1 m above the sediment. M. methanica wasobserved at every sampling station and M. trie. -osporium was observed at only two, but a sea-sonal winter variation in the former, but not inthe latter, correlated with the laboratory obser-vation that M. methanica was cold sensitive.This study supports the view that methane oxi-dizers grow primarily just above the anaerobicenvironments, in areas of low oxygen tensions.

It may be that some methanotrophs, althoughhaving an obligate requirement for reduced Clcompounds, commonly co-utilize ancillary car-bon and energy sources. This is discussed furtherin Carbon Metabolism. It is even possible thatsome species have an obligate requirement formethane plus another organic substrate. To ourknowledge there have been no attempts to iso-late such strains. Ecological aspects are not dis-cussed in detail here, since a highly authoritativereview ofmethylotrophic ecology by Hanson hasrecently been published (50).

Morphology and PhysiologyAn outstanding morphological feature of

methanotrophs is the ability to develop exten-sive intracytoplasmic membrane structures.These have been described by several groups ofworkers in many species (195) and, as discussedabove, are used as a taxonomic criterion. Similarmembranes are found in photosynthetic bacte-ria, ammonia and nitrite oxidizers, blue-greenalgae (cyanobacteria), and some higher hydro-carbon utilizers, and they contain lipids that areunusual in bacteria (195).Accepting that these membranes need not

fulfil the same role in these different groups, inthe current context, what is their role in meth-anotrophs? They are clearly not necessary forgrowth on methanol, since facultative methylo-

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trophs are devoid of them. It has therefore beenassumed that the membranes are associated spe-cifically with the initial metabolism of methane,and this contention would appear to be sup-ported by the fact that facultative methano-trophs only contain such membranes whengrown on methane (94, 123, 195).Fine structure oftype I methane-utilizing

bacteria. Type I methanotrophs possess uni-form arrays of membranes, distributed evenlythroughout the cytoplasm, consisting of astacked series of flattened discoidal vesicles (28,30, 57, 76, 126, 150). Early studies suggested auniformity of morphology in these arrays, eachsaccule being limited by a unit membrane 7.5 to8.0 nm in width and adjacent saccules beingseparated by a 2.0-nm gap. The saccule widthvaried between 15.0 and 20.0 nm (150). Conti-nuity between these intemal membrane arraysand the cytoplasmic membrane has been noted(28, 30, 46). In species subject to detailed exam-ination (M. capsulatus, Methylomonas sp.), theinternal membrane system is extensive, occu-pying most of the cell volume (30, 76, 150).Branching of these internal membranes has alsobeen noted (30).

Recently, however, it has been demonstratedthat the internal morphology of these type Imethane utilizers is radically affected by cultureage and condition (76). Exponentially growingorganisms exhibit the classical orderly parallelstacks of membrane sacs described above. How-ever, as the culture enters the early stationaryphase, these membrane arrays become less nu-merous and less orderly, and many organismscontain electron-lucent droplets of unknowncomposition. In the late stationary phase thisdisordered state becomes even more pro-nounced, the membrane sacs being widely dis-tended, if present at all, and many spheroidalinclusions are found. In addition to this degen-eration of the regular membrane arrays in oldercultures, other morphological pecularities havebeen reported. In a Methylomonas sp. two typesof polar organelle have been described, bothapparently contiguous with the cytoplasmicmembrane. One is 20.0 nm wide, from the cyto-plasmic to the limiting membrane, the inter-space being filled with "spikes" perpendicular tothe bordering layer. The second possesses twobordering layers, each 7.0 nm thick and 1.5 nmapart, parallel to and 45.0 nm from the cytoplas-mic membrane. The intervening space is filledwith spikes 30.0 nm in length. Older cells alsocontain myelin-like structures, consisting of avariable number of electron-dense and electron-transparent striated layers with a periodicity of3.2 nm. It has been suggested that lipid material

from degenerating membranes is transformedinto these structures (30).

Later stationary-phase cultures of M. capsu-latus often possess crystalloid inclusions, pre-sumably protein, with regular 10-nm spacing(76). In addition to the uniform membranestacks observed in Methylococcus mobilis sp.nov. (57), regular patterns of tubular structures22 nm in diameter have been observed when thisspecies is grown with peptone as the organicnitrogen source. The significance of these var-ious structures and their prevalence are un-known.

All detailed studies of type I methanotrophshave involved organisms from batch cultures inshake flasks. Little is known of their morphologyunder carefully defined growth conditions, i.e.,in a fermentor, although De Boer and Hazeu(30) noted that the membrane stacks were lessextensive under these conditions, more areas oflow electron density being scattered throughoutthe cytoplasm, with the outer double layer ofthe cell wall forming unusual projections.The presence of membranes in methanol-

grown type I species has also been reported (30,46, 76). However, organisms are largely filledwith electron-lucent droplets, but transfer backto growth on methane induces membrane for-mation, concomitant with a reduction in thenumber of electron-lucent droplets (76). It hasthus been concluded that intracytoplasmicmembranes are induced by methane per se. In-temal membranes have, however, been observedduring the growth of Methylococcus strainNCIB 11083 on methanol (91), although thepossibility that they have been induced by am-monia, a methane analog, could not be com-pletely ruled out.The behavior ofthe stacks ofintracytoplasmic

membranes during binary fission has been de-scribed as both an arbitrary division and redis-tribution of the membranes between daughtercells (30, 46, 98) and invagination of the cyto-plasmic membrane. Monosov (98) concludedthat the processes involved in membrane genesisare connected with the accumulation of phos-pholipid micelles in growing organisms. The cen-ters of micelle fornation are closely associatedwith the cytoplasmic membrane and may beprecursors of the intracytoplasmic bilayers.Fine structure of type II methane-utiliz-

ing bacteria. Davies and Whittenbury (28) firstdescribed the two types of membrane organiza-tion in methane-oxidizing bacteria. They re-ported that membranes from methane- andmethanol-grown type II methanotrophs weresimilar in appearance to the cytoplasmic mem-brane, but were less ordered than type I mem-

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brane arrays. Membranes in type II species werealways paired and enclosed variable-sized lu-mens. Although they were usually distributedthroughout the cytoplasm, in some sections theyappeared to be mainly peripheral. Extensive pe-ripheral "stacks" of membranes have also beenobserved in various type II methanotrophs, e.g.,Methanomonas methanooxidans (Methylo-monas methanooxidans) (149) and M. tricho-sporium OB3b (181); flattened vesicles havebeen seen in the latter strain (180, 181), andlarge vesicles have been seen in Methanomonasmargaritae (106, 162).M. triVhosporium OB3b also shows marked

changes in intracytoplasmic membrane contentand internal morphology which are dependenton growth conditions (10, 13, 62, 146). Duringgrowth in shake flasks, the membrane contentappeared to be related to the amount of oxygenin the gas phase (146). Organisms grown withlittle air or oxygen showed extensive pairedmembranes (Fig. 1A), whereas correspondinglyless membranes were observed in those grownwith more air or oxygen (Fig. 1B) Growth incontinuous culture at 300C under a variety ofconditions, for example, methane, oxygen, ornitrogen limitation or at maximum growth rate,yielded organisms lacking parallel paired mem-branes and containing only membrane-boundvesicles (Fig. 1C) Interestingly, the appearanceof membranes in shake flask-grown organismscorrelated with the particulate location ofMMO(146) (see Energy Metabolism). Organisms withextensive membrane arrays were obtained re-cently by growth in continuous culture at lowcell density with excess nitrate and low methaneand oxygen partial pressures (D. Scott, unpub-lished data).M. trichosporium OB3b, grown on methanol,

also undergoes morphological changes withvarying growth conditions (10). Cells grown inbatch culture display many electron-lucent ves-icles throughout the early logarithmic and mid-logarithmic phase. Towards stationary phase,vesicles no longer prevail and irregular mem-brane arrays are seen. The vesicular morphologyis retained during continuous culture at 300C,but at 220C well-ordered paired membranes arepresent.There are reports of surface structures similar

to those of some type I species in two Methylo-cystis strains. Spiral, tubular appendages per-pendicular to and distributed over the outersurface have been observed in Methylocystisechinoides (sp nov.) (160) and Methylocystis sp.strain IC493S/5 (55).Three facultative methanotrophs have been

examined for intracytoplasmic membranes aftergrowth on different substrates. M. organo-

philum exhibits typical type II membranes onlywhen grown on methane, not when methanol orglucose are carbon sources (123). Similarly, bothMethylobacterium ethanolicum and Methylo-bacterium hypolimneticum have membranesduring methane growth, but not when hetero-trophically grown (94).Effect of growth conditions on intracy-

toplasmic membranes. It is now clear that theinternal morphology of methanotrophs is de-pendent upon growth conditions. As previouslymentioned, M. organophilum contains intracy-toplasmic membranes only during growth onmethane (123), and the membrane content isdependent on dissolved oxygen tension. At verylow dissolved oxygen tension, three to four layersof paired membranes are present, whereas athigher dissolved oxygen tension, typically onlyone peripheral paired membrane is observed.Similar low dissolved oxygen tension duringmethanol, glucose, or succinate growth does notinduce membranes. Synthesis of membranes isalso encouraged by low temperature and lowgrowth rates. Nevertheless, the nature of regu-lation of membrane synthesis remains unclear.In several methanotrophs it appears that meth-ane per se does not induce membrane formation,for methanol-grown organisms under certainconditions possess arrays of intracytoplasmicmembranes (10, 28, 30, 91). It is conceivable thatunder these latter circumstances, membranesare synthesized in response to a methane analog,ammonia or methanol (10, 91).Recent studies indicate that such membrane

arrays are not static assemblies, but are in adynamic state of flux, and changes in the arraysoccur in response to environmental changes (10,76, 106, 146, 162). Degradation of intracyto-plasmic membranes occurs in stationary-phasebatch cultures, yielding distended membrane lu-mens. Paired membranes are seldom present inorganisms grown in continuous culture under avariety of conditions. The predominant mor-phology is represented by either distended mem-brane stacks or vesicles in type I or type IImethanotrophs, respectively. These structuresprobably arise from lipid accumulation in theorganism during unbalanced growth. Over-qtili-zation of carbon source occurs in many meth-anotrophs, with the concomitant accumulationof intracellular storage polymers (90). Thus, inthe first instance, morphology may be deter-mined by the relative levels of carbon (methaneor methanol) and nitrogen sources. The subse-quent extent of membrane synthesis may thenbe controlled by the dissolved oxygen tension(123, 146), which may reflect the specific locali-zation of electron transfer proteins (146).Phospholipid and fatty acid compositions

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A

C.

C

IFIG. 1. Effect ofgrowth conditions on internal morphology ofM. trichosporium OB3b (146): transmission

electron micrographs ofthin sections ofM. trichosporium OB3bprepared aftergrowth ofthe bacterium underdifferent conditions, i.e., (A) oxygen-limited shake flask culture, (B) shake flask culture with high oxygenlevels, (C) chemostat cultivation (specific growth rate, 0.02/h; cell density, 0.2 glliter). Bar = 0.2 um.

of methanotrophs. It has been suggested thatthe fatty acid composition of methane-oxidizingbacteria is probably correlated with the struc-ture of the membrane system (46). Type I meth-anotrophs, as exemplified by M. capsulatus, ex-

hibit a typical gram-negative bacterial phospho-lipid pattern (95), the major component is phos-phatidylethanolamine with, in addition, phos-phatidylglycerol, phosphatidylcholine, and car-diolipin. A characteristic of type I methano-

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trophs is that the predominant esterified fattyacid has a chain length of 16, both the saturated(C160) and the monounsaturated (C16:1) acidsbeing found (1, 95, 150). Makula (95) describedthe predominant positional isomers of themonounsaturated acid as A9, A10, and A1".

Descriptions of the phospholipid patterns intype II methanotrophs are at variance, whichmight be ascribed to differences in methods ofcultivation (95, 182). For M. trichosporium thepredominant phospholipid was reported as phos-phatidylglycerol (182) or the methyl esters ofphosphatidylethanolamine (95), small amountsof cardiolipin and lysophosphatidylglycerolbeing found in the latter case. One striking fea-ture is the similarity of the ratio of acidic phos-pholipids to phosphatidylethanolamine and itsderivatives (about 1:4) in all strains. The sameratio was observed in both whole organisms andmembrane preparations, indicating that the in-ternal membrane composition is not unusual(182). The major esterified fatty acid of type IImethanotrophs is C18:1 (95, 149, 182), and thepredominant positional isomer was identified asA10 (95).

It has been suggested that the similarity infatty acid composition of phospholipids mightbe advantageous in facilitating the synthesis ofvarious phosphatides via a common precursor,and this might be especially important to orga-nisms with complex intracytoplasmic membranesystems (182). However, the biochemical signif-icance of this singularity of composition remainsto be elucidated.A comparative study of the phospholipid com-

positions of the facultative methanotroph M.organophilum grown under different conditionswas performed in an attempt to identify lipidsspecifically associated with intracytoplasmicmembranes (123). The amount of total lipidcorrelated with the amount of membrane mate-rial observed in thin sections, and the amount ofneutral lipids in methane-grown cells was twicethat found in methanol-grown cells. No qualita-tive difference was observed; however, phospha-tidylcholine, phosphatidylethanolamine, andmethyl esters of phosphatidylethanolaminewere present under all growth conditions.Growth on methane resulted in a decrease in theratio ofphosphatidylcholine to phosphatidyleth-anolamine. Several sterols, differing from thoseidentified in M. capsulatus (11), were found,most notably in organisms grown on methane atlow dissolved oxygen tension, conditions favor-ing membrane formation. Sterols are believed toplay an important structural role in membraneformation, but in this study the sterols were notconfined to the intracytoplasmic membrane ma-

terial. This fact, together with the low levelspresent, contraindicates a structural role inmethanotroph membranes.Exposure and cyst formation by meth-

anotrophs. Three varieties of resting stage, ex-ospores and two types of cysts, were originallyidentified in methanotrophs (187).Exposure formation has been described in de-

tail in strains of M. trichosporium (135, 136,187). Spore formation is initiated during theearly stationary phase. The onset is character-ized by the appearance at the polar tip, distalfrom the holdfast, of a second capsule. Thedevelopment of this second capsule may be mon-itored by fluorescent antibody staining tech-niques (135) by determining the increase in theintensity of fluorescence relative to the stage ofsporulation. Antigens that are different fromvegetative cell antigens are thus present on thesurfaces of the exospores and exospore regionsof vegetative cells. Both immature and maturestages of spore development have now beenidentified (136). Cup- and crescent-shaped im-mature exospores are found, indicating that aninvaginated sphere is formed at one end of thevegetative mother cell at the onset of sporula-tion. The normal spherical exospores representa mature stage. The fine structures of the differ-ent types of spores are essentially similar. Thethick, electron-dense exospore wall is sur-rounded by the cell wall and its associated cap-sule. Extensive intracytoplasmic membraneshave been found along the interior periphery ofthe exospore wall (136).

Spores are heat and desiccation resistant, sur-viving at least up to 18 months in the dried statein the absence of methane, and have no detect-able respiratory activity (187). Germination oc-curs after aerobic exposure to methane. Thethick exospore wall disappears, and a germ tubedevelops, which lacks the extensive fibrillar cap-sule and into which the intracytoplasmic mem-brane extends. Freshly formed spores germinatewithin 2 to 3 days, whereas mature spores (7days to 18 months old) take 7 to 15 days.

"Lipid cyst" formation has been described foranother type II species, Methylocystis parvus,which, on entering stationary phase, forms largelipid inclusions, mainly poly-,8-hydroxybutyrate(PHB), and loses its internal membrane system(187).Type I methanotrophs, in general, form Azo-

tobacter-like cysts that are resistant to desicca-tion, but not heat. Encystment appears to bepromoted by conditions of low oxygen tension.Strains of Methylomonas and Methylococcusspecies form immature cysts, with structuresintermediate between vegetative cells and ma-

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ture cysts, that are not desiccation resistant butthat will survive the absence of methane forlonger periods than will vegetative cells (187).Ultrastructure of methane-utilizing

yeasts. The ultrastructure of two yeast strainscapable of growth on methane was studied byWolf and Hanson (194). Both S. roseus strain Yand R. glutinis strainY when grown on methanecontained microbodies which were sites of cata-lase activity. Their rare occurrence in glucose-,ethanol-, or acetate-grown cells suggests thatmicrobodies play a role in methane metabolismin these strains. Similar organelles were alsofound in R. glutinis C7 cells grown on hexadec-ane (194).

CARBON METABOLISMObligate methanotrophs that use the serine

pathway to incorporate carbon at the oxidationlevels of formaldehyde and carbon dioxide intocell constituents have been isolated less fre-quently than those that employ the RMP path-way. This may reflect the greater efficiency ofthe latter cycle (159) in respect to growth rateson methane alone, although it does not neces-sarily reflect the abundance of either type oforganism in the environment. The possibilitythat type II species with the serine pathway andan intact tricarboxylic acid (TCA) cycle maygrow better on mixtures of C, substrates andmore complex carbon sources is discussed below.

The Serine PathwayDetails of the serine pathway have been elu-

cidated in facultative methylotrophs and havebeen authoritatively reviewed (20, 128, 195). Thecarbon assimilation pathways in type II meth-anotrophs, classified according to morphologicalcharacteristics, were first studied by Quayle andco-workers, who demonstrated that such bacte-ria possess high activities of the soluble enzymehydroxypyruvate reductase (87, 88). High activ-ities of this key enzyme have subsequently beentaken to indicate the pathway's operation in avariety oftype II methanotrophs, but little efforthas been devoted to confirming the details ofthe pathway in most of these species. The fol-lowing are species for which there is evidence foroperation of the pathway: M. trichosporiumstrains PG and OB3b (88), Methylosinus spor-ium (88), M. parvus OBBP (88), M. methanoox-idans (88), Methylovibrio soehngenii strains Aand B (58), strains 1 and NIG 3.1 (56), Methy-losinus sp. strains, CRL15 and CRL16 (119),Methylocystis sp. strain CRL18 (119), strainM102 (99) M. trichosporium subsp. methanoli-cum (171), M. parvus subsp. fuscus (171), M.

organophilum (122), isolate R6 (118), M. ethan-olicum (94), and M. hypolimneticum.The original studies of Lawrence et al. (87) on

methane-grown M. methanooxidans describedearly labeling of C4 carboxylic acids, serine, gly-cine and "unknown compounds" when the cellswere incorporating [14C]methanol at 300C. Theunknown compounds were tentatively identifiedas folate derivatives. At lower temperatures, theearly label resided predominantly in serine andglycine, and the labeling pattern suggested thatthe assimilation pathway was similar to thatdescribed for Pseudomonas strain AM1 (86).Str0m et al. (159) could not detect the key

enzymes of the RMP pathway in M. trichospo-rium. Other enzymes associated with pentosesugar rearrangement had low activities com-pared with those in type I methane utilizers.

All facultative methane-utilizing bacteria pos-sess high levels of hydroxypyruvate reductaseand are assumed to be serine pathway species(26, 117, 128). Formaldehyde, a highly reactivemolecule generated by oxidation of methane viamethanol, enters the pathway by reacting chem-ically with tetrahydrofolate to produce N5,N10-methylenetetrahydrofolate (Fig. 2). The first en-zymatic step, catalyzed by serine transhydroxy-methylase (serine hydroxymethyltransferase,EC 2.1.2.1), involves the condensation of themethylene tetrahydrofolate with glycine to formserine. This enzyme is present in most bacteria,being involved in the synthesis of serine andpurines, but in methylotrophic bacteria usingthe serine pathway, it assumes a more centralrole in metabolism.O'Connor and Hanson (102) purified two ser-

ine transhydroxymethylase activities from thefacultative methane utilizer M. organophilumXX. One activity predominated during growthon methanol, and the other predominated duringgrowth on succinate. These two isoenzymes werepurified, and their properties differed substan-tially. The methanol-induced enzyme had twicethe molecular weight of the succinate-inducedenzyme, possessed different electrophoretic andsedimentation properties, and, unlike the otherisoenzyme, was not stimulated by the metal ionsMg2e and Zn2+. Both activities were competi-tively inhibited by glycine, but only the metha-nol-induced enzyme was stimulated by glyoxyl-ate. This result is not surprising in view of thefact that glyoxylate is the precursor of glycinevia the action of serine-glyoxylate aminotrans-ferase.There is little infornation concerning regula-

tion of the channeling of formaldehyde intoeither oxidative or assimilative metabolism(132). Harder and Attwood (51) proposed that

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the reactivity of free formaldehyde with tetra-hydrofolate is such that it is only when all thecoenzyme is derivatized that formaldehyde isavailable for oxidative energy generation. Con-trol would hence be effected through the relativelevels of reduced nicotinamide adenine di-nucleotide (NADH), adenosine 5'-triphosphate(ATP), and tetrahydrofolate. Quayle (132)pointed out, however, that a tetrahydrofolate-mediated mechanism can participate in the oxi-dation of formaldehyde to formate and bypasssuch control.

In facultative methanotrophs, the followingkey serine pathway enzymes are inducible bygrowth on methanol: serine-glyoxylate amino-transferase, glycerate kinase, and glyoxylate-stimulated serine transhydroxymethylase (102).These activities are not repressed during growthon methanol ifsuccinate is added to the medium.Half of the 2-phosphoglycerate generated by

formaldehyde fixation is converted to the netproduct of the pathway, 3-phosphoglycerate.The remainder is converted to phosphoenolpyr-uvate, the acceptor molecule for the second im-portant carbon fixation step in this pathway(Fig. 2). Phosphoenolpyruvate is carboxylatedto form oxaloacetate, which is then reduced tomalate (85). Carbon dioxide is derived mainlyfrom the oxidation of reduced C, substrates. Ithas been calculated that approximately 50% ofthe carbon derived from methanol is assimilatedas carbon dioxide by the facultative methylo-troph Pseudomonas AM1 (84). The originalshort-term labeling experiments implicated thecarboxylation of phosphoenolpyruvate, whichwas found to be irreversible (85). Str0m et al.(159) predicted a higher carbon dioxide require-ment for serine pathway organisms than forRMP pathway organisms, and the fact that car-boxylase activities in the former are an order ofmagnitude higher than those in the latter hasreinforced this view (92, 171). However, the ex-act nature of the carboxylase systems of theserine pathway methane utilizers is unclear.Phosphoenolpyruvate carboxylase plays a key

role in methylotrophic metabolism. The studiesof Newaz and Hersh (100) suggested that thisenzyme was at the crossroads between carbon

FIG. 2. Serine pathway. a, Serine transhydroxy-methylase; b, serine-glyoxylate aminotransferase; c,

hydroxypyruvate reductase; d, glycerate kinase; e,enolase; f, phosphoenolpyruvate carboxylase; g, mal-ate dehydrogenase; h, malate thiokinase; i, malyl-CoA lyase; j, isocitrate lyase. ICL+, Isocitrate lyasepathway; ICL- possible alternatives to ICL+. FAD,Flavin adenine dinucleotide; FADH2, reduced FAD;ADP, adenosine 5'-diphosphate.

assimilation and energy generation. The phos-phoenolpyruvate carboxylases of methylo-trophic strains examined to date are predomi-nantly of the class 3 type, subject neither toactivation nor to inhibition. Studies of methyl-amine-grown Pseudomonas strain MA (ATCC23319) (100) indicated, however, that the car-boxylase is activated by NADH and, in thisorganism at least, thus becomes a focal point forregulation of metabolism.

In contrast, O'Connor (101) described two iso-enzymes of phosphoenolpyruvate carboxylase inM. organophilum. The isozyme of methanol-grown cells is insensitive to both acetyl coen-zyme A (acetyl-CoA) and NADH, but succinate-grown organisms possess an acetyl-CoA-stimu-lated activity. The two activities fractionate atdifferent ammonium sulfate concentrations, in-dicating two different forms of the enzyme.Loginova and Trotsenko (92) found high ac-

tivities of phosphoenolpyruvate carboxylase intwo strains of M. trichosporium. Activity wasgreater in the presence of Mg2e than in thepresence of Mn2+. A similar activity has beenidentified in a type II methanotroph, strainM102 (99). It had previously been suggested thatcarbon dioxide fixation by this strain dependedupon the presence of methane to furnish intra-cellular acceptor molecules (108). Radiolabelingstudies in both whole organisms and extracts ofthis strain have led to the postulation of theinduction of several carboxylating systems,namely, phosphoenolpyruvate carboxylase,NADH (-phosphate) [NAD(P) H]-dependent"malic enzyme," phosphoenolpyruvate carboxy-kinase, phosphoenolpyruvate carboxytransphos-phorylase, and pyruvate carboxylase (99).There is only one other report of the presence

of phosphoenolpyruvate carboxykinase in amethylotroph (82), and other workers havefound the complete absence of other carboxylat-ing systems in serine pathway strains (92, 171,172). Naguib (99) also described variable loca-tions of carboxylating enzymes according to thephysiological condition of the organism, a situ-ation reminiscent of that reported for the meth-ane monooxygenase in M. trichosporium OB3band discussed in Comparison of methane mon-ooxygenases from different species (14, 146).However, Naguib (99) ascribed this variation tostripping of the activities from the membranesduring cell breakage, implying that they arepredominantly membrane bound in vivo. Hecontends that the operative carboxylating sys-tems depend on both the physiological statusand the mode of metabolic control in these or-ganisms. On the basis of the published data, itappears to be difficult to assess these enzyme

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activities in methylotrophs, and therefore thispostulate awaits confirmation.The various carboxylating activities found in

serine pathway organisms can be summarized asfollows:

(i) "malic enzyme"pyruvate + NAD(P)H + H+ + C02

z± malate + oxidized NADP (NADP+)

(ii) pyruvate carboxylase

pyruvate + C02 + ATP a± oxaloacetate+ adenosine 5'-diphosphate

+ inorganic phosphate

(iii) phosphoenolpyruvate carboxylase

phosphoenolpyruvate + C02* oxaloacetate + inorganic phosphate

(iv) phosphoenolpyruvate carboxykinase

phosphoenolpyruvate + C02+ adenosine 5'-diphosphate w± oxaloacetate

+ ATP

(v) phosphoenolpyruvate carboxytransphos-phorylase

phosphoenolpyruvate + C02+ inorganic phosphate w± oxaloacetate

+ pyrophosphate

In the present context it is relevant to examinethe role of pyruvate in the metabolism of type IImethanotrophs. Quayle (129) presented a gen-eralized scheme for the serine pathway whichincluded pyruvate carboxylase. The reports ofpyruvate-carboxylating systems are, as de-scribed above, at variance, and Trotsenko andShishkina (172) concluded that known pathwaysfor using pyruvate for gluconeogenesis are notphysiologically functional in M. trichosporiumstrains and Methylocystis minimus. Further-more, they suggested that the formation of ace-tyl-CoA from pyruvate is of minor importancein these species, since the activity of the E1component of the pyruvate dehydrogenase com-plex is very low. In contrast, early reports sug-gested the pyruvate dehydrogenase activity intype II methane utilizers was easily detected incell-free extracts (27). A definitive comparativesurvey would help to clarify the situation. Thefacultative methylotroph Hyphomicrobiumstrain X possesses limited ability to utilize py-ruvate (52), lacking most enzymes associatedwith pyruvate metabolism. Mutants of Pseu-domonas AM1 lacking the E2 component ofpyruvate dehydrogenase have been described(12). Because of the loss of this enzyme, the

mutants had the properties of restricted facul-tative methylotrophy, similar to the hyphomi-crobia.Malate dehydrogenase has been reported in

both obligate and facultative methanotrophs(117, 122). Reported specific activities are in theranges 108 to 480 and 14 to 1,410 nmol/min permg of protein in obligate and facultative meth-ylotrophs, respectively.The generation and cleavage of malyl-CoA is

another important step in the serine pathway(Fig. 2). Extracts of methane-grown M. tricho-sporium OB3b contain no detectable ATP- andCoA-dependent malate lyase, but do catalyzeMg2e-dependent cleavage of CoA (144). Thissuggests that malate thiokinase, the enzyme re-sponsible for malyl-CoA generation, might belimiting in the first case, and hence any malyl-CoA generated would be rapidly hydrolyzed bythe lyase. Alternatively, an acyl-CoA transferasemight be involved. Malyl-CoA has been detectedat high activities in methanol-grown M. organ-ophilum (101).The action of malyl-CoA lyase regenerates

one acceptor molecule of glyoxylate and onemolecule of acetyl-CoA. The problem encoun-tered by all serine pathway organisms is theregeneration of a second glyoxylate acceptormolecule from acetyl-CoA. Three biochemicalsolutions have been proposed, but only two canbe applied to type II methanotrophs, for theylack the key enzyme involved in the first solution(below), namely, isocitrate lyase (117, 171, 172).

(i) The isocitrate lyase pathway, reviewedelsewhere (20, 128, 195), uses enzymes of theTCA cycle in conjunction with isocitrate lyaseto cleave isocitrate to succinate and glyoxylate.

(ii) Studies of mutants of the facultativemethylotroph Pseudomonas AM1 (2) suggestedthat in this organism, at least, enzymes may bepresent that cause acetyl-CoA to be directlyoxidized to glyoxylate, with glycolate as a pos-sible intermediate (Fig. 2). Bellion (6) eliminatedany other known biochemical routes by whichthis might occur by failing to demonstrate anyof the key enzymes of either the glycerate (tar-tronic semialdehyde) pathway or the /B-hydrox-yaspartate pathway in Pseudomonas strainPAR, a non-isocitrate lyase serine pathway or-ganism.

(iii) There is little direct biochemical evi-dence to support the direct oxidation proposal(ii), and work by Kortstee (83) was based on thesupposition that acetyl-CoA might be oxidizedsubsequent to its condensation with an unknowncompound which was then cleaved to yield gly-oxylate. It was not possible to demonstrate thedirect oxidation of acetate or acetyl-CoA in ex-

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tracts of the facultative methylotroph Pseudom-onas strain 80 or the presence of glycolate duringconversion of acetyl-CoA to glyoxylate. Kortsteediscovered that only one of a large number ofcompounds would condense with acetyl-CoAand produce glyoxylate as an end product whenincubated with cell-free extracts. This com-pound was 2-oxoglutarate, and its addition toextracts effected an almost complete conversionof acetyl-CoA to glyoxylate.

Labeling studies demonstrated that the twocarbon atoms of glyoxylate were derived fromacetyl-CoA. Incubation of extracts with bothhomocitrate and homoisocitrate resulted in theformation of products whose 2,4-dinitrophenyl-hydrazones were indistinguishable from authen-tic glyoxylate and glycerate derivatives. On thebasis of these experimental data, Kortstee pro-posed the homoisocitrate pathway in which ace-tyl-CoA condensed with 2-oxoglutarate and,with subsequent rearrangement, produced ho-moisocitrate (Fig. 2). Cleavage of this C7 mole-cule regenerated one molecule of glyoxylate andone of glutarate. The mechanism ofregenerationof 2-oxoglutarate from glutarate is tentativelyproposed to parallel the oxidation of succinateto oxaloacetate in the TCA cycle. At the time ofwriting, however, this pathway has not beendescribed in any serine pathway methanotroph,and it awaits unambiguous confirmation.

The Ribulose Monophosphate (Quayle)Cycle and Other Aspects of Carbon

Assimilation by Type I MethanotrophsThe biochemistry of formaldehyde assimila-

tion by type I methylotrophs (M. capsulatus[Texas] [89], M. capsulatus [Bath] [89], Meth-ylococcus ucrainicus [96], Methylococcus ther-mophilus [96], Methylococcus albus [96], Meth-ylococcus minimus [96], M. mobilis [57], M.methanica [89], Methylomonas rubrum [96],Methylomonas agile [88], Methylomonas rosa-ceus [88], Methylomonas sp. [30], Methylobac-ter capsulatus [88], Methylobacter vineclandii[88], and Methylobacter bovis [96]) has been thesubject of much more intense study that that oftheir serine pathway counterparts. Since theearly 1960s, workers, notably at Sheffield andWarwick, United Kingdom, have meticulouslydocumented the details of the RMP pathway offormaldehyde assimilation (Fig. 3), and this areahas been adequately reviewed elsewhere (20,128, 133, 195). The details of the cycle differsomewhat between species, and the three var-iants are shown in Fig. 3. It was suggested thatthis pathway may have been the progenitor ofthe closely related ribulose diphosphate cycle(133). Quayle (132) recently discussed aspects of

regulation in the RMP pathway.For some years it was thought that the clas-

sification of methylotrophs as type I or type IIbased upon morphological criteria rigorouslycorresponded to their modes of carbon assimi-lation (128). In every methanotroph examined,the dual presence of key enzymes of both assim-ilation pathways could not be demonstrated(88). Recently, however, it has become increas-ingly clear that some methanotrophs which as-similate carbon via the RMP pathway containsignificant activities of enzymes involved inother assimilation pathways. This has been mostpersuasively elucidated in the case of M. cap-sulatus (Bath).The low activities of hydroxypuruvate reduc-

tase (159) and malyl-CoA lyase (144) originallyfound in both M. capsulatus (Texas) and M.methanica were attributed to involvement inprotein and prophyrin synthesis. Significantlyhigher incorporation of label from formalde-hyde, as compared with that from methane ormethanol, into serine and glycine in M. capsu-latus (Texas) suggested that its total carbonassimilation might not be due entirely to theRMP pathway (40). Entry of formaldehyde intocellular carbon via purine synthesis would notaccount for the observed variation in labelingpattern; similar results were obtained with for-mate. At the time, assimilation via CO2 wasthought to be negligible (40).

In the light of quantitative differences ingrowth yields of the type I strains Methylo-monas albus and M. capsulatus (Bath), whichpresumably possessed the same single carbonassimilation pathway, the key enzymes of bothassimilation pathways of both strains were reex-amined (186). M. albus, with the greater growthyield, contained only enzymes of the RMP path-way, whereas M. capsulatus (Bath) possessed,in addition, hydroxypyruvate reductase. Hy-droxypyruvate reductase activity was detectedat 450C, but not at 300C, and a similar hightemperature optimum for this enzyme in othertype I methanotrophs was noted by Malashenko(96), who also demonstrated that his isolatescontained serine-glyoxylate aminotransferase.Pulse-labeling studies in continuous cultures ofM. capsulatus (Bath), using [14C]methanol as acarbon source, demonstrated the incorporationof label into sugar phosphates of the RMP path-way at both 30 and 450C, but label was found toreside also in serine and glycine at 450C. Labelwas also incorporated from formate at 450C,formate acting as a source of carbon dioxide(184).Among the 12 type I isolates examined by

Trotsenko (171), only 1 was found to contain

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solely the RMP pathway. Low activities of hy-droxypyruvate reductase and serine-glyoxylateaminotransferase were found in the others. Sup-plementing the growth medium with formatecaused a substantial increase in the levels ofhydroxypyruvate reductase. Formate was thusconsidered to be an inducer for the serine path-way in these isolates.Malashenko (96) and Colby et al. (20) referred

to the caution required to assess the roles ofcertain enzymes of the serine pathway that weredetected in these organisms and stressed thatthe enzymes may not be indicative of the com-plete cycle itself, but may merely be involved ina serine exchange reaction sequence. The evi-dence in the case of M. capsulatus (Bath) ap-pears to be rather more substantial in this re-gard, although the exact contribution made bythis minor pathway has not been elucidated.However, Colby et al. (20) suggested that it maybe "significant under certain environmental con-ditions" and, indeed, "of crucial importance tosurvival."

In addition to the two carbon assimilationpathways discussed above, the presence of thekey enzymes of the Calvin (ribulose diphos-phate) cycle, ribulose diphosphate carboxylaseand phosphoribulokinase, was demonstrated incell-free extracts of M. capsulatus (Bath) (164,165). The rate of C02 fixation by intact orga-nisms was low, however, and contributed onlyabout 2.5% (wt/wt) of the total cell carbon.Fixation depended upon the presence of meth-ane, indicating an energy requirement for incor-poration (165). Radiotracer studies demon-strated early incorporation of label into 3-phos-

FIG. 3. RMP pathway (adapted from Colby et al.

[20]). Ru5P, Ribulose-5-phosphate; Hu6P, D - erythro-L-glycero-3-hexulose-6-phosphate; F6P, fructose-6-phosphate; FDP, fructose-1,6-diphosphate; G3P,glyceraldehyde 3-phosphate; DHA, dihydroxyace-tone; DHAP, DHA-phosphate; E4P, erythrose 4-phosphate; Xu5P, xylulose-5-phosphate; S7P, sedo-heptulose-7-phosphate; SDP, sedoheptulose-1,7-di-phosphate; R5P, ribose-5-phosphate; G6P, glucose-6-phosphate; 6PG, 6-phosphogluconate; PYR, pyru-

vate. 1, 3-Hexulose phosphate synthase; 2, phospho-3-hexuloisomerase; 3, 6-phosphofructokinase; 4, fruc-tosediphosphate aldolase; 5, transketolase; 6, tran-saldolase; 7, ribulosephosphate epimerase; 8, ribulo-sephosphate isomerase; 9, sedoheptulose diphospha-tase; 10, fructose diphosphatase; 11, triosephosphateisomerase; 12, glucosephosphate isomerase; 13, glu-cose-6-phosphate dehydrogenase; 14, 6-phosphoglu-conate dehydratase +phospho-2-keto-3-deoxyglucon-ate aldolase; 15, transketolase + triokinase. (Repro-duced with permission from the Annual Review ofMicrobiology.)

phoglycerate and also aspartate, citrate, andmalate, the latter three arising from carboxyla-tion of C3 metabolites. Ribulose diphosphatecarboxylase thus functions in vivo in this meth-ane utilizer, generating labeling patterns similarto those observed for autotrophic bacteria grownheterotrophically. Nevertheless, despite thepresence of a complete Calvin cycle, M. capsu-latus (Bath) could not be grown autotrophicallyon C02.

Ribulose diphosphate carboxylase from thisorganism requires a divalent cation for activity,has an alkaline pH optimum, is inhibited by 6-phosphogluconate, and possesses an oxygenaseactivity (165). This secondary activity of theenzyme generates phosphoglycolate, which maybe subsequently cleaved by a specific phospho-glycolate phosphatase, recently discovered inextracts of M. capsulatus (Bath) (166). Themetabolic fate of the glycolate may possiblyinvolve incorporation via glyoxylate, using theserine pathway, thus providing a role for thispathway in a type I strain, which incorporateslabel from ['4C]glycolate into serine and glycine(166). The presence of ribulose diphosphate car-boxylase was demonstrated in only two strainsof M. capsulatus (Bath).

Fixation of CO2 by type I methanotrophs canoccur by a variety of different mechanisms (92,171). Certain type I methanotrophic strains(Methylobacter chrococcum 90, M. methanica12, M. bovis 90) do not possess pyruvate carbox-ylase, and activities ofphosphoenolpyruvate car-boxylase, found only with Mn2+ ions in this case,are an order of magnitude lower than thosefound in serine pathway organisms. Phospho-enolpyruvate carboxykinase and "malic en-zyme" activities have also been detected. Thevariation in activities of carboxylases betweentype I methanotrophs is thought to reflect dif-ferences in the contribution made to the overallcarbon balance by the serine pathway (171). Forinstance, carboxylases are present in M. bovis90, which possesses key enzymes of the serinepathway which are not found in M. methanica12, possessing solely the RMP pathway. Thus,one might place methanotrophs in groups basedon CO2 assimilation which directly correlateswith their primary assimilation pathways: serine-* RMP + serine -- RMP. No account is takenhere of the contribution made by the recentlydiscovered presence of the Calvin cycle in somemethanotrophs.

The Dissimilatory RibuloseMonophosphate Pathway

Gluconate 6-phosphate dehydrogenase ispresent in methane-utilizing bacteria that pos-

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sess the assimilatory RMP pathway with theEntner-Doudoroff variant of glyceraldehyde 3-phosphate production (20, 27, 131, 159). Thepresence of this enzyme permits cyclic oxidationof formaldehyde to C02 by the RMP pathwaywithout the intermediacy of formate. The path-way fiumishes NADPH, which is not otherwisegenerated by the oxidation of methane to C02(195), although an NADP+-linked formaldehydedehydrogenase was recently described in M.capsulatus (Bath) (155). The occurrence of thiscycle is often accompanied by low or undetect-able activities of formaldehyde dehydrogenaseand formate dehydrogenase (131). The operationof this cyclic oxidative pathway could not bedetected in M. methanica, which possesses ahighly active formaldehyde dehydrogenase(173).

Intracellular and Extraceflular PolymersProduced by Methane-Utilizing Bacteria

Early studies revealed sudanophilic granules,identified as PHB, deposited in large quantities(25% of the dry weight of 4-day-old cultures) inM. methanica (53, 81). Many of the methane-utilizing strains identified by Whittenbury et al.(188) were packed with lipid inclusions, mainlyPHB. In addition, M. parvus forms large PHBinclusions on entering stationary phase (187).Two strains of the methane-using vibrio M.soehngenii (58) and cysts of thermophilic meth-anotrophs (96) were found to possess variousamounts of PHB. Lipid inclusions identified asPHB have also been noted in the type II meth-anotroph M. trichosporium OB3b (10, 167, 182)at levels from 2.4% of total dry weight underunspecified growth conditions (87) to 30% inmethane excess continuous culture (10). An en-zyme involved in PHB mobilization, f)-hydrox-ybutyrate dehydrogenase, was found in extractsof methylotrophs, and acetone accumulationconcomitant with PHB degradation was dem-onstrated (167). Acetone production, probablycatalyzed by acetoacetate decarboxylase, oc-curred during the cooxidation of ethane by thisstrain. It is unlikely that the production wasrelated to acetate formation, since acetone alsoaccumulates during other cooxidative reactions,such as epoxidation, catalyzed by this strain (D.J. Best, unpublished data). The precise reasonsfor this mobilization of PHB reserves and theseemingly wasteful excretion of acetone underthese conditions are not clear. Degradation ofPHB to acetoacetic acid would provide reducingequivalents via the action of the oxidized nico-tinamide adenine dinucleotide (NAD+)-linkedBl-hydroxybutyrate dehydrogenase. Subsequent

metabolism of acetoacetic acid might prove en-ergetically expensive under conditions where theorganism is challenged with an assimilable sub-strate, and thus acetone may be excreted tomaintain NADH levels. Clearly, a more exten-sive survey of PHB metabolism is called for inorder to clarify its role as a storage polymer inmethanotrophs and the regulation of its synthe-sis and subsequent motilization.Accumulation and excretion of polysaccha-

rides has been noted in a number of methano-trophs (38, 73, 90, 151). Methylococcus NCIB11083, grown in continuous culture on methaneunder ammonia limitation, accumulates abranched polysaccharide, composed almost en-tirely of glucose residues, with a-i -* 4 and a-i-- 6 linkages and with an average molecularweight of 2 x 105 to 4.5 x 105. Accumulation,which reaches 32.5% of the dry weight at lowdilution rates in continuous culture, is inverselyproportional to the dilution rate. Polyglucose israpidly metabolized at 420C in the absence ofCH4 without any change in culture viability.This catabolism is accompanied by the uptakeof 35S, indicative of protein synthesis, and theyield of organism has been estimated to be 0.26g/g of polyglucose metabolized. Chemical char-acterization of the polymer showed that it pos-sessed an absorption spectrum similar to that ofamylopectin when complexed with iodine, butthe extent of attack by a- and 8l-amylases sug-gested it was similar in chain branch length toglycogen (90). Accumulation of this polymer,which performs an energy storage function, isnot confined to conditions of low growth rateunder nitrogen limitation, but may also occurwhen a rapidly growing culture is limited bysome other medium constituent. Linton andCripps (90) suggested that the polymer accu-mulates due to "over-utilization" of methane todeal with the problem of nonregulated genera-tion of excess carbon and reducing equivalents.Similarly, the type II methanotroph M. tricho-sporium OB3b accumulates PHB in the earlylogarithmic phase of growth on methanol (10).Whether such over-utilization of carbon sourcewill result in excretion or accumulation of stor-age materials seems to depend upon the partic-ular species.

Excretion ofpolysaccharide by a type II meth-anotroph, N. parvus OBBP (67), is surprising inview of the fact that type II species possess fewenzymes of carbohydrate metabolism (27, 159,171). After growth on methanol, this strain con-tains 62% polysaccharide, consisting of the mon-omers D-glucose (82%) and L-rhamnose (14%)together with minor quantities of L-glucose, D-rhamnose, and D-galactose. Further investiga-

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tion of such polysaccharide synthesis in type IImethanotrophs is of vital importance with re-gard to establishing the overall carbon balanceof these strains. A recent report on polysaccha-ride production in a putative M. methanicastrain described differences in polymer structuredependent on whether the enrichment culturewas methane or methanol grown; the polymercontained glucose, galactose, mannose, fucose,and rhamnose in the proportion 6:2:1:2:1 or 6:4:0:0, respectively (73). This led to the tantalizingsuggestion that the isolate was a symbiotic as-sociation of a methane utilizer and a methanolutilizer and that, when grown on methane, themethane utilizer deoxygenated the galactose andmannose residues of the polysaccharide pro-duced by the methanol utilizer, as some form ofoxygen conservation mechanism. The situation,however, is far from clear. Methanotrophs arealso known to excrete a variety of colored pig-ments, largely uncharacterized, but presumed tobe carotenoids.

Intermediary Metabolism, theTricarboxylic Acid Cycle, and Utilization

of Ancillary Carbon SourcesSerine pathway methanotrophs do not possess

some of the enzymes of carbohydrate metabo-lism, i.e., glucose-6-phosphate dehydrogenase orgluconate 6-phosphate dehydrogenase (27, 159,171). This is also true for methane-grown facul-tative methane utilizers. Glucose-6-phosphatedehydrogenase is induced only when the orga-nisms are grown on glucose (117, 122), and theactivity is either NAD or NADP linked. In con-trast, type I methanotrophs possess hexokinase(171), NADP-linked glucose-6-phosphate dehy-drogenase, and gluconate 6-phosphate dehydro-genase (27, 159, 171). Nevertheless, despite earlyreports of a stimulation of growth by glucose inM. methanica (39), most workers have con-cluded that type I methanotrophs do not assim-ilate more than trace amounts of glucose (40).The enzymatic activities associated with pyru-vate metabolism in these strains were discussedin previous sections.One of the most significant biochemical differ-

ences between type I and type II methanotrophsis that the TCA cycle is incomplete in the for-mer, but complete in the latter. The crucialenzymatic lesion is the absence of 2-oxoglutaratedehydrogenase (27, 171). All other enzymaticactivities associated with the cycle have beendemonstrated, albeit at relatively low levels. Iso-citrate dehydrogenase in M. capsulatus (Texas)is strictly NAD dependent (27).Complete TCA cycles have been reported in

facultative methanotrophs (26, 94, 117, 122).Specific activities of the enzymes of the cyclewere severalfold higher in succinate-grown thanin methane-grown organisms (117). Two TCAcycle enzymes, citrate synthase and 2-oxoglutar-ate dehydrogenase, were repressed in methanol-grown M. organophilum, suggesting that theTCA cycle is not as important to this speciesduring growth on C1 compounds as duringgrowth on succinate (103).The classical experiments to further elucidate

the nature and role of the TCA cycle in meth-anotrophs involved measurements of the incor-poration of radioactive acetate into cellular con-stituents. In the representative type I strain, M.capsulatus (Texas), both 2-oxoglutarate dehy-drogenase and aconitate hydratase are unde-tectable and the endogenous rate of acetate up-take is stimulated by methane, methanol, orformate (114, 115). Label is incorporated mainlyinto lipids, but 25% resides in only four aminoacids, glutamate, proline, arginine, and leucine.The incorporation of label into glutamate indi-cates that aconitate hydratase is, in fact, present.In contrast, similar experiments with the type IImethane utilizer M. methanooxidans demon-strated not only that the addition of acetate tothe medium enhanced the growth yield but alsothat both acetate carbons were incorporated intoamino acids of the glutamate and aspartate fam-ilies, as well as those derived from pyruvate.Radioactive CO2 was also released, which is en-tirely consistent with a functional TCA cycle invivo in a type II methanotroph (177).

Oxidation of C1 compounds and primary al-cohols supports the assimilation of acetate bycell suspensions of M. methanica (Texas) andM. trichosporium Pa, with similar patterns andrates of incorporation in both strains (113). As-similation of acetate coupled with primary al-cohol oxidation suggests that this oxidation fur-nishes the energy necessary for acetate uptake.The role of the TCA cycle in methanotrophs

is not fully elucidated or understood, but it wassuggested that type II organisms, with their highdemand for NAD(P)H for MMO and the serinepathway, could not exist without a fully func-tional TCA cycle (184). Lack of detailed knowl-edge in this area may be responsible for theconclusion that the TCA cycle is not particularlyimportant in methylotrophic metabolism (132),which is probably true of type I species, but thepresence of a catabolic TCA cycle in type IIorganisms requires an explanation.Many specialist bacteria lack a complete TCA

cycle (148). The absence of 2-oxoglutarate de-hydrogenase confers a purely biosynthetic roleon the other enzymes of the cycle rather than

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the dual roles of biosynthesis and catabolism. Ingeneral, the mode of regulation of citrate syn-thase is a good indication of the presence orabsence of a complete cycle. NADH inhibits theenzyme when the cycle is primarily catabolic,whereas 2-oxoglutarate will inhibit the enzymein strains that use the incomplete cycle in apurely biosynthetic capacity. Although this cor-ollary appears to hold for specialist strains, suchas phototrophs and lithotrophs, it does not holdfor methanotrophs examined to date. The citratesynthases of both M. trichosporium and M. al-bus are insensitive to NADH and 2-oxoglutarate(22). The absence of feedback control is consist-ent with a catabolic function of the TCA cyclein these species. Inhibition of citrate synthaseby high levels ofATP occurs, but is regarded asphysiologically unimportant (22). Further stud-ies of TCA cycle regulation are required to clar-ify its role in methanotrophs.A second approach to the evaluation of the

cycle in specialist strains, as suggested by Smithand Hoare (148), now seems especially applica-ble to type II methanotrophs, where compari-sons can be made between versatile species iden-tical to specialist strains in all respects otherthan the capacity for growth on organic com-pounds. The identification and rigorous charac-terization offacultative type II methane utilizersnow makes possible such an approach to theproblem.A third approach also asks important ques-

tions concerning the role of the TCA cycle, withspecial reference to type II methanotrophs witha complete cycle. There are many reports ofbroad-specificity oxidoreductases present inmethanotrophic bacteria, and these are dis-cussed in Energy Metabolism. Of particular in-terest is the oxidation of alkanes to alkanoicacids, which may be f8-oxidized to acetate, andthe implications that such activity raises con-cerning the utilization of ancillary carbonsources and the nature of obligate methanotro-phy itself. The precise contribution to the overallcarbon assimilation by methanotrophs still re-mains to be elucidated, but it has been pointedout elsewhere that "co-oxidation is not someaberrant and useless activity but can be directlybeneficial to methanotrophs provided methaneis also available" (184). The simplest example isthe oxidation of ethane to acetate, which mightprovide the organism with reducing power, ATP,and acetate, that may be used as a carbon (andenergy) source (via the TCA cycle) (184). Thisconcept of metabolically useful co-oxidized sub-strates was expanded upon by Higgins and co-workers (60, 61, 64), who observed the oxidationand subsequent degradation of radioactive hex-

adecane to radioactive C02 and certain phen-ylalkanes by M. trichosporium OB3b. Such ox-idative processes would most probably yield ace-tate, which could be assimilated under condi-tions where the primary growth substrate, meth-ane, is limiting. Certainly the role of acetate asa central metabolite in such a type II strainshould not be doubted, for it stands at a meta-bolic crossroads between the serine pathway,the TCA cycle, and carbon and energy storagein the form of PHB, a polymer that can only bemobilized via acetyl-CoA (Fig. 4). Thus, it hasbeen argued that the ability to use co-oxidizablesubstrates via conventional acetate metabolismconfers an advantage on methanotrophic orga-nisms that possess a complete TCA cycle (60-61, 64). Further evidence in support of this hy-pothesis is found from observations that, undercarbon limited conditions, hexadecane will sig-nificantly increase the cell yield ofM. trichospo-rium OB3b (F. Taylor, personal communica-tion). It was suggested that in their naturalenvironments, methylotrophs are unlikely tohave access to a continuous supply of their spe-cific energy source; their survival will then de-pend on an ability to obtain metabolically usefulenergy from energy sources by means other thanspecialized oxidation reactions (148). It is likelythat methane is usually present in the naturalhabitats of methanotrophs, but it may often begrowth limiting, thereby offering an advantageto species that can co-utilize ancillary carbonand energy sources.Such approaches as these do, however, once

again beg the question of the biochemical basisof obligate methanotrophy. Although it wasdemonstrated that the mutational loss of 2-ox-oglutarate dehydrogenase does confer obligacyupon an otherwise facultative methylotroph(163), the absence of this enzyme does not offera rational explanation for methylotrophic fastid-iousness. Many strains with the same enzymiclesion are capable of heterotrophic growth (148).It certainly does not offer any explanation forthe obligacy of type II methane utilizers, and, todate, only hypotheses are available to explainthis dietary particularity. These include (i) sub-strate toxicity; (ii) substrate impermeability oran energetic requirement for substrate uptake asdemonstrated for acetate; (iii) coarse metaboliccontrol by enzyme induction or repression, en-zymes being strictly regulated at levels that al-low efficient growth on one substrate only; (iv)unusual or inadequately coupled electron trans-port mechanisms and energy transduction; (v)specific biosynthetic deficiencies; (vi) absolutedependence on a metabolite available only dur-ing growth on the characteristic carbon source;

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(vii) failure to oxidize NADH, although this isnot tenable in the case of methanotrophs, sincethey possess NADH oxidase activities (27); and(viii) multiple enzymic lesions (172). For a muchmore detailed discussion of this problem, thereader is once again referred to the excellentreview of Smith and Hoare (148).

Methane-Utilizing YeastsMethanol-utilizing yeasts incorporate formal-

dehyde via a xylulose-5-phosphate cycle, par-tially reminiscent of the RMP pathway (132).However, due to the difficulty in obtaining suf-ficient biomass, very little is known of the path-ways of carbon incorporation of the recentlyisolated methane-utilizing yeasts. Large, yeast-like microorganisms were observed in methano-trophic enrichment cultures from shield lakes(143), but the isolation of pure methane-utilizingyeast strains was first reported by Wolf andHanson (192). Every isolate oxidized methane toC02, but such intermediates as methanol, form-aldehyde, and formate would not supportgrowth. The greatest cell yields were found incultures supplied with an atmosphere containing10% (vol/vol) C02. In this case no radioactivelabel from methane was incorporated into TCA-precipitable material. If the C02 level was low-ered, however, label was found in cellular com-ponents, suggesting that these yeasts assimilatecarbon as C02 and not as one of the intermedi-ates ofmethane oxidation. These isolated strainshave a wide nutritional capacity. They use meth-ylamine, butane, octane, some higher hydrocar-bons, ethanol and other alcohols, acetate, andsome sugars. Further elucidation of the path-ways involved in these eucaryotic organisms iskeenly awaited.

ENERGY METABOLISMThe pathway of methane oxidation to carbon

dioxide via methanol, formaldehyde, and for-mate in methanotrophic bacteria has been wellestablished for many years. Although consider-able attention has been focused on the enzymesof this oxidative sequence, until recently littlewas known of the mechanisms involved. Signif-icant advances have now been made towardselucidating the enzymic mechanisms ofmethaneand methanol oxidation at the molecular level.

Methane MonooxygenaseEarly reports of cell-free methane- and

NADH-dependent oxygen consumption in par-ticulate preparations of M. capsulatus (Texas)(137, 138) and M. methanica (43) were followedby descriptions of the properties of cell-freeMMO from M. methanica (19), M. trichospo-

rium OB3b (170) and M. capsulatus (Bath) (16).Methane monooxygenase activity was purifiedand studied in detail from the latter two strainsand, at present, is best characterized in M. cap-sulatus (Bath).A three-componentMMO fromM. capsulatus

(Bath) was purified by Dalton and co-workers(24) by conventional chromatographic tech-niques (Fig. 5). Component A has a molecularweight of about 220,000, with two nonheme ironatoms and two acid-labile sulfur atoms per mol-ecule. This oxygen-labile protein is purified un-der anaerobic conditions and is thought to bethe hydroxylase component ofthe enzyme. Com-ponent B, a colorless protein with a molecularweight of about 15,000, is stable in the presenceof phenylmethylsulfonyl fluoride. Although itsrole in the oxygenase reaction is uncertain, it isessential for activity. Component C has a molec-ular weight of 44,000, and each molecule con-tains one molecule of flavin adenine dinucleo-tide, two atoms of nonheme iron, and two atomsof acid-labile sulfur. The ability to be reducedby NADH (17) and the ability to catalyze thereduction by NADH of other acceptors (18)indicate that component C is the reductase com-ponent of the monooxygenase complex. Com-ponent A is not directly reduced by NADH, butgives an electron paramagnetic resonance signalon reduction with sodium dithionite. This signalis enhanced in the presence of a substrate, eth-ylene, suggesting that component A binds sub-strate when in a reduced form (20). Thus, com-ponent C transfers electrons from NADH tocomponent A, which then binds the substrate,allowing oxidation (Fig. 6) (24).The methane-oxidizing enzyme from M. trich-

osporium OB3b was purified (169), althoughrecently this purification procedure has notyielded active enzyme in our laboratory. Thethree-component enzyme comprised a solubleCO-binding cytochrome c, a copper-containingprotein (protein 1), and a small protein, withmolecular weights of 13,000, 47,000, and 9,400,respectively (169). The soluble CO-binding cy-tochrome c has an oxidase function (170), andalthough evidence suggests that it binds MMOsubstrates, such as methane and ethylene, (49)as well as oxygen, it was also thought to be thereductase component (169). The roles of theother two components remain unclear, althoughprotein 1 may be involved in oxygen binding(169).More recent work by Dalton's group and in

our own laboratory revealed MMO activities incell-free extracts of M. trichosporium OB3b ofa substantially different nature from those pub-lished earlier for both the cell-free enzyme and

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574 HIGGINS ET AL.

o X

_ -

-W

'S..,.a

,o la0

ofS-

4c

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a

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METHANE-OXIDIZING MICROORGANISMS

0.5M NaCl

FRACTION C

5'AMPSepharose 4B

Unboundmaterialdiscarded

Sephadex G-100

Component C

CRUDE EXTRACT

DEAE-Cel lulose

FRACTION A

pH fractionation

Ultrogel ACA34

DEAE-Se harose

Sephadex G200

I

Coei?onent A

0.2M NaCl

31(NH4)2S04 fractionation

Sephadex G-100

DEAE - Sephadex

Sephadex G-50

Component B

FIG. 5. Purification scheme for the three components ofMMO from M. capsulatus (Bath) (from Dalton[24]).

the purified enzyme (62, 146, 151). Stirling andDalton (157) resolved the MMO of this speciesinto two fractions by ion-exchange chromatog-raphy. Although reduced activity was observedwhen the two fractions were reconstituted, fullactivity was restored to fraction 1 by addition ofpurified components B and C from M. capsula-tus (Bath). No complementation of activity wasobserved with fraction 2 of M. trichosporiumOB3b extracts, but these results suggest thatfraction 1 contains the hydroxylase componentof MMO and that there may be some homoge-neity between the two enzyme systems (157).

Discrepancies in the nature of the M. tricho-sporium MMOs found by different workers maybe due partly to different growth and prepara-tion procedures (see Comparison of methanemonooxygenases from different species).Comparison of methane monooxygen-

ases from different species. The ability ofMMO to oxygenate a wide range of substratesin vitro was demonstrated with cell-free extractsof M. capsulatus (Bath) (21), M. trichosporiumOB3b, and M. methanica (153). Substrates in-cluded n-alkanes, n-alkenes, ethers, and ali-

FIG. 4. Central position of acetyl-CoA in the me-

tabolism of type II methanotrophs. HICL, Homoiso-citrate lyase; PEP, phosphoenol-pyruvate; ADP,adenosine 5-diphosphate; AL, alanine; VAL, valine;LEU, leucine; ASP, aspartic acid; LYS, lysine; MET,methionine; THR, threonine; ISO, isoleucine; ARG,arginine; GLN, glutamine; HIS, histidine; PRO, pro-

line.

cyclic, aromatic, and heterocycic hydrocarbonsfor M. capsulatus (Bath) and M. trichosporiumOB3b. M. methanica had a more limited sub-strate specificity: n-hexane and n-heptane andall cyclic compounds were not oxygenated. Thefacultative methane-oxidizing bacteria appear tohave a similar varied substrate range, althoughits extent is unclear, since oxidation of aromatic,alicycic, and other complex hydrocarbons havenot been reported. M. organophilum CRL26, forinstance, oxidizes C2 to C4 n-alkanes and n-al-kenes at rates similar to those of obligate meth-anotrophs (68, 69), and both M. ethanolicumand M. hypolimneticum yield propene oxide andethanol from propylene and ethane, respec-tively (94). Although some reactions catalyzedby MMO are analogous to the oxidation of itsnatural substrate (e.g., formation of primary al-cohols from n-alkanes and 1-phenylalkane sidechains) many of the substrates attacked do notresemble methane. Hydroxylation of aromatic,heterocyclic, and multi-ring compounds may,therefore, indicate a free radical reaction (74) orsome other reactive oxygen mechanism.

It is particularly interesting that broad sub-strate oxygenation also occurs in whole organismsuspensions of M. trichosporium OB3b (60, 62,64) and M. capsulatus (Bath) (24), althoughsome products of MMO activity are subject tofurther attack by other nonspecific oxidoreduc-tases (see below).MMOs from most sources appear to be obli-

gatorily NAD(P)H linked. The enzymes fromM. methanica (19, 43), M. capsulatus (Texas)

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reduced acceptor B ?

NADH+H+ C.FADFeS25 A.Fe2 + CH °2

NAD+ Reduced A.Fe3 CH OH+H 0C.FADFe S2 C3O+20

DCPIPFerricyanideCytochrome c

FIG. 6. Pathway of electron transfer between the MMO components in the oxidation of methane. Theprosthetic groups of components A and C are indicated as Fe and Flavin adenine dinucleotide (FAD)-Fe2S2.The role ofB is uncertain. DCPIP, dichlorophenol indophenol. (From Dalton [24].)

(138), M. capsulatus (Bath) (16), M. trichospo-rium OB3b (62, 146, 153, 157), and M. organo-philum CRL26 (118) all require reduced pyridinenucleotides for in vitro activity. Tonge et al.(170) originally reported cell-free MMO from M.trichosporium OB3b to utilize ascorbate andmethanol, as well as NAD(P)H as an electrondonor. Later, ascorbate-mediated cell-freeMMOwas also described in M. capsulatus grown at450C (125). It is unclear whether NAD(P)H isthe direct electron donor in vivo in some meth-ane-oxidizing bacteria, for there is evidence thatintermediate electron transfer protein(s) may beinvolved in extracts of M. trichosporium OB3bgrown under oxygen-limiting conditions (146)(see below).A study of the location of MMO activity in

cell-free extracts of M. trichosporium OB3b re-vealed an interesting correlation with the en-zyme's sensitivity to inhibitors (146). Duringoxygen-limiting growth conditions in shakeflasks, particulate MMO activity sensitive tochelating agents, thiol reagents, amytal, andKCN is present. Similar inhibitors were reportedfor particulate MMOs of M. capsulatus (Texas)(137), M. methanica (19, 44), Methylosinus sp.CRL15 (118), and, originally, for M. trichospo-rium OB3b (169). Recently, it was found thatgrowth of M. trichosporium OB3b in a chemo-stat under low methane and oxygen partial pres-sures with excess dissolved nitrate yields orga-nisms that are packed with intracytoplasmicmembranes and that show only particulateMMO activity (146).

In extracts prepared from M. trichosporiumOB3b grown under all conditions other thanoxygen limitation, MMO was present only in thesoluble fraction. Activity was insensitive to che-lating and thiol reagents and amobarbital, asshown earlier for soluble activities in this strain(157) and M. capsulatus (Bath) (16, 154).

It is possible, therefore, that particulateMMOs retain a close association with electrontransfer proteins and that NAD(P)H may be anindirect electron donor in vivo. Soluble MMO,however, shows no sensitivity to electron trans-port inhibitors or chelating agents, suggestingthat NAD(P)H interacts directly with the en-zyme. Further evidence for the latter case isprovided by the properties of component C ofthe soluble M. capsulatus (Bath) enzyme (17,18) (see above).Hou and co-workers have reported the ab-

sence of MMO activity in whole organism sus-pensions of methanol-grown obligate and facul-tative methanotrophs (119, 168) and suggest,therefore, that MMO is only induced duringgrowth on methane. Although this certainly ap-pears to be the case for the facultative methaneutilizers M. ethanolicum and M. hypolimneti-cum (94), MMO activity was demonstrated inthe following methanol-grown obligate methan-otrophs: Methylococcus NCIB 11083 (91), M.capsulatus (Texas) (76), and M. trichosporiumOB3b (10). In these strains, at least, MMO isapparently constitutive.Mechanisms of oxidation by methane

monooxygenase. In view of the substrate spec-ificity of the MMOs studied so far, the involve-ment of free radicals appears likely. Hutchinsonet al. (74) proposed a free radical mechanism formethane oxidation involving generation of amethyl radical, stabilized by coordination to aniron atom, which reacts with molecular oxygento produce a methylperoxy-iron derivative. Ad-dition of two methyl peroxide radicals with sub-sequent loss of an oxygen molecule and furtherproton loss or addition generates methanol orformaldehyde. Direct experimental evidence forradical involvement is, however, lacking. At-tempts to detect radicals by using spin-trappingagents have proved unsuccessful (D. J. Best and

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D. Scott, unpublished data). Although the the-oretical calculations of barrier heights for meth-ane oxidation by a P450 hydroxylase favor aconcerted "attachment-rearrangement" mecha-nism involving 's oxygen atoms (barrier height- 2 kcal/mol) (127), the experimental activationenergy of methane oxidation by cell-free MMOfrom M. capsulatus at temperatures above 200Cwas determined to be about 18 kcal/mol (125).This is, however, close to the theoretical barrierheight of about 14 kcal/mol for a triplet hydrox-ylation with 3p oxygen atoms involving a two-step "abstraction-recombination" mechanismwith the formation of radical intermediates.Nuclear magnetic resonance studies of MMO

in M. capsulatus (Bath) showed the operationof an "NIH shift" mechanism during the ringhydroxylation of 1-phenylethane (25), thus:

D 02 HO OHH H HD

H H HHHC2H6 NADH+H+ NAD+ C2H6

This is analogous to the classical NIH shiftmechanisms found for aromatic hydroxylases(78, 168).

Methanol DehydrogenaseThe broad-specificity NAD(P)+-independent

methanol dehydrogenase of the type first de-scribed by Anthony and Zatman (5) is commonto all methane- and methanol-utilizing bacteriastudied so far. Methanol dehydrogenases havebeen characterized from several methane oxidiz-ers and, in general, appear to be closely similar(110-112, 176). In vitro, methanol dehydroge-nase can be coupled to phenazine (m)ethosulfatein the presence of ammonia or a primary amineas activator and will oxidize many primary al-cohols.The enzyme is thought to be coupled to the

electron transport chain at the level of cyto-chrome c in vivo. Recently, Duine et al. (35)reported the isolation of methanol dehydroge-nase from Hyphomicrobium X with a functionalcoupling to cytochrome c. Potassium ferricya-nide-oxidized cytochrome c shows a character-istic reduced spectrum on addition of methanolto the complex. Furthermore, this coupling isonly functional in anaerobically prepared ex-tracts and is irreversibly destroyed on exposureto oxygen, yielding the conventional, NH4+-ac-tivated, dye-linked dehydrogenase.A major contribution to our understanding of

the biochemistry of all methane- and methanol-utilizing bacteria is the elucidation of the struc-ture of the prosthetic group of methanol dehy-

drogenase. Evidence had accumulated that thecoenzyme is a novel, nitrogen-containing o-qui-none (29, 34, 37, 183).

Salisbury et al. (145) used X-ray crystallog-raphy to determine the structure of a purifieddegradation product of the cofactor and pro-posed the trivial name "methoxatin" for thecofactor (I in Fig. 7). Confirmation of the struc-ture came from Duine et al. (36), and theyproposed the name "pyrrolo-quinoline quinone,"since the o-quinone structure is essential foractivity. Although the prosthetic group of meth-anol dehydrogenase cannot, at present, be recon-stituted with its own apoenzyme, a functionalholoenzyme can be reconstituted with glucosedehydrogenase, indicating that this group of"quinoproteins" have similar cofactors (36).

Forrest et al. (45) have proposed a mechanismfor the oxidation of methanol involving meth-oxatin (I) (Fig. 7), which forms an addition com-pound with the amino group of a lysine residue(or with ammonia or a primary amine) at the C4position (II). 1,4-Elimination of water producesthe quinone analog (HI) in the active enzyme.1,4-Addition of primary alcohol allows a cyclicrearrangement with release of the oxidized prod-uct.

Oxidation of primary alcohols by the fac-ultative methane utilizer MethylobacteriumCRL26 is also catalyzed by an ammonium ion-requiring, phenazine (m)ethosulfate-dependentmethanol dehydrogenase (117). Similar activitywas also reported for M. organophilum XX(191); additionally, this enzyme oxidized second-ary and tertiary alcohols at low activities. Am-monium ions were not required, but were stim-ulating for newly purified enzyme, although am-monium ions were essential for activity afterstorage. The newly isolated species M. ethanol-icum also has an ammonium ion-requiring,phenazine methosulfate-linked methanol dehy-drogenase when grown on methane, but thisactivity is about sevenfold lower in extracts ofethanol-grown organisms (94). The presence ofan alternative ethanol-oxidizing enzyme was notreported, but this may be similar to the situationduring growth of two facultative methylotrophson methanol and ethanol (51). During methanolgrowth of strain PAR, only a typical methanoldehydrogenase with a molecular weight of112,000 was detected, which oxidized primaryand secondary alcohols. In ethanol-grown orga-nisms, however, this enzyme was absent and wasreplaced by an NAD-dependent alcohol dehy-drogenase with a molecular weight of 150,000.The latter enzyme was not found during growthon methanol, and although it, too, showed broadsubstrate specificity, it did not oxidize methanol(7).

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RH2COOH

-4 COOH HN

OH

HOOC

rnnm .H flZ., I NHR

II

III

NHR

NHR

OH

FIG. 7. Proposed mechanism for the role ofmethoxatin in methanol oxidation by methanol dehydrogenase(adapted from Forrest et al. [45]).

Formaldehyde Dehydrogenase

Ci-utilizing bacteria contain several differenttypes of formaldehyde dehydrogenase activity(2, 155). These can be classified into two groups:(i) NAD(P)+-linked enzymes and (ii) NAD(P)+-independent enzymes that require an artificialelectron acceptor (e.g., phenazine methosulfateor dichlorophenol indophenol) for in vitro activ-ity. Both groups encompass enzymes which arespecific for formaldehyde and nonspecific alde-hyde dehydrogenases.

Stirling and Dalton (155) purified a NAD(P)+-linked formaldehyde dehydrogenase from M.capsulatus (Bath) and subsequently reportedpreliminary evidence for an NAD+-linked en-zyme in cell-free extracts of M. trichosporium

OB3b (157). In addition, a heme-containing al-dehyde dehydrogenase was purified from M.trichosporium PG which required phenazinemethosulfate for in vitro activity (116). Thisactivity was demonstrated in cell-free extracts ofseveral type I, type II, and facultative methane-utilizing strains, although extracts of M. capsu-latus (Texas and CRL24) and Methylosinussporium 5 showed very low activity (6 to 7 nmol/min per mg of protein). Cyclic oxidation of form-aldehyde of the type discussed in Carbon Me-tabolism, which would yield two NAD(P)H mol-ecules per C-1 unit, has yet to be demonstratedin methanotrophs.The ability to derive readily available reduc-

ing equivalents in the form of NAD(P)H fromformaldehyde dehydrogenases may be impor-

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tant in influencing the yield during growth onmethane (3, 4) and the usefulness of supplemen-tary carbon sources (see Carbon Metabolism).

Formate Dehydrogenase

A typical NAD-linked formate dehydrogenasehas been found in several methylotrophs andthree methane oxidizers (80, 112, 139).

Secondary Alcohol Dehydrogenase

Phenazine methosulfate-dependent methanoldehydrogenase from methane-utilizing bacteriahas a broad substrate specificity for C1 to C10primary alcohols, but shows either little or noactivity towards secondary alcohols (see above).Recently, however, a secondary alcohol-specificdehydrogenase activity was observed in cell-freeextracts and whole organism suspensions of sev-eral types of obligate and facultative methaneand methanol utilizers (71). Both primary andsecondary alcohols are the products of short-chain n-alkane oxidation by MMO (21, 119).Hou and co-workers described the accumulationof methylketones from secondary alcohols (re-flecting secondary alcohol dehydrogenase activ-ity) (71) and from n-alkanes (reflecting MMOand secondary alcohol dehydrogenase activities)(72, 120) by intact methanotrophs. The activitycould not be demonstrated in extracts of eitherof the facultative methanotrophs M. ethanoli-cum or M. hypolimneticum, although whole or-ganisms oxidized 2-butanol (94). Although it hasbeen known for some time that n-alkanes are co-

oxidized to methylketones byM. methanica (89,93), the Exxon group showed that this is due toNAD+-dependent secondary alcohol dehydro-genase present in cell-free extracts, in additionto MMO.Secondary alcohol dehydrogenase was puri-

fied from methanol-grown Pseudomonas sp.

ATCC 21439 (70) and has a molecular weight ofabout 95,000, comprising two identical subunits.Although the dehydrogenase was specific forsecondary alcohols, it did not oxidize C7 to C10alcohols and showed maximum activity towards2-butanol. Hou et al. (70) do not ascribe a phys-iological function to secondary alcohol dehydro-genase activity in C1 metabolism, but do suggestthat this NAD+-linked activity would be advan-tageous for methanotrophic organisms, whichmay be NAD(P)H limited (4). However, the factthat the oxidation of n-alkanes via secondaryalcohols to methylketones yields no net increasein NADH or other reducing agent seems to havebeen overlooked. Although some secondary al-cohols are present in the environment, the rela-tively specific secondary alcohol dehydrogenase

shows no activity towards secondary alcoholsabove 2-hexanol, i.e., it is unlikely to attacksecondary alcohols that might be derived fromsubstances common in the environment.

Oxidation of Multicarbon CompoundsThe capacity of methane-oxidizing bacteria to

metabolize an extensive and varied range oforganic compounds has prompted considerableresearch effort and much speculation over themetabolic significance of these processes (60, 61,158) to methanotrophs. Although MMO hasbeen at the forefront of this attention and isalmost always responsible for the initial oxida-tive attack on these compounds, many transfor-mation products are the result of further oxida-tion by other oxidoreductases. Whole organismsuspensions of M. trichosporium OB3b and M.capsulatus (Bath) oxidize n-alkanes, n-alkenes,aromatic and alicyclic hydrocarbons, phenols,long-chain and alicyclic alcohols, and hetero-cyclic, multi-ring, and halogenated hydrocar-bons (24, 60). The rate of these oxidations canbe similar to the methane oxidation rate (e.g.,ethane, propane, propene) or, in the case ofmorecomplex compounds, an order ofmagnitude less.An important factor in determining the rate andextent of whole organism biotransformations isthe level of endogenous energy reserves. Orga-nisms harvested from carbon excess, nitrogen-limited cultures ofM. trichosporium OB3b havehigh levels (20 to 30%) of PHB (146) and showconcomitantly faster propene oxidation andgreater product accumulation than those orga-nisms with low levels of PHB from carbon-lim-ited, nitrogen excess growth conditions (Bestand Scott, unpublished data). A similar situationarises in M. capsulatus (Bath) (24), with accu-mulation of storage polymers in low specific-growth-rate cultures (0.03/h). This fact mayhave been overlooked in some cases and shouldbe carefully considered when comparing the invivo oxidation abilities of methane oxidizers.

Stirling and Dalton (156, 158) defined theoxidation of nongrowth substrates catalyzed byMMO and subsequent enzymes as "fortuitous,"suggesting that these activities are a conse-quence ofthe mechanistic requirement for meth-ane, methanol, and formaldehyde oxidation.Higgins and co-authors (60, 61), in contrast, ar-gued that as carbon (acetate) and energy(NADH and ATP) can be derived from oxida-tion of some environmentally important hydro-carbons (e.g., n-alkanes) a selection pressuremay have existed to retain or develop nonspe-cific enzymes, and that this metabolism shouldbe termed "supplementary" (60). (This is dis-cussed further in Evolution of Methanotrophs.)

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Energy Transduction and Growth YieldsThe electron transfer and energy transducing

systems in methane-oxidizing bacteria are notfully understood, but our current knowledge wasrecently thoroughly reviewed by Higgins (59)and Wolfe and Higgins (195). Anthony (3, 4)discussed the prediction of, growth yields inmethylotrophic bacteria and identified severalimportant considerations. First, yields for mostmethylotrophic bacteria are determined byNAD(P)H supply as well as ATP, and, in partic-ular, methane-oxidizers may be NAD(P)H lim-ited. The extent of this limitation depends pri-marily upon the mechanism of supply of thereducing source to the MMO. Bacteria that ox-idize methane, utilizing NADH as the directelectron donor to the monooxygenase, have pre-dicted yields in the range of 9.2 to 12.0 g (drywt)/mol of substrate, depending on the type ofcarbon incorporation pathway, and are NADHlimited. Alternatively, strains with a monooxy-genase system coupled to reduced methanol de-hydrogenase as the source of reducing equiva-lents, via CO-binding cytochrome c (65), havepredicted yields as high as 17.4 g (dry wt)/molofsubstrate and are predominantlyATP limited.If NADH is generated by reverse electron trans-port, yields would be between those for NADH-coupled and methanol dehydrogenase-coupledsystems if ATP was required, and the same asthe latter system if ATP was not required.For organisms with an NADH-coupled mono-

oxygenase system, the yield may be further af-fected by the type of reduced cofactor used inthe formaldehyde dehydrogenase reaction.NAD(P)+-linked formaldehyde dehydrogenasestrains will have slightly higher yields than thosewhich use an NAD(P)+-independent enzyme.The experimental determination of growthyields on methane is difficult, but apparentlyreliable figures have been reported that are sub-stantially greater than the theoretical predic-tions (3, 4). In addition, molar yields on methaneand methanol are commonly found to be similar.This has been taken to indicate that MMO maybe unusual among monooxygenases in being en-ergetically conservative (65), but there are alter-native explanations. Reconciliation of theoryand practice awaits a more complete under-standing of methanotrophic energetics.

Methanotrophic Yeasts and AnaerobicMethane Oxidation

The likelihood that methanotrophic yeastsobtain energy by oxidizing methane to C02 andassimilate carbon from C02 (192) is discussed inCarbon Metabolism. This apparently inefficientmechanism may be the reason for the very slow

generation times (2 to 7 days). As yet, nothing isknown of the enzymology of methane oxidationin these yeasts.

It was for many years widely assumed thatmethane is biologically inert in anaerobic envi-ronments, but recent geochemical evidence hasindicated that methane does disappear in anaer-obic environments (50). Zehnder and Brock(198), however, showed low levels of methaneoxidation during anaerobic growth of somemethanogenic bacteria. The ratio of methaneoxidized to methane formed was between 0.001and 0.3%, but preliminary evidence suggests thatthe route of methane oxidation is not simplyreversal of the methane formation process. An-aerobic oxidation of methane also occurred innonmethanogenic bacteria isolated from the sed-iment surface of Lake Mendota, Wis. (109). Sul-fate, acetate, and methane were all required foranaerobic growth of the isolates, and it appearsthat acetate is directly assimilated and methaneis oxidized to CO2 without incorporation. Again,nothing is known of the nature of the enzymesinvolved.

NITROGEN METABOLISM

Methane-utilizing microorganisms play a rolein the nitrogen cycle not only by virtue of thenecessary incorporation of nitrogen into theirbiomass but also by transformation of inorganicnitrogen compounds and dinitrogen (195) and,perhaps, of complex, nitrogen-containing organ-ics (21, 64). The ability of many methanotrophsto fix atmospheric nitrogen in addition to usingnitrate or ammonia as a nitrogen source hasbeen long established (128). Some informationon pathways of ammonia assimilation by obli-gate methanotrophs is now available. Daltonand Whittenbury (26) detected glutamate de-hydrogenase and glutamine synthetase activitiesin M. capsulatus (Bath). The activities of thesetwo enzymes varied with different nitrogensources, glutamate dehydrogenase having thehighest activity in ammonia-grown organisms,whereas glutamine synthetase was the more ac-tive enzyme after growth under nitrogen-fixingconditions. Drozd et al. (33) found that an ap-parently different strain of Methylococcus didnot possess glutamate dehydrogenase but wasconstitutive for enzymes of the glutamine syn-thetase-glutamine 2-oxoglutarate aminotrans-ferase system (15).

In a more thorough study of seven obligatemethanotrophs representing both type I andtype H, Shishkina and Trotsenko (147) foundthat the type I strains tested, except M. capsu-latus (Foster and Davis strain), had glutamate(or alanine) dehydrogenase and glutamine syn-

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thetase activities. Type II methane utilizers hadenzyme activities typical of the glutamine syn-thetase-glutamine 2-oxoglutarate aminotrans-ferase system. M. capsulatus (Foster and Davis),a type I methanotroph, not only had alanine(but not glutamate) dehydrogenase and gluta-mine synthetase activities but in addition hadglutamate synthase activity. Thus, this strainmay possess two pathways for ammonia assimi-lation. In this study, enzyme activities variedaccording to the nitrogen source (NH4', N03,or N2).

Nitrogen fixation by Methylosinus strain 41(32) is accompanied by hydrogenase activitywhich can support acetylene reduction (31). Ni-trite is known to be reduced by methanotrophs(188), as it must be, prior to assmilation asammoma.Other transformations of nitrogen compounds

reflect the activity of the broad-substrate-speci-ficity MMO. Oxidation of ammonia to nitrite bymethane utilizers was observed by Hutton andZobell (75) and by Whittenbury and his col-leagues (188). Initiation of this process is veryprobably due to MMO activity, for ammonia isa competitive inhibitor of methane oxidation (asmethane is of ammonia oxidation in Nitrosom-onas [107, 161]. Dalton (23) detected hydroxyl-amine as an oxidation product of ammonia byM. capsulatus (Bath), as did Higgins et al. (64)working with M. trichosporium OB3b.

Recently a specific hydroxylamine:cyto-chrome-c oxidoreductase, distinct from metha-nol dehydrogenase, has been described in M.ethylococcus thermophilus (152). Under someconditions, ammonia oxidation may well serveas a supplementary energy source (97), a conceptdiscussed in Energy Metabolism. A number ofgroups, in monitoring the wide substrate speci-ficity of MMO (Energy Metabolism), haveshown that pyridine is a substrate, being oxi-dized to pyridine-N-oxide (21, 64). When suchnitrogen compounds are available in the naturalhabitats of methanotrophs, it seems reasonableto suppose that they will be transformed bythese organisms, which may thus play severalroles in the nitrogen cycle (60).

GENETICS OF METHANOTROPHSProgress in our understanding of the genetics

of methane utilizers has been hampered by thedifficulty ofobtaining phenotypic mutants. Suchmutants arose with a frequency of about lo-3 inM. capsulatus and M. albus treated with achemical mutagen, such as N-methyl-N-nitro-N-nitrosoguanidine, but generally reverted athigh frequency (54, 190). Other chemical muta-gens, such as alkylmethane-sulfonates, and ir-

radiation with ultraviolet and y sources wereunsuccessful in mutating obligate methano-trophs, although both N-methyl-N-nitro-N-ni-trosoguanidine and ultraviolet mutagenesis gaverise to auxotrophic mutants of the facultativemethane utilizer M. organophilum (105). In thisstudy, exogenous wild-type DNA transformedthe mutants to prototrophy at a frequency of 5x 10-3, competence occurring at late logarithmicphase.

It seems likely that the inability to isolatemutants of obligate methanotrophs is not due toa fundamental difference in the genetics of thesemutants compared with enterobacteria, for ex-ample, but is perhaps due to the use of inappro-priate techniques. For example, at least somemethanotrophs are probably not mutable byultraviolet due to lack of error-prone "SOS"DNA repair mechanisms (189). Methanotrophsresemble other microorganisms in the possessionof lytic bacteriophages (174, 175) and extrachro-mosomal DNA, at least in facultative methano-trophs, whose MMOs may be plasmid encoded(50, 196). A major aim of genetic studies is, ofcourse, to understand the organization and reg-ulation of the processes of metabolism, and it isnot surprising that studies of Cl-oxidative andCl-assimilative pathways are furthest advancedin facultative methylotrophs. In the facultativemethanotroph M. organophilum, O'Connor(101) and O'Connor and Hanson (103, 104)showed by DNA transfornation experimentsthat many of the key enzymes of C, oxidationand aimilation (with the notable exception offormate dehydrogenase) are coordinately regu-lated and appear to be located in a single operon.The use of promiscuous plasmids to both gen-

erate mutants by transposon mutagenesis (9)and map mutants by chromosome mobilization(66, 124) in methanotrophs offers the best hopefor rapid elucidation of genetic systems in thesespecies. Transfer of the antibiotic resistancemarkers carried by such plasmids to facultativemethylotrophs has been reported by severalgroups (8, 47, 79, 178). In addition, Warner et al.(179) reported transfer of the plasmid R68.45 tothe methanotroph M. trichosporium OB3b.

EVOLUTION OF METHANOTROPHSThe possibility that the present-day aerobic

methane utilizers might represent those specieswhich evolved the capacity to oxidize and growon methane when the earth's atmosphere be-came sufficiently oxidized is set in context byQuayle and Ferenci (133). They discuss the cur-rent relationships between autotrophy andmethylotrophy, of which methanotrophy is aspecific case, and suggest evolutionary routes for

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the development of similar carbon assimilationpathways in these two groups of species.There are, for instance, considerable morpho-

logical similarities between methane utilizersand nitrifying bacteria in their intemal mem-brane arrays; the two groups initiate the oxida-tion of their respective energy sources, CH4 andNH3, by a monooxygenase, and both nitrifiersand type I methanotrophs lack a complete TCAcycle. In addition, some methanotrophs appearto be capable of partial lithotrophic metabolism,deriving energy from ammonia oxidation. Car-bon assimilation pathways are not the same,however, though Quayle and Ferenci (133) havesuggested possible evolutionary sequences bywhich the type I methanotrophic RMP pathwayof carbon assimilation might be linked with theclosely related ribulose diphosphate cycle of thenitrifying bacteria.

Shishkina and Trotsenko (147) found that M.capsulatus, used in their studies of nitrogenassimilation (see above), possessed metabolicactivities typical of both type I and type IImethanotrophs and suggested that the speciesmight represent an intermediate between thetwo types. Further evidence to this effect is citedby Taylor et al. (166), who detected the enzymesof the autotrophic Calvin cycle in M. capsulatus(Bath) as well as [1-14C]glycolate incorporationby a pathway probably involving enzymes foundin the type II serine pathway for carbon assimi-lation. Our present state of knowledge of meth-anotrophs is far from complete, and the alter-native suggestions of reclassifying M. capsulatus(Bath) as a type X methylotroph (185) or thedivision of Methylococcus into two genera (141)highlight our lack of understanding of thoserelationships within this group ofmethylotrophswhich is necessary for detailed comparison withother microbial groups.Of particular interest in the present context is

the evolution of MMO. Although there is con-siderable controversy as to the protein chemistryofMMO (132), the remarkable scope of MMOsfrom different sources to effect oxidations ofmany organic compounds (see above) is worthyof further mention. This phenomenon has beenvariously terned "cometabolism" and "fortui-tous metabolism" (156) and Quayle (130) hasproposed that the term "cometabolism" be con-fined to the oxidation of substrates, such as CO,for which reducing power must be provided bythe addition of a suitable cosubstrate. It shouldnot be overlooked, however, that the cosubstrateneed not be exogenously supplied, and reducingpower can be generated from endog)nous stor-age polymers. "Fortuitous metabolisn" wouldthen describe oxidation of such a substrate asethane. In this case, the reducing power for the

initial oxidative attack by MMO is replenishedby further oxidation of the product of the MMOreaction.

It has been suggested (60) that there mayhave been selection pressures for evolution ofbroad-specificity MMOs and other oxidoreduc-tases or for the retention of such enzymes, sincethey may confer some physiological advantageson methanotrophs. Although either case defiesconclusive proof, some possible products ofMMO activity, such as ethanol derived fromethane oxidation or alcohols derived from long-chain alkanes, can be incorporated by methan-otrophs after further oxidation by nonspecificoxidoreductases (see above and reference 60)and appear to increase cell yield (177) (Taylor,personal communication). In this context theterm "supplementary metabolism" seems moreappropriate.

It is possible to speculate on the developmentof MMO. As discussed by Quayle and Ferenci(133), oxygen appeared in the atmosphere aftera considerable period of biotic evolution. Onemight consider a "primitive" anaerobe, ferment-ing fornaldehyde, confronted with a developingaerobic environment. The ability to supplementdwindling supplies of formaldehyde by oxidizing(though perhaps also dwindling) methane andmethanol would have conferred considerable ad-vantage. A relatively inefficient MMO with poorsubstrate specificity would have been the suffi-cient outcome of such selection pressure at theoriginal event, especially since other previouslyunavailable reduced organic compounds mightalso then be oxidized, yielding perhaps energyand carbon for incorporation. Subsequent spe-cialization of the methane utilizers to their pres-ent habitats must have been accompanied byselection pressure for development of a highlyactive MMO, not subject to competitive inhibi-tion by other commonly occurring compounds,unless the ability to oxidize those other com-pounds conferred an advantage slightly out-weighing that of improved catalysis of methaneoxidation. That the latter case seems to existtoday, at least in those few MMOs so far exam-ined, suggests that there is indeed some advan-tage to maintaining a broad-substrate-specificityMMO. It would appear, however, that faculta-tive methane utilizers could be at an advantageover obligate strains in situations where carbonsources other than methane are available.Perhaps facultative methane utilizers find lit-

tle advantage in an MMO with catholic specific-ity because they can avail themselves of othercarbon sources. Whole methane-grown cells ofMethylobacterium strain R6, the facultativemethane utilizer studied by the Exxon group,can oxidize carbon monoxide, ethane, propane,

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ethylene, and propylene (117), and the two fac-ultative isolates of Lynch et al. (94) can oxidizeethylene and propylene if grown on methane.More detailed studies of the substrate specifici-ties of the MMOs of facultative methane utiliz-ers are unfortunately not available.A particularly fascinating question is whether

the isolation of facultative methanotrophs,which has led only to species similar to type IIobligate organisms, is fortuitous or truly repre-sentative. This can only be resolved by the iso-lation of more species, but it would not be sur-prising if it proved invariably to be the lattercase. The ability of type II obligate strains tometabolize non-Cl substrates makes the basis oftheir obligacy and their relationship to faculta-tive strains more difficult to define. It does,superficially, suggest a close evolutionary link,which is not, however, supported by DNA hy-bridization studies (50).

BIOTECHNOLOGICAL APPLICATIONSOF METHANOTROPHS

For many years, methanotrophs have beenconsidered for single-cell protein processes, anddevelopment in this area continues (48). Thediscovery of the extensive range of chemicaltransformations catalyzed by methanotrophshas opened up the possibility of their exploita-tion for biocatalysis, and several patents havebeen filed on this subject (60). It seems likelythat such processes will be developed within thenext few years. They may also prove useful as

sources of biopolymers.To capitalize fully on the biotransformational

capacities of these bacteria, it may well be nec-

esary to develop reliable techniques for theirgenetic manipulation. Successful developmentof such techniques would also facilitate expres-

sion of foreign DNA coding for products of in-dustrial or medical importance. There may besignificant advantages in using methanotrophsfor synthesis of foreign proteins, as they appear

to have low protease activities, as exemplifiedby long-tern stability of enzyme activities innongrowing organisms. The fact that many en-

zymes of C, metabolism are apparently consti-tutive in obligate methanotrophs suggests thatthey possess operons suitable for cloning and thesubsequent expression of foreign DNA in thesebacteria.

LITERATURE CITED1. Andreev, L. V., Y. A. Trotsenko, and V. F.

Galchenko. 1977. Fatty acid composition ofmethylotrophic bacteria, p. 12-15. In G. K.Skryabin, M. V. Ivanov, E. N. Kondratjeva, G.A. Zavarzin, Yu. A. Trotsenko, and A. I. Nes-terov (ed.), Microbial growth on Cl-com-

pounds. Abstracts of the Second InternationalSymposium on Microbial Growth on Cl-com-pounds. USSR Academy of Sciences, Push-chino, USSR.

2. Anthony, C. 1975. The biochemistry of meth-ylotrophic bacteria. Sci. Prog. (London) 62:167-206.

3. Anthony, C. The prediction of growth yields inmethylotrophs. J. Gen. Microbiol. 104:91-104.

4. Anthony, C. 1980. Methanol as substrate: theo-retical aspects, p. 35-57. In D. E. F. Harrison,I. J. Higgins, and R. Watkinson (ed.), Hydro-carbons in biotechnology. Heyden & Son, Ltd.,London.

5. Anthony, C., and L. J. Zatman. 1964. Themicrobial oxidation of methanol. 2. The meth-anol-oxidising enzyme of Pseudomonas sp.M27. Biochem. J. 92:614-621.

6. Bellion, E. 1977. Investigations into the ICL-serine pathway, p. 36-37. In G. K. Skryabin,M. V. Ivanov, E. N. Kondratjeva, G. A. Zavar-zin, Yu. A. Trotsenko, and A. I. Nesterov (ed.),Microbial growth on C,-compounds. Abstractsof the Second International Symposium onMicrobial Growth on C,-Compounds. USSRAcademy of Sciences, Pushchino, USSR.

7. Bellion, E., and G. T.-S. Wu. 1978. Alcoholdehydrogenases from a facultative methylo-trophic bacterium. J. Bacteriol. 135:251-258.

8. Berger, H., and D. Blohm. 1978. A geneticsystem for methylotrophic bacteria using theR68.45 plasmid, p. 2606. In Abstracts ofposters, 12th FEBS Meeting, Dresden.

9. Beringer, J. E., J. L. Beynon, A. V. Bu-chanan-Wollaston, and A. W. B. Johnston.1978. Transfer of the drug-resistance transpo-son Tn5 to Rhizobium. Nature (London) 276:633-634.

10. Best, D. J., and I. J. Higgins. 1981. Methane-oxidising activity and membrane morphologyin a methanol-grown obligate methanotroph,Methylosinus trichosporium OB3b. J. Gen.Microbiol. 125:73-84.

11. Bird, C. W., J. M. Lynch, S. J. Pirt, W. W.Reid, C. J. W. Brooks, and B. S. Middle-ditch. 1971. Steroids and squalene in Meth-ylococcus capsulatus grown on methane. Na-ture (London) 230:473-474.

12. Bolbot, J. A., and C. Anthony. 1980. The me-tabolism of pyruvate by a facultative meth-ylotroph Pseudomonas AML. J. Gen. Micro-biol. 120:233-244.

13. Brannan. J., and I. J. Higgins. 1978. Effect ofgrowth conditions on the intracytoplasmicmembranes of Methylosinus trichosporiumOB3b. Proc. Soc. Gen. Microbiol. 5:69.

14. Brannan, J., D. Scott, and I. J. Higgins. 1980.The effect of growth- and post-growth-condi-tions on intracytoplasmic membrane contentand location of methane mono-oxygenase ac-tivities in Methylosinus trichosporium OB3b,p. 11-13. In G. M. Tonge (ed.), Abstracts ofposters, Third International Symposium onMicrobial Growth on C, Compounds. Univer-sity of Sheffield Press, Sheffield, United King-dom.

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15. Brown, C. M., D. S. Macdonald-Brown, andJ. L. Meers. 1974. Physiological aspects ofmicrobial inorganic nitrogen metabolism. Adv.Microb. Physiol. 11:1-52.

16. Colby, J., and H. Dalton. 1976. Some propertiesof a soluble methane mono-oxygenase fromMethylococcus capsulatus strain Bath. Bio-chem. J. 157:495497.

17. Colby, J., and H. Dalton. 1978. Resolution ofthe methane mono-oxygenase of Methylococ-cus capsulatus (Bath) into three components.Purification and properties of Component C, aflavoprotein. Biochem. J. 171:461-468.

18. Colby, J., and H. Dalton. 1979. Characterisa-tion of the second prosthetic group of the fla-voenzyme NADH-acceptor reductase (Com-ponent C) of the methane mono-oxygenasefrom Methylococcus capsulatus (Bath). Bio-chem. J. 177:903-908.

19. Colby, J., H. Dalton, and R. Whittenbury.1975. An improved assay for bacterial methanemono-oxygenase: some properties of the en-zyme from Methylomonas methanica. Bio-chem. J. 151:459-462.

20. Colby, J., H. Dalton, and R. Whittenbury.1979. Biological and biochemical aspects of mi-crobial growth on C, compounds. Annu. Rev.Microbiol. 33:481-517.

21. Colby, J., D. L Stirling, and H. Dalton. 1977.The soluble methane mono-oxygenase ofMethylococcus capsulatus (Bath). Its abilityto oxygenate n-alkanes, n-alkenes, ethers, andalicyclic, aromatic and heterocyclic com-pounds. Biochem. J. 165:395-402.

22. Colby, J., and L J. Zatman. 1975. Regulationof citrate synthase activity in methylotrophsby reduced NAD+,adenine nucleotides and 2-oxoglutarate. Biochem. J. 150:141-144.

23. Dalton, H. 1977. Ammonia oxidation by themethane oxidising bacterium Methylococcuscapsulatus strain Bath. Arch. Microbiol. 114:273-279.

24. Dalton, H. 1980. Transformations by methanemono-oxygenase, p. 85-97. In D. E. F. Harri-son, I. J. Higgins and R. Watkinson (ed.), Hy-drocarbons in biotechnology. Heyden & Son,Ltd., London.

25. Dalton, H. 1981. Methane mono-oxygenasesfrom a variety of microbes, p. 1-10. In H.Dalton (ed.), Microbial growth on C, com-pounds. Proceedings ofthe Third InternationalSymposium on Microbial Growth on Cl Com-pounds. Sheffield, United Kingdom. Heyden& Son, Ltd., London.

26. Dalton, H., and R. Whittenbury. 1975. Nitro-gen metabolism in Methylococcus capsulatusstrain Bath, p. 379-388. In H. G. Schlegel, G.Gottschalk, and N. Pfennig (ed.), Symposiumon microbial production and utilisation ofgases(H2, CH4, CO), Gottingen. Akademie der Wi-ssenschaften, Gottingen.

27. Davey, J. F., R. Whittenbury, and J. F. Wilk-inson. 1972. The distribution in the methylo-bacteria of some key enzymes concerned withintermediary metabolism. Arch. Mikrobiol. 87:359-366.

28. Davies, S. L., and R. Whittenbury. 1970. Finestructure of methane and other hydrocarbonutilising bacteria. J. Gen. Microbiol. 61:227-232.

29. De Beer, R., D. Van Ormondt, M. A. Van Ast,R. Banen, J. A. Duine, and J. Frank. 1979.ESR andENDOR at 9 and 35 GHz on a powderof the enzyme methanol dehydrogenase fromHyphomicrobium X. J. Chem. Phys. 70:4491-4495.

30. De Boer, W. E., and W. Hazeu. 1972. Obser-vations on the fine structure of a methane-oxidising bacterium. Antonie van Leeuwen-hoek J. Microbiol. Serol. 38:33-47.

31. De Bont, J. A. M. 1976. Hydrogenase activity innitrogen fixing, methane-oxidising bacteria.Antonie van Leeuwenhoek J. Microbiol. Serol.42:255-259.

32. De Bont, J. A. M., and E. G. Mulder. 1974.Nitrogen fixation and co-oxidation of ethyleneby a methane-utilising bacterium. J. Gen. Mi-crobiol. 83:113-121.

33. Drozd, J. W., J. D. Linton, J. Downs, R.Stephenson, M. L. Bailey, and S. J. Wren.1977. Growth energetics in methylotrophic bac-teria, p. 91-94. In G. K. Skryabin, M. V. Ivanov,E. N. Kondratjeva, G. A. Zavarzin, Yu. A.Trotsenko, and A. I. Nesterov (ed.), Microbialgrowth on Cl-compounds. Abstracts of the Sec-ond International Symposium on MicrobialGrowth on Cl-Compounds. USSR Academy ofSciences, Pushchino, USSR.

34. Duine, J. A., and J. Frank. 1980. The pros-thetic group of methanol dehydrogenase. Pu-rification and some of its properties. Biochem.J. 187:221-226.

35. Duine, J. A., J. Frank, and L. G. De Ruiter.1979. Isolation of a methanol dehydrogenasewith a functional coupling to cytochrome c. J.Gen. Microbiol. 115:523-526.

36. Duine, J. A., J. Frank, and P. E. J. Verwiel.1980. Structure and activity of the prostheticgroup of methanol dehydrogenase. Eur. J. Bio-chem. 108:187-192.

37. Duine, J. A., J. Frank, and J. Westerling.1978. Purification and properties of methanoldehydrogenase from Hyphomicrobium X.Biochim. Biophys. Acta 524:277-287.

38. Dworkin, M., and J. W. Foster. 1956. Studieson Pseudomonas methanica (S6hngen) nov.comb. J. Bacteriol. 72:646-659.

39. Dworkin, M., and J. W. Foster. 1958. Experi-ments with some micro-organisms which utiliseethane and hydrogen. J. Bacteriol. 75:592-603.

40. Eceleston, J., and D. P. Kelley. 1973. Assimi-lation and toxicity of some exogenous C1 com-pounds, alcohols, sugars and acetate in themethane oxidising bacterium, Methylococcuscapsulatus. J. Gen. Microbiol. 75:211-221.

41. Ehhalt, D. H. 1976. The atmospheric cycle ofmethane, p. 13-36. In H. G. Schlegel, G. Gotts-chalk, and N. Pfennig (ed.), Symposium onmicrobial production and utilisation of gases(H2, CH4, CO), G6ttingen. Akademie der Wi-ssenschaften, Gottingen.

42. Enebo, L 1967. A methane-consuming green

MICROBIOL. REV.


mbr.asm

.org/D

ownloaded from



algae. Acta Chem. Scand. 21:625-32.43. Ferenci, T. 1974. Carbon monoxide-stimulated

respiration in methane-utilising bacteria.FEBS Lett. 41:94-98.

44. Ferenci, T., T. Strom, and J. R. Quayle. 1975.Oxidation of carbon monoxide and methane byPseudomonas methanica. J. Gen. Microbiol.91:79-91.

45. Forrest, H. S., S. A. Salisbury, and C. G.Kilty. 1980. A mechanism for the enzymicoxidation of methanol involving methoxatin.Biochem. Biophys. Res. Commun. 97:248-251.

46. Galachenko, V. F., and N. E. Suzina. 1977.Peculiarities of membrane apparatus organi-sation of methanotrophic bacteria, p. 12-15. InG. K. Skryabin, M. V. Ivanov, E. N. Kon-dratjeva, G. A. Zavarzin, Yu. A. Trotsenko, andA. I. Nesterov (ed.), Microbial growth on Cl-compounds. Abstracts of the Second Interna-tional Symposium on Microbial Growth on C1-Compounds. USSR Academy of Sciences,Pushchino, USSR.

47. Gautier, F., and R. Bonewald. 1980. The useof plasmid R11.62 and derivatives for genecloning in the methanol-utilising Pseudomo-nas AML. Mol. Gen. Genet. 178:375-380.

48. Hamer, G., and D. E. F. Harrison. 1980. Singlecell protein: the technology, economics andfuture potential, p. 59-73. In D. E. F. Harrison,I. J. Higgins, and R. Watkinson (ed.), Hydro-carbons in biotechnology. Heyden & Son, Ltd.,London.

49. Hammond, R. C., F. Taylor, and L. J. Higgins.1979. Studies on the mechanism of methanemono-oxygenase of Methylosinus trichospo-rium OB3b. Soc. Gen. Microbiol. Q. 6:89.

50. Hanson, R. S. 1980. Ecology and diversity ofmethylotrophic organisms. Adv. Appl. Micro-biol. 26:3-39.

51. Harder, W., and M. M. Attwood. 1978. Biology,physiology and biochemistry of Hyphomicro-bia. Adv. Microb. Physiol. 17:303-359.

52. Harder, W., A. Martin, and M. M. Attwood.1975. Studies on the physiological significance

of the lack of pyruvate dehydrogenase complexin Hyphomicrobium sp. J. Gen. Microbiol. 86:319-326.

53. Harrington, A. A., and R. E. Kallio. 1960.Oxidation of methanol and formaldehyde byPseudomonas methanica. Can. J. Microbiol.6:1-7.

54. Harwood, J. H., E. Williams, and B. W. Bain-bridge. 1972. Mutation of the methane oxidis-ing bacterium, Methylococcus capsulatus. J.Appl. Bacteriol. 35:99-108.

55. Haubold, R. 1978. Two different types of surfacestructures of methane-utilising bacteria. Z.AUg. Mikrobiol. 18:511-515.

56. Hazeu, W. 1975. Some cultural and physiologicalaspects of methane-utilising bacteria. Antonievan Leeuwenhoek J. Microbiol. Serol. 41:121-134.

57. Hazeu, W., W. M. Batenburg-van der Egte,and J. C. de Bruyn. 1980. Some characteris-tics of Methylococcus mobilis sp nov. Arch.Microbiol. 124:211-220.

58. Hazeu, W., and P. J. Steenis. 1970. Isolationand characterisation of two vibrio shapedmethane oxidising bacteria. Antonie van Leeu-wenhoek J. Microbiol. Serol. 36:67-72.

59. Higgin, I. J. 1980. Respiration in methylo-trophic bacteria, p. 187-221. In C. J. Knowles(ed.), Diversity of bacterial respiratory sys-tems, vol. 1. CRC Press Inc., Boca Raton, Fla.

60. Higgins, I. J., D. J. Best, and RK C. Hammond.1980. New findings in methane-utilising bacte-ria highlight their importance in the biosphereand their commercial potential. Nature (Lon-don) 286:561-564.

61. Higgins, L. J., D. J. Best, and R. C. Hammond.1981. Reply to Stirling and Dalton. Nature(London) 291:169-170.

62. Higgins, I. J., D. J. Best, and D. Scott. 1981.Hydrocarbon oxidation by Methylosinus trich-osporium, metabolic implications of the lack ofspecificity of methane mono-oxygenase, p. 11-20 In H. Dalton (ed.), Microbial growth on C,compounds. Proceedings of the Third Inter-national Symposium on Microbial Growth onC, Compounds, Sheffield, United Kingdom.Heyden & Son, Ltd., London.

63. Higgins, L. J., and R. C. Hammond. 1981.Transformation of C1 compounds by micro-or-ganisms. In D. T. Gibson (ed.), Biochemistryof microbial degradation. Marcel Dekker, Inc.,New York. (In press.)

64. Higgins, L J., R. C. Hammond, F. S. Sarias-lani, D. Best, M. M. Davies, S. E. Tryhorn,and F. Taylor. 1979. Biotransformation ofhydrocarbons and related compounds by wholeorganism suspension of methane-grown Meth-ylosinus trichosporium OB3b. Biochem. Bio-phys. Res. Commun. 89:671-677.

65. Higgins, I. J., and J. R. Quayle. 1970. Oxygen-ation ofmethane by methane-grown Pseudom-onas methanica and Methanomonas metha-nooxidans Biochem. J. 118:210-218.

66. Hofloway, B. W., V. Krishnapillai, and A. F.Morgan. 1979. Chromosomal genetics ofPseu-domonas. Microbiol. Rev. 43:73-102.

67. Hou, C. T., A I. Laskin, and R. N. PateL 1978.Growth and polysaccharide production byMethylocystis parvus OBBP on methanol.Appl. Environ. Microbiol. 37:800-804.

68. Hou, C. T., R. Patel, A. L Laskin, and N.Barnabe. 1979. Microbial oxidation of gaseoushydrocarbons: epoxidation of C2 to C4 n-al-kenes by methylotrophic bacteria. Appl. Envi-ron. Microbiol. 38:127-134.

69. Hou, C. T., R. N. Patel, A. I.Laskin, and N.Barnabe. 1980. Microbial oxidation of lowern-alkenes and n-alkanes by resting cell suspen-sions of various methylotrophic bacteria, andthe effect ofmethane metabolites. FEMS (Fed.Eur. Microbiol. Soc.) Microbiol. Lett. 9:267-270.

70. Hou, C. T., R N. Patel, A. L Laskin, N. Bar-nabe, and L. Marezak. 1979. Identificationand purification of a nicotinamide adenine di-nucleotide-dependent secondary alcohol de-hydrogenase from C1-utilising microbes. FEBSLett. 101:179-183.

VOL. 45, 1981


mbr.asm

.org/D

ownloaded from


586 HIGGINS ET AL.

71. Hou, C. T., R. N. Patel, A. I. Laskin, N. Bar-nabe, and I. Marczak. 1979. Microbial oxi-dation of gaseous hydrocarbons: production ofmethyl ketones from their corresponding sec-ondary alcohols by methane- and methanol-grown microbes. App. Environ. Microbiol. 38:135-142.

72. Hou, C. T., R. N. Patel, A. I. Laskin, I. Mar-czak, and N. Barnabe. 1981. Microbial oxi-dation of gaseous hydrocarbons: production ofalcohols and methyl ketones from their corre-sponding n-alkanes by methylotrophic bacte-ria. Can. J. Microbiol. 27:107-115.

73. Huq, M. N., B. J. Ralph, and P. A. D. Rickard.1978. The extracellular polysaccharide of amethylotrophic culture. Aust. J. Biol. Sci. 31:311-316.

74. Hutchinson, D. W., R. Whittenbury, and H.Dalton. 1976. A possible role of free radicals inthe oxidation of methane by Methylococcuscapsulatus. J. Theor. Biol. 58:325-335.

75. Hutton, W. E., and C. E. Zobell. 1953. Produc-tion of nitrite from ammonia by methane oxi-dizing bacteria. J. Bacteriol. 65:216-219.

76. Hyder, S. L., A. Mayers, and M. L. Cayer.1979. Membrane modulation in a methylo-trophic bacterium Methylococcus capsulatus(Texas) as a function of growth substrate. Tis-sue Cell 11:597-610.

77. Ivanov, M. V., S. S. Belyaev, and S. S. Laui-navichus. 1976. Methods of quantitative in-vestigation of microbiological production andutilisation of methane, p. 63-67. In H. G. Schle-gel, G. Gottschalk, and N. Pfennig (ed.), Sym-posium on microbial production and utilisationof gases (H2, CH4, CO), Gottingen. Akademieder Wissenschaften, Gottingen.

78. Jerina, D. M., and J. W. Daly. 1974. Areneoxides: a new aspect of drug metabolism. Sci-ence 185:573-582.

79. Jeyaseelan, K., and J. R. Guest. 1979. Transferof antibiotic resistance to facultative methylo-trophs with plasmid R68.45. FEMS (Fed. Eur.Microbiol. Soc.) Microbiol. Lett. 6:87-89.

80. Johnson, P. A., and J. R. Quayle. 1964. Micro-bial growth on C, compounds. 6. Oxidation ofmethanol, formaldehyde and formate by meth-anol-grown Pseudomonas AML. Biochem. J.93:281-289.

81. Kallo, R. E., and A. A. Harrington. 1960.Sudanophilic granules and lipids of Pseudom-onas methanica. J. Bacteriol. 80:321-324.

82. Kirikova, N. N., and A. K. Romanova. 1972.Carbon dioxide assimilation by Pseudomonassp. 2 cells which use methanol and formate.Microbiology (USSR) 41:173-177.

83. Kortatee, G. J. J. 1980. The homoisocitrate-glyoxylate cycle in pink, facultative methylo-trophs. FEMS (Fed. Eur. Microbiol. Soc.) Mi-crobiol. Lett. 8:59-65.

84. Large, P. J., D. Peel, and J. R. Quayle. 1961.Microbial growth on C, compounds. 2. Synthe-sis of cell constituents by methanol- and for-mate-grown Pseudomonas AM1 and metha-nol-grown Hyphomicrobium vulgare. Bio-chem. J. 81:470-480.

85. Large, P. J., D. Peel, and J. R. Quayle. 1962.Microbial growth on C, compounds. 3. Distri-bution of radioactivity in metabolites of meth-anol-grown Pseudomonas AM1 after incuba-tion with ['4C]methanol and ['4C]bicarbonate.Biochem. J. 82:483-488.

86. Large, P. J., and J. R. Quayle. 1963. Microbialgrowth on Cl compounds. 5. Enzyme activitiesin extracts of Pseudomonas AM1. Biochem. J.87:386-396.

87. Lawrence, A. J., M. B. Kemp, and J. R.Quayle. 1970. Synthesis of cell constituents bymethane-grown Methylococcus capsulatus andMethanomonas methanooxidans. Biochem. J.116:631-636.

88. Lawrence, A. J., and J. R. Quayle. 1970. Al-ternative carbon assimilation pathways inmethane-utilising bacteria. J. Gen. Microbiol.63:371-374.

89. Leadbetter, E. R., and J. W. Foster. 1960.Bacterial oxidation of gaseous alkanes. Arch.Mikrobiol. 35:92-104.

90. Linton, J. D., and R. E. Cripps. 1978. Theoccurrence and identification of intracellularpolyglucose storage granules in MethylococcusNCIB 11083 grown in chemostat culture onmethane. Arch. Microbiol. 117:41-48.

91. Linton, J. D., and J. Vokes. 1978. Growth ofthe methane-utilising bacterium Methylococ-cus NCIB 11083 in mineral salts medium withmethanol as the sole source of carbon. FEMS(Fed. Eur. Microbiol. Soc.) Microbiol. Lett. 4:125-128.

92. Loginova, N. V., and Y. A. Trotsenko. 1979.Pyruvate and phosphoenolpyruvate carboxy-lases in methylotrophs. Microbiology (USSR)48:202-207.

93. Lukins, H. B., and J. W. Foster. 1963. Methylketone metabolism in hydrocarbon-utilizingmycobacteria. J. Bacteriol. 85:1074-1087.

94. Lynch, M. J., A. E. Wopat, and M. L.O'Connor. 1980. Characterization of two newfacultative methanotrophs. Appl. Environ. Mi-crobiol. 40:400-407.

95. Makula, R. A. 1978. Phospholipid compositionofmethane-utilizing bacteria. J. Bacteriol. 134:771-777.

96. Malashenko, Y. R. 1976. Isolation and charac-terisation of new species (thermophilic andthermotolerant ones) of methane-utilisers, p.293-300. In H. G. Schlegel, G. Gottschalk, andN. Pfennig (ed.), Symposium on microbial pro-duction and utilisation of gases (H2, CH4, CO),Gottingen. Akademie der Wissenschaften, Got-tingen.

97. Malashenko, Y. R., L. G. Sokolov, V. A. Ro-manovskaya, and Y. B. Shkurko. 1979. Ele-ments of lithotrophic metabolism in the obli-gate methylotroph Methylococcus thernophi-lus. Microbiology (USSR) 84:468-474.

98. Monosov, E. Z. 1977. Membrane genesis inmethane oxidising bacteria, p. 25-27. In G. K.Skryabin, M. V. Ivanov, E. N. Kondratjeva, G.A. Zavarzin, Yu. A. Trotsenko, and A. I. Nes-terov (ed.), Microbial growth on Cl-com-pounds. Abstracts of the Second International

MICROBIOL. REV.


mbr.asm

.org/D

ownloaded from



Symposium on Microbial Growth on C,-Com-pounds. USSR Academy of Sciences, Push-chino, USSR.

99. Naguib, M. 1979. Alternative carboxylation re-actions in Type II methylotrophs and the lo-calisation of carboxylase activities in the intra-cytoplasmic membranes. Z. Allg. Mikrobiol.19:333-340.

100. Newaz, S. S., and L. B. Hersh. 1975. Reducednicotinamide adenine dinucleotide-activatedphosphoenolpyruvate carboxylase in Pseu-domonas MA: potential regulation betweencarbon assimilation and energy production. J.Bacteriol. 124:825-33.

101. O'Connor, M. L. 1981. Extension of the modelconcerning linkage of genes coding for C-1-re-lated functions in Methylobacterium organo-philum. Appl. Environ. Microbiol. 41:437-441.

102. O'Connor, M. L, and R. S. Hanson. 1975.Seine transhydroxymethylase isoenzymesfrom a facultative methylotroph. J. Bacteriol.124:985-996.

103. O'Connor, M. L., and R. S. Hanson. 1977.Enzyme regulation in Methylobacterium or-ganophilum. J. Gen. Microbiol. 101:327-332.

104. O'Connor, M. L, and R. S. Hanson. 1978.Linkage relationship between mutants ofMethylobacterium organophilum impaired intheir ability to grow on one-carbon compounds.J. Gen. Microbiol. 104:105-111.

105. O'Connor, M. L., A. E. Wopat, and R. S.Hanson. 1977. Genetic transformation inMethylobacterium organophilum. J. Gen. Mi-crobiol. 98:265-272.

106. Ohtomo, T., H. lizuka, and K. Takeda. 1977.Effect of copper sulphate on the morphologictransition of a strain of Methanomonas mar-

garitae. Eur. J. Appl. Microbiol. 4:267-272.107. O'Neill, J. G., and J. F. Wilkinson. 1977. Oxi-

dation of ammonia by methane-oxidising bac-teria and the effects of ammonia on methaneoxidation. J. Gen. Microbiol. 100:407-412.

108. Overbeck, J. 1976. Some remarks on the ecologyof CO2 metabolism of heterotrophic and meth-ylotrophic bacteria, p. 263-266. In H. G. Schle-gel, G. Gottschalk, and N. Pfennig (ed.), Sym-posium on microbial production and utilisationof gases (H2, CH4, CO), Gottingen. Akademieder Wissenschaften, Gottingen.

109. Panganiban, A. T., T. E. Patt, W. Hart, andR. S. Hanson. 1979. Oxidation of methane inthe absence of oxygen in lake water samples.Appl. Environ. Microbiol. 37:303-309.

110. Patel, R. N., H. R. Bose, W. J. Mandy, and D.S. Hoare. 1972. Physiological studies of meth-ane- and methanol-oxidizing bacteria: compar-

ison of a primary alcohol dehydrogenase fromMethylococcus capsulatus (Texas strain) andPseudomonas species M27. J. Bacteriol. 110:570-577.

111. Patel, R. N., and A. Felix. 1976. Microbialoxidation of methane and methanol: crystalli-zation and properties of methanol dehydro-genase from Methylosinus sporium. J. Bacte-riol. 128:413-424.

112. Patel, R. N., and D. S. Hoare. 1971. Physiolog-

ical studies of methane and methanol-oxidizingbacteria: oxidation of C-1 compounds by Meth-ylococcus capsulatus. J. Bacteriol. 107:187-192.

113. Patel, R. N., S. L. Hoare, D. S. Hoare. 1979.[1-"C]-acetate assimilation by obligate meth-ylotrophs, Pseudomonas methanica and Meth-ylosinus trichosporium. Antonie van Leeuwen-hoek J. Microbiol. Serol. 45:499-511.

114. Patel, R., S. L. Hoare, D. S. Hoare, and B. F.Taylor. 1975. Incomplete tricarboxylic acid cy-cle in a type I methylotroph, Methylococcuscapsulatus. J. Bacteriol. 123:382-384.

115. Patel, R. N., D. S. Hoare, and B. F. Taylor.1969. Biochemical basis of obligate autotrophyin Methylococcus capsulatus (Texas). Bacte-riol. Proc., p. 128.

116. PateL R. N., C. T. Hou, P. Derelanko, and A.Felix. 1980. Purification and properties of aheme-containing aldehyde dehydrogenasefrom Methylosinus trichosporium. Arch. Bio-chem. Biophys. 203:654-662.

117. Patel, R. N., C. T. Hou, and A. Felix. 1978.Microbial oxidation of methane and methanol:isolation of methane-utilizing bacteria andcharacterization of a facultative methane-uti-lizing isolate. J. Bacteriol. 136:352-358.

118. Patel, R. N., C. T. Hou, A. L. Laskin, A. Felix,and P. Derelanko. 1979. Microbial oxidationof gaseous hydrocarbons. II. Hydroxylation ofalkanes and epoxidation of alkenes by cell-freeparticulate fractions of methane-utilizing bac-teria. J. Bacteriol. 139:675-679.

119. PateL R. N., C. T. Hou, A. L. Laskin, A. Felix,and P. Derelanko. 1980. Microbial oxidationofgaseous hydrocarbons: production ofsecond-ary alcohols from corresponding n-alkanes bymethane-utilizing bacteria. Appl. Environ. Mi-crobiol. 39:720-726.

120. Patel, R. N., C. T. Hou, A. I. Laskin, A. Felix,and P. Derelanko. 1980. Microbial oxidationof gaseous hydrocarbons: production of meth-ylketones from corresponding n-alkanes bymethane-utilizing bacteria. Appl. Environ. Mi-crobiol. 39:727-733.

121. PateL R. N., W. J. Mandy, and D. S. Hoare.1973. Physiological studies of methane- andmethanol-oxidizing bacteria: immunologicalcomparison of a primary alcohol dehydro-genase from Methylococcus capsulatus andPseudomonas sp. M27. J. Bacteriol. 113:937-945.

122. Patt, T. E., G. C. Cole, J. Bland, and R. S.Hanson. 1974. Isolation and characterizationof bacteria that grow on methane and organiccompounds as sole sources of carbon and en-ergy. J. Bacteriol. 120:955-964.

123. Patt, T. E., and R. S. Hanson. 1978. Intracyto-plasmic membrane, phospholipid, and sterolcontent of Methylobacterium organophilumcells grown under different conditions. J. Bac-teriol. 134:636-644.

124. Pearce, L E., and E. Meynell. 1968. Specificchromosomal affinity of a resistance factor. J.Gen. Microbiol. 50:159-172.

125. Pilayashenko-Novokhatnyi, A. I., A. N. Gri-

VOL. 45, 1981


mbr.asm

.org/D

ownloaded from


588 HIGGINS ET AL.

goryan, A. P. Kovalev, V. S. Belova, andR. L. Gvozdev. 1979. The kinetic isotope effectand the temperature dependence of the meth-ane mono-oxygenase reaction. Dokl. Biochem.245:147-150.

126. Proctor, H. M., J. R. Norris, and D. W. Rib-bons. 1969. Fine structure of methane-utilisingbacteria. J. Appl. Bacteriol. 32:118-121.

127. Pudzianowski, A. T., and G. H. Loew. 1980.Quantum-chemical studies of model cyto-chrome P450 hydrocarbon oxidation mecha-nisms. 1. A MINDO/3 study of hydroxylationand epoxidation pathways for methane andethylene. J. Am. Chem. Soc. 102:5443-5449.

128. Quayle, J. R. 1972. The metabolism of one-car-bon compounds by micro-organisms. Adv. Mi-crob. Physiol. 7:119-203.

129. Quayle, J. R. 1975. Unsolved problems in themicrobial metabolism of methane and metha-nol, p. 59-65. In G. Terui (ed.), Microbialgrowth on C, compounds. Proceedings of theFirst International Symposium on MicrobialGrowth on Cl-Compounds, Tokyo. Society ofFermentation Technology, Osaka.

130. Quayle, J. R. 1980. Historical perspectives, p. 1-9. In D. E. F. Harrison, I. J. Higgins, and R.Watkinson (ed.), Hydrocarbons in biotechnol-ogy. Heyden & Son, Ltd., London.

131. Quayle, J. R. 1980. Microbial assimilation of C,-compounds. Biochem. Soc. Trans. 8:1-10.

132. Quayle, J. R. 1980. Aspects of the regulation ofmethylotrophic metabolism. FEBS Lett. 117(Suppl):K16-K27.

133. Quayle, J. R., and T. Ferenci. 1978. Evolution-ary aspects of autotrophy. Microbiol. Rev. 42:251-273.

134. Reed, W. M., and P. R. Dugan. 1978. Distri-bution ofMethylomonas methanica and Meth-ylosinus trichosporium in Cleveland Harbor asdetermined by an indirect fluorescent anti-body-membrane filter technique. Appl. Envi-ron. Microbiol. 35:422-430.

135. Reed, W. M., and P. R. Dugan. 1979. Study ofdevelopmental stages of Methylosinus tricho-sporium with the aid of fluorescent-antibodystaining techniques. Appl. Environ. Microbiol.38:1179-1183.

136. Reed, W. M., J. A. Titus, P. R. Dugan, and R.M. Pfister. 1980. Structure of Methylosinustrichosporium exospores. J. Bacteriol. 141:908-913.

137. Ribbons, D. W. 1975. Oxidation of C, com-pounds by particulate fractions from Methylo-coccus capsulatus: distribution and propertiesof methane-dependent reduced nicotinamideadenine dinucleotide oxidase (methane hy-droxylase). J. Bacteriol. 122:1351-1363.

138. Ribbons, D. W., and J. L. Michaelover. 1970.Methane oxidation by cell-free extracts ofMethylococcus capsulatus. FEBS Lett. 11:41-44.

139. Ribbons, D. W., and A. M. Wadzinski. 1976.Oxidation of C, compounds by particulate frac-tions from Methylococcus capsulatus, p. 359-369. In H. G. Schlegel, G. Gottschalk, and N.Pfennig (ed.), Symposium on microbial produc-

tion and utilisation of gases (H2, CH4, CO),Gottingen. Akademie der Wissenschaften, Got-tingen.

140. Romanovskaya, V. A. 1978. Nomenclature ofobligate methylotrophs. Microbiology (USSR)47:1063-1072.

141. Romanovskaya, V. A. 1980. Certain aspects ofclassification of methane-oxidising bacteria, p.89-91. In G. M. Tonge (ed.), Abstracts of pos-ters, Third International Symposium on Micro-bial Growth on C, Compounds. University ofSheffield Press, Sheffield, United Kingdom.

142. Romanovskaya, V. A., Y. S. Sadovmikov,and Y. R. Malashenko. 1978. Formation oftaxons of methane-oxidising bacteria using nu-merical analysis methods. Microbiology(USSR) 47:120-130.

143. Rudd, J. W. M., A. Furutani, R. J. Flett, andR. D. Hamilton. 1976. Factors controllingmethane oxidation in shield lakes: the role ofnitrogen fixation and oxygen concentration.Limnol. Oceanogr. 21:357-364.

144. Salem, A. R., A. J. Hacking, and J. R. Quayle.1973. Cleavage of malylcoenzyme A into acetyl-coenzyme A and glyoxylate by PseudomonasAM1 and other Ci-unit-oxidising bacteria. Bio-chem. J. 136:89-96.

145. Salisbury, S. A., H. S. Forrest, W. B. T. Cruse,and 0. Kennard. 1979. A novel coenzymefrom bacterial dehydrogenases. Nature (Lon-don) 280:843-844.

146. Scott, D., J. Brannan, and I. J. Higgins. 1981.The effect of growth conditions on intracyto-plasmic membranes and methane mono-oxy-genase activities in Methylosinus trichospo-rium OB3b. J. Gen. Microbiol. 125:63-72.

147. Shishkina, V. N., and Y. A. Trotsenko. 1979.Pathways of ammonia assimilation in obligatemethane utilisers. FEMS (Fed. Eur. Microbiol.Soc.) Microbiol. Lett. 5:187-191.

148. Smith, A. J., and D. S. Hoare. 1977. Specialistphototrophs, lithotrophs and methylotrophs: aunity among a diversity of procaryotes? Bac-teriol. Rev. 41:419-448.

149. Smith, U., and D. W. Ribbons. 1970. Finestructure of Methanomonas methanooxidans.Arch. Mikrobiol. 74:116-122.

150. Smith, U., D. W. Ribbons, and D. S. Smith.1970. The fine structure of Methylococcus cap-sulatus. Tissue Cell 2:513-520.

151. Sohngen, N. L. 1906. Uber Bakterien, welcheMethan als Kohlenstoffnahrung und Energie-quelle gebrauchen. Centr. Bakt. Parasitenk.15:513-517.

152. Sokolov, L. G., V. A. Romanovskaya, Y. B.Shkurko, and Y. R. Malashenko. 1980.Comparative characterisation of the enzymesystems of methane-utilising bacteria that ox-idise NH20H and CH30H. Microbiology(USSR) 49:142-148.

153. Stirling, D. I., J. Colby, and H. Dalton. 1979.A comparison of the substrate and electron-donor specificities of the methane mono-oxy-genases from three strains of methane-oxidis-ing bacteria. Biochem. J. 177:361-364.

154. Stirling, D. I., and H. Dalton. 1977. Effect of

MICROBIOL. REV.


mbr.asm

.org/D

ownloaded from



metal-binding agents and other compounds on

methane oxidation by two strains of Methylo-coccus capsulatus. Arch. Microbiol. 114:71-76.

155. Stirling, D. I., and H. Dalton. 1978. Purificationand properties of an NAD(P)+-linked formal-dehyde dehydrogenase from Methylococcuscapsulatus (Bath). J. Gen. Microbiol. 107:19-29.

156. Stirling, D. I., and H. Dalton. 1979. The for-tuitous oxidation and cometabolism of variouscarbon compounds by whole-cell suspensionsof Methylococcus capsulatus (Bath). FEMS(Fed. Eur. Microbiol. Soc.) Microbiol. Lett. 5:315-318.

157. Stirling, D. L, and H. Dalton. 1979. Propertiesof the methane mono-oxygenase from extractsof Methylosinus trichosporium OB3b and evi-dence for its imilarity to the enzyme fromMethylococcus capsulatus (Bath). Eur. J. Bio-chem. 96:205-212.

158. Stirling, D. L, and H. Dalton. 1981. Fortuitousoxidations by methane-utflising bacteria. Na-ture (London) 291:169.

159. Strom, T., T. Ferenei, and J. R. Quayle. 1974.The carbon asimilation pathways of Meth-ylococcus capsukatu, Pseudomonas methan-ica and Methylosinus trichosporium (OB3b)during growth on methane. Biochem. J. 144:465-476.

160. Suzina, N. E., and B. A. Fikhte. 1977. Finestructure of tubular formations found out on

the surface of cells Methylocystis echinoidessp. nov., p. 23-25. In G. K. Skryabin, M. V.Ivanov, E. N. Kondratjeva, G. A. Zavarzin, Yu.A. Trotsenko, and A. I. Nesterov (ed.), Micro-bial growth on Cl-compounds. Abstracts of theSecond International Symposium on MicrobialGrowth on Cl-Compounds. USSR Academy ofSciences, Pushchino, USSR.

161. Suzuki, I., S.-C. Kwok, and U. Dular. 1976.Competitive inhibition of ammonia oxidationin Nitrosomonas europaea by methane, car-bon monoxide or methanol. FEBS Lett. 72:117-120.

162. Takeda, K., and K. Tanaka. 1980. Ultrastruc-ture of intracytoplasmic membranes of Meth-anomonas margaritae cells grown under dif-ferent conditions. Antonie van LeeuwenhoekJ. Microbiol. Serol. 46:15-25.

163. Taylor, L J., and C. Anthony. 1976. A biochem-ical basis for obligate methylotrophy: proper-

ties of a mutant of Pseudomonas AMi lacking2-oxoglutarate dehydrogenase. J. Gen. Micro-biol. 93:259-265.

164. Taylor, S. 1977. Evidence for the presence ofribulose 1,5-bisphosphate carboxylase andphosphoribulokinase in Methylococcus capsu-

latus (Bath). FEMS (Fed. Eur. Microbiol.Soc.) Microbiol. Lett. 2:305-307.

165. Taylor, S., H. Dalton, and C. Dow. 1980. Pu-rification and initial characterisation of ribu-lose 1,5-bisphosphate carboxylase from Meth-ylococcus capsulatus (Bath). FEMS (Fed. Eur.Microbiol. Soc.) Microbiol. Lett. 8:157-160.

166. Taylor, S. C., H. Dalton, and C. S. Dow. 1981.Ribulose 1,5-bisphosphate carboxylase/oxy-

genase and carbon aimilation in Methylococ-cus Cap8Ulatus (Bath). J. Gen. Microbiol. 122:89-94.

167. Thompson, A. W., J. G. O'Neill, and J. F.Wilkinson. 1976. Acetone production bymethylobacteria. Arch. Mikrobiol. 109:243-246.

168. Tomaszewski, J. E., D. M. Jerina, and J. W.Daly. 1975. Deuterium isotope effects duringformation of phenols by hepatic monooxygen-ases. Evidence for an altemative to the areneoxide pathway. Biochemistry 14:2024-2031.

169. Tonge, G. M., D. E. F. Harrison, and L J.Higgins. 1977. Purification and properties ofthe methane mono-oxygenase enzyme systemfrom Methylo8inus trichosporium OB3b. Bio-chem. J. 161:333-344.

170. Tonge, G. M., D. E. F. Harrison, C. J.Knowles, and L J. Higgins. 1975. Propertiesand partial purification of the methane-oxidis-ing enzyme system from Methylosinus tricho-spor ium. FEBS Lett. 58:293-299.

171. Trotsenko, Y. A. 1976. Isolation and character-isation of obligate methanotrophic bacteria, p.329-336. In H. G. Schlegel, G. Gottschalk, andN. Pfennig (ed.), Symposium on microbial pro-duction and utilisation of gases (H2, CH4, CO),Gottingen. Akademie der Wissenschaften, Got-tingen.

172. Trotsenko, Y. A., and Y. A. Shishkina. 1980.Multiple metabolic lesions in obligate meth-anotrophic bacteria, p. 64-65. In G. M. Tonge(ed.), Abstracts of posters, Third InternationalSymposium on Microbial Growth on C, Com-pounds. University of Sheffield Press, Shef-field, United Kingdom.

173. Trotsenko, Y. A., and A. P. Sokolov. 1980.Disimilatory hexulose-phosphate cycle and itscontrol in various methylotrophs, p. 62. In G.M. Tonge (ed.), Abstracts of posters, ThirdInternational Symposium on Microbial Growthon C, Compounds. University of SheffieldPress, Sheffield, United Kingdom.

174. Tyutikov, F. M., N. N. Belyaevf, L. C. Smir-nova, A. S. Tikhonenko, and A. S. Kri-visky. 1976. Isolation of bacteriophages ofmethane-oxidising bacteria. Mikrobiologiya 6:1056-1062. (In Russian.)

175. Tyutikov, F. M., L. A. Bespalova, B. A. Re-bentish, N. N. Aleksandrushkina, and A.S. Krivisky. 1980. Bacteriophages of meth-anotrophic bacteria. J. Bacteriol. 144:375-381.

176. Wadzinski, A. M., and D. W. Ribbons. 1975.Oxidation of C1 compounds by particulate frac-tions from Methylococcus capsulatus: proper-ties of methanol oxidase and methanol dehy-drogenase. J. Bacteriol. 122:1364-1374.

177. Wadzinski, A. M., and D. W. Ribbons. 1975.Utilization of acetate by Methanomonas meth-anooxidans. J. Bacteriol. 123:380-381.

178. Warner, P. J., L. J. Higgins, and J. W. Drozd.1979. Transfer of antibiotic resistance into afacultative methylotroph and an obligatemethylotroph by conjugation. Soc. Gen. Micro-biol. Q. 6:71-72.

179. Warner, P. J., L. J. Higgins, and J. W. Drozd.

VOL. 45, 1981


mbr.asm

.org/D

ownloaded from


590 HIGGINS ET AL.

1980. Conjugative transfer of antibiotic resist-ance to methylotrophic bacteria. FEMS (Fed.Eur. Microbiol. Soc.) Microbiol. Lett. 7:181-185.

180. Weaver, T. L., and P. R. Dugan. 1974. Intra-cytoplasmic membrane function in the meth-ylotrophic bacterium Methylosinus trichospo-rium. In Abstr. Annu. Meet. Am. Soc. Micro-'biol. 74:173.

181. Weaver, T. L., and P. R. Dugan. 1975. Ultra-structure ofMethylosinus trichosporium as re-vealed by freeze etching. J. Bacteriol. 121:704-710.

182. Weaver, T. L., M. A. Patrick, and P. R. Du-gan. 1975. Whole-cell and membrane lipids ofthe methylotrophic bacterium Methylosinustrichosporium. J. Bacteriol. 124:602-605.

183. Westerling, J., J. Frank, and J. A. Duine.1979. The prosthetic group of methanol dehy-drogenase from Hyphomicrobium X: electronspin resonance evidence for a quinone struc-ture. Biochim. Biophys. Acta 87:719-724.

184. Whittenbury, R., J. Colby, H. Dalton, and H.C. Reed. 1976. Biology and ecology of methaneoxidisers, p. 281-292. In H. G. Schlegel, G.Gottschalk, and N. Pfennig (ed.), Symposiumon microbial production and utilisation of gases(H2, CH4, CO), Gottingen. Akademie der Wis-senschaften, Gottingen.

185. Whittenbury, R., and H. Dalton. 1981. Themethylotrophic bacteria. In M. P. Starr, H.Stolp, H. G. Truper, A. Balows, and H. G.Schlegel (ed.), The prokaryotes. Springer-Ver-lag KG, Berlin, Federal Republic of Germany.(In press.)

186. Whittenbury, R., H. Dalton, M. Eccleston,and H. L. Reed. 1975. The different types ofmethane oxidising bacteria and some of theirmore unusual properties, p. 1-9. In G. Terui(ed.), Microbial growth on C, compounds. Pro-ceedings of the International Symposium onMicrobial Growth on C, Compounds, Tokyo.Society of Fermentation Technology, Osaka.

187. Whittenbury, R., S. L. Davies, and J. F.Davey. 1970. Exospores and cysts formed by

methane-utilising bacteria. J. Gen. Microbiol.61:219-226.

188. Whittenbury, R., K. C. Phillips, and J. F.Wilkinson. 1970. Enrichment, isolation andsome properties of methane-utilising bacteria.J. Gen. Microbiol. 61:205-218.

189. Williams, E., and M. A. Shimmin. 1978. Radia-tion-induced filamenation in obligate methy-lotrophs. FEMS (Fed. Eur. Microbiol. Soc.)Microbiol. Lett. 4:137-141.

190. Williams, E., M. A. Shimin, and B. W. Bain-bridge. 1977. Mutation in the obligate meth-ylotrophs Methylococcus capsulatus andMethylomonas albus. FEMS (Fed. Eur. Micro-biol. Soc.) Microbiol. Lett. 2:293-296.

191. Wolf, H. J., and R. S. Hanson. 1978. Alcoholdehydrogenase from Methylobacterium organ-ophilum. Appl. Environ. Microbiol. 36:105-114.

192. Wolf, H. J., and R. S. Hanson. 1979. Isolationand characterisation of methane-utilisingyeasts. J. Gen. Microbiol. 114:187-194.

193. Wolf, H. J., and R. S. Hanson. 1980. Identifi-cation of methane-utilising yeasts. FEMS (Fed.Eur. Microbiol. Soc.) Microbiol. Lett. 7:177 -179.

194. Wolf, H. J., and R. S. Hanson. 1980. Ultrastruc-ture of methanotrophic yeasts. J. Bacteriol.141:1340-1349.

195. Wolfe, R. S., and L. J. Higgins. 1979. Microbialbiochemistry of methane-a study in contrasts.Int. Rev. Biochem. 21:267-353.

196. Wopat, A. E., and M. L. O'Connor. 1980. Plas-mids in Methylobacterium ethanolicum, a newfacultative methanotroph, p. 16. In G. M.Tonge (ed.), Abstracts of posters, Third Inter-national Symposium on Microbial Growth onC, Compounds. University of Sheffield Press,Sheffield, United Kingdom.

197. Zajic, J. E., B. Volesky, and A. Wellman.1969. Growth of Graphium sp. on natural gas.Can. J. Microbiol. 15:1231-1236.

198. Zehnder, A. J. B., and T. D. Brock. 1979.Methane formation and methane oxidation bymethanogenic bacteria. J. Bacteriol. 137:420-432.

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Methane-Oxidizing Microorganisms › content › mmbr › 45 › 4 › 556.full.pdf · METHANE-OXIDIZING MICROORGANISMS 557 rized in Table 1, whichwascompiled fromthe data of Ehhalt