Metabolismo n Alcanos Review

8/3/2019 Metabolismo n Alcanos Review

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Dedicated to Professor Wolfgang BABEL on the occasion of his 65th birthday

Bacterial Metabolism ofn-Alkanes and Ammonia

under Oxic, Suboxic and Anoxic Conditions

BERTHE-CORTI,* L., FETZNER, S.

Carl von Ossietzky Universitt Oldenburg * Corresponding authorFachbereich Biologie, Geo- und Umweltwissenschaften Phone: + 49 441 798 3290Ammerlnder Heerstrae 114118 Fax: + 49 441 798 325026111 Oldenburg, Germany E-mail: [email protected]

Summary

n-Alkanes are widespread in the biosphere. Due to the lack of functional groups, these alkanes exhibitlow chemical reactivity. However, many microorganisms have evolved pathways to utilise n-alkanesas a growth substrate, and moreover, fortuitous alkane oxidation may play an important role in alkanedegradation. This review discusses the ecology of n-alkane-degrading and ammonia-oxidising bacte-ria with a focus on alkane metabolism in the transition from oxic to anoxic conditions, the pathwaysofn-alkane and ammonium oxidation, and the enzymes catalysing n-alkane and ammonia activation.

n-Alkane degrading bacteria occur in oxic as well as strictly anoxic environments, and they live invery diverse habitats, including marine or fresh water, soils, sediments or aquifers. Aerobic ammo-nium-oxidising as well as methanotrophic bacteria are often found in stratified habitats such as bio-films and sediments.Aerobic pathways involving oxygenases that catalyse the initial activation of n-alkanes and ammo-nium are well known. However, anaerobic ammonium oxidation as well as anaerobic utilisation ofhydrocarbons have been demonstrated only in the past decade and are the subject of current researchefforts. Enzyme systems that catalyse aerobic alkane oxidation involve a number of well-character-ised monooxygenases such as cytochrome P450 monooxygenases, multi-component alkane mono-oxygenases (also known as -hydroxylase systems), methane monooxygenases, and ammoniamonooxygenase. Alternative enzymes, for example an n-alkyl hydroperoxide-forming dioxygenase,have also been postulated, but contrary to the monooxygenases, an n-alkane oxidising dioxygenasehas not yet been biochemically characterised. The oxygenase components of soluble methanemonooxygenase and alkane monooxygenase contain binuclear iron centres that mediate dioxygenactivation, whereas particulate methane monooxygenase, ammonia monooxygenase, and presumably

distinct butane monooxygenases are copper-containing enzymes.Little is known about the impact of the oxygen concentration on bacterial alkane degradation, and ithas not yet been investigated which pathways and enzymes are active in bacteria which utilise alkanesat suboxic or even quasi-anoxic conditions. Methane monooxygenase as well as ammoniamonooxygenase have low half-saturation constants for oxygen and, in addition, both have an amplesubstrate spectrum. Activation ofn-alkanes by cooxidation has been demonstrated for both types ofenzymes. In suboxic to quasi-anoxic habitats, in which alkane, ammonium and methane oxidisingbacteria as well as other organotrophic microorganisms live in close vicinity, a cooperative effectwith respect to n-alkane degradation may occur.

WILEY-VCH Verlag Berlin GmbH, 13086 Berlin, 2002 0138-4988/02/03-407-0299 $ 17.50+.50/0

Acta Biotechnol. 22 (2002) 3--4, 299--336


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Introduction

Alkanes as constituents of natural gas, petroleum, petrochemical products and coal [1,2] and as metabolites from organisms [3] are widespread compounds in terrestrial andmarine environments. The aerobic biodegradation of these compounds is well docu-mented [47]. Numerous hydrocarbon-degrading strains are described, which belong tovery different taxonomic groups (for a list, see [8]). Recently, novel alkane-degradingstrains, such as Marinobacter hydrocarbonoclasticus [9] Fundibacter jadensis [10],and Alcanivorax borkumenis [11] have been isolated, especially from marine habitats.Hydrocarbon-polluted sites are often deficient in molecular oxygen or are even com-pletely anoxic. In the last decade, the alkane metabolism of denitrifying and of sulphate-reducing bacteria under anoxic conditions has been the subject of intensive investigation

[1215]. A few papers also describe hexadecane degradation under methane evolution[16, 17]. Several metabolic mechanisms of anaerobic oxidation of alkanes are currentlybeing discussed [14, 18, 19]. Little is known, however, about hydrocarbon degradationin the transition from oxic to anoxic conditions.This review discusses the bacterial degradation of n-alkanes under oxic, suboxic andanoxic conditions, with a focus on the processes occurring in the transition between oxicand anoxic conditions. In order to explain bacterial alkane metabolism under dioxygen-deficient conditions, first the ecology of alkane-degrading bacteria as well as of ammo-nium-oxidising bacteria is described, as the latter are assumed to also be involved inalkane metabolism under dioxygen-depleted conditions (see section EnvironmentalAspects). To elucidate the role of dioxygen in alkane metabolism, a survey of the varietyof metabolic pathways and of the enzymes involved in bacterial alkane, methane andammonia oxidation is provided (see sections Pathways of Alkane and Ammonium Oxida-tion and Enzymes Catalysing Alkane and Ammonia Oxidation). Methane and ammonia-oxidising enzymes are included, since these are known to mediate the oxidation of anumber of alkanes. Thus, methanotrophs and nitrifiers might well contribute to alkanedegradation in natural habitats. Most of the enzymes, which catalyse the initial activa-tion of alkanes, require molecular oxygen as a cosubstrate, regardless of whether theenzymes have a high specificity or act fortuitously. Therefore, the question arises,which strategies and pathways of alkane utilisation are used by the indigenous bacterialcommunities in suboxic and quasi-anoxic habitats? This question is especially ad-dressed in the section Alkane Degradation under Oxygen Deficient Conditions of thisreview.

Environmental Aspects

Ecology of Alkane-Degrading Bacteria

Due to spills or, in some places, due to natural activity, alkanes are present in such dif-ferent environments as marine or fresh water, in terrestrial sites or in aquifers, as well asin different climates. Because alkane degradation is a common phenomenon in allinvestigated habitats, it is impossible to address a specific ecology of alkane-degradingbacteria.

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When investigating alkane degradation in soil, most authors adjust the soil humidity tooptimum conditions (e.g., 50% water-holding capacity [20]). Microbial activity is con-siderably lower in dry soils than in humid soils; accordingly RADWAN et al. [21]reported that the alkane concentration in oil-polluted Kuwaiti desert soils remainedfairly constant during the dry hot season, but decreased considerably (about 50%) dur-ing the rainy months. In most studies, alkane degradation is reported to occur at a moreor less neutral pH. Articles on alkane degradation refer only very superficially to the pHvalue as a factor, which influences degradation rates. This is not surprising as manyhabitats have a more or less neutral pH (seawater and marine sediments, for example,have a pH of about 7 to 8). Industrial fermentations using hydrocarbons as a carbonsource are often carried out at neutral conditions. However, fresh water and fresh-watersediments are not always neutral; they are sometimes acidic. The pH of soil according

to the soil composition and history ranges from about pH 2.5 to about 10 [22]. SCHAUERpoints out in his review that the oxidation rates of hydrocarbons vary only slightly at pHvalues of between 5 and 8, whereas the oxidation and the toxicity of the organic acidswhich are products of microbial hydrocarbon metabolism are clearly dependent on thepH [23].The influence of temperature on hydrocarbon/alkane degradation has not been investi-gated in much detail, with the exception of low temperatures. Most publications onhydrocarbon/alkane degradation are based on experiments performed at moderate tem-peratures between 20 and 30 C. Some exceptions exist: for example, RUETER et al. [13]described a slightly thermophilic anaerobic strain (TD3) that degrades alkane under sul-phate-reducing conditions; KLUG and MARKOVETZ [24] reported on a thermophilic bac-terium which is able to degrade n-tetradecane; and BERTHE-CORTI and MORELLI isolatedan aerobic thermotolerant bacterium that grows and degrades hexadecane up to 60 C(unpublished results). Industrial alkane degradation at high temperatures (e.g., 65 to70 C) has also been reported [25, 26].WALKER and COLWELL [27] demonstrated alkane degradation by microbial populationsenriched from water and sediment from the Chesapeake Bay at low temperatures(010 C), using a model petroleum containing 85.3% n-alkanes, 3.9% pristane andcyclic alkanes, as well as 6.39% mono- and polynuclear aromatics. Chesapeake Bay,however, is not a constantly cold region. The average temperature is below 10 C foronly 6 months a year. The authors isolated the genera Vibrio, Aeromonas, Pseudomonasand Acinetobacter, which are well-known alkane degraders at temperate conditions [8,23]. WHYTE et al. [2830] reported the ability of psychrotrophic microorganisms, espe-cially ofRhodococcus sp., to degrade alkanes of variable chain-length at temperaturesof 0 to 5 C.

Alkane degradation in other environments, such as extremely acidic or hot habitats,seems to be rare or has not been investigated thus far.

Electron Acceptors for Alkane Degradation

One important factor for hydrocarbon degradation in polluted habitats is the presence ofan adequate electron acceptor. This is in the optimal case molecular oxygen. How-ever, wet soils, swamps, fresh water or marine sediments and aquifers are microbial

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habitats, which are often subject to hydrocarbon pollution and which are frequentlyoxygen limited. Apart from the aquifers, the habitats are characterised by a thin (afew mm to a few cm) oxic surface layer, an oxygen and redox stratification zone, whosemagnitude depends on the type of the sediment and on the biological activity, and ananoxic zone. Examples of such stratification in fresh-water sediments and marine sedi-ments have been published by SRENSEN et al., JRGENSEN and REVSBECH, and SWEERTSet al. [3133] and for wet soils by FRENZEL et al. [34]. The different zones contain char-acteristic electron acceptors such as dioxygen, nitrate and sulphate as well as Fe(III),Mn(IV) and CO2, which can be used for a respirative metabolism of organic matter [35,36]. Despite the different physicochemical conditions in biofilms and microbial mats,stratification and correlation with the metabolism have also been reported for biofilm-habitats (for more information, see [3740]).

Fig. 1. Mass ratio of hexadecane uptake into cells to hexadecane mineralisation at dif-ferent dissolved oxygen tensions (DOT) in sediment-seawater suspensions with autoch-thonous microbial populations (adapted from [46]).

Depending on the spatial distribution of the electron acceptors in these habitats, micro-organisms with very different metabolic capabilities are to be expected. In the oxic



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layer, genera and species, which use dioxygen as an electron acceptor, predominate per-forming aerobic alkane metabolism (see [6, 7, 41, 42], and section Oxidation of Alkanesby Aerobic Bacteria). In anoxic environments, alkane degradation has been demonstratedfor sulphate-reducing bacteria and denitrifying species as well as for methane-generating species (see [1219, 43], and section Oxidation of Alkanes by Anaerobic Bacte-ria). SO and YOUNG [44] reported anaerobic alkane degradation in enrichment culturesfrom a marine estuarine sediment. The cultures were able to utilise alkanes anaerobi-cally under sulphate-reducing, denitrifying, iron-reducing and methanogenic conditions.The sediment had a long history of hydrocarbon pollution. ZHANG and YOUNG [45] alsoisolated microorganisms able to anaerobically degrade polyaromatic hydrocarbons fromthe same site. In contrast to metabolism under oxygen saturation or in complete absenceof oxygen, little is known about alkane degradation by microaerophilic or facultative

anaerobic species under oxygen-deficient conditions.MICHAELSEN et al. [46] investigated hexadecane mineralisation by an indigenousmicrobial community, which was taken from intertidal sediment. Hexadecanemineralisation decreased and production of metabolites increased when the dissolvedoxygen concentration was reduced from oxic (approx. 168 mol O2/l) to suboxic(approx. 0.8 mol O2/l) conditions (Fig. 1). This well agrees with data ofBONIN andBERTRAND [47], who demonstrated that the ratio of CO2 production to heptadecaneconsumption in cultures of Pseudomonas nautica decreased when the oxygen concen-tration was reduced to suboxic conditions.

Alkane-Degrading Communities

In suspension cultures containing intertidal sediment, BERTHE-CORTI and co-workershave demonstrated that bacterial communities derived from intertidal sediment are ableto degrade hexadecane under oxic, suboxic ( 0.8 mol O2/l) and quasi-anoxic condi-tions (


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for bacteria of the Cytophaga-Flavobacteria-Bacteroides (CFB) group [53]. LLOBET-BROSSA et al. [54] determined nearly the same community composition in samples takenfrom the upper few cm of intertidal sediment from the German North Sea coast.However, the community composition of the suspension culture investigated byBERTHE-CORTI and BRUNS [49] clearly changed in the transition from suboxic to anoxicconditions. The amount of Eubacteria, which could be hybridised with the probeEUB338 comprised 7080% of the total cell number (DAPI-stained cells) undersuboxic conditions and decreased to 4050% under quasi-anoxic conditions, when theoxygen supply was extremely low. The sum of the other detected phylogenetic groups(-, -, -Proteobacteria and CFB), which under suboxic conditions comprisedapproximately 100% of the total number of Eubacteria, decreased to about 30% underquasi-anoxic conditions [49].

Microaerobic growth of pure cultures is described for Pseudomonas nautica, a denitri-fying marine bacterium, which grows at low oxygen concentrations of about 22 mol ofO2/l (10% of air saturation) using heptadecane, irrespective of whether nitrate is presentin addition to oxygen or not [55]. Pseudomonas aeruginosa grows using n-hexadecaneunder low-oxygen concentrations and under anoxic denitrifying conditions [56]. Fromsuboxic enrichment cultures containing intertidal sediment, BRUNS and BERTHE-CORTIhave isolated a new alkane-degrading species, Fundibacter jadensis [10], which is ableto grow and mineralise hexadecane under oxic and suboxic, but not under completelyanoxic conditions (unpublished data).

Alkane Degradation under Oxygen Deficient Conditions

The oxygophily of bacteria, which, among other factors, is a result of the oxygenaffinity of the oxidising enzyme systems (vmax/kM) and the half-saturation constant forbacterial growth (kdo), determines how a single bacterium or a population can grow atcertain oxygen concentrations. The published kdo values for bacteria are in the range of afew moles of oxygen per litre (Tab.1). It is difficult to compare the kdo values ofdifferent authors because the experimental settings used vary considerably, differingfrom batch to continuous cultures. Nevertheless, the data suggest that bacteria whichgrow with alkanes as a carbon source can compete with bacteria growing on othersubstrates at very low oxygen concentrations.Half-saturation constants of cultures do not imply that each individual cell or eachspecies in a community has the same substrate dependence. Using difficult carbon

sources, especially xenobiotics, one has to assume cooperative metabolic effects suchas cometabolism or cooxidation, concerted metabolic attack, and modification ofindividual growth parameters, etc. (for a review, see [57]). As a consequence, half-saturation constants are multifactorial values that depend on the intrinsic factors of cells,the actual environment and the history of a culture [58]. BERTHE-CORTI and EBENHH[59] demonstrated that in a marine bacterial community the half-saturation constantdecreased when the oxygen concentration in the culture had been reduced. LAANBROEKet al. [60] showed a similar adaptation for the oxygen affinity (vmax/kdo ) of theammonium-oxidising bacterium Nitrosomonas and the nitrite oxidising bacterium



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Nitrobacter in mixed cultures (see also section Ammonia Oxidation under Oxygen Defi-cient Conditions).The typical stratification of most oxygen-deficient habitats is characterised by a zone, inwhich molecular oxygen enters by diffusion and is simultaneously consumed bymicrobial activity, so that its concentration is so low that it cannot be analysed byoxygen electrodes. Consequently, it is not possible to calculate half-saturation constantsfor oxygen. The relationship between the oxygen supply and the metabolic activity ofthe cells in these quasi-anoxic habitats is determined by the flux rate of oxygen (oxygensupply rate, OSR). BERTHE-CORTI and BRUNS [49] demonstrated that in quasi-anoxiccontinuous sediment-suspension cultures with hexadecane as the sole carbon source, thedependence of the growth rate of a bacterial community (analysed as protein productionrate) and the hexadecane consumption rate on the oxygen supply can be characterised

by a hyperbola analogous to the MONOD kinetics. When bacterial mud-flat communitiesare cultivated at quasi-anoxic conditions and the OSR is reduced stepwise to aminimum of 0.06 mmol O2/l h, a threshold value of 0.035 mmol O2/l h was esti-mated for the OSR at which the aerobic hexadecane metabolism of bacterial mud-flatcommunities ceased and anaerobic metabolism predominated.

Tab. 1. Half-saturation constants of oxygen for growth of some bacteria on differentcarbon sources_____________________________________________________________________________________________________________________________________________

Organism Kdo Carbon source Reference(mol O2/l)

_____________________________________________________________________________________________________________________________________________

Escherichia coli 0.41 Glucose [61]

Azotobacter vinelandii 0.53 Glucose [61]Sphaerotilus natans 0.44 Glucose-mineral

Sphaerotilus sp. 1.03 base medium [62]

Mycobacterium sp. 3.8 Acetate [63]

Mycobacterium sp. 5.9 Pyrene [63]

Enrichment culture 37.5 2,4-Dichloro- [64]from industrial sewage phenoxyacetate

Fundibacter jadensis approx. 0.6 Hexadecane Authors(unpublisheddata)

Bacterial community 0.4 0.6 Hexadecane [59]from intidal sediment_____________________________________________________________________________________________________________________________________________

Ecology of Ammonium-Oxidising Bacteria

Ammonium can be used as a source of energy by various autotrophic and heterotrophicnitrifying bacteria [65]. The nitrosobacteria oxidise the ammonium to nitrite, while thenitrobacteria oxidise nitrite to nitrate. Autotrophic nitrifying bacteria are found in soil,in marine and fresh water as well as in sediments and wastewater [6670]. The known



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autotrophic nitrifying genera are affiliated with different phylogenetic subdivisions ofthe Proteobacteria, such as the -, -, - and -subdivisions as well as the newlydescribed phylum Nitrospira, which comprises the genera Nitrospira marina andN. moscoviensis [71, 72]. Ammonium oxidising bacteria are abundant at oxic-anoxicinterfaces, which have a high flux of ammonia from the anoxic phase and a lowconcentration of molecular oxygen. Environmental factors such as pH, temperature,substrate-(NH3/NH4

+) and oxygen concentration control the bacterial activity [7375].JENSEN et al. [76] reported that high oxygen concentrations ( 200 mol/l) even seemedto inhibit nitrification in sediments. The authors investigated the coupling of nitrifica-tion and denitrification as well as the regulation of the process by oxygen in a fresh-water model sediment.Several chemoorganotrophic bacteria capable of oxidising ammonium have been

described. They comprise strains ofPseudomonas putida [77], Alcaligenes faecalis [65,78], Arthrobacter globiformis [65], Aerobacter aerogenes, Mycobacterium phlei, Strep-tomyces sp. and Thiosphaera pantotropha [65] as well as Nocardioides sp. strain CF8[79]. These bacteria might also contribute to the initial oxidation of alkanes via theactivity of ammonia monooxygenase (AMO, see section Ammonia Monooxygenase).Their contribution to alkane mineralisation in nature, however, may vary considerably.P. putida, for example, is well known for its high capacity to degrade hydrocarbons,including alkanes; and the Nocardioides sp. strain CF8 oxidises butane. For otherstrains, however, there is no information about their hydrocarbon-degrading capacity.In anoxic habitats, which have high ammonium concentration, classical autotrophicnitrifiers are frequently found and might possibly contribute to anaerobic ammonia oxi-dation [65, 8082] (see section Microaerophilic and Anaerobic Activities of the AutotrophicNitrifiers). However, anaerobic ammonium oxidation via the so-called anammox process

(see section Anaerobic Oxidation of Ammonium: The Anammox Reaction) is also performedby newly described bacteria, which group into the order of the Planctomycetales[8385]. The strain, which was first described, is a yet uncultured chemolithotrophicanammox strain and had been designated Candidatus Brocadia anammoxidans [83, 84].Another anammox strain within the Planctomycetales isolated from biofilm material ismore similar to a third strain named Candidatus Kuenenia stuttgartiensis than to Candi-datus Brocadia anammoxidans [84, 85]. The anammox process has a great potential forremoving nitrogen from wastewater and allows for efficient denitrification in the ab-sence of available carbon sources [82, 86].

Ammonia Oxidation under Oxygen Deficient Conditions

Chemoautotrophic ammonium-oxidising and nitrite-oxidising bacteria need oxygen inorder to produce nitrite and nitrate. In the natural habitats of these organisms, in mostcases molecular oxygen is limited, forcing both types of bacteria to compete with eachother as well as with other oxygen-consuming bacteria.The effect of the dissolved oxygen concentration on pure nitrifying cultures has beeninvestigated since the early 1960s [87]. STENSTROM (1980) provides an overview ofthese early papers and demonstrates that the bacterial capacity of nitrification is depend-ent on the ammonia or nitrogen concentration, the dissolved oxygen concentration and



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the cell mass in a culture [88]. (In this paper, cell mass is expressed in terms of retentiontime of continuous cultures). At high cell retention times (high biomass), nitrificationproceeds at dissolved oxygen concentrations of 0.51.0 mg O2/l (15 mol/l31 mol/l).At lower cell retention times, higher oxygen concentrations are needed. The authorestimates a critical dissolved oxygen concentration of 0.3 mg/l (9 mol/l), below whichno nitrification occurs. However, the half-saturation constants for oxygen represent onlyone aspect of the bacterial oxygen dependence. As will be explained in the section Am-monia-Oxidising Nitrifiers, oxygen is merely the cosubstrate of ammonia oxidation. In1988, JAYAMOHAN et al. [89] investigated the effect of the dissolved oxygen concentra-tion in cultures of nitrifying bacteria. The authors show that the ammonia-oxidisingbacteria have lower half-saturation constants for dioxygen (kdo = 0.63 mg O2/l, 20 molof O2/l) than the nitrite-oxidising bacteria (1.32 mg O2/l, 41 mol/l), but the half-satura-

tion value for ammonia is higher (kS = 3.59 mg N/l) than the kS values for nitrite oxida-tion (1.55 mg N/l). HANAKI et al. [90] investigated the combined effect of the organicloading and the dissolved oxygen concentration on the NH3 uptake and the growth ofnitrifying bacteria (ammonia oxidisers) as well as of nitrite oxidisers. In systems whereno organic loading had been performed, low dissolved oxygen concentrations(15.6 mol/l) did not affect the total turnover of ammonia in ammonia-oxidising bacte-rial cultures. The reason was that, on the one hand, the ammonia oxidation rate of thecells was reduced and, on the other, the growth yield of the bacteria was doubled at lowoxygen concentrations, which compensated for the reduced ammonia oxidation rate ofthe cells. However, high organic loading reduced ammonia oxidation. The inhibitoryeffect was enhanced by low dissolved oxygen tensions. The ammonia oxidation had ahalf-saturation constant at an oxygen concentration of 0.32 mg O2/l (10 mol/l). Incontrast, nitrite oxidation was strongly inhibited at low oxygen concentrations. The

growth yield of nitrite-oxidising bacteria, however, was unaffected. The effect of theoxygen concentration on the mixotrophic growth of the nitrite-oxidising bacteriumNitrobacter hamburgensis and the ammonium-oxidising bacterium Nitrosomonas euro-paea was investigated by LAANBROEK et al. [60]. The authors showed that the specificaffinity of the strains for oxygen is not a constant but, instead, depends on the actualoxygen concentration.In 1980, GOREAU et al. [75] published a paper on the production of NO2

and N2O bynitrifying bacteria at reduced oxygen concentrations. The authors cultivated pure cul-tures of the ammonium-oxidising bacterium Nitrosomonas sp. and the nitrite-oxidisingbacterium Nitrobacter sp. at oxygen saturation (approx. 7 mg O2/l, equivalent to218 mol O2/l) and under suboxic conditions (0.18 mg O2/l, 5.6 mol O2/l). The authorscompared the data with other ammonia oxidisers and demonstrated that, except for

Nitrobacter sp., the N2O production of the strains increased from 0.3% to nearly 10%(moles of N in N2O per mole of NO2 ) when the oxygen concentration was reduced.

Oxidation of Alkanes by Ammonium-Oxidising Bacteria

Bioremediation of hydrocarbon-polluted sites in soil is often improved by adding nutri-ents, especially nitrogen, in the form of ammonium salts. However, about 10 years ago,the assumption was verified that ammonium-oxidising bacteria can also oxidise a wide



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variety of hydrocarbons, as has been described for Nitrosomonas europaea. The hydro-carbon substrates converted by AMO (see section Ammonia Monooxygenase) includealkanes and alkenes as well as chlorinated aliphatics and aromatic compounds [9194].It is assumed that the oxidation of hydrocarbons catalysed by AMO leads to products,which are not further metabolised as carbon sources but which feed into a cometabolicpathway in the soil community. In bacterial communities with a considerable abundanceof ammonium-oxidising bacteria, initial activation of aliphatics may occur via anunspecific activity of ammonium-oxidising bacteria. Due to the low oxygen half-saturation constant of ammonia oxidation (see section Ammonia Oxidation under OxygenDeficient Conditions), hydrocarbon oxidation by AMO can also occur in the transitionfrom oxic to anoxic conditions. However, the relationship between ammonia oxidationand hydrocarbon degradation is very complex. Experiments with soil contaminated with

diesel fuel [95] or contaminated with diesel fuel and hexadecane [96] show that thestimulation of hydrocarbon degradation by the addition of ammonium is only possibleunder very specific soil conditions.

Emulsification and Uptake of Liquid or Solid Alkanes

Released into the environment, liquid and solid alkanes tend to adsorb to particles, e.g.,in soil and sediment. The degree of sorption depends on the physicochemical character-istics (e.g., grain size, chemical composition and charge) of the particles as well as onthe type of the alkane. In aqueous media, liquid alkanes form droplets, whose sizedepends on the mechanical energy that acts on the water/hydrocarbon mixture, as well

as on the concentration of surface-active substances. Microorganisms that use alkanes asa carbon source have to cope with these factors.Generally, the use of alkanes as a carbon source involves specific processes whichinclude: (i) interaction of cells with hydrocarbon dissolved in the aqueous phase;(ii) adherence of the cells to the surface of hydrocarbon droplets or particles which areconsiderably larger than the microbial cell, e.g. by means of fimbriae [97]; (iii) pro-duction and excretion of bioemulsifiers to enhance the solubilisation of the hydrocarbonand, consequently, the uptake of the solubilised substrate [98]. An excellent insight intothe mechanisms of alkane uptake and growth on hydrocarbons in laboratory cultures ispresented by MIURA [99].Several hydrocarbon-degrading bacteria produce biotensides, which are soluble or par-ticulate high molecular mass molecules consisting of a polar and an apolar part (for an

overview see [100, 101]). Biotensides improve the use of hydrocarbons mainly by sta-bilisation of hydrocarbon-water emulsions and by facilitating the adherence of cells tothe droplet surface [102]. Much of the work on the production and function of bio-tensides was done with Acinetobacter strains [102]. But also other hydrocarbon-degrading genera, such as Pseudomonas, Achromobacter, Arthrobacter, Brevibac-terium, Corynebacterium, Rhodococcus and Alcanivorax are known to comprise strains,which produce biosurfactants [11, 100, 101, 103]. Biotensides can serve as a carbonsource for other bacteria of a community and so increase cometabolic effects in alkane-degrading bacterial communities.



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Production of biotensides depends, among other factors, on the type of the availablenitrogen source and presumably also on the oxygen concentration in the habitat. In thecase ofAcinetobacter, it has been shown that the biotenside production is accompaniedby the production of a vesicle-like structure and/or by the formation of biofilms towhich bacteria adhere [102, 104, 105]. This biofilm formation has different effects. Onthe one hand, it improves the contact between cells and hydrocarbon and helps the cellsto remain fixed on the soil or sediment particles, which facilitates alkane degradation.On the other hand, cell growth in the biofilm increases its thickness, causing problemsin the bioavailability of gases and nutrients such as dioxygen, N- and P-compounds.This effect slows down alkane degradation.

Pathways of Alkane and Ammonium Oxidation

Oxidation of Alkanes by Aerobic Bacteria

n-Alkanes are saturated, non-polar compounds lacking functional groups. At 20 C andatmospheric pressure, they are gaseous (C1 to C4), liquid (C5 to C16) or solid (> C16).There are also cycloalkanes, which in most cases are liquid, and branched alkanes [2,106]. Alkane-oxidising bacteria can be divided into three groups: the methane-oxidisingbacteria, the bacteria utilising other gaseous alkanes and those growing on the liquidand/or solid alkanes.

Oxidation of Liquid and Solid Alkanes

Oxidation of medium-chain or long-chain alkanes by aerobic bacteria occurs viamonoterminal, biterminal or subterminal pathways (Fig. 2) [107, 108]. The initial oxi-dation of alkanes to the corresponding primary or secondary alcohols is a monooxy-genation involving either a cytochrome P450 [109, 110, 111] (for a review, see [112]),or a rubredoxin-dependent hydroxylase (see section Alkane Oxygenases). n-Alkanolsresulting from terminal oxidation are oxidised via aldehydes to fatty acids. The fattyacids produced by terminal alkane oxidation can enter further metabolic pathways(Fig. 3). Mineralisation proceeds via -oxidation and the tricarboxylic acid cycle, inwhich four reducing equivalents are released per cycle. The reducing equivalents areintroduced into the respiratory chain, which (depending on the predominant redoxpotential in the cell environment) contains distinct electron carriers and uses differentterminal electron acceptors (see section Electron Acceptors for Alkane Degradation). Theelectron transport systems, which are involved in the transformation of the alkanes tofatty acids, are common cell compounds and not specific for the alkane metabolism.Thus far, nothing is known about which metabolic step might be a bottleneck of aerobicalkane mineralisation under oxygen-deficient conditions, although BONIN and BER-TRAND [47] assume that under oxygen limitation, oxygen is used for the initial oxidationof the alkane heptadecane; and is thus lacking as a terminal electron acceptor. Valuesfor the oxygen affinity or kMO values of alkane mono- or dioxygenases have notbeen reported so far. The only exception is methane monooxygenase (MMO):GREEN and DALTON [113] observed a kMO value 16.8 mol O2/l for the soluble MMO of



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Fig. 2. Metabolic pathways for the microbial degradation of n-alkanes: terminal (1),biterminal (2) and subterminal (3) alkane oxidation [108]

Methylococcus capsulatus (BATH ); and JOERGENSEN [114] denotes a kMO value of0.3 mol O2/l for the in vivo activity of MMO of the methanotrophic bacterium strainOU-4-1.

Oxidation of Gaseous Alkanes

Methane oxidation by aerobic methanotrophic bacteria has been well characterised,whereas the bacterial degradation of the other gaseous alkanes (ethane, propane andbutane) has received comparatively less attention. It appears that most of the bacterialisolates oxidising short-chain alkanes belong to the GRAM-positive Corynebacteria,Nocardia sp., Mycobacterium sp. and Rhodococcus (Dietzia) sp., but Pseudomonasstrains have also been reported [115117]. However, the methanotrophs might also beinvolved in the oxidation of short-chain alkanes, since their soluble methane monooxy-genase has an extremely broad substrate specificity and catalyses the oxidation of a



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number of gaseous and liquid alkanes as well as of ammonia (see section MethaneMonooxygenases).

Fig. 3. Diagram of metabolic fluxesThe solid lines represent the carbon fluxes, the dotted lines show the flux of molecularoxygen and of the electron acceptors (adapted from [59]).

For propane-oxidising bacteria, both terminal and subterminal oxidations have beenproposed [115, 117, 118]. Butane was found to undergo terminal oxidation via1-butanol and butyraldehyde to butyrate in Pseudomonas butanovora [116]. The capac-ity to oxidise ammonia in addition to gaseous alkanes has been demonstrated not onlyfor methane-oxidising bacteria, but also for butane-grown bacteria. HAMAMURA et al.[119] examined three butane-degrading bacteria, all of which were able to oxidise am-monia. Pseudomonas butanovora oxidised ammonia to hydroxylamine, while iso-late CF8 (Nocardioides sp.) and Mycobacterium vaccae JOB5 produced nitrite. Theinvestigated bacteria were not able to oxidise methane [119]. This indicates a possible

interaction between ammonia-oxidising and alkane-degrading activity in bacterialcommunities able to degrade gaseous alkanes.

Oxidation of Alkanes by Anaerobic Bacteria

For many decades, alkanes were thought to undergo biodegradation only in the presenceof molecular oxygen. However, anaerobic metabolism of long-chain alkanes in sedi-ments had already been discussed in the 1970s [120]. In the past 10 years, a number of



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bacterial communities [16, 17, 44, 121] as well as isolates [1215, 43, 122], whichdegrade hydrocarbons under strictly anoxic conditions, have been described. Anaerobicalkane degradation has been demonstrated to occur under sulphate-reducing and nitrate-reducing conditions. Recently, strictly anaerobic bacterial formation of CH4 and CO2from long-chain alkanes has been reported [16, 17]. This was assumed to involve(i) acetogenic bacteria which decompose the alkane to acetate and H2, (ii) a group ofarchaea which form CH4 and CO2 from acetate and (iii) another group of archaea con-verting CO2 and H2 to CH4 [16].A key publication by AECKERSBERG et al. [12] describes the isolation of a sulphate-reducing bacterium, designated strain Hxd3 that utilises C12 to C20 alkanes for growth.Later, the moderately thermophilic sulphate-reducing strain TD3 growing on C6 to C16alkanes was isolated by RUETER et al., who also demonstrated that sulphate-reducing

bacteria can utilise aliphatic and aromatic hydrocarbons directly from crude oil underanoxic conditions [13]. C13 to C18 alkanes supported the growth of the sulphate-reducingstrain AK-01 which had been isolated from an estuarine sediment [122]. Another sul-phate-reducer, strain Pnd3, utilised only C14 to C17 alkanes [43]. It appears that eachindividual strain utilises a distinct and, in most cases, narrow range of n-alkanes.Besides the sulphate-reducing strains mentioned above, three denitrifying strains,HxN1, OcN1 and HdN1, have been isolated, which are capable of utilising C6 to C8, C8to C12 and C14 to C20 alkanes, respectively [15]. Strain HxN1 does not grow on aromatichydrocarbons, but it is closely related to the Azoarcus species, which utilise alkylben-zenes. Cell extracts of strain HxN1 contain a protein, whose N-terminus is similar to thesmall subunit (BssC) of benzylsuccinate synthase from denitrifying bacteria that utilisetoluene. This observation could hint at a mechanistic similarity between alkane activa-

tion and toluene activation in anaerobes. Presumably via a radical mechanism, benzyl-succinate synthase catalyses the addition of toluene at its methyl group to fumarate inorder to form benzylsuccinate [18, 19]. Substituted succinates with alkane-derived alkylchains were indeed formed by the denitrifying strain HxN1 oxidising n-hexane and alsoby a sulphate-reducing enrichment culture growing on n-dodecane [19]. Other resultsalso suggest that activation of alkanes may occur by the addition of a carbon compound.During growth with hexadecane and heptadecane, cellular fatty acids in the sulphate-reducing strain Hxd3 were mainly C-odd and C-even, respectively, indicating an altera-tion of the carbon chain length by a C-odd carbon unit during the initial reactions ofalkane metabolism. In contrast, fatty acid analyses of the sulphate-reducer strain Pnd3after growth on alkanes did not indicate an alteration of the carbon chain by a C-oddcarbon unit, suggesting a reaction different from the one in strain Hxd3 [43]. In strain

AK-01, another sulphate reducer, 2-, 4- and 6-methyl-branched fatty acids were identi-fied when the strain was grown on n-alkanes. Isotope-labelling experiments suggestedthat exogenous carbon is added subterminally at the C-2 position of the alkanes. Theoriginal terminal carbon of the alkane thus transforms into a methyl group on the subse-quently formed fatty acid. The carbon addition reaction, however, does not appear to bea direct carboxylation of inorganic bicarbonate; the nature of the carbon compoundadded onto the alkane chain is not yet known [14]. At present, the pathway (or path-ways) of anaerobic alkane degradation and the enzyme(s) involved in alkane activationare subjects of intense research efforts [18, 19].



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Ammonia-Oxidising Nitrifiers

Biochemistry of Nitrification

Ammonium can be used as a source of energy by various autotrophic and heterotrophicnitrifying bacteria. The autotrophic nitrosobacteria oxidise the ammonium to nitrite,while the nitrobacteria oxidise nitrite to nitrate. Ammonia oxidation is initiated by theenzyme ammonia monooxygenase (AMO, see section Ammonia Monooxygenase), whichutilises ammonia (rather than ammonium) and dioxygen as substrates and whichrequires two electrons to reduce one atom of oxygen to water, while the second oxygenatom is incorporated into ammonia to form hydroxylamine. Hydroxylamine is subse-quently oxidised to nitrite by the periplasmic multiheme-enzyme hydroxylamine oxi-doreductase (HAO). Oxidation of hydroxylamine in the periplasm generates the protongradient, which drives ATP synthesis.

AMO: NH3 + 2 H+ + 2 e + O2 NH2OH + H2OHAO: NH2OH + H2O NO2

+ 5 H+ + 4 e

Fig. 4. Electron transfer in Nitrosomonas europaea

Solid lines and dashed lines represent established and hypothetical electron transport pathways,respectively. NADH dh, NADH dehydrogenase; AMO, ammonia monooxygenase; HAO, hydroxy-lamine oxidoreductase; C-P460, cytochrome P460; Q, ubiquinone-8; Cu NiR, copper-containing nitritereductase; NOR, nitric oxide reductase; Cu aa3, aa3-type cytochrome c oxidase; CcM552, membrane-associated cytochrome c552; Cc552, cytochrome c552; Cc554, cytochrome c554. Alternativesubstrates are substrates other than ammonia; their oxidation does not produce hydroxylamine.(Re-drawn from [126]).

The electron acceptor of HAO in N. europaea is cytochrome c554, which possiblydonates electrons to the membrane-associated cytochrome cM552. Cytochrome cM522 car-



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ries the electrons to ubiquinone, thus mediating between the periplasm and the cyto-plasmic membrane [123126]. Two of the four electrons resulting from hydroxylamineoxidation must be routed back to the membrane-bound AMO in order to activate O 2 andto maintain steady-state rates of ammonia oxidation. Ubiquinol is thought to be theelectron donor to the AMO protein. Under steady state conditions, 1.65 of the remainingtwo electrons would pass down the electron transfer chain from ubiquinol to a cyto-chrome aa3 oxidase, presumably via a bc1 complex and periplasmic cytochrome c552;0.35 electrons would be used to generate NADH [124, 126]. It has been proposed thatthe NADH dehydrogenase mediates reverse electron flow from ubiquinol to NAD+[123] (Fig. 4).In the presence of an organic source of energy, ammonia is also oxidised to nitrite (andnitrate) by many heterotrophic bacteria and the methanotrophs [72, 127, 128]. Hetero-

trophic nitrifiers may simultaneously perform aerobic denitrification, converting mostof their oxidation products (nitrite) into the gaseous products N2O and/or N2 [65, 72,129]. However, N2O emission may also directly result from hydroxylamine oxidation,as has been suggested for Alcaligenes faecalis strain TUD [130]. Heterotrophic nitrifi-cation is thought to dissipate rather than conserve energy. The physiological role ofammonia oxidation in heterotrophic bacteria is not fully understood [65, 72, 129,131, 132].Physiological and molecular biological evidence suggests that heterotrophic nitrifierspossess an AMO, which is homologous to the AMO of the autotrophic nitrifiers[127129]. In both cases, ubiquinol is the electron donor for the monooxygenase reac-tion [128]. In contrast, the HAO enzymes from the heterotrophic nitrifiers Pseudomo-nas sp. strain PB16 [133], Alcaligenes faecalis strain TUD [130], Thiosphaera

pantotropha strain LMD82 [132] and Paracoccus denitrificans strain GB17 (Thio-sphaera pantotropha GB17) [134] are entirely different from the HAO of the autotro-phic nitrifier N. europaea. HAO from P. denitrificans (strain GB17) does not containheme groups, but has non-heme, non-iron-sulphur iron centres; the possibility of a binu-clear iron (FeIII-FeIII) centre has been discussed [134]. The catalytic mechanisms of theenzymes must also differ, since in contrast to nitrite formation by the enzyme fromN. europaea, production of nitrite by HAO of P. denitrificans requires molecular oxy-gen. Under anoxic conditions, the P. denitrificans enzyme catalyses the formation ofN2O as the sole product. The first reaction intermediate in the reaction of hydroxy-lamine with the HAO ofP. denitrificans might be a molecule of nitroxyl (NOH) boundto the FeII-FeII binuclear site. Aerobic formation of nitrite is probably the result of thereaction of enzymatically formed nitroxyl with dioxygen, whereas under anoxic condi-tions, two molecules of nitroxyl may combine to N2O, releasing H2O [134].In P . denitrificans (Thiosphaera pantotropha) GB17, the electrons resulting fromhydroxylamine oxidation are assumed to be transferred from the Fe II-FeII site viacytochrome c550 or pseudoazurin to the oxidase [128, 132, 134]. Cytochrome c550 andpseudoazurin can donate electrons to the denitrification enzymes, but not to the quinonepool, thereby forming a possible link between heterotrophic nitrification and aerobicdenitrification [72, 132, 134]. The ubiquinol necessary to restart the ammonia oxida-tion must be produced from the oxidation of the carbon source or from reverse electronflow [128].



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Microaerophilic and Anaerobic Activities of the Autotrophic Nitrifiers

Nitrosomonas europaea and other classical aerobic nitrifiers are known to be able togrow at low oxygen tensions [75, 82, 135, 136]. Growth is accompanied by theproduction of N2O, NO, and/or N2. However, different nitrifying strains may vary withrespect to the gaseous nitrogen compounds generated [82, 135138]. RITCHIE andNICHOLAS [139] described the production of N2O from nitrite by N. europaea cells understrictly anoxic conditions. In principle, N2O and NO can be produced by a variety ofmechanisms [135, 139]. They may occur as products of the reaction catalysed by HAO[139, 140]. On the other hand, a nitrite reductase may catalyse the reduction of nitrite toN2O and NO [135, 139, 140]. Kinetic analysis of the production of N2O by cells ofNi-trosomonas europaea in the presence of14N-ammonia and 15N-nitrite indicated that the

N from nitrite, but neither nitrate, nor ammonium, is incorporated into N 2O [135]. InNitrosomonas europaea, most of the produced NO was due to denitrification of nitriterather than to chemical decomposition of nitrite or ammonium oxidation [138]. Hence,the autotrophic nitrifiers in the presence of a suitable electron donor utilise nitrite as aterminal electron acceptor, i.e., they denitrify under oxygen-deficient conditions.Under oxygen deficiency and/or strictly anoxic conditions, pyruvate [141], hydroxyl-amine [139, 142], hydrazine [138] and molecular hydrogen [137] have been shown tobe utilised as electron donors by Nitrosomonas europaea for nitrite respiration. Underanoxic conditions, even ammonium was an electron donor for nitrite reduction by Nitro-somonas eutropha. Thus, the aerobic nitrifiers catalyse anaerobic ammonium oxida-tion (see section Anaerobic Oxidation of Ammonium: The Anammox Reaction). Ammoniumand nitrite disappeared slowly at a molar ratio of approximately one, but growth ofN. eutropha

was not measurable under these conditions [137]. In contrast, anaerobicgrowth of N. eutropha was observed with ammonia as electron donor and nitrogendioxide (NO2) as electron acceptor [143]. The anaerobic ammonia oxidation rate in thepresence of CO2 and 25 ppm NO2 was approximately 10 times lower than in the pres-ence of air. Ammonia and NO2 were consumed at a ratio of approximately 1:1, and anequivalent amount of NO was produced. Besides NO, nitrite was formed; between 40and 60% of the produced nitrite was denitrified to N2 and, to a lesser extent, to N2O.Hydroxylamine, a typical intermediate of aerobic ammonia oxidation, seems to beformed from ammonia under anoxic conditions as well [143]. If this is the case, thequestion arises as to whether this reaction is also catalysed by ammonia monooxygenase(AMO), and which source the oxygen atom is derived from. It could be argued that thenormal AMO reaction with dioxygen as cosubstrate might have occurred at O 2 partialpressures near zero. However, the dioxygen concentration used in these experimentswas extremely low (less than 100 ppb, 3.1 mol O2/l) [143]. If the available amount ofmolecular oxygen had been used for nitrification, the calculated ammonia oxidationwould not account for the much higher rates of ammonia conversion observed. Thus, ifhydroxylamine formation is indeed catalysed by AMO, the oxygen source for thereaction is unknown. One may assume that nitrite, NO2 and NO could have been used.However, as pointed out by the authors of this study, direct incorporation of oxygenfrom NO2, and especially from NO, into ammonia to form hydroxylamine seemsthermodynamically improbable [143].



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HOOPER et al. [125] considered one possible role of NO in anaerobic ammonia oxidationby the aerobic autotrophic nitrifiers to be a condensation of NO and ammonia,catalysed by AMO or an AMO-like enzyme, leading to the formation of hydrazine,which thereafter could be converted into dinitrogen by HAO (see section AnaerobicOxidation of Ammonium: The Anammox Reaction). The oxidation of hydrazine would yieldthe reducing equivalents required to combine NO and ammonia, or to reduce nitrite toNO.In the anammox process, hydroxylamine is most likely derived from nitrite. Ammoniais thought to be oxidised with hydroxylamine to hydrazine, which is then oxidised to N 2.The oxidation of hydrazine is assumed to provide the reducing equivalents for nitritereduction to hydroxylamine (see section Anaerobic Oxidation of Ammonium: The AnammoxReaction). As mentioned above, externally added hydrazine under anoxic conditions

may also serve as an electron donor for nitrite reduction by Nitrosomonas europaea,resulting in N2O and NO formation from nitrite [138]. However, hydroxylamineformation from nitrite has not been reported for Nitrosomonas.

Anaerobic Oxidation of Ammonium: The Anammox Reaction

In 1977, BRODA published a theoretical paper entitled Two kinds of lithotrophs missingin nature, describing the potential existence of chemolithotrophic bacteria able to oxi-dize ammonia to dinitrogen with dioxygen, nitrate or nitrite as electron acceptor [144].18 years later, MULDER et al. [145] observed that ammonium had disappeared from theeffluent of a denitrifying fluidised bed reactor at the expense of nitrate with a concomi-tant increase in dinitrogen gas production. The observed ammonium loss was explained

by anaerobic ammonium oxidation (anammox). In this obligatorily anaerobic process,it was suggested that ammonium had been oxidised, with nitrate serving as the electronacceptor, producing dinitrogen:

5 NH4+ + 3 NO3

4 N2 + 9 H2O + 2 H+ G0 = 297 kJ per mol NH4

+ [145]

The production of N2 from ammonium and nitrate was verified by feeding 15N-labelledammonium and 14N-labelled nitrate. Mixed-labelled nitrogen gas, 15-14N2, was the domi-nant product (98.2% of the total labelled dinitrogen) [146]. However, from the reactionsuggested by MULDER et al. [145], it was to be expected that 75% of the labelled dini-trogen would be 14-15N2, and 25% would be 15-15N2. The observed labelling percentagesuggested that nitrite, rather than nitrate, is the direct oxidising agent in the anammoxreaction [146]. The free energy balance for this reaction is just as favourable as when O2accepts the electrons:

NH4+ + NO2 N2 + 2 H2O G0 = 358 kJ per mol NH4+

8 NH4+ + 6 O2 4 N2 + 12 H2O + 8 H+ G0 = 315 kJ per mol NH4

+ [146]

After having realised that nitrite rather than nitrate might be the electron acceptor, asynthetic medium was composed for the selective enrichment of anammox microor-ganisms. The anoxic medium containing ammonium and nitrite as the only electron do-nor and electron acceptor, respectively, was fed into a fluidised bed reactor, usingbicarbonate as the only carbon source. On a feed of 30 mM ammonium, conversionrates of 3 kg NH4

+-N/m3 day were reached, equivalent to a specific anaerobic ammo-



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nium oxidation rate of 1,0001,100 nmol NH4+ per h and mg volatile solids. Maximumspecific conversion rates obtained were 1,3001,500 nmol NH4

+ per h and mg volatilesolids [81]. However, instead of the expected 1:1 removal of ammonium and nitritepredicted by the equation ofBRODA [144], a 1.3:1 ratio for the NO2

: NH4+ removal was

found. This appeared to be due to the formation of nitrate (about 10% of the N-feed, or17% of the nitrite, was converted to nitrate). The oxidation of nitrite to nitrate couldgenerate the reducing equivalents necessary for CO2 fixation [81, 82, 147] (Fig. 5).

Fig. 5. Possible anammox pathwayAmmonium is oxidised by hydroxylamine to form hydrazine. Reducing equivalentsderived from hydrazine could reduce nitrite to form hydroxylamine. Nitrate formationfrom nitrite might generate reducing equivalents for CO2 fixation and growth [86, 147].

Labelling experiments with 14CO2 confirmed that the anammox process is autotrophic,and that incorporation of CO2 is dependent on the presence of both ammonium andnitrite. It is interesting to note that in the fluidised bed reactor under conditions ofenrichment for anammox microorganisms, low numbers of aerobic nitrifiers were pre-sent in the sludge at all times, suggesting that they can survive long periods of anaero-biosis [81] (see section Ecology of Ammonium-Oxidising Bacteria and section Microaero- philoic and Anaerobic Activities of the Autotrophic Nitrifiers). However, even if theiranaerobic nitrification rate were the same as under oxic conditions, their activity wouldbe orders of magnitudes too low to explain the observed ammonium oxidation rate of800 nmol NH4

+ per h and mg volatile solids [81]. From the rates of anaerobic ammo-nium oxidation reported from several experiments, it is evident that the specific rates for

anaerobic ammonium oxidation byNitrosomonas

cultures are about 25-fold lower thanthe rates observed in the anammox process [82].The anammox pathway has been verified using 15N-labelled nitrogen compounds [146,147]. It has been proposed that ammonium is biologically oxidised by hydroxylamine toform hydrazine, which is then oxidised to dinitrogen [147]. Since the anammox processis only possible under strictly anoxic conditions and is strongly inhibited by dioxygen,an AMO-like hydroxylamine formation seems improbable. Other mechanisms for theformation of hydroxylamine include the incomplete reduction of nitrite by a cytochromec-type nitrite reductase. The reducing equivalents for the reduction of nitrite to hydro-



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xylamine are presumably generated in the conversion of hydrazine to N2 (Fig. 5) [147,148]. The observation by STROUS et al. [149] that nitrite inhibition of the anammoxprocess could be overcome by adding trace amounts of hydrazine or hydroxylamineconfirms that hydrazine is an intermediate in the anammox reaction.HAO from the aerobic nitrifier N. europaea is capable of catalysing the oxidation ofhydrazine, presumably to N2 [125, 150]. It can therefore be assumed that similar reac-tions (and similar HAO enzymes) are operative in nitrite respiration with hydrazine aselectron donor and in the anammox process. A novel type of HAO which catalyses theoxidation of hydroxylamine and hydrazine, and which, in terms of its amino acidsequence of several peptide fragments seems to be quite different from the HAO ofN. europaea, was purified from an anammox enrichment culture. However, similar tothe Nitrosomonas HAO, but different from the HAO of the heterotrophic nitrifiers (see

section Biochemistry of Nitrification), the enzyme from the anammox culture containedseveral c-type cytochromes [151]. HAO was found to be present inside a cytoplasmicmembrane-bounded structure, the anammoxosome [152].

Enzymes Catalysing Alkane and Ammonia Oxidation

Alkane Oxygenases

The first step in the bacterial oxidation of medium- and long-chain alkanes to the corre-sponding alcohols requires molecular oxygen and is usually catalysed either byP450 monooxygenases (for a review, see WONG [112]), or by multiprotein monooxy-genase systems. The OCT plasmid-encoded alkane monooxygenase from Pseudomonas

oleovorans, which catalyses the oxidation of medium-chain-length alkanes from hexaneto dodecane, has been studied with great intensity. It is a three-component enzymesystem consisting of a soluble NADH-rubredoxin reductase (AlkT), a solublerubredoxin (AlkG) and a particulate oxygenase component AlkB referred to as - oralkane hydroxylase [153156].The oxygenase component AlkB is an integral membrane protein which is assumed tocontain six transmembrane segments and two hydrophilic loops which, as well as theN-terminus and a large C-terminal domain, are located in the cytoplasm. Only threevery short loops appear to be exposed to the periplasm [157]. AlkB contains apresumably oxo-bridged diiron(III) cluster similar to the one found in the hydroxylasecomponent of soluble methane monooxygenase (sMMO) [158]. However, despite theapparent spectroscopic similarity of the hydroxylase components of sMMO and AlkB,

the AlkB metal centre is distinct from the carboxylate-rich diiron site of the sMMOhydroxylase. AlkB contains a conserved set of eight histidines common to nonhemeintegral-membrane desaturases and hydroxylases. These histidines are required foractivity in at least one member of this family [158] and might serve as ligands to thediiron cluster.The alkane hydroxylase, besides carrying out n-alkane hydroxylation, also catalyses theoxidation of substituted cyclic alkanes, the hydroxylation of the alkyl substituent ofethylbenzene and several other alkylbenzenes, stereospecific epoxidation of terminalalkenes, oxygenative O-demethylation of methyl ethers, oxygenative formation of alde-



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hydes from terminal alcohols and the stereoselective sulphoxidation of methyl thioethersubstrates [155, 159]. It has a substrate specificity, which is rather low, but showsconsiderable regio- and stereospecificity [159].The genetics of an alkane monooxygenase system from the dodecane-degradingstrain Acinetobacter calcoaceticus ADP1 has been studied by HILLEN and co-workers[160163]. Similar to the P. oleovorans enzyme, this monooxygenase system is thoughtto contain, besides the hydroxylase, a rubredoxin and a rubredoxin-reductase [160, 163].The hydroxylase component (AlkM) also contains the eight-histidine motif that ischaracteristic of nonheme integral-membrane hydroxylases [162]. However, the geneticorganisation of the genes which encode proteins involved in alkane oxidation differmarkedly in the OCT plasmid of P . oleovorans and in A . calcoaceticus ADP1[160163].

In 1988, FINNERTY [164] postulated a unique pathway ofn-alkane degradation via ann-alkyl hydroperoxide, which is metabolised via the corresponding peroxy acid andaldehyde to a carboxylic acid and an alcohol [164, 165] (Fig.6). The first step in thishypothetical pathway could be catalysed by a dioxygenase. Actually, an alkane-oxidising enzyme which is independent of reducing equivalents and which requiresdioxygen as the sole cosubstrate was purified from Acinetobacter sp. strain M-1. It is ahomodimeric, Cu2+-dependent flavoprotein, which accepts C10 to C30 alkanes as sub-strates. The enzyme presumably catalyses the formation of n-alkyl hydroperoxide,which was observed as an unstable, transient oxidation product in cell extractsmediating alkane oxidation. The detection of this enzyme supports the FINNERTY path-way; however, the stoichiometry of n-alkane oxidation catalysed by the presumeddioxygenase could not be established due to the instability of the product [165].Different Acinetobacter strains have been shown to use different strategies for alkane

degradation. Multi-component alkane monooxygenase, cytochrome P450 and thedioxygenase activity catalysing long-chain alkane oxidation in strain M-1 are alternativesystems.In contrast to the diiron alkane hydroxylase of P. oleovorans and Acinetobacter sp.strain ADP1, butane monooxygenase from Nocardioides sp. strain CF8 (a heterotrophicnitrifier) has been suggested to be a copper-containing monooxygenase similar topMMO (see section Methane Monooxygenases) and AMO (see section Ammonia Monooxy-genase) [79]. However, there are apparently different types of butane monooxygenasesystems in different butane-utilising bacteria [119]. Notably, AMO and both sMMO andpMMO catalyse the oxidation of butane (and propane), although these alkanes do notsupport growth of ammonia-oxidising bacteria or methanotrophs. With MMO andAMO, both terminal and subterminal oxidations of short-chain alkanes were observed

(see sections Methane Monooxygenases and Ammonia Monooxygenase). However, geneticdata on butane monooxygenases, which would allow for comparative sequence analysisare not yet available.

Anaerobic Oxidation of an Alkyl Substituent: Ethylbenzene Dehydrogenase

Direct oxidation not only of alkanes, but also of alkyl substituents was previouslythought to generally require molecular oxygen. Recently, KNIEMEYER and HEIDER [166]



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purified an ethylbenzene dehydrogenase from the denitrifying Azoarcus sp. strainEbN1, which catalyses the dioxygen-independent hydroxylation of ethylbenzene to(S)-1-phenylethanol (Fig. 7). This is the first enzyme known to oxidise a nonactivatedhydrocarbon without using molecular oxygen as cosubstrate. The enzyme was identifiedas a periplasmic molybdenum/iron-sulphur/heme b protein consisting of three subunits( structure). It also shows activity with n-propylbenzene, but does not catalyse theoxidation of toluene [166]. The reaction involves the introduction of an oxygen atomderived from water into the substrate with concomitant reduction of an electronacceptor. This type of reaction is characteristic of the molybdenum enzymes containinga molybdenum pyranopterin cofactor [167170].

Fig. 6. The FINNERTY pathway for long-chain n-alkane oxidationIn Acinetobacter sp. strain HO1-N, alkane oxidation to the corresponding alcohol andfatty acid is presumed to occur via alkane hydroperoxide, peroxy acid and aldehyde[164, 165].



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There are apparently different molybdenum- and iron-sulphur-containing ethylbenzenedehydrogenases in different strains ofAzoarcus, since in cell extracts of strain EB-1, anethylbenzene dehydrogenase activity was detected that is p-benzoquinone-dependentand membrane-bound [171]. It is a heterotrimeric protein related to molybdenumenzymes of the DMSO reductase family containing several [4Fe4S] clusters and amolybdenum pyranopterin cofactor. Based on sequence analysis, the subunit isthought to be a membrane anchor, whereas the and subunits contain the molybde-num pyranopterin cofactor and iron-sulphur clusters, respectively [172].

Fig. 7. Hydroxylation of ethylbenzene to (S)-1-phenylethanol catalysed by ethylbenzenedehydrogenase

Methane Monooxygenases

Methane is wide-spread in nature. Methane monooxygenase (MMO), which catalysesthe first step in the oxidation of methane by methanotrophs, exists in two forms: the

cytoplasmic, soluble methane monooxygenase (sMMO); and the membrane-bound,particulate methane monooxygenase (pMMO).Soluble MMO has extremely broad substrate specificity and catalyses the adventitiousoxidation of saturated, unsaturated, linear, branched and cyclic hydrocarbons up toapproximately eight carbons. CO, ammonia, aromatic compounds, heterocycles, halo-genated alkenes and ethers are also oxidised [173178]. Not all methanotrophs possesssMMO. The sMMO enzymes from Methylococcus capsulatus (BATH) and Methylosinustrichosporium OB3b have been the subjects of intense research efforts (for reviews, see[179182]). sMMO consists of three components: a 251 kDa hydroxylase; a 15.8 kDaprotein B or coupling protein which has a regulatory function; and a 38.5 kDareductase component (protein C). Multi-component monooxygenases exhibitingsequence similarities to sMMO are the toluene monooxygenases, phenol hydroxylase

and alkene monooxygenase [183]. The hydroxylase component has an 222 configu-ration. Its subunit contains a diiron centre, which is the site of catalysis. The structureof the carboxylate-bridged diiron centre has been solved by X-ray crystallography. Bothirons are bridged by two OH (or one OH and one H2O), as well as both carboxylateoxygens of E144 (bis-hydroxo-bridged diamond core structure) [184186]. How-ever, the number of water/hydroxo/oxo ligands coordinated to the site can alternatebetween different forms, allowing the site to generate new open coordination sites onthe diiron centre, where dioxygen and substrate can interact [185]. The regulatory pro-tein B appears to play an essential role in facilitating electron transfer between the



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reductase and the hydroxylase components and in accelerating the formation of severalcatalytic intermediates [187189]. The two formal redox potentials of the complex ofhydroxylase and component B are substantially more negative than the values for thehydroxylase [190, 191]. Protein C is a [2Fe2S]- and FAD-containing reductase compo-nent that transfers electrons from NADH to the diiron sites of the hydroxylase. Thereductase also influences the midpoint redox potential of the hydroxylase, raising theredox potential for the second electron transfer to the diiron cluster [179, 190, 192].This provides a larger driving force for electron transfer from the reductase and favoursthe transfer of two rather than one electron, as required [192]. Upon formation of thecomplex of a hydroxylase dimer and two copies of each the reductase and protein B, thehydroxylase undergoes a dramatic conformational change which allows for close inter-action between the active centres of the reductase and hydroxylase components and

which affect the environment of the diiron centre [187]. Thus, dynamic binding interac-tions of the hydroxylase with both the reductase and component B during catalysisinfluence rate and efficiency of the sMMO-catalysed methane hydroxylation [187].The exact identity of some intermediate species and the precise details of the mecha-nism of dioxygen activation and substrate oxidation by sMMO are currently subjects ofdebate [180, 182, 193196]. Several possible mechanisms have been suggested [182,193, 197]. One unresolved issue is whether the hydroxylation mechanism involves theformation of discrete radicals [179, 195, 198, 199] or whether a non-radical pathwayoccurs [193, 200202].In contrast to sMMO, which is expressed under low-copper growth conditions andwhich is not present in all methanotrophs, pMMO is expressed under growth conditionsin which the copper-to-biomass ratio is high; and it is seemingly present in all metha-notrophs. The expression of pMMO is accompanied by the extensive formation of intra-

cytoplasmic membranes, where the pMMO resides. Compared to sMMO, pMMO ismuch less characterised, mainly due to the difficulty of studying membrane-bound pro-teins as well as the fact that pMMO, once solubilised, is sensitive to oxygen and light[203, 204], similar to AMO. pMMO from Methylococcus capsulatus (BATH) consists ofthree subunits of 45, 26, and 23 kDa [205]. In its active form, the enzyme prepared byZAHN and DISPIRITO [203] contained 23 iron and approximately 15 copper atoms permol. ZAHN and DISPIRITO suggested that the catalytic site might involve both iron andcopper. However, pMMO purified from the same organism by NGUYEN et al. [204]contained only copper (1215 Cu ions per heterotrimer). The copper ions in pMMO arepresumed to group into two distinct classes which play a role in either electron transfer(weakly bound Cu(I) ions) or catalysis (Cu(II) ions) [204]. pMMO purified from Methy-losinus trichosporium OB3b contained 13 copper atoms and 1 iron atom per molecule.

Based on EPR data, the iron atom was thought to be contained in the active site; thecopper was identified as a type 2 Cu2+ centre [205]. EPR parameters of pMMO in Methylomicrobium albumBG8 and Methylococcus capsulatus (BATH) were also charac-teristic of a type 2 Cu2+, arranged in a square planar or square pyramidal configuration[204, 206]. The Cu2+ is presumably bound to four nitrogens, and three or four of thesenitrogens are histidine imidazoles [207]. However, the structure of the metal site(s) inpMMO is still a subject of speculation [204, 206, 207].It is interesting to note that, similar to the amoCAB genes encoding AMO (see sectionAmmonia Monooxygenase), the pmoCAB genes encoding pMMO are present twice in the



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genome of Methylococcus capsulatus (BATH), and that, also similar to amoC, a thirdcopy ofpmoC has been identified [208, 209]. In M. trichosporium OB3b and Methylo-cystis sp. strain M, two virtually identical copies of pmoCAB were present, but thereseems to be no third copy of pmoC[209]. The role of these multiple gene copies has notbeen well understood in either methanotrophs or nitrifiers. In M. capsulatus (BATH),both sets of the pmoCAB genes seem to be functionally equivalent, but their relativeexpression alters depending on the copper concentration [210]. Comparison of the pmoand amo genes indeed suggests that pMMO and AMO may be evolutionarily related(4287% identity at the amino acid level) [181].In contrast to AMO and sMMO, which exhibit broad substrate specificity, pMMO hasrelatively narrow substrate specificity, catalysing the oxidation of alkanes and alkenesup to five carbons, but not oxidising aromatic compounds or cyclohexane [176].

Ammonia Monooxygenase

The extreme instability of ammonia monooxygenase (AMO) activity from Nitroso-monas europaea [211, 212] has so far precluded purification of catalytically activeprotein; as a result, little is known regarding its structure, enzymatic mechanism, or theidentity and function of its cofactors. AMO is a membrane protein. As predicted by theamino acid sequence, the active site-containing subunit A consists of five transmem-brane segments and a large periplasmic loop. Subunit B has two transmembranedomains and two large periplasmic domains [125]. Indirect evidence indicated afunctional role for copper in AMO of the autotrophic nitrifiers [211213]. However, theAMO activity from the heterotrophic nitrifier Paracoccus denitrificans (Thiosphaera

pantotropha) is apparently more stable than AMO from N. europaea; MOIR et al. [214]succeeded in purifying a catalytically active enzyme from this heterotrophic nitrifier.The purified AMO is composed of two subunits and presumably contains a labilecopper centre.Particulate MMO from Methylococcus capsulatus (BATH), which is a copper protein(see section Methane Monooxygenases), and the AMOs from N . europaea andP. denitrificans have a remarkably similar susceptibility to inhibitors, including copper-chelating agents [212, 214].An interesting property of AMO ofN. europaea is its broad substrate specificity, whichhas been deduced largely from the activities in whole cells. It catalyses the oxidation ofa variety of nonphysiological substrates including methane, methanol, CO, alkanes (upto C8), cyclohexane, alkenes (up to C5), halogenated alkanes, halogenated alkenes, and

a number of aromatics (for a review, see [125]). When the hydrocarbons were longerthan ethane, both 1-ol and 2-ol oxidation products were observed. The alkenes gave riseto either epoxides or unsaturated alcohol products [91]. Chloromethane was oxidised toformaldehyde [92]. For monohalogenated ethanes, acetaldehyde was the major organicproduct, indicating that the halogenated carbon was the preferred position for oxidation.In contrast, the major products of chloropropane oxidation were the alcohols 1-chloro-2-propanol and 3-chloro-1-propanol, followed by propionaldehyde. For n-chlorinatedalkanes, the rate of dechlorination was greatest with chloroethane, decreasing in the caseof chloropropane and chlorobutane [92].



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With aromatic substrates, AMO of N. europaea catalyses alkyl substituent hydroxyla-tions, styrene epoxidation, ethylbenzene desaturation to styrene, and aniline oxidation tonitrobenzene. Alkyl substituents were the preferred oxidation sites, but the ring was alsooxidised to produce phenolic compounds. Some halobenzenes were oxidised to halo-phenols [94].The products of the oxidation of substrates other than ammonia are not assimilated byN. europaea. Since aliphatic halogenated hydrocarbons are industrial chemicals consid-ered to be priority pollutants (U.S. Environmental Protection Agency) [92]; theiroxidation and/or dehalogenation by AMO is of considerable interest. AMO, in fact,could transform many pollutant chemicals that may contaminate soils and aquifers. Theubiquity of nitrifying bacteria may facilitate the biodegradation of these and othercompounds (such as aromatics), since the activity of AMO may provide intermediatesfor heterotrophic bacteria. Nitrifiers may even be useful in bioremediation applications.As mentioned above, AMO is similar to pMMO in cellular location, inhibitor profile,and the putative involvement of copper. However, the hydrocarbon substrate range ofAMO appears to be more similar to that of sMMO rather than to that of pMMO [125].N. europaea contains two nearly identical copies of the operon amoCAB, which encodesthe AMO enzyme [215218]. There is a third copy of amoC (amoC3), which is notassociated with amoA or amoB and which reveals an approximately 60% similarity tothe other copies ofamoC [215]. The hao gene and the cyc or hcy gene are also presentin three copies in N. europaea [216]; cyc/hyc encodes cytochrome c554, an electronacceptor of HAO. Mutagenesis experiments have shown that the both amoCAB operonsare expressed at different levels and that no single copy of amo or hao is essential [218,219]. In the absence of ammonium, a single 1.1 kb transcript derived from amoC1,2 wasobserved, whereas in the presence of ammonium, both the 1.1 kb transcript and a 3.5 kb

transcript representing the full amoCAB operon were detected [216]. Synthesis ofmRNA detected by hao probes also required the presence of ammonium [220]. Sincethe two operons ofamo are expressed at different levels, it had been assumed that theyare transcribed from different promoters. However, the DNA sequences of the twoputative promoters were found to be identical; therefore, they cannot account for theobserved transcriptional differences [216].Most am o sequences known are from -Proteobacteria. The AMO polypeptides of Nitrosomonas, Nitrosolobus and Nitrosospira are highly homologous [72]. The am ogenes ofNitrosococcus oceani and Nitrosococcus sp. strain C-113 (-Proteobacteria)were 8088% identical to each other, 4953% identical to the pmo genes which encodethe pMMO ofMethylococcus capsulatus (BATH) and only 3942% identical to the amogenes of the -subdivision autotrophic ammonia-oxidising bacteria [221].

Concluding Remarks

An attempt has been made in this paper to discuss the complexity ofn-alkane metabo-lism by bacteria in natural habitats ranging from oxic to anoxic conditions; the enzymesystems involved in alkane activation are also described. As bacterial metaboliccapabilities in natural habitats are the result of a very complex web of cooperations, wefocused not only on bacteria that use n-alkanes as a carbon and energy source, but alsoon bacteria which fortuitously oxidise n-alkanes.



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Alkanes are widespread in the biosphere, and it is known that n-alkane-degrading bacte-ria are present in all kinds of aquatic as well as terrestrial habitats. Both aerobic andanaerobic alkane degradation have been described, but thus far, only few publicationshave focused on alkane degradation in the transition from oxic to anoxic conditions.There is not enough information to decide whether there is a threshold value for oxygenat which enzymes of the aerobic alkane metabolism are switched off and anaerobicalkane metabolism starts, or whether there are enzymes with a very high oxygen affinitythat allow for special microaerobic alkane metabolism at suboxic to quasi-anoxicconditions.In most studies, pure cultures are used to investigate metabolic routes. Indeed, this clas-sical approach permits the detailed biochemical and genetic characterisation ofpathways and enzyme systems of alkane degradation. However, in natural habitats,

microorganisms live in communities, whose metabolic capacities are characterised byintensive cooperation among the community members. In addition to microorganismsthat utilise n-alkanes as a source of carbon and energy, there are also other bacteriawhich co-metabolically oxidise alkanes, contributing to alkane degradation.Distinct types of oxygenases catalyse the first step in the alkane oxidation by aerobicbacteria. The mechanisms of anaerobic alkane activation are completely different fromthose of aerobic pathways, involving a radical reaction of the hydrocarbon with fu-marate, or maybe other modification reactions. This, of course, raises the questions ofhow alkane-degrading bacteria cope with extremely low oxygen concentrations, andwhich alkane activating mechanism(s) they use. Hexadecane-degrading bacterial com-munities from intertidal sediment have been demonstrated to adapt to extreme limita-tions in the oxygen supply. These communities showed only minor changes in their

composition after the oxygen supply had been reduced from oxic to suboxic conditions,but showed considerable changes in the transition from suboxic to quasi-anoxic condi-tions. This suggests that, within a community, several parallel biochemical pathwaysexist. Some of these pathways may be performed by species, which possess alkane oxy-genases with a high oxygen affinity, as is described by KUKOR and OLSEN in the case ofthe catechol 2,3-dioxygenase of the Pseudomonas isolate PKO1 [222]. Others may pos-sess pathways which fortuitously co-oxidise alkanes and thus generate substrates forheterotrophic bacteria. Alternative pathways might also be possible but have not beencharacterised so far. However, in bacterial communities which contain ammonium-oxi-dising and methanotrophic bacteria, the enzymes, ammonia monooxygenase and meth-ane monooxygenase, which both have very low kMO values, might be involved in alkaneoxidation at low oxygen concentrations. These enzymes fortuitously co-oxidise alkanesto products which might readily be utilised by other members of the community.The investigation of the pathways and of the enzymes of alkane metabolism in the tran-sition from oxic to anoxic conditions signifies a contribution to a comprehensiveunderstanding of how microorganisms adapt to distinct environmental conditions.

Received 8 October 2001Received in revised form 15 April 2002Accepted 8 May 2002



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