Survey
* Your assessment is very important for improving the workof artificial intelligence, which forms the content of this project
* Your assessment is very important for improving the workof artificial intelligence, which forms the content of this project
Bacterial cell structure wikipedia , lookup
Microorganism wikipedia , lookup
Human microbiota wikipedia , lookup
Disinfectant wikipedia , lookup
Marine microorganism wikipedia , lookup
Magnetotactic bacteria wikipedia , lookup
Phospholipid-derived fatty acids wikipedia , lookup
FEMS Microbiology Reviews 63 (1989) 235-264 Published by Elsevier 235 FEMSRE 00124 Microbial metabolism of short-chain unsaturated hydrocarbons S. Hartmans, J.A.M. de Bont and W. Harder 1 Division of Industrial Mierobiology, Department of Food Seience, Agricultural University, Wageningen, and 1 Department of Microbiology, State University of Groningen, Haren, The Netherlands Received 10 February 1989 Accepted 17 April 1989 Key words: Ethene; Acetylene; Alkenes; Epoxides 1. INTRODUCTION The microbial degradation of hydrocarbons has been studied extensively during the last decades. In general, reviews of such studies grouped the hydrocarbons in classes of compounds for collective discussion. Microbial degradation of aromatic hydrocarbons has been reviewed several times [1,2] which is in keeping not only with the often detailed physiological and genetic studies undertaken but also with the importance of the mineralization of these compounds in the carbon cycle in nature. Aliphatic long chain hydrocarbons have mostly been dealt with as a distinct group of hydrocarbonous compounds that enter the environment through unintentional and undesirable spillage from the petrochemical industries [3]. They have also been considered as cheap substrates in biotechnological processes [4]. Methane utilization is generally treated as a specific topic, mainly because the methane-utilizing bacteria comprise a distinct group of organisms with rather specific biochemical characteristics [5]. Terpenes have also been reviewed as a separate class of compounds. Correspondence to: S. Hartmans,Division of Industrial Microbiology, Departmentof Food Science,AgriculturalUniversity, P.O. Box 8129, 6700 EV Wageningen,The Netherlands. These natural products have practical applications especially in the flavour and fragrance industries and their potential as substrates in biotechnological processes employing micro-organisms as biocatalysts has been reviewed [6]. Until recently, the microbial metabolism of unsaturated short chain hydrocarbons received considerably less attention. Nonetheless, this group comprises the important naturally occurring plant hormone ethene (ethylene). Furthermore, many of the gaseous alkenes and alkynes are used extensively in the petrochemical industry. The recent interest in organisms degrading gaseous alkenes and alkynes stems mainly from applied aspects. The organisms involved may be employed either in the treatment of waste gases or in the biotechnological production of epoxides. However, the ecological implications are also important. The fate of gaseous hydrocarbons is studied with increased interest because of their contribution to the production of carbon monoxide and their involvement in the production and degradation of ozone [7]. The microbial degradation of the heterogeneous group of short chain unsaturated hydro-. carbons and its implications has received only limited attention in reviews on other classes of hydrocarbons [8,4]. The intention of the present article is to focus on the short chain unsaturated 0168-6445/89/$03.50 © 1989 Federationof European MicrobiologicalSocieties 236 h y d r o c a r b o n s . It will c o n c e n t r a t e on the b i o d e g r a d a t i v e routes e m p l o y e d b y the various microo r g a n i s m s involved a n d b o t h a p p l i e d a n d ecological aspects will b e considered, 2. P R O P E R T I E S , S O U R C E S A N D SHORT-CHAIN OLEFINS SINKS OF Some relevant p r o p e r t i e s of the h y d r o c a r b o n s u n d e r review have b e e n c o m p i l e d in T a b l e 1. Several c h l o r i n a t e d alkenes are also i n c l u d e d because of their extensive use as i n d u s t r i a l solvents. F r o m the table it is obvious t h a t all c o m p o u n d s are gaseous or volatile at a m b i e n t c o n d i t i o n s with the exception of some of the c h l o r i n a t e d c o m p o u n d s . G a s e o u s s u b s t r a t e s which are p o o r l y solu- Table 1 Physical properties of selected olefins Compound Ethyne (acetylene) Ethene (ethylene) Propene (propylene) 1-Butene cis-2-Butene trans-2-Butene Butadiene Isoprene (2-methylbutadiene) Styrene (phenylethene) Chloroethene (vinyl chloride) 1,1-Dichloroethene (vinylidene chloride) Boiling point ( o C at 1 Atm) Explosion limits in air (V//V%)b - 84.0 1.15 c 2.4-83 - 103.7 9.5 c 2.7-34 - 47.6 -6.3 - 138.9 - 105.5 - 4.4 8.9 c 10 c 4.5 c cis-l,2-Dichloroethene trans- l,2-Dichloro- ethene Trichloroethene (trichloroethylene) Tetrachlorethene (perchloroethylene) a mol m Partioning coefficient (air/water at 30 ° C) a 3 gas phase (mol 34.1 2.0-11.1 1.6-10 1.4-16.3 - 1.0- 9.7 145.2 0.2 d 1.1- 8.8 - 13.4 1.26 e 3.6-31 37 60.3 1.25 e 0.19 e 6.5-15.5 _ 47.5 0.46 e _ 87 0.49 e 121.1 0.91 e m -3 - water p h a s e ) - 1 ; b [12]; c [13]; d determined experimentally; e [14]. b l e in w a t e r offer c e r t a i n a d v a n t a g e s w h e n studying their m i c r o b i a l m e t a b o l i s m . A t low s u b s t r a t e c o n c e n t r a t i o n s a n d at low cell densities gas chrom a t o g r a p h i c h e a d - s p a c e analysis a f f o r d s very sensitive, r a p i d a n d reliable m o n i t o r i n g of biological processes in c o m p a r i s o n w i t h o t h e r s u b s t r a t e s that dissolve well in water. H o w e v e r , the d i s a d v a n t a g e s of using gaseous r a t h e r t h a n solid or liquid substrates have b e e n m o r e o b v i o u s to microbiologists. A s a consequence, research on the m e t a b o l i s m of gaseous s u b s t r a t e s has b e e n p e r f o r m e d in only a few research groups. W o r k in the field of gaseous c o m p o u n d s therefore lags b e h i n d that involving the m o r e usual substrates. A m i c r o b i o l o g i s t is g e n e r a l l y m o r e f a m i l i a r with h a n d l i n g glucose t h a n with h a n d l i n g a s u b s t r a t e such as ethene. A d i s a d v a n t a g e of a p o o r l y w a t e r - s o l u b l e gaseous substrate is the special a t t e n t i o n t h a t needs to be given to the kinetics of gas transfer to the a q u e o u s phase. T h e i n c u b a t i o n systems used o b v i o u s l y need special c o n s i d e r a t i o n s , not o n l y to keep the substrate a v a i l a b l e to the o r g a n i s m s b u t s o m e t i m e s also to allow r e m o v a l of gaseous r e a c t i o n p r o d u c t s such as c a r b o n dioxide. A n o t h e r i m p o r t a n t aspect is that certain m i x t u r e s o f gas a n d air m a y explode. E t h e n e in air for i n s t a n c e forms explosive m i x t u r e s in c o n c e n t r a t i o n s b e t w e e n 2.7% a n d 34%. Nevertheless, m i c r o - o r g a n i s m s have not o n l y been grown on gaseous s u b s t r a t e s in sealed b a t c h cultures b u t also in o p e n c h e m o s t a t cultures. W o r k with c h e m o s t a t cultures using gaseous substrates has m a i n l y b e e n p e r f o r m e d with m e t h a n e - u t i l i z i n g b a c t e r i a [9,10] a n d m o r e recently also with ethenea n d p r o p e n e - u t i l i z i n g b a c t e r i a [11]. T h e c o m p o u n d s listed in T a b l e 1 are p r e d o m i n a n t l y of a n t h r o p o g e n i c origin with the n o t a b l e e x c e p t i o n of ethene a n d isoprene. E t h e n e is a special a n d i n t e r e s t i n g case. It is a simple, rather reactive m o l e c u l e t h a t can u n d e r g o various reactions. F o r this r e a s o n it is p r o d u c e d b y m a n in massive q u a n t i t i e s f r o m fossil reserves a n d is used as a b u i l d i n g b l o c k for synthesis in the p e t r o c h e m ical industry. E t h e n e also p l a y s an u n e x p e c t e d l y i m p o r t a n t role in p l a n t physiology. It is a gaseous h o r m o n e t h a t acts o n all p l a n t s a n d is not only p r o d u c e d b y p l a n t s b u t also b y b a c t e r i a a n d fungi. T h e v a r i o u s effects o f ethene o n p l a n t s have b e e n s t u d i e d a n d d e s c r i b e d in detail [15,16] b u t the 237 molecular basis for the observed phenomena is still unresolved [17]. It remains to be shown how this simple unsaturated hydrocarbon exerts its drastic effects on plants at concentrations well below 1 ppm. The relative contributions to ethene emissions into the atmosphere from biogenic and anthropogenic sources have been estimated by Sawada and Totsuka [7]. Biogenic sources taken into account were both plants and micro-organisms from terrestrial as well as aquatic ecosystems. The conclusions of Sawada and Totsuka are that most ethene released is of biogenic origin (74%), while the anthropogenic ethene (26%) originates mainly from burning biomass and to a lesser extent from the combustion of coal and fuel oil [7]. The total amount of ethene released into the atmosphere by both biogenic and anthropogenic sources was estimated at 35 x 1 0 6 tonnes per year which is approximately half the quantity produced and processed annually by the petrochemical industry. The volatile compound isoprene is also excreted by many plants [18]. Emission rates are dependent on temperature and light intensity and can account for 0.1 to 2% of the carbon fixed during photosynthesis by oak plants [19]. Isoprene emissions from natural vegetations have also been quantified and were shown to be significantly higher than the contribution from forest fires [20]. In contrast to the situation for isoprene and ethene, the other compounds in Table 1 are predominantly of anthropogenic origins. The sources of such compounds sometimes are directly linked to a specific human activity. For instance chlorinated hydrocarbons are used on a large scale as solvents. Many of these toxic compounds, including chlorinated ethenes, are persistent in the environment and consequently are widely distributed in ground water. In industrial areas concentrations up to 100 ~tg per 1 have been detected [21,22]. Recently however, there have been several reports on the biotransformation of chlorinated ethenes which will be discussed in more detail in sections 4.7 and 6.1.2. The gaseous, non-chlorinated compounds may also be attributed to specific sources. For example, more than 90% of the acetylene generated originates from automobile exhaust, making it a useful tracer gas in air pollution studies [23]. The atmospheric concentrations of the non-methane hydrocarbons ( N M H C ) have been monitored several times at different locations. In urban areas the N M H C - c o n c e n t r a t i o n was in the range of 250-1000 ppbC while concentrations in samples from rural locations were 10-100 fold lower. At both types of locations parafins are the major class of compounds detected while olefins were usually present in the range of 6-12% [24,25]. In several rural air samples ethene and isoprene were the major olefins detected [24] although in other cases isoprene was not present in sufficient quantities to guarantee correct identification [25]. Short-chain unsaturated hydrocarbons have a short atmospheric lifetime. Ethene is predominantly destructed in the troposphere by reactions with O H radical or with ozone, while only a small percentage diffuses into the stratosphere [7]. It is unlikely that the non-methane hydrocarbons from anthropogenic sources that are released into the atmosphere will affect or will be affected by micro-organisms in either terrestrial or aquatic ecosystems. Nevertheless, as will be discussed in section 4, several bacterial species have the ability to completely metabolize these compounds. In section 5 some considerations will be presented which could explain these metabolic capabilities. An important part of the reasoning will be that traces of the so called anthropogenic volatile hydrocarbonS are also formed by microbial activities. 3. B I O T R A N S F O R M A T I O N OF UNSATURATED HYDROCARBONS: GENERAL ASPECTS Unsaturated hydrocarbons m a y be metabolized by an initial attack on the unsaturated moiety of the molecule (Fig. 1) or by an attack elsewhere on the molecule. The following reactions may be observed at the unsaturated moiety of the molecule: (i) oxidation with hydrogen peroxide and halide ions by haloperoxidases, (ii) oxidation by mono-. oxygenases using molecular oxygen as oxidant, (iii) addition of water to the unsaturated bond and (iv) reduction of the alkene or alkyne. T h e addition of ammonia to an unsaturated carbon- 238 0 /\ R ---HC--CH--R' R--HC~CH--R'~ OH H I I R--HC--CH--R' R-- H2C--CH2--R' Fig. 1. Biotransformations of the alkenic bond. carbon bond, e.g. the formation of aspartate from fumarate and ammonia by aspartate ammonialyase, will not be discussed. 3.1. Oxidation of unsaturated carbon-carbon bonds 3.1.1. Oxidation by haloperoxidases Haloperoxidases c a t a l y z e the formation of a,fl-halohydrins from alkenes [26] and a-halogenated ketones from alkynes [27] in the presence of halide ion and hydrogen peroxide. In the absence of halide ions, the action of a chloroperoxidase may also result in the formation of epoxides [28]. At higher concentrations of halide ions, vicinal dihalogenated products are also formed from alkenes [29]. The role of these enzymes in nature is not always clear [30] and they presumably are not involved in the metabolism of short chain unsaturated hydrocarbons. 3.1.2. Oxidation by monooxygenases Alkane metabolism has been studied for many years and several times alkenes have been proposed as intermediates in the degradation ot long-chain alkanes. This assumption was based on the detection of trace amounts of 1-alkenes after growth with the corresponding alkanes (see [31] for references) and the formation of 1-decene from decane by cell-free extracts of Gandida rugosa [32]. Reduction of N A D + in the presence of alkane by crude extracts of Candida tropicalis was later shown to be caused by impurities in the alkanes used [31]. Arguments against the involvement of 1-alkenes in the degradation pathway of alkanes to primary alcohols and subsequently carboxylic acids are however, abundant [33] and it is now generally accepted that degradation of aliphatic hydrocarbons in general proceeds via initial oxidation by a monooxygenase yielding primary or secondary alcohols [8]. The first evidence for oxidation of the double bond of an alkene was the isolation of 1,2-dihydroxyhexadecane from cultures of Candida lipolytica growing on 1-hexadecene [34]. Later it was shown that a large amount of the oxygen in the diol was derived from molecular oxygen [35]. Using heptane-grown Pseudomonas cells van der Linden [36] demonstrated the formation of 1,2-epoxyheptane from 1,heptene. Subsequently many more bacteria were found that possessed monooxygenases capable of epoxidating alkenes (see [37] for a review). Although alkane- and alkene-grown cells can generally epoxidate alkenes, reports concerning the metabolism of 1-alkenes with a chain-length longer than C 5 show that the epoxide is not a major intermediate in the degradative pathways, which generally proceed via oxidation of the saturated terminal methyl group. The first reports of an epoxidation reaction actually participating in the complete metabolism of an alkene were on ethene metabolism in Mycobacterium E20 [38,39]. 3.2. Hydratation of unsaturated carbon-carbon bonds Hydratation of fumarate to malic acid in the citric acid cycle is probably the best known biochemical hydratation of a double carbon-carbon bond. A similar hydratation reaction was reported by Wallen et al., who demonstrated formation of 10-hydroxystearic acid from oleic acid by a Pseudomonas sp. [40]. Further work by Schoepfer [41] revealed that the 10-hydroxystearic acid produced was optically active and had the R-configuration. In a chloroacrylic acid-degrading bacterium, we have recently detected two chloroacrylic acid hydratases, acting on either the cis- or the trans-isomer of this acid (unpublished results). Hydratases acting on unsubstituted alkenes are less well studied. Iida and Iizuka reported the enzymatic conversion of 1-decene to 1-decanol in crude extracts of a decane-grown Candida rugosa, although it is not clear if this reaction participates in decane catabolism [42]. To our knowledge the only report on degradation of an aliphatic alkene involving a hydratase is the degradation of 1hexadecene by a methanogenic enrichment culture 239 [43]. It is implicated that anaerobic 1-hexadecene metabolism proceeds via an initial hydratation to 1-hexadecanol. Degradation of acetylenic bonds is associated with hydratation of the triple bond. These hydratation reactions are discussed in more detail in section 4.1 of this review. 3.3. Reduction of unsaturated carbon-carbon bonds Apart from the reduction of the 2,3-trans double bond in fatty acid synthesis, examples of reductases acting on double bonds are scarce. One example is the hydrogenation of unsaturated fatty acids by anaerobic rumen bacteria [44]. Until now there is no evidence that such a reduction of unsaturated carbon-carbon bonds is involved in the mineralization of unsaturated hydrocarbons. Although reduction of chlorinated ethenes under methanogenic conditions results in sequential dechlorination, it is not clear if the double bond itself is ultimately reduced [45]. The enzymatic reduction of acetylene and several other unsaturated compounds by the action of a nitrogenase is a special case involving a special enzyme. 4. SPECIFIC WAYS BIODEGRADATION PATH- 4.1. Ethyne and acetylenic compounds 4.1.1. Ethyne (acetylene) The first report on the biodegradation of ethyne (acetylene) by aerobic bacteria was published in 1932 by Birch-Hirschfeld [46] who described growth of Myobacterium lacticola with ethyne as sole carbon and energy source. Almost fifty years later a Nocardia rhodochrous, which requires the pyrimidine moiety of thiamine for growth [47], and a Rhodococcus A1 [48] were isolated with ethyne as carbon source. The Rhodococcus A1 also utilized propyne as growth substrate. The description of the strain designated M. lacticola could fit both the Nocardia and the Rhodococcus A1. The Mycobacterium was grown in the presence of soil extract, which could fulfill the growth factor requirement of the Nocardia rhodochrous. None of the above strains were capable of growth with ethene. Other aerobic acetylene-utilizing bacteria have been isolated [49,50], but these organisms were not studied in detail. The observation of Birch-Hirschfeld that acetaldehyde accumulated when ethyne-grown cells were incubated in the presence of ethyne was confirmed by the Bont and Peck [48]. Kanner and Bartha [51] also identified acetaldehyde as an intermediate in ethyne metabolism. They also showed that growth of the N. rhodochrous with ethyne and ethanol resulted in similar activities of acetaldehyde dehydrogenase, acetothiokinase and isocitrate lyase. However, ethyne hydratase, the enzymic activity that would explain acetaldehyde formation from ethyne, could not be detected in cell-free extracts of ethyene grown N. rhodochrous [51]. In cell-free extracts of ethyne grown Rhodococcus A1 ethyne consumption and acetaldehyde formation were detected when the extract was incubated under an atmosphere of nitrogen [48]. Aerobic incubation of the extract resulted in irreversible inactivation of ethyne hydratase activity. Using dialyzed cell-free extracts of Rhodococcus A1 a stoichi0metric conversion of ethyne to acetaldehyde was observed. In crude extracts the K m of the hydratase for ethyne was calculated to be 0.6 m M [48]. The concentration of ethyne in the gas phase which corresponds with this value is 1.5% [v/v]. In order to grow at ethyne concentrations far below the K m of the hydratase, Rhodococcus A1 probably synthesizes large amounts of the enzyme to ensure a sufficient carbon flux. When these cells subsequently are exposed to high concentrations of e t h y n e an u n b a l a n c e d o v e r p r o d u c t i o n of acetaldehyde results. It should however be borne in mind that the K m was determined with crude extracts and not with whole cells. The K m for these two situations need not necessarily be the same. Anaerobic degradation of ethyne has also been observed [52]. With sulphate as an electron acceptor an enrichment culture which grew anaerobically at the expense of ethyne was obtained. Acetate was identified as an intermediate in the degradation of ethyne to carbon dioxide [53]. Schink recently reported the isolation of strictly 240 anaerobic acetylene-utilizing bacteria [54]. These isolates were assigned to a new species in the genus Pelobacter, namely P. acetylenicus. Acetylene was fermented by disproportionation to acetate and ethanol. Acetylene hydratase was not detected in cell-free extracts but the enzymes necessary for acetaldehyde metabolism were present in high activities. 4.1.2. Other acetylenic compounds To study enzymatic transformations of the acetylenic bond, Yamada and Jakoby [55] enriched for organisms capable of utilizing the nongaseous and water-soluble acetylenedicarboxylic acid. From a Pseudomonas isolated on acetylene dicarboxylic acid a more than 100-fold purified enzyme preparation was obtained which transformed acetylenedicarboxylic acid to pyruvate and carbon dioxide in a cofactor-independent reaction (EC 4.2.1.72). The mechanism by which this reaction proceeds was not elucidated. Neither of the potential intermediates, oxaloacetate (which would result from a hydratation) or acetylene monocarboxylic acid (which would result from an initial decarboxylation of the acetylene dicarboxylic acid), resulted in pyruvate formation with the partially purified protein preparation. To provide further information on the mechanism of this reaction a Pseudomonas was isolated with acetylene monocarboxylic (propynoic) acid as the sole carbon source [56]. From extracts of this strain an enzyme was partially purified (93fold) which catalyzed the hydratation of propynoic acid to malonic semialdehyde (3-oxo-propionic acid) without the addition of cofactors under an atmosphere of helium (EC 4.2.1.71). Acetylenedicarboxylic acid and propynol did not serve as substrates. Also using propynoic acid, de Bont et al. isolated a Gram-negative and a Gram-positive bacterium [57]. Interestingly, when propynol was used as a carbon-source in enrichment cultures only fungi were isolated. Using 3-butyn-l-ol as carbon source both fungi and Gram-negative bacteria were isolated [57]. From these isolates the facultative methylotroph Pseudomonas BB1 was chosen for further study of 3-butyn-l-ol metabolism [58]. Cell-free extracts of 3-butyn-l-ol grown Pseudomonas BB1 contained a phenazine methosulphate-dependent alcohol dehydrogenase and 3-butynoic acid hydratase activity yielding succinic semialdehyde (4-oxo-butyric acid). With propynoic acid as a substrate malonic semialdehyde was formed in cell-free extracts. Although Yamada and Jakoby [56] did not test 3-butynoic acid as a substrate for their enzyme preparation, it probably resembles the hydratase activity of 3-butyn-l-ol grown Pseudomonas BB1. The metabolism of acetylenic compounds thus seems to be catalyzed by hydratases, and until now at least three different enzymes capable of hydratating the acetylenic bond in different molecules have been described: (i) an oxygen sensitive ethyne hydratase [48], (ii) an acetylene dicarboxylate hydratase (EC 4.2.1.72) yielding pyruvate and CO 2 [55] and (iii) an acetylenecarboxylate hydratase (EC 4.2.1.71) yielding malonic semialdehyde [58,56]. 4.2. Ethene Ethene (ethylene), the simplest olefin, can be oxidized to epoxyethane by several types of microorganisms, including methanotrophs [59], alkaneutilizers [60] and Nitrosomonas europaea [61]. These organisms, however, are not able to grow with ethene, but other bacteria have been isolated which utilize ethene as the sole source of carbon and energy. Ishikura and Foster [35] isolated an orange-yellow pigmented 'ethylene bacterium' from soil. It was a Gram-positive, motile, non-sporulating rod that was also capable of growth with ethanediol. In 1976 both Heyer [62] and de Bont [63] reported the isolation of several Mycobacteria capable of growth with ethene as sole carbon and energy source. Subsequent enrichment cultures with ethene have always resulted in the isolation of Mycobacteria, although very recently non-mycobacterial ethene utilizers have been isolated by Mahmoudian and Leak [64]. Using propene or 1-butene as the carbon source in enrichment cultures, van Ginkel et al. [65] in most cases isolated Xanthobacters. These Xanthobacter strains, along with Nocardia H8 which was isolated with 1- 241 hexene as a carbon source, were all capable of growth with ethene. However, the growth rates of these strains with ethene as substrate were lower than those of the ethene-utilizing Mycobacteria. This could explain why Mycobacteria are almost always isolated when ethene is used as carbon source in batch-type enrichment cultures. Growth of the 'ethylene bacterium' in the presence of labelled oxygen on ethene resulted in incorporation of significant amounts of 180 in cell material as compared to growth with acetate under the same conditions [35]. This would implicate the involvement of a monooxygenase type of reaction in the assimilation of ethene into cell material. More specific evidence for the involvement of such an enzyme was obtained in studies with Mycobacterium E20. In this ethene-utilizing Mycobacterium it is possible to accumulate epoxyethane from ethene when whole cells are incubated in the presence of fluoroacetate [38], implicating that ethene is metabolized via epoxyethane in a reaction catalyzed by a monooxygenase. When similarly inhibited cells were incubated with either 1SO2 o r H2180 it was established that the oxygen atom in epoxyethane was indeed derived from molecular oxygen [39]. Monooxygenase activity with ethene was confirmed using cell-free extracts of ethene-grown Mycobacterium E20. The reaction was shown to be O 2- and N A D H - d e p e n dent. N A D H could be replaced by N A D P H , although this resulted in a lower specific activity. Whole cells of ethene-grown Mycobacterium E20 also oxidized ethane to some extent but this capacity was not present in crude extracts of ethene-grown cells. Ethane-grown ceils oxidized ethane and ethene, but it was not possible to detect any monooxygenase activity in crude extracts of these cells. Apparently Mycobacterium E20 is capable of synthesizing two different monooxygenases: a soluble alkene monooxygenase after growth with ethene and an unstable alkane monooxygenase after growth with ethane. This dependency of the monooxygenase induction on the growth substrate was further confirmed when the enantiomeric composition of the epoxypropane produced by ethene- and ethane-grown Mycobacterium E20 was compared (see 6.2.3). Although it has been known for some time that microorganisms can oxidize alkenes to the corresponding epoxyalkanes [36], very little is known about the further metabolism of these compounds. Microbial metabolism of epoxides has recently been reviewed by Weijers et al. [66]. The metabolism of epoxyethane, the most simple epoxide, was studied in Mycobacterium E20. When ethane-grown Mycobacterium E20 was incubated with ethane, in the presence of fluoroacetate, acetate accumulated analogous to epoxyethane accumulation from ethene. This could implicate that epoxyethane, analogous to acetate in ethane metabolism, is metabolized in a CoA-dependent reaction. Cell-free extracts of ethenegrown cells of Mycobacteriurn E20 were able to catalyze the oxidation of epoxyethane. The reaction was completely dependent upon the presence of N A D + and CoA, and the epoxyethane degradation rate was approximately doubled by adding F A D to the reaction mixture [38]. Besides N A D +, CoA and FAD, a fourth unknown dissociable cofactor was involved in the enzymic conversion of epoxyethane. This dialysable, heat stable cofactor was present in ethene-grown cells, but not in ethanol-grown cells. The nature of the unknown cofactor was not elucidated. Evidence that the product of the epoxyethane dehydrogenase reaction was acetyl-CoA was sought in experiments using [14C]epoxyethane. Incubation of cell-free extract with the radioactive epoxide and the required cofactors along with citrate synthase, oxaloacetate and fluorocitrate resulted in radioactivity in ether-extracts which cochromatographed with citrate. Omission of citrate synthase or oxaloacetate from the complete reaction mixture resulted in almost no radioactivity in the citrate spot. It is not clear whether one single enzyme or an enzyme complex is responsible for the oxidation of epoxyethane and much remains to be elucidated concerning this novel enzymic activity. Anaerobic degradation of ethene has until now not been reported. The strict oxygen-dependency of ethene-degradation strongly suggests that ethanol formation due to hydration of the double-bond does not occur. The activation of the double-bond apparently is only brought about by a monooxygenase. 242 4.3. Propene and 1-butene Analogous to the situation with ethene, many bacteria are capable of oxidizing propene to 1,2-epoxypropane [59,60]. In contrast to ethene, however, propene is an asymmetric molecule with an, unsaturated and a saturated carbon-carbon bond, allowing more than one possibility for initial enzymic attack. With propane-grown cells of Mycobacterium convolutum acrylic acid was identified as oxidation product of propene, indicating the initial formation of 3-hydroxy-l-propene [67]. But with other bacteria grown on propane, epoxypropane was reported as an oxidation product of propene with only trace amounts of 3-hydroxy-l-propene being detected [60]. With Pseudomonas fluorescens N N R L B-1244 the propene consumption rate was 20% higher than the 1,2-epoxypropane formation rate. The 1,2-epoxypropane degradation rate was not reported so that only a minimal ratio of epoxidation versus hydroxylation of 6 can be calculated. Although the presence of two different enzymes could not be ruled out, both hydroxylation and epoxidation of propene to 3-hydroxy-1propene and 1,2-epoxypropane, respectively, is probably effected by the same enzyme system in both Pseudomonas fluorescens N R R L B-1244 and Brevibacterium sp. strain CRL56 [60]. This situation has also been encountered in Nitrosomonas europaea [68]. The utilization of propene as a carbon and energy source is less frequently described. The first report of an organism capable of growth with this gaseous compound was a 'methanbakterium' isolated in 1930 [69]. Cerniglia et al. [67] also isolated a propene-utilizing organims. They proposed, on the basis of isocitrate lyase activities and the fatty acid composition of their strain PL-1 after growth with different substrates, that propene was metabolized via initial attack at the double bond resulting in a C 2 + C 1 cleavage. Experiments with Mycobacterium Pyl [70,71] and Xanthobacter Py2 [72], which were both isolated with propene as carbon source, and Nocardia By1 and Xanthobacter By2 [65] which were enriched with 1-butene, revealed that in these strains 1-alkenes were epoxidized to the corresponding 1,2- epoxyalkanes. In the strains Pyl and Py2, NADH-dependent propene-monooxygenase activity was detected in crude ceil-free extracts. 1,2epoxypropane, the product of the monooxygenase reaction, was also utilized as growth substrate. Both strains can also utilize 1-butene as a growth substrate. In contrast to strain Pyl, strain Py2 also grows on ethene although the growth rate is low. In Mycobacterium Pyl isocitrate lyase activity was induced after growth on propene, 1,2-propanediol and acetate, indicating that these substrates are metabolized via acetyl-CoA. G r o w t h on 1-butene and propionic acid did not result in isocitrate lyase induction. These results correspond to the results obtained by Cerniglia et al. [67], but further indications as to how 1,2epoxypropane is metabolized are still lacking. An analogous reaction as was proposed in ethenemetabolism [38] in which epoxyethane is oxidized to acetyl-CoA seems very unlikely in view of the increased isocitrate lyase activities after growth with propene. Another possibility for the further metabolism of 1,2-epoxypropane in Mycobacterium Pyl via 1,2-propanediol and propionaldehyde as was shown in Nocardia A60 [73] was ruled out by carrying out simultaneous adaptation experiments with propene- and 1,2-propanediol-growth cells (Hartmans, unpublished results). 1,2-Propanediol metabolism in Mycobacterium Pyl proceeds via acetol which is subsequently cleaved into acetate and formaldehyde by acetol monooxygenase [74]. Although this explains the induction of isocitrate lyase after growth with 1,2-propanediol in Mycobacterium Pyl, the metabolic pathway of propene degradation via 1,2-epoxypropane to acetyl-CoA still remains to be elucidated. 4.4. 2-Butene In contrast with ethene, propene and 1-butene, 2-butene is an internal alkene thus possessing only saturated terminal carbon atoms. This difference in chemical structure is reflected in the ability of organisms to degrade the compound. It is not utilized as a growth substrate by at least six different 1-alkene-utilizers tested, including ethene-, propene- or 1-butene-utilizers [65]. Fujii et al. 243 however, have described two Mycobacteria which were isolated with propene and 1-butene, respectively, that were also able to grow on the C z to Ca saturated hydrocarbons and both isomers of 2butene [75]. Both strains grew very poorly with ethene and did not grow with 1,3-butadiene as a carbon source. Enrichment cultures with trans-2 butene as the carbon-source resulted in three bacterial isolates, two strains of the genus Nocardia and one Mycobacterium [65]. One of the Nocardia strains, a red-pigmented bacterium designated as Nocardia TB1 was chosen to study trans-butene degradation [76]. Strain TB1 was also capable of growing on the C 3 to C 6 alkanes but did not grow with methane, ethane, 1,3-butadiene or the C 2 to C 6 1-alkenes. Growth with cis-butene was extremely slow and doubling times on butane and transbutene were 6 and 30 hours, respectively. Using arsenite as inhibitor, butyric acid and crotonic acid accumulated when butane- or transbutene-grown cells were incubated with their respective growth substrates. Surprisingly, trans-butene- and butane-grown cells oxidized trans-butene at a higher rate than butane, the substrate which supports the faster growth, cis-butene was degraded at the same rate as trans-butene. Based on enzymic activities of trans-butene, butane and succinate grown cells a degradative pathway as shown in Fig. 2 was proposed [70]. Although 2,3-epoxybutane was degraded by trans-butene-grown cells of strain TB1, it was not considered an intermediate in trans-butene metabolism as its degradation did not result in increased CO2-formation by washed cells, whereas the oxidation of crotonic alcohol and trans-butene itself did. Degradation of 2,3-epoxybutane resuited in the excretion of an unidentified product, probably originating from a hydroxylation reaction by the monooxygenase. Further evidence that Nocardia TB1 contains an alkane-type monooxygenase was obtained when the enantiomeric composition of epoxides formed by both butane- and trans-butene grown cells was analyzed (Table 4). 4.5. Butadiene and isoprene H3C-CH2~CHT CH3 02~ NADH 02tNADH H3C-CH : CH-CH3 % H3C-CH2-CH2-CH2 I x~.-NADH °° "H I NAD NADH L H3C- Cl4=CH -(H 2 ['~NADH °° NADH ~'-~NADH H3E-CH2-IEH2-C~0 H3C-CH ~CH-C ~0 ATP+ ~oA ATP~-- CoA ~OH \OH ! H~E-CH2-CH2-C~ ~ "CoA H3C-CH =CH-C ~0 "CoA 13-oxidation Fig. 2. Proposed degradative pathway of butane and transbutene in Nocardia TB1 [76]. Isoprene (2-methylbutadiene) is a naturally occurring compound, whereas its non-methylated analogue butadiene, to our knowledge, is not formed biologically [77]. Butadiene can be compared to ethene, with respect to the unsaturated character of its carbon atoms, with the two double bonds probably behaving as a conjugated system. Microbial utilization of butadiene has been reported by Watkinson and Somerville who isolated a Nocardia species from enrichments with butadiene as the sole carbon and energy source [78]. Respiration rates of butadiene-grown cells with butadiene and 1,2-epoxy-3-butene as substrate were similar. This oxidative capacity was absent from acetate-grown cells. Based on isocitrate lyase activities butadiene metabolism in Nocardia sp. 249 was thought to proceed via acetate. The degradation p a t h w a y that was p r o p o s e d for butadiene metabolism was very speculative, and not based on measurement of enzymic activities or identification of possible intermediates. It was suggested that butadiene is epoxidized to 1,2- 244 epoxy-3-butene, which subsequently would be hydrolyzed to the corresponding diol and oxidized to 2-oxo-3-butenoic acid. Oxidative decarboxylation to acrylic acid followed by hydratation to lactate and oxidation to pyruvate would after decarboxylation, eventually result in acetate formation. Interestingly, oxidation of racemic 1,2-epoxy-3butene was not complete, and it was shown that the remaining epoxide material was optically active, thus implicating that the epoxide degrading enzymic activity was stereoselective. Based on this observation it was concluded that the epoxidation of butadiene by the Nocardia species is stereospecific [78]. Enrichments using different soil samples and butadiene or isoprene as a carbon source in all cases resulted in the isolation of pink-pigmented bacteria belonging to the genus Nocardia [65,77]. All isolates were capable of growth on both substrates, possibly suggesting a connection between the degradation pathway of the two alkadienes. In cell-free extracts of alkadiene-grown Nocardia IP1 oxidation of these compounds was N A D H - and oxygen-dependent, indicating that these compounds are degraded by a monooxygenase [77]. Incubation of washed cell suspensions of alkadiene grown Nocardia TB1 with the respective growth substrates in the presence of 1,2-epoxyalkanes as competitive inhibitors of epoxide degradation, all the possible mono- and diepoxides of butadiene and isoprene could be detected. Although it was proposed that the initial step in the metabolism of both compounds Nocardia TB1 probably was the formation of an epoxide, the degradation pathway of these alkadienes remains to be elucidated. 4.6. Styrene Although styrene contains an aromatic nucleus, it can also be classified as a substituted alkene. Styrene can be oxidized by Methylosinus trichosporium OB3b [79] and Nocardia corallina B-276 [80]. In both cases the alkenic moiety of the molecule is attacked, resulting in the formation of styrene oxide (7,8-epoxyethylbenzene or phenyloxirane). Omori et al. [81] were, to our knowledge, the first to attempt the isolation of styrene-utilizers. They tested 101 soil samples without success, probably because the concentration of styrene that was used in the enrichment cultures was too high (2% w / v ) . Sielicki et al. [82] using a concentration of 1% (w/v), which is still much more than the solubility of styrene in water, obtained a mixed culture utilizing styrene. In ether extracts from styrene cultures phenylacetate and 2-phenylethanol were identified. Using a pure culture of a styrene-utilizing Pseudomonas Shirai and Hisatsuka [83] also demonstrated accumulation of 2phenylethanol from both styrene and styrene oxide. Based on these results it was proposed that styrene oxide is an intermediate in the transformation of styrene to 2-phenylethanol. Baggi et al. [84] isolated phenylacetic acid and 2-hydroxyphenylacetic acid from styrene-grown cultures of a Pseudomonas fluorescens, once more indicating that styrene metabolism involves initial attack of the ethylenic bond. Using low concentrations of styrene it was recently shown (van der Werf and Hartmans, unpublished results) that styrene utilizers are very abundant. F r o m all soil samples tested, bacterial strains were isolated that could utilize styrene as a sole carbon and energy source. In several of these strains an oxygen- and N A D H - d e p e n d e n t styrenedegrading enzymic activity was present in cell-free extracts after growth with styrene. The further metabolism of the probable oxidation product styrene oxide, was in some cases via phenylacetaldehyde which was formed by a styrene oxide isomerase activity. Reduction or oxidation of the phenylacetaldehyde thus formed would then have resulted in either the formation of 2-phenylethanol or phenylacetic acid, the intermediates previously isolated by other groups [83,84]. 2-phenylethanol formation from styrene could also result from a hydratation reaction. Indeed very recently anaerobic isolates with styrene as sole carbon and energy source were shown to produce 2-phenylethanolas intermediate in styrene degradation [85]. F r o m the above we can conclude that there are at least two different modes of initial attack of styrene, both involving the alkenic bond. 245 4. 7. Chlorinated ethylenic compounds Vinyl chloride (chloroethene), the simplest chlorinated ethylenic compound is aerobically utilized as a sole carbon and energy source by Mycobacterium L1 [86]. Ethene, but not propene, also supports growth of this strain. Growth on vinyl chloride and ethene both result in alkene monooxygenase induction. Oxidation of vinyl chloride by vinyl chloride- and ethene-grown cells can be competitively inhibited by ethene or propene, indicating that in vivo vinyl chloride is degraded by the alkene monooxygenase. The product of the monooxygenase reaction with vinyl chloride has not been identified, but probably is chloroepoxyethane (chlorooxirane). The metabolism of chlorooxirane, which is a very unstable compound that rearranges to chloroacetaldehyde, has not been studied. However, using washed cell-suspensions it was possible to detect chlorideion formation after incubation with vinyl chloride, but not with chloroacetaldehyde, indicating that this compound is not an intermediate in vinyl chloride metabolism. Although vinyl chloride grown Mycobacterium L1 cells were capable of oxidizing dichloroethene isomers, strain L1 did not grow with chloroethenes other than vinyl chloride. Until now, no other bacterial strains are known that can grow with chlorinated ethenes. Biotransformation of ethenes with more than one chloride substituent, especially trichloroethene and tetrachloroethene, however, has been extensively studied the last decade. Both oxidative and reductive transformations have been reported (see [87] for a recent review). Oxidative transformation of di- and trichloroethenes by mixed- and pure-cultures of methane utilizers has been reported by several authors [21,88,89]. It is generally accepted that the initial oxidation step is brought about by the a-specific methane monooxygenase resulting in the formation of the relatively unstable epoxides. Any further transformations that take place are still obscure, although, using labeled trichloroethene, it was possible to detect the formation of labeled carbon dioxide in a pure culture of a methaneoxidizing bacterium [89]. Tetrachloroethene was not degraded by the methane-utilizing cultures. Nelson et al. [90] recently reported that toluene stimulates the trichloroethene-degradative ability of the natural microflora of environmental water samples. Several pure cultures capable of utilizing toluene also degrade trichloroethene. These authors also presented evidence that a toluene dioxygenase is involved in trichloroethene degradation by one of the pure cultures. Studies with toluene induced cells of Pseudomonas putida F1 [91] revealed that trichloroethene degradation by these cells was at a significantly greater initial rate than by Methylosinus trichosporium OB3b. Mutants of Pseudomonas putida F1 defective in the gene encoding for the oxygenase component of toluene dioxygenase failed to degrade trichloroethene, whereas a spontaneous revertant simultaneously regained the ability to oxidize toluene and to degrade trichloroethene, thus confirming the role of the oxygenase component of toluene dioxygenase in trichloroethene degradation. The three isomeric dichloroethenes were also degraded by Pseudomonas putida F1, but tetrachloroethene, chloroethene (vinyl chloride) and ethene were not. Until now, tetrachloroethene degradation under aerobic conditions has not been reported. Under methanogenic conditions, however, a sequential transformation of tetrachloroethene via trichloroethene and dichloroethenes to chloroethene (vinyl chloride) was observed in anaerobic laboratory columns fed with an acetate-containing medium [45]. The transformation of tetrachloroethene to trichloroethene is also accomplished by pure cultures of several Methanosarcina spp., although the transformation rate differs significantly between strains [92]. 5. E C O L O G I C A L ASPECTS The interesting question arises as to how the organisms described above acquired their metaboric potential to degrade the various gaseous and volatile olefins. Several of these compounds are assumed to have entered the environment during this century due to human activities. It would be interesting to know if the microbes capable of metabolizing these compounds have also only re- 246 cently acquired this capacity; or have these degradative capacities evolved over a much longer period of time as a result of the presence of these supposedly anthropogenic compounds arising from natural sources? Before speculating on these aspects, it seems worthwhile to first examine the situation for the natural compound ethene. The other short chain unsaturated hydrocarbons, with the exception of isoprene, are generally considered to be anthropogenic and will be discussed subsequently. Another interesting aspect of the short chain unsaturated hydrocarbons is their inhibitory effect on certain metabolic processes. These effects will be discussed in section 5.3 and attention will be given to the use of acetylene as a tool to determine in situ metabolic activities. 5.1. Ethene." formation and degradation in nature Ethene presents a special ecological case because the gas has such pronounced effects on plants and fruits at very low concentrations. In soil it may affect plant life when accumulating due to the production by microorganisms or to a lesser extent by plants. It also accumulates during storage of harvested fruits, vegetables and flowers through the endogenous production by the plant material. Several relevant aspects of ethene in nature were discussed 10 years ago by Primrose [93] and his review presently still offers a solid basis for our understanding of the role of microbes in the production and utilization of ethene. New insights, however, have emerged during the last decade. Especially on the biosynthesis of the gas, substantial progress has been made. It has now been clearly established that at least three different routes for the biosynthesis of ethene exist. In plants, methionine serves as the precursor and yields ethene through S-adenosyl-methionine and 1-aminocyclopropane-l-carboxylic acid [94,17]. In bacteria ethene is also formed from methionine but in a different fashion. In E. coli [94] and other microorganisms [96] 2-keto-4-methylthiobutyric acid is an intermediate which is subsequently converted into ethene and methanethiol [97]. In the fungus Penicillium digitatum ethene is formed from H~N-CH-COOH CH2 EhH2 $1 rl2N-CH-EOOH i EH 2 CH2 *~-Adenosyt Plants EH] ~ H~N • ,, / COOH sCx ~ H2C=CH~ H2C-- [H2 O2-dependent CH~ H2N- CH -COOH i CH2 5H 2 ~ S [0OH O=C Ecoli !H 2 CH? ~ H2C:CH 2 Oz-dependenf i [H~ EOOH O=C Pemolhum [H2 0 3- ~H~ ~lgltafum depend enf ~ H2[zCH 2 EOOH Fig. 3. Biosynthetic pathways of ethene formation. a-ketoglutarate as was shown using a combination of partially purified enzyme preparations [98]. The reaction also required Fe2+-ions and L-arginine but the carbon atoms of ethene were derived from a-ketoghitarate as shown with 14C-labelled substrates. In a later communication it was stated that oxygen is also essential for ethene formation [99]. The three biosynthetic pathways leading to ethene formation are shown in Fig. 3. It is interesting to note that in all three routes enzymic reactions are implicated that require molecular oxygen. Since ethene formation has also been observed under strict anaerobic conditions, it is therefore obvious that other pathways leading to ethene formation remain to be discovered. The rate at which ethene is produced by either plants or microorganisms varies considerably. For plants this rate varies from species to species and organ to organ and it also depends on the stage of development of the plant [15]. For microorganisms the rate of ethene production depends very much on the organism tested, and also on the culture conditions applied. Ethene-producing microorganisms have been compiled from the literature by Primrose [93] and his conclusions were that such organisms are ubiquitous in soil and water. This concept was supported by Fujii et al. [75] who screened 296 microorganisms of which 51 produced ethene. The positive strains included bacteria as well as fungi and yeasts. The best producing organism in terms of quantities formed, 247 was a Penicillium digitatum strain. By mutant selection it was eventually possible to obtain a growth-suppressed mutant that was able to convert 2.1% of the carbon present in glucose into ethene-carbon [99]. In soil the situation is clearly different and here only very minute quantities are produced. It is very difficult to determine the absolute quantity of ethene produced in soil since under aerobic conditions the gas is also oxidized by ethene-utilizers [100,101,63] resulting in a considerably lower net observed production, or in no apparent production at all. This point has not always been kept in mind by many researchers studying the production of ethene in soil and consequently has resuited in some in interesting theories [102-108]. Another favoured approach in the area of ethene formation in soil has been to amend soil with different substrates that are potentially able to generate ethene [102,106-108]. The substrates were generally added in relatively high concentrations (mM-range). Obviously, when such excessive amounts, of for instance methionine, are added to aerobic soil, the many different organisms able to form ethene from this compound in only a few enzymic steps will do so immediately, resulting in unusually high ethene evolution rates. The ethene-utilizers present clearly cannot immediately oxidize this excess of ethene formed, resulting in a transient accumulation of ethene. Such experiments have little bearing on the acutal situation in soil; they can only give indications as to potential precursors for ethene formation. Realistic rates for ethene formation can only be obtained under anaerobic conditions. Such rates, when recalculated on a basis of ng ethene (g soil) -a day 1 vary from 0.01-10 [109] and 0.06-2.5 [107] for various soils to 600 for pine-needle litter [104] and to a difficult to explain high rate of 7000 in a silt loam soil [105]. Estimation of ethene formation rates in soil is interesting to know, but more important for the ecosystem are the in situ concentrations of the gas. Several measurements in field situations have been reported by K.A. Smith and colleagues [110,111]. From their results it can be generalized that under anaerobic conditions ethene may accumulate to 10 ppm (determined in the gas phase) or sometimes even higher, whereas under aerobic conditions the gas is almost never present at concentrations of more than 0.1 ppm. This concentration incidently is approximately the threshold concentration for the action of ethene as a plant hormone. The ethene-utilizing bacteria, in keeping the concentrations below this level in aerobic soil, thus hold a key position in maintaining the soil environment suitable for plant roots. When assessing the effects of ethene production in soil on plant growth [112], the significance of these bacteria should be given explicit attention. The implicit premise made on the functioning of the ethene-utilizers in soil is that they are able to degrade ethene at the extremely low concentrations prevailing in soil. A concentration of 0.1 ppm in the gas-phase corresponds to a concentration of 0.45 nM in the water-phase. The Michaelis-Menten constant of the ethene-utilizing Mycobacterium E3 has been determined and was estimated to be 100 ppm [113] ethene in the gas-phase corresponding to 0.45 ~M in the water-phase. For comparison the K m for methane of the purified methane monooxygenase from Methylococcus capsulatus (Bath) is 3 /~M [114]. This observation does not make it likely that such organisms are actively engaged in the oxidation of ethene at the concentrations present in aerobic soils. Nevertheless, van Ginkel et al. [115], using a compost filter with its indigenous population in a packed bed through which 2 ppm ethene in air was passed continuously, demonstrated ethene degradation at this low concentration. They also demonstrated that the organisms responsible for ethene degradation were actually able to grow under these conditions, as evidenced by an increase in ethene-utilization rate in time. It would seem that this remarkable capacity of the ethene-utilizers to metabolize ethene at these low concentrations, and to obviously benefit from this, is only possible when they grow mixotrophically. Components from the compost almost certainly will have functioned as supplementary carbon and energy sources for the bacteria. Plants not only produce ethene, they also metabolize ethene and during the last decade considerable progress has been made concerning the metabolism of ethene in plants. Until recently it 248 was believed that higher plants were not able to metabolize the hormone, but by using 14C-labeled ethene it has now been firmly established that different products are formed in plants [116,117]. The rates at which ethene is metabolized are, however, extremely low and in the order of 0.5 nmol (kg dry weight)-1 hr-1. This process therefore will not be significant as an ethene sink in nature. Interestingly, in plants ethene is attacked in the same way as in bacteria. In both cases the initial step in ethene metabolism is the oxidation to epoxyethane. The values reported for the K m of ethene oxidation by the plant monooxygenases are, depending on the plant being studied, in the range of 1 nM to 1 /*M [117]. 5.2. Supposedly anthropogenic short chain unsaturated hydrocarbons 5.2.1. Biogenic formation The generally accepted view on atmospheric hydrocarbons with respect to their origin is that a division can be made in biogenic and anthropogenic hydrocarbons [7,24,25]. The biogenic hydrocarbons are methane, ethene, isoprene and the terpenes, but these hydrocarbons may of course also be found in anthropogenic emissions. The anthropogenic hydrocarbons supposedly enter the biosphe, re only as a result of human activities and thus should be 'new' compounds for microorganisms. This division on the basis of the source of the hydrocarbons may be useful to determine to what extent hydrocarbons in air can be attributed to certain emissions. But the division on biogenic versus anthropogenic is misleading in understanding the metabolic potential of micro-organisms. Indeed several reports are available which demonstrate the biological formation of most compounds considered in this section. The production of ethane, propane, butanes, ethene, propene and 1-butene from soil have been reported [118,119]. The gases were observed under anaerobic conditions, which would indicate the involvement of anaerobic bacteria. However, a more pronounced formation of the gases was observed when soil initially kept anaerobic was subsequently made aerobic. Also, amending soil with various substrates enhanced the formation of the gases. These observations led Goodlass and Smith [119] to the hypothesis that substrates first mobilized under anaerobic conditions stimulate the formation of gases under aerobic conditions. The involvement of aerobic organisms as anticipated by Goodlass and Smith [119] has found strong support by interesting work published by a group of Japanese researchers [120-123]. They screened m a n y bacteria, yeasts and fungi under aerobic conditions and m a n y strains were found that produced ethane, ethene, acetylene, propane, propene, butanes, butenes and pentanes. Even aerobic methane formation was detected. Recently it was shown that acetylene, contrary to the general belief, is not exclusively connected with human activities. It is apparently also produced by natural processes in seawater [124]. The quantities that are formed by the various processes described above are minute and consequently concentrations of these gases are always low in nature. But from time to time, depending on the local conditions in for instance soil, the compounds will be present in appreciable quantities with concentrations in the gas-phase reaching the ppm-range [119]. The presence of these compounds in these concentrations implies that microorganisms have been exposed to these substrates in the course of evolution and it therefore is quite understandable that so many organisms have acquired the capability to grow on short chain unsaturated hydrocarbons. 5.2.2. Enzymes with relaxed substrate specificity The capability of various organisms to grow on substrates not abundantly present in nature could also be the result of the action of certain enzymes with relaxed substrate specificity. As will be discussed in more detail in the following section on applied aspects, enzymes involved in the initial biodegradation of hydrocarbons generally oxidize a broad range of substrates. An organism able to grow on a natural substrate may thus also be able to grow on a xenobiotic compound with similar chemical or structural properties. Such an example has been described by van Ginkel et al. [77] who isolated bacteria on the biogenic isoprene (2methyl-l,3-butadiene) and on the anthropogenic 1,3-butadiene. Strains isolated on isoprene or on 249 butadiene strongly resembled each other and moreover, all strains isolated were able to grow on both substrates. It was thus anticipated that a connection existed between the pathways for isoprene and butadiene metabolism and that the ability to degrade the anthropogenic 1,3-butadiene was a mere reflection of the ability to degrade the biogenic isoprene. A similar mechanism was also envisaged by Schink [54] when explaining the ecological significance of acetylene fermentation in anaerobic environments. He speculated thai the observed hydratation of acetylene to acetaldehyde in Pelobacter acetylenicus was due to an unspecific hydratase that in natural environments would be involved in the detoxification of not only acetylenic compounds, but also nitriles and cyanides. 5.2.3. Recently evolved enzymes and biodegradative routes A prerequisite for growth with novel compounds is that the necessary degradative enzymic activities are present. In some cases effective enzymic activities may already be present in the microorganism thus only requiring an appropriate regulation of enzyme synthesis to allow degradation of the novel compound. Very often a mutation resulting in constitutive synthesis of the required enzyme is sufficient. If sufficiently effective enzymes are not at hand, microorganisms can acquire new metabolic activities as a result of mutations affecting the rates of enzyme synthesis or the structure of enzyme proteins [125,126]. These studies share the fact that microorganisms were challenged to develop new pathways by altering their own genetic information due to mutations in structural a n d / o r regulatory genes, or perhaps by recruitment of single 'silent genes'. In nature however, much more genetic information is present than in a single microorganism. Another strategy therefore is, that the degradation capabilities of an organism can be expanded by recruitment of genetic information from the environment. An example is the in vivo construction of an organism with the novel capacity to degrade 4-chlorobenzoate. In Pseudomonas sp. B13 the first step in 3-chlorobenzoate degradation is the oxidation of 3-chlorobenzoate by a very specific 1,2-dioxygenase. This dioxygenase does not oxidize 4-chlorobenzoate and hence strain B13 cannot grow with 4-chlorobenzoate as substrate. By direct transconjugant mating between the TOL plasmidcontaining Pseudomonas putida PaW1 as donor of the non-specific toluate 1,2-dioxygenase and strain B13 as recipient it was possible to select 4-chlorobenzoate-degrading derivatives. The enzymatic, regulatory and genetic aspects concerning this and other genetically manipulated strains degrading chlorinated aromatic compounds have been reviewed by Reineke [127] and others [128]. Interestingly, 4-chlorobenzoate-degrading organisms isolated directly from nature do not degrade 4-chlorobenzoate via 4-chlorocatechol but via a hydrolytic reaction to 4-hydroxybenzoate (see [127] for references). As has already been pointed out in the previous sections the unsaturated short chain hydrocarbons, with the exception of the chlorinated analogues, cannot generally be considered as true xenobiotics, thus making it difficult to judge whether the degradative pathways involved have evolved very recently or have existed in nature for a long time. Chloroethene (vinyl chloride) can be considered as a true xenobiotic which has only recently entered the environment, directly, and indirectly via the anaerobic transformation of other chlorinated ethenes (see section 4.7), from anthropogenic sources. It is the only chlorinated ethene which, until now has been shown to support aerobic growth, but the metabolic pathways still needs elucidating. It is therefore difficult to speculate on the evolutionary events that took place resulting in the capacity to degrade vinyl chloride. With other chlorinated hydrocarbons, which also have just recently entered the environment, more progress has been made. The aerobic degradative pathways of 1,2-dichloroethane and dichloromethane metabolism have been described [129,130] and the enzymes performing the initial dehalogenation step purified [131,132]. The dichloromethane dehalogenases (group A) from several dichloromethane utilizers isolated from different environments have been characterized and were shown to have the same immunological properties and identical N-terminal amino acid 250 sequences [133]. This could be an indication that this enzyme has evolved only very recently. A novel dichloromethane dehalogenase (group B) has recently been characterized [134] which had only one of the 15 amino acids of the N-terminus in the same position as in the group A enzyme. The group B enzyme has a higher specific activity than the group A enzyme but represents a lower fraction of total protein when compared with organisms having the group A enzyme. Nevertheless, the new isolate grew at a higher rate (0.22 h -~) with dichloromethane than the strains with the group A enzyme (0.07-0.10 h - l ) . The K m of the group B enzyme was however, also somewhat higher so that it will be very interesting to see which enzyme type will predominate under different ecological conditions and in what way any further evolution will take place. 5. 3. Inhibitory effects on metabolic functions Ethene and its multiple functions as a hormone in plant physiology has of course attracted the attention of researchers in areas other than plant physiology. It has been attempted to assess a role for ethene in the production of cultivated mushrooms [135] and the production of secondary metabolites by Aspergilli and Streptomycetes [136,137] but no direct evidence concerning an effect of ethene was given. Arguments indicating a role for ethene as a critical regulator in soil [103] and a causative agent in fungistatis [138] were refuted in later publications [109,139]. An effect on methanogenic bacteria has only been noted at elevated levels. Oremland and Taylor [140] showed that in anaerobic sediments methanogenesis was reversibly inhibited by 5% ethene in the gas-phase. Schink [141] later confirmed that inhibition of methanogenesis is due to a direct inhibition of the methanogens, with 0.5% ethene in the gas-phase already resulting in more than 98% inhibition of methane formation with several different substrates. In conclusion, however, it may be stated that the biological effects of ethene are confined to plants and that microorganisms are affected by the gas in isolated cases only. Interestingly this situation is reversed in the case of acetylene. This gas has only limited effects on the physiology of whole plants but it is known that m a n y metabolic processes in bacteria are blocked by acetylene. The effects of acetylene on enzymes has very recently been reviewed by Hyman and Arp [142]. They have collected detailed information on the effects of acetylene on the enzymes nitrogenase, hydrogenase, ammonia monooxygenase, methane monooxygenase, nitrate reductase and nitrous oxide reductase. Therefore, only limited attention has to be given here to the effects of the gas. In the present context, only the use of acetylene in ecological studies will be briefly considered. Three activities can be monitored in situ in ecosystems by employing acetylene as a selective inhibitor i.e. nitrogen fixation, denitrification and methane oxidation. The reduction of acetylene to ethene has been used extensively in ecological studies to assess nitrogen fixation rates. The standard assay employs 10% acetylene in the gas-phase. The merits of the technique, as well as the precautions to be taken when applying it in ecosystems, have been discussed in detail in the literature. A recent, very interesting observation is that under certain conditions acetylene is not only reduced to ethene but also to traces of ethane [1431. Acetylene is also used for in situ measurements of denitrification rates, generally resulting in a complete inhibition of N20-reduction activity at acetylene concentrations of 0.1%. The merits of the acetylene method for assessing denitrification rates have been summarized by Knowles [144]. Much less attention has been given to acetylene as a useful tool in the determination of methaneoxidizing bacteria in situ. Methane formation and methane consumption are interrelated in nature at interfaces where oxygen meets anaerobic zones. In the anaerobic zone methane is produced and at the interface methane is oxidized. Such interfaces occur for instance in stratified lakes, in the upper mud-layer of ditches or at the root-soil interface of plants growing in submerged soils. Using acetylene it is possible to determine methane oxidation rates in such complex systems since the inhibition of methane oxidation is complete at 10 p p m acetylene, whereas methane formation is only inhibited at acetylene concentrations of approximately 100 p p m as determined for pure cultures 251 [145] or higher as determined for marine sediments [146]. The difference in net methane accumulation in the presence and absence of acetylene in the lower ppm-range therefore is equal to the amount of methane oxidized in the system. Methane oxidation rates using acetylene as an inhibitor have been determined in a marine environment [147] and in rice paddy fields [148] although it has not always been realized that acetylene has a dual function as inhibitor of both methane formation and methane oxidation [149]. As a general rule, it should be kept in mind that acetylene has m a n y different effects on microorganisms present in an ecosystem under study, and that therefore artifacts may be detected rather than quantifying the metabolic rates of the microbial p o p u l a t i o n present in the ecosystem [150,49,145]. 6. A P P L I E D ASPECTS 6.1. Biodecontamination of Unsaturated compounds In several instances the presence of alkenic compounds is a problem in air or water environments. Low concentrations of ethene in the atmosphere of fruit storage facilities can have devastating effects. The presence of the carcinogen vinyl chloride or the odorous styrene can be a serious problem in the atmosphere of the polymer industry. The less volatile chlorinated ethenes like the suspected carcinogens tetrachloroethene and trichloroethene are frequently found as contaminants in drinking-water aquifers [21]. 6.1.1. Removal of gaseous contaminants Gaseous contaminants may be effectively removed through biofiltration using undefined support materials for the active biomass. Such systems are operated successfully when very dilute and undefined mixtures of odorous gaseous compounds are to be eliminated. The microbial population in such system is very heterogeneous and is not only thriving on the gaseous substrates to be removed but also on components of the support as for instance compost [151]. More defined systems however, may also be of use whenever a defined gas mixture is to be treated. In such cases pure cultures, immobilized on a defined support, may be applied. Several of such systems have been studied recently, including the removal of aldehydes and ketones [152] and ethene [113]. To prevent the accumulation of ethene in storage places for fruit and flowers ventilation of the atmosphere is often applied. However, ventilation is impracticable when the temperature a n d / o r the gas composition in the warehouse differ from the open air. As an alternative for chemical or physical methods to remove ethene from the storage atmosphere the possibility of applying a bioreactor has been investigated [113]. It was shown that immobilization of ethene-utilizing cells of Mycobacterium E3 on lava or perlite did not result in a decrease in the maximal specific oxidation rate or in an increase in the apparent K m in comparison with free cells. However, the cell densities employed were quite low, thus requiring residence times of 15 to 30 s to decrease the ethene concentration from 3.2 p p m to 1 ppm. The Vma~ of Mycobacterium E3 is 50 nmol ethene rain -1 (rag protein) -~ and the K m for ethene is 100 p p m in the gas phase [113]. Although the results seem promising it would be advantageous to have a biocatalyst with a significantly higher affinity towards ethene. Application of Mycobacterium L1 to remove vinyl chloride from waste gases has also been proposed recently [86]. The stability of vinyl chloride degradation by washed cells was, however, not very high and depended strongly on the biomass concentration. It was shown later (Hartmans, unpublished results) that this inactivation is caused by the excretion of a toxic product, probably chloroepoxyethane. Using growing cultures in a chemostat it was possible to degrade vinyl chloride quite efficiently, but also under these conditions autoinactivation of the vinyl chloridedegrading population sometimes occurred when there were irregularities in the vinyl chloride supply. Due to the involvement of the highly toxic chloroepoxyethane in the degradative pathway of vinyl chloride, Mycobacterium L1 does not seem to be a suitable biocatalyst for the removal of vinyl chloride from waste gases. 252 Styrene removal from waste gases has also been reported [153] using a biofilter which was inoculated with a Nocardia sp. The styrene in styrene containing air (1.3 g m 3) was completely degraded by passing the air through a 2.5 meter biofilter at a superficial gas flow of 100 m h -1. However, no comments were made on the stability of the system. Before becoming applicable in a practical situation the stability of this type of reactor with a monoculture immobilized on a defined cartier will have to be enhanced. It is reasonable to expect that further research in this field will lead to successful bioreactors for the effective removal of polluting gaseous compounds. 6.1.2. Removal of chlorinated ethenes with methane-utilizers The increasing frequency with which chlorinated ethenes are detected in ground water has stimulated research concerning the degradation and removal of these compounds. Currently, treatment of ground water containing chlorinated hydrocarbons consists of pumping the water to the surface and stripping out the components in aeration towers or removing the pollutants with a sorbent [21]. Recently it was shown that exposure of a soil column to natural gas resulted in mineralization of trichloroethene [22]. Since then there have been several reports concerning the degradation of chlorinated ethenes by mixed and pure methane-utilizing cultures [21,89,88]. The degradation of trichloroethene is proposed to proceed via the epoxide which spontaneously breaks down to other products [89]. Using trans-l,2-dichloroethene it was shown that both mixed and pure cultures of methane-utilizers transformed this compound to the relatively stable epoxide [89]. Application of methane-utilizing cultures to treat ground water does not seem very realistic as one of the most important pollutants, tetrachloroethene, is not degraded under these conditions. 6.2. Biotechnological production of epoxides from alkenes Epoxides are reactive molecules and there are many chemical syntheses in which epoxides can function as intermediates. In the past ten years the possibility of using bacteria to transform alkenes to the corresponding epoxides has been suggested by several authors. This is also reflected by the patient literature (see [4,154]). The capacity to epoxidate alkenes is not limited to alkene-utilizing bacteria. In theory any microorganism containing a monooxygenase with activity towards alkenes could be exploited. Furthermore, a process has been considered utilizing isolated enzymes (haloperoxidase, halohydrin epoxidase and pyranose-2oxidase) in which D-fructose is a co-product in the transformation of an alkene to an epoxyalkane [1551. 6.2.1. Bacteria that can epo,,idate alkenes Microbial epoxidation of 1-alkenes was first demonstrated by van der Linden [36] who detected the formation of 1,2-epoxyoctane from loctene by heptane-grown Pseudomonas aeruginosa. The capacity to epoxidate 1-octene was only present in alkane-grown cells and it was suggested that the epoxidation of the 1-alkene was effected by the alkane-hydroxylating activity. 2octene was not epoxidated under the same conditions. In 1963 Coon and co-workers demonstrated octanol-formation in crude extracts of an octanegrown Pseudomonas. It was shown that N A D H and molecular oxygen were required for activity. Further work by the same group revealed that the hydroxylation system of Pseudomonas oleovorans consisted of three proteins (see review of May [37]). Subsequently M a y and Abbott [156] showed that the enzyme system purified from Pseudomonas oleovorans also catalyzes the epoxidation of 1-octene to 1,2-epoxyoctane. It was thus confirmed that the hydroxylation and epoxidation reactions are accomplished by the same monooxygenase system. Further work [157] revealed that with 1-decene as substrate, 1,2-epoxydecane and 9-decene-l-ol were formed as oxidation products. The epoxide formation however, predominated. Using alkadienes it was shown that with decreasing chain length the epoxidation rate decreases rapidly. In contrast, the hydroxylation rate of the methyl-group of an alkane is less affected by the chain length. As a consequence, 253 propene and 1-butene are only hydroxylated by the enzyme system from Pseudomonas oleovorans [1571. The methane monooxygenase from Methylococcus capsulatus (Bath), which is also a multicomponent enzyme and which has been very thoroughly studied by the group of Dalton (see [158] for a recent publication), exhibits a different substrate specificity. Unlike the terminal alkane hydroxylase from Pseudomonas oleovorans the methane monooxygenase oxidizes n-alkanes to mixtures of the corresponding 1- and 2-alcohols [159]. Using soluble extracts of Methylococcus capsulatus (Bath) Dalton and co-workers also showed that the short-chain 1- and 2-alkenes were epoxidated, in contrast to the hydroxylation reactions which were observed with extracts from Pseudomonas oleovorans [157]. Of the 31 different compounds oxidized by cell free extract from Methylococcus capsulatus (Bath) [159] only 12 were oxidized by whole cells [59]. With the exception of chloromethane and bromomethane the oxidation of these compounds, including the alkenes, was dependent on, or significantly stimulated by, formaldehyde. It was suggested that the oxidation of formaldehyde supplied the reducing power necessary for the in vivo oxidation of the various compounds by methane monooxygenase. With chloromethane and bromomethane the probable reaction product of monooxygenase activity is formaldehyde, thus explaining the inability of exogenously supplied formaldehyde to stimulate the oxidation of these halomethanes. The restricted range of compounds oxidized by whole cells, compared with cell-free extracts of Methylococcus capsulatus (Bath), was not further discussed [59]. With Methylosinus trichosporium OB3b Higgins et al. found that with whole cells of this methanotroph a much larger range of compounds was oxidized [79]. However, no oxidation rates were presented. The apparent differences in the range of compounds oxidized by whole cells of Methylococcus capsulatus (Bath) and Methylosinus trichosporium OB3b may, in part, be a result of the different incubation conditions used. Hou et al. compared three different methanotrophs with respect to their capacity to epoxidate various alkenes, and also reported that methane metabolites stimulated epoxide formation rates [160]. There were no significant differences between the tested methanotrops. Besides methanotrophs, various bacteria isolated with short-chain alkanes have also been screened for their capacity to epoxidate short-chain alkenes [60]. In contrast with octane-grown Pseudomonas eleovorans, enrichments with propane as carbon-source resulted in organisms which formed 1,2-epoxypropane from propene. Although traces a 3-hydroxy-l-propene were detected after incubation of propane-grown cells with propene, 1,2epoxypropane was the major oxidation product. Hyman and Wood [61] demonstrated that Nitrosomonas europaea cells containing ammonia monooxygenase activity were capable of forming epoxyethane from ethene. From inhibition and competition experiments they concluded that the epoxidation of ethene is probably accomplished by ammonia monooxygenase. The highest rate of epoxyethane formation was obtained by adding hydrazine as donor of reducing power. Ammonia monooxygenase also oxidized many other alkenes and alkanes [68]. Although epoxides are intermediates in the alkene-metabolism of short-chain alkene-utilizing bacteria, and therefore also substrates for these bacteria, it is possible to produce epoxyalkanes with alkene-utilizing bacteria by exploiting the substrate specificity of the epoxide-degrading enzymic activities of these bacteria [161]. In this way for example, it is possible to form 1,2-epoxypropane with ethene-grown Mycobacteria and epoxyethane with propene-grown cells [162]. Furthermore, Furuhashi et al. [163] reported the accumulation of 1,2-epoxypropane during growth of Nocardia corallina B-276 on propene. A general conclusion concerning the specificity of the monooxygenases from the different types of organisms can be that alkane- and methane-grown bacteria can perform epoxidation as well as hydroxylation reactions, whereas alkene-grown bacteria can only perform the epoxidation of alkenes (Fig. 4). 6.2.2. Biotechnological production of short-chain epoxides The biotechnological production of epoxides 254 Atkene-grown bocterio H]C-( CH2)n- CH= CH2 ~ /0\ HI C-(CHz)n- CH- C H2 0H i H2C ~ (ell2) n- CH = EH2 H~C-(CHz)-CH: CH2 / / , F , /0\ _ n - - ~ ~ . H3C-(CH2)n CH-CHz AlkcLne- g r own bacferlo Fig. 4. General modes of oxidative attack of hydrocarbons by alkene- and alkane-grown bacteria. can be envisaged in several ways. Application of enzyme preparations seems very unrealistic due to the low stability of the monooxygenases in vitro. Employment of whole cells containing m o n o oxygenase activity seems a much more attractive alternative. Two possible modes of operation, using whole cells containing monooxygenase activity towards alkenes, have been described. Furuhashi et al. [163] described the production of 1,2-epoxypropane using Nocardia corallina B-276 growing on propene. Apparently in this organism the oxidation of propene proceeds at a higher rate than the consumption of the product 1,2-epoxypropane. After 5 days of incubation of Nocardia corallina B-276 growing on propene a 1,2-epoxypropane formation of 0.6 grams per litre was reported [163]. The specific rate of 1,2-epoxypropane formation was not reported, but can be estimated to have been approximately 1 nmol rain i (mg dry weight)-1. Alternatively, epoxides can also be produced with non-growing cells in which a monooxygenase activity is present. 6.2.2.1. Epoxidation by non-growing cells. By using non-growing cells for the production of epoxides much higher yields, based on the alkene substrate, can be realized. Indeed most of the literature concerning the biotechnological production of epoxides has focussed on systems using non-growing cells. In Table 2 some typical initial 1,2-epoxyalkane formation rates by resting cellsuspensions of different bacteria are shown. To realize the biotechnological production of short-chain epoxides several novel bioreactor configurations have been proposed. The aim of these gas-solid and multiphase bioreactors is to facilitate the supply of the relatively low water-soluble gaseous substrates (oxygen and alkene) and to efficiently remove the inhibitory product [71,164, 165]. Theoretically the gas-solid bioreactor, with a minimal amount of water surrounding the immobilized biocatalyst, would seem ideal to facilitate mass-transfer of gaseous substrates and product. However, this minimizing of the water-phase can become critical, as for several alkene-utilizers it was shown that lowering the water-activity to 99% resulted in a 70% decrease in the alkene oxidation rate [166]. The multiphase bioreactor proposed for the production of 1,2-epoxypropane [164] has the disadvantage that the biocatalyst has to be immobilized to facilitate separation of the biocatalyst and the organic phase, although for the production of 1,2-epoxyoctane the classical stirred tank reactor with an organic phase has also been studied [167]. The most essential component of a biotechnological process however, is not the bioreactor but the biocatalyst. If non-growing cells are to be used as biocalyst for the biotechnological process of epoxide production two major problems, besides the stability and initial activity of the monooxygenase, are cofactor regeneration and product inhibition. 6.2.2.2. Cofactor regeneration and co-substrate. With Methylococcus capsulatus strains it was shown Table 2 Epoxide formation rates by resting cell-suspensions of various bacteria Organism Growth substrate Formation rate nmol min m g protein ] 1,2epoxyethane 1,2epoxypropane 1,2epoxybutane methane 33 33 11 methane 16 30 4 propane 16 40 5 0 16 14 46 0 0 Methylococcus capsulatus C R L M1 [160] Methylosinus trichosporium OB 3b [160] Brevibacterium CRL56 [60] Mycobacterium E3 [161] ethene Xanthobacter Py2 [162] propene 255 that a co-substrate could enhance initial epoxide formation rates [160,59]. The variations in the magnitude of the reported enhancement can possibly be ascribed to the different cultivation conditions used resulting in different endogenous respiration rates. Although, due to its extremely low growth rate, Nitrosomonas europaea does not seem a suitable biocatalyst for the production of epoxides it is interesting to note the effect of an external source of reducing power on epoxyethane production by this organism. Hyman and Wood [61] reported a 10 to 60-fold increase in the epoxyethane formation rate by whole cells of Nitrosomonas europaea by adding hydrazine as reductant. The highest rate of epoxyethane formation that was reported was 250 nmol min-1 (mg protein) -1. In this case the transformation rate of ethene to epoxyethane was apparently completely determined by the supply of reducing power. In addition to increasing the initial epoxide formation rate a co-substrate can also result in an increased stability of the epoxide formation. Using a small gas-solid bioreactor with Mycobacterium Pyl immobilized on sand it was shown that the addition of propionaldehyde as co-substrate resuited in an increased epoxyethane production over a period of several days [71]. Working with several different ethene-utilizing Mycobacteria immobilized on lava it was found that the different strains did not all react in the same manner with respect to the addition of ethanol or ethylacetate as co-substrate during the oxidation of propene to 1,2-epoxypropane [168]. With strains 2W and Eul the rate of 1,2-epoxypropane formation increased significantly when ethanol or ethylacetate were simultaneously supplied with the substrate propene. Furthermore, with ethylacetate as co-substrate biocatalysis was possible for a prolonged period of time with these two strains. With strain E3 co-substrates had no effect on 1,2-epoxypropane formation. The most obvious co-substrate, the growth substrate ethene, cannot be applied during the 1,2-epoxypropane production for several reasons. Ethene and propene are transformed by the same monooxygenase, so that the presence of ethene results in a lower propene turnover and furthermore the presence of 1,2-epoxypropane competitively inhibits the degradation of epoxyethane [161] and hence the production of the desired reducing equivalents. An alternative is to operate the gas-solid bioreactor in a mode in which ethene and propene are supplied in alternating cycles of 15 h. In this manner 1,2-epoxypropane production, although at low rates, was maintained for 20 days [168]. Besides replenishment of reducing equivalents during the ethene cycle, presumably, de novo synthesis of the alkene monooxygenase also takes place. 6.2.2.3. Product toxicity. The lower epoxides are reactive molecules with the capacity to alkylate biological macromolecules (see [169] for references). Epoxyethane for example, due to its gaseous nature at room temperature, is often used to sterilize heat sensitive materials. Besides high conversion rates the economics of a biotechnological process for the production of epoxides to a large extent also depends on the product concentration that can be achieved. Higher product concentrations result in lower purification costs. For this reason, when comparing potential biocatalysts for the production of epoxides it is relevant to know the epoxide tolerance. Habets-Criitzen and de Bont [169] have compared the short-term effect of 1,2epoxypropane on the propene oxidation rate of an ethene-utilizer, an ethane-utilizer and a methanotroph (Table 3). The 1,2-epoxypropane concentrations resulting in approximately 50% inhibition of the propene oxidation rate by whole cells were 20 mM, 1 mM and 5 mM for the ethene-, ethane- and methaneutilizer respectively. When Mycobacterium E20 was grown on ethene instead of ethane, an inhibition pattern comparable to that of the ethene-grown Mycobacterium E3 was obtained, indicating that Mycobacterium E20 possesses two distinct monooxygenase activities, as previously assumed [39]. By performing respiration experiments with other readily oxidizable substrates at the same 1,2epoxypropane concentrations, it was shown that the inhibition was specific for propene oxidation. Furthermore, using strain E3 it was shown that the inhibition of propene oxidation was irreversible. Apparently the epoxide reacts preferentially with the monooxygenase. Surprisingly, the inhibi- 256 Table 3 Inactivation by 1,2-epoxypropane of propene oxidation by whole cell suspensions and cell-free extracts of different bacteria [169] Organism Mycobacterium E3 (ethene grown) Mycobacterium E20 (ethane grown) 1,2-epoxypropane concentration (mM) Propene oxidation rate (relative activities) Whole Cell-free cells extract 0 5 10 50 100 500 0 0.03 0.09 0.4 100 75 63 38 15 1 1130 84 49 29 1 (methane grown) tion of p r o p e n e - 15 - 7 2.5 Methylosinus trichosporium OB3b 100 100 98 97 91 33 - 0 1O0 1O0 0.5 1 5 10 25 90 78 39 22 5 100 95 72 56 27 oxidation by whole cells of Mycobacterium E3 was m u c h s t r o n g e r t h a n the i n h i b i t i o n o f cell-free extract. H o w e v e r , the results o b t a i n e d with cell-free extracts m a y b e m i s l e a d i n g b e c a u s e u p o n s o n i c a t i o n m o r e t h a n 90% of the in vivo activity is lost. A l k e n e m o n o o x y g e n a s e is a m u l t i c o m p o n e n t e n z y m e [169], so thot it c o u l d b e p o s s i b l e t h a t the 1 , 2 - e p o x y p r o p a n e reacts with the c o m p o n e n t w h i c h is n o t rate l i m i t i n g in vitro. Based o n the relative e p o x i d e t o l e r a n c e of the alkene-utilizing M y c o b a c t e r i a , o n e c o u l d c o n c l u d e that these b a c t e r i a are m o r e s u i t a b l e for the p r o d u c t i o n of e p o x i d e s t h a n m e t h a n e - a n d ethaneutilizing bacteria. A l t h o u g h several p a t e n t s have b e e n registered for the b i o t e c h n o l o g i c a l p r o d u c t i o n of epoxides, e c o n o m i c factors still f a v o u r the use of c h e m i c a l processes, as extensively d o c u m e n t e d b y D r o s z d [171]. H e h o w e v e r also c o n c l u d e d , t h a t the biot e c h n o l o g i c a l p r o d u c t i o n of e p o x i d e s e n r i c h e d in a n o p t i c a l i s o m e r m i g h t be interesting. E p o x i d e s o f three or m o r e c a r b o n a t o m s c o n t a i n at least one a s y m m e t r i c c e n t r e (Fig. 5). If it is p o s s i b l e to p r o d u c e these e p o x i d e s stereospecifically such a p r o d u c t w o u l d have a higher value t h a n the racemic e p o x i d e a n d c o u l d f o r m the s t a r t i n g p o i n t of s o m e interesting s t e r e o c h e m i c a l syntheses. 6.2.3. Stereospecific production of epoxides M a y a n d c o - w o r k e r s were the first to r e p o r t on the e n a n t i o m e r i c c o m p o s i t i o n of b i o l o g i c a l l y p r o d u c e d epoxides. T h e 1 , 2 - e p o x y o c t a n e p r o d u c e d f r o m 1-octene b y Pseudomonas oleovorans was 92% in the R - f o r m a n d 8% in the S - f o r m (see [172]). W i t h several o t h e r b a c t e r i a 1,2-epoxyoctane, 1,2e p o x y d e c a n e , 1 , 2 - e p o x y t e t r a d e c a n e a n d 1,2-epoxyh e x a d e c a n e were also p r e d o m i n a n t l y p r o d u c e d in the R - f o r m (see [172]). T h e s e strains were n o t active with s h o r t - c h a i n alkenes. T h e 1,2-epoxytet r a d e c a n e p r o d u c e d with Nocardia corallina B-276 was m o r e t h a n 86% in the R - f o r m [163]. A l t h o u g h they d i d n o t d e t e r m i n e the e n a n t i o m e r i c c o m p o s i tion F u r u h a s h i et al. a s s u m e d t h a t the 1,2-epoxyp r o p a n e p r o d u c e d b y the s a m e o r g a n i s m was also p r e d o m i n a n t l y of the R - f o r m . H a b e t s - C r i i t z e n et al. [172] a n a l y z e d the e n a n t i o m e r i c c o m p o s i t i o n of several e p o x i d e s f o r m e d b y a l k e n e - u t i l i z i n g bacteria. W h e n g r o w n on alkenes all these strains also p r e d o m i n a n t l y f o r m e d the R - f o r m o f 1,2e p o x y p r o p a n e a n d 1 , 2 - e p o x y b u t a n e . O n e strain, Mycobacterium E20, w h e n g r o w n o n ethane, p r o d u c e d r a c e m i c 1 , 2 - e p o x y p r o p a n e [172]. This conf i r m e d the a s s u m p t i o n that Mycobacterium E20 c o n t a i n s different m o n o o x y g e n a s e s d e p e n d i n g on the g r o w t h s u b s t r a t e used. This o b s e r v a t i o n with H R 0 HI 1 ~ 1 1 H 0 R H R-1,2-epoxyaqkane HI 1 ~ 1 1 H H R .S' 1 , 2 - e p o x y a l k a n e Fig. 5. Epoxidation of 1-alkenes can result in two stereoisomers of the 1,2-epoxyalkanes. 257 Table 4 Enantiomeric composition of epoxyalkanes produced by various bacteria a Bacterial strain Methylococcus capsulatus (Texas) Methylosinus trichosporium OB3b Pseudomonas sp. P9y Nocardia sp. TB1 Nocardia sp. TB1 Mycobacterium parafortuTtum E3 Mycobacterium aurum L1 Mycobacterium aurum L1 Xanthobacter sp. Py2 Nocardia sp. By1 Nocardia sp. IP1 Nocardia sp. IP1 Nocardia sp. IP1 Nocardia corallina B-276 Nocardia corallina B-276 Nitrosomonas europaea ATCC 19178 growth substrate methane methane propane butane trans-butene ethene ethene vinyl chloride propene 1-butene butane 1,3-butadiene isoprene ethene propane CO2/NH 4 1,2epoxypropane R S 1-chloro-2,3epoxypropane R S 1,2epoxybutane R S 56 54 45 68 60 91 99 99 91 90 91 86 87 32 52 48 nd nd 45 69 63 84 90 87 44 46 55 32 40 9 1 1 9 10 9 14 13 68 48 52 - - 59 60 99 98 97 41 40 1 2 3 1 99 1 99 99 99 99 83 89 14 1 1 1 17 11 86 trans-2,3epoxybutane R S nd nd 55 31 37 16 10 13 - - 87 95 99 84 84 29 13 5 1 16 16 71 51 50 49 50 - - 85 87 82 89 78 81 81 80 80 73 - 15 13 18 11 22 19 19 20 20 27 - a [173] relative values in %, +2%. Mycobacterium E20 prompted further investigations concerning the enantiomeric composition of short-chain epoxyalkanes produced by different bacteria [173]. The epoxides formed by methane utilizers tested were racemic (Table 4). Propaneand butane-grown strains tested also produced racemic epoxides. In contrast all strains grown on unsaturated hydrocarbons formed optically active epoxyalkanes. Two Nocardia species, strains IP1 and B-276 differed in that they also produced predominantly the R-form of the optically active unsubstituted epoxides, after growth on alkanes. Interestingly, the ammonia monooxygenase from Nitrosomonas europaea exhibited an opposite stereospecificity compared with all the other strains that produced optically active epoxides. The S-form of 1-chloro-2,3-epoxypropane has the same steric conformation as the R-form of 1,2-epoxypropane. Of the above epoxides, 1chloro-2,3-epoxypropane is the most interesting chiral starting material for the synthesis of optically active compounds due to the presence of two different reactive groups. Unfortunately, 1-chloro2,3-epoxypropane is also the most inhibitory of the above epoxides, so that even though the optical purity can be very high, it will prove difficult to produce it economically in a biotechnological process. Recently several other interesting epoxides for the synthesis of chiral products, nave been produced using alkene-utilizers [174], but unfortunately the optical purity was not as high as with 1-chloro-2,3-epoxypropane. Stereospecific epoxidation of substituted alkenes has also been reported [175]. Using bacteria belonging to the genera Rhodococcus, Mycobacterium, Nocardia and Pseudomonas it was possible to epoxidate 4-(2methoxyethyl)phenylallylether stereospecifically yielding optically pure epoxides in the S-config0 / Hu ~ l l CH3 CH3 X0 H3C t l ~ u H H H CH3 (2S.3S) (2R,3R) Degradation products Fig. 6. Stereoselective degradation of 2S,3S-epoxybutane by Xanthobacter Py2 resulting in optically pure 2R,3R-epoxybutane [176]. 258 uration. These epoxides were subsequently chemically transformed to the r - b l o c k e r S ( - ) Metoprolol in optical purities ranging from 95 to 99%. An alternative approach leading to optically active epoxides is stereoselective degradation of one of the isomers. It was recently shown that using the propene-grown Xanthobacter Py2 it was possible to produce enantiomerically pure 2r,3repoxybutane from a racemic mixture of trans-2,3epoxybutane due to the stereoselective degradation of only the 2s,3s-isomer (Fig. 6) [176]. Although very high enantiospecificities have been observed we know of only one strain which is used commercially to produce an optically active 1,2-epoxide. Serva (Heidelberg, F.R.G.) supplies 1,2-epoxytetradecane consisting of at least 97% R-isomer which has been produced using Nocardia corallina B-276 according to the procedure described by Furuhashi and Takagi [177]. It is not quite clear how this value of 97% relates to the value reported earlier [163] of 1,2-epoxytetradecane produced by the same strain containing more than 86% of the R-isomer. Summarizing one can conclude that the choice of organism is crucial when considering the biotechnological production of optically active epoxides. General rules are difficult to make. One could be that methane-utilizers form epoxides racemically whereas alkene-utilizers generally produce optically active epoxides. Alkane-utilizers form a less homogenous group with respect to the stereospecificity of performed epoxidation reactions. When considering the production of novel epoxides it is generally not possible to predict which bacterial strain will produce the epoxide in the highest optical purity [174]. Therefore a screening programme is necessary to select the best organism for a specific epoxidation reaction. 7. C O N C L U D I N G R E M A R K S Summarizing the previous sections one can conclude that a surprising amount of work concerning the microbial degradation and formation of short-chain unsaturated hydrocarbons has been published during the last few decades. The earlier work mainly concerned the role of ethene in soil ecosystems in relation to its plant-hormone activity. Subsequent work focused on the degradation of the various short-chain olefins and questioned the previously assumed anthropogenic nature of many of the compounds under review. The capacity to degrade the short-chain olefins is apparently limited to a few genera, e.g. Mycobacterium, Nocardia, Rhodococcus and Xanthobacter. It is noteworthy that Pseudomonads, which are generally considered to be metabolically very versatile, were only isolated in a few cases e.g. with olefins of intermediate chain length (1-hexene) or substituted olefins (styrene and butynol). The possibility of producing (optically active) epoxides with (alkene-utilizing) microorganisms has inspired several research groups during the last decade. Other recent publications concerning applied aspects bacterial alkene metabolism focus on the potential of using microorganisms to produce olefins from renewable sources and the possibilities of biologically degrading environmental contaminants like chlorinated ethenes. In spite of the significant research efforts described above, many questions remain to be answered. A detailed understanding of many degradative pathways is still lacking and the significance of the olefin-degrading organisms in the natural ecosystem still needs quantitation. Future work could thus lead to the discovery of novel enzymatic activities and should result in a better understanding as to how these olefin degrading microorganisms have evolved and of their function in natural ecosystems. REFERENCES [1] Dagley, S. (1985) Microbial metabolism of aromatic compounds, in Comprehensive Biotechnology, Vol. 1, (Bull, A.T. and Dalton, H., eds.), pp. 483-505, Pergamon Press, Oxford. [2] Gibson, D.T. and Subramanian, V. (1984) Degradation of aromatic hydrocarbons, in Microbial Degradation of Organic Compounds (Gibson D.T., ed.), pp. 181-252, Marcel Dekker, New York. [3] Atlas, R.M. (1981) Microbial degradation of petroleum hydrocarbons: an environmental perspective. Microbiol. Rev. 45, 180-209. [4] Btihler, M. and Schindler, J. (1984) Aliphatic hydro- 259 [5] [6] [7] [8] [9] [10] [11] [12] [13] [14] [15] [16] [17] [18] [19] [20] [21] carbons, in Biotechnology, Vol. 6A, Biotransformations (Kieslich, K., ed.), pp. 370-372, Chemie Verlag, Weinheim, F.R.G. Anthony, C. (1986) Bacterial oxidation of methane and methanol. Adv. Microbiol. Physiol. 27, 113-210. Krasnobajew, V. (1984) Terpenoids, in Biotechnology, Vol. 6A, Biotransformations (Kieslich, K., ed.), pp. 97-125, Chemie Verlag, Weinbeim, F.R.G. Sawada, S. mad Totsuka, T. (1986) Natural and anthropogenic sources and fate of atmospheric ethylene. Atmos. Environ. 15, 821-832. Britton, L.N. (1984) Microbial degradation of aliphatic hydrocarbons, in Microbial Degradation of Organic Compounds (Gibson, D.T. ed.), pp. 89-129, Marcel Dekker, New York. Barnes, L.J., Drodz, J.W., Harrison, D.E.F. and Hamer, G.J. (1976) Process considerations and techniques specific to protein production from natural gas, in Microbial Production and Utilization of Gases. (Proc. Symp. 1975) (Schlegel, H.G., Gottschalk, G. and Pfennig, N., eds.), pp. 301-315, E. Goltse KG, Goettingen, F.R.G. Drozd, J.W. and McCarthy, P.W. (1980) Mathematical model of microbial hydrocarbon oxidation, in Microbial Growth on C 1 Compounds (Dalton, H., ed.), Heyden and Son, London. Ginkel, C.G. van, Habets-Criitzen, A.Q.H., Last, A.R.M. van der and Bont, J.A.M. de (1987) A description of microbial growth on gaseous alkenes in a chemostat culture. Biotechnol. Bioeng. 30, 799-804. Chemiekaarten (1988) Stuurgroep-chemiekaarten (NVVK, VI, VNCI). Landolt-BiSrnstein (1976) Gleichgewicht der Absorption von Gasen in Fliissigkeiten, IV. Band, 4. Teil, Bandteil c, Springer-Verlag, Berlin. Gossett, J.M. (1987) Measurement of Henry's law constant for C 1 and C 2 chlorinated hydrocarbons. Environ. Sci. Technol. 21,202-208. Abeles, F.B. (1973) Ethylene in Plant Biology, Academic Press, New York. Fuchs, Y. and Chalutz, E. (1984) Ethylene: Biochemical, Physiological and Applied Aspects. Martinus Nijhoff, The Hague. McKeon, T.A. and Yang, S.F. (1987) Biosynthesis and metabolism of ethylene, in Plant hormones and their role in plant growth and development (Davies, P.J., ed.), pp. 94-112, Martinus Nijhoff Publishers, Dordrecht, The Netherlands. Rasmussen, R.A. (1970) Isoprene: Identified as a foresttype emission to the atmosphere. Environ. Sci. Technol. 4, 667-671. Tingey, D.T., Manning, M., Grothaus, L.C. and Burns, W.F. (1979) The influence of light and temperature on isoprene emission rates from live oak. Physiol. Plant. 47, 112-118. Ayers, G.P. and Gillett, R.W. (1988) Isoprene emissions from vegetation and hydrocarbon emissions from bushfires in tropical Australia. J. Atmos. Chem. 7, 177-190. Fogel, M.M., Taddeo, A.R. and Fogel, S. (1986) Biode- gradation of chlorinated ethenes by a methane-utilizing mixed culture. Appl. Environ. Microbiol. 51, 720-724. [22] Wilson, J.T. and Wilson, B.H. (1985) Biotransformation of trichloroethylene in soil. Appl. Environ. Microbiol. 49, 242-243. [23] Whitby, R.A. and Altwicker, E.R. (1978) Acetylene in the atmosphere: sources, representative ambient concentrations and ratios to other hydrocarbons. Atmos. Environ. 12, 1289-1296. [24] Arnts, R.R. and Meeks, S.A. (1981) Biogenic hydrocarbon contribution to the ambient air of selected areas. Atmos. Environ. 15, 1643-1651. [25] Sexton, K. and Westberg, H. (1984) Nonmethane hydrocarbon composition of urban and rural atmospheres. Atmos. Environ. 6, 1125-1132. [26] Geigert, J., Neidleman, S.L., Dalietos, D.J. and DeWitt, S.K. (1983) Haloperoxidases: enzymatic synthesis of a,/3-hatohydrins from gaseous alkenes. Appl. Environ. Microbiol. 45, 366-374. [27] Geigert, J., Neidleman, S.L. and Dalietos, D.J. (1983) Novel haloperoxidase substrates. J. Biol. Chem. 258, 2273-2277. [28] Geigert, J., Lee, T.D., Dalietos, D.J., Hirano, D.S. and Neidleman, S.L. (1986) Epoxidation of alkenes by chloroperoxidase catalysis. Biochem. Biophys. Res. Commun. 136, 778-782. [29] Geigert, J., Neidleman, S.L., Dalietos, D.J. and DeWitt, S.K. (1983) Novel haloperoxidase reaction: synthesis of dihalogenated products. Appl. Environ. Microbiol. 45, 1575-1581. [30] Wever, R., Boer, E. de, Plat, H. and Krenn, B.E. (1987) Vanadium: An element involved in the biosynthesis of halogenated compounds and nitrogen fixation. FEBS Lett. 216, 1-3. [31] Gallo, M., Bertrand, J.C., Roche, B. and Azoulay, E. (1973) Alkane oxidation in Candida tropicalis. Biochim. Biophys. Acta 296, 624-638. [32] Iizuka, H., Iida, M., Unami, Y. and Hoshino, Y. (1968) n-Decane dehydrogenation by a cell-free extract of Candida rugosa. Z. Alg. Mikrobiol. 8, 145-149. [33] Klug, M.J. and Markovetz, A.J. (1971) Utilization of hydrocarbons by micro-organisms. Adv. Microbiol. Physiol. 5, 1-39. [34] Bruyn, J. (1954) An intermediate in the oxidation of hexadecene-1 by Candida lipolytica. Koninkl. Akad. Wetenschap. Proc. Ser. C. 57, 41-45. [35] Ishikura, T. and Foster, J.W. (1961) Incorporation of molecular oxygen during microbial utilization of olefins. Nature 192, 892-893. [36] Linden, A.C. van der (1963) Epoxidation of a-olefins by heptane-grown Pseudomonas cells. Biochim. Biophys. Acta 77, 157-159. [37] May, S.W. (1979) Enzymatic epoxidation reactions. Enzyme Microbiol. Technol. 1, 15-22. [38] Bont, J.A.M. de and Harder, W. (1978) Metabolism of ethylene by Mycobacterium E20. FEMS Microbiol. Lett. 3, 89-93. [39] Bont, J.A.M. de, Attwood M.M., Primrose, S.B. and 260 [40] [41] [42] [43] [44] [45] [46] [47] [48] [49] [50] [51] [52] [53] [54] [55] [56] Harder, W. (1979) Epoxidation of short chain alkenes in Mycobacterium E20: the involvement of a specific mono-oxygenase. FEMS Microbiol. Lett. 6, 183-188. Wallen, L.L,, Benedict, R.G. and Jackson, R.W. (1962) The microbiological production of 10-hydroxystearic acid from oleic acid. Arch. Biochem. Biophys. 99, 249-253. Schoepfer, G.J. (1966) Stereospecific conversion of oleic acid to 10-hydroxystearic acid. J. Biol. Chem. 241, 5441-5447. Iida, M. and Iizuka, H. (1971) Enzymatic conversion of 1-decene to decyl alcohol by Candida rugosa JF 101. Z. Alg. Mikrobiol. 11, 301-305. Schink, B. (1985) Degradation of unsaturated hydrocarbons by methanogenic enrichment cultures. FEMS Microbiol. Ecol. 31, 69-77. Kellens, M.J., Goderis, H.L. and Tobback, P.P. (1986) Biohydrogenation of unsaturated fatty acids by a mixed culture of rumen microorganisms. Biotechnol. Bioeng. 28, 1268-1276. Vogel, T.M. and McCarty, P.L. (1985) Biotransformation of tetrachloroethylene to trichloroethylene, dichloroethylene, vinyl chloride and carbon dioxide under methanogenic conditions. Appl. Environ. Microbiol. 49, 10801083. Birch-Hirschfeld, L. (1932) Die Umsetzung yon Acetylen durch Mycobacterium lactieola. Zentralbl. Bakteriol. II Abt. 86, 113-128. Kanner, D. and Bartha, R. (1979) Growth of Nocardia rhodochrous on acetylene gas. J. Bact. 139, 225-230. Bont, J.A.M. de and Peck, M.W. (1980) Metabolism of acetylene by Rhodococcus A1. Arch. Microbiol. 127, 99-104. Topp, E. and Germon, J.-C. (1986) Acetylene metabolism and stimulation of denitrification in an agricultural soil. Appl. Environ. Microbiol. 52, 802-806. Tam, T.Y., Mayfield, C.I. and Inniss, W.E. (1983) Aerobic acetylene utilization by stream sediment and isolated bacteria. Curr. Microbiol. 8, 165-168. Kanner, D. & Bartha, R. (1982). Metabolism of acetylene by Nocardia rhodochrous, J. Bact. 150, 989-992. Watanabe, I. and Guzman, M.R. de (1980) Effect of nitrate on acetylene disappearance from anaerobic soil. Soil Biol. Biochem. 12, 193-194. Culbertson, C.W., Zehnder, A.J.B., and Oremland, R.S. (1981) Anaerobic oxidation of acetylene by estuarine sediments and enrichment cultures. Appl. Environ. Microbiol. 41, 396-403. Schink, B. (1985) Fermentation of acetylene by an obligate anaerobe, Pelobacter acetylenicus sp. nov. Arch. Microbiol. 142, 295-301. Yamada, E.W. and Jakoby, W.B. (1958) Enzymatic utilization of acetylenic compounds (I) An enzyme converting acetylene-dicarboxylicacid to pyruvate. J. Biol. Chem. 233, 706-711. Yamada, E.W. and Jakoby, W.B. (1959) Enzymatic utilization of acetylenic compounds (II) Acetylenemonocarboxylic acid hydrase. J. Biol. Chem. 234, 941-945. [57] Bont, J.A.M. de, Scholten, J. and Tweel, W.J.J, van den (1985) Isolation of microorganisms on 3-butyn-l-ol and other acetylenic compounds. Curr. Microbiol. 12, 267-272. [58] Tweel, W.J.J. van den and Bont, J.A.M. de (1985) Metabolism of 3-butyn-l-ol by Pseudomonas BB1. J. Gen. Microbiol. 131, 3155-3162. [59] Stirling, D.I. and Dalton, H. (1979) The fortuitous oxidation and cometabolism of various carbon compounds by whole-cell suspensions of Methylo¢occus capsulatus (Bath). FEMS Microbiol. Lett. 5, 315-318. [60] Hou, C.T., Patel, R., Laskin, A.I., Barnabe, N. and Barist, I. (1983) Epoxidation of short-chain alkenes by resting-cell suspensions of propane-grown bacteria. Appl. Environ. Microbiol. 46, 171 177. [61] Hyman, M.R. and Wood, P.M. (1984) Ethylene oxidation by Nitrosomonas europaea. Arch. Microbiol. 137, 155-158. [62] Heyer, J. (1976) Mikrobi~lle Verwertung von Athylen. Z. Allg. Mikrobiol. 16, 633-637. [63] Bont, J.A.M. de (1976) Oxidation of ethylene by soil bacteria. Antonie van Leeuwenhoek 42, 59-71, [64] Mahmoudian, M. and Leak, D.J. (1988) Personal communication. [65] Ginkel, C.G. van, Welten, H.G.J. and Bont, J.A.M. de (1987) Oxidation of gaseous and volatile hydrocarbons by selected alkene-utihzing bacteria. Appl. Environ. Microbiol. 53, 2903-2907. [66] Weijers, C.G.A.M., Haan, A. de and Bont, J.A.M. de (1988) Microbial production and metabolism of epoxides. Microbiol. Sci. 5, 156-159. [67] Cerniglia, C.E., Blevins, W.T. and Perry, J.J. (1976) Microbial oxidation and assimilation of propylene. Appl. Environ. Microbiol. 32, 764-768. [68] Hyman, M.R., Murton, I.B. and Arp, D.J. (1988) Interaction of ammonia monooxygenase from Nitrosomonas europaea with alkanes, alkenes and alkynes. Appl. Environ. Mirobiol. 54, 3187-3190. [69] Tausz, J. and Donath, P. (1930) Ueber die Oxydation des Wasserstoffs und Kohlenwasserstoffe mittels Bakterien. Hoppe-Seyler's Z. Physiol. Chem. 190, 141-168. [70] Bont, J.A.M. de, Primrose, S.B., Collins, M.D. and Jones, D. (1980) Chemical studies on some bacteria which utilize gaseous unsaturated hydrocarbons. J. Gen. Microbiol. 117, 97--102. [71] Bont, J.A.M. de, Ginkel, C.G. van, Tramper, J. and Luyben, K.Ch.A.M. (1983) Ethylene oxide production by immobilized Mycobacterium Pyl in a gas/solid bioreactor. EnZyme Microbiol. Technol. 5, 55-60. [72] Ginkel, C.G. van and Bont, J.A.M. de (1986) Isolation and characterization of alkene-utilizing Xanthobacter spp. Arch. Microbiol. 145, 403-407. [73] Bont, J.A.M. de, Dijkenl J.P. van and Ginkel, C.G. van (1982) The metabolism of 1,2-propanediol by the propylene oxide utilizing bacterium Nocardia A60. Biochim. Biophys. Acta 714, 465-470. [74] Hartmans, S. and Bont, J.A.M. de (1986) Acetol mono- 261 oxygenase from Mycobacterium Pyl cleaves acetol into acetate and formaldehyde. FEMS Microbiol. Lett. 36, 155-158. [75] Fujii, T., Ogawa, T. and Fukuda, H. (1985) A screening system for microbes which produce olefin hydrocarbons. Agfic. Biol. Chem. 49, 651-657. [76] Ginkel, C.G. van, Welten, H.G.J., Hartmans, S. and Bont, J.A.M. de (1987) Metabolism of trans-2-butene and butane in Nocardia TB1. J. Gen. Microbiol. 133, 1713-1720. [77] Ginkel, C.G. van, Jong, E. de, Tilanus, J.W.R. and Bont, J.A.M. de (1987). Microbial oxidation of isoprene, a biogenic foliage volatile and of 1,3-butadiene, an antropogenic gas. FEMS Microbiol. Ecol. 45, 275-279. [78] Watkinson, R.J. and Somerville, H.J. (1976) The microbial utilization of butadiene, in Proceedings International Biodegradation Symposium (3rd) Applied Science, Essex, pp. 35-42. [79] Higgins, I.J., Hammond, R.C., Sariaslani, F.S., Best, D., Davies, M.M., Tryhorn, S.E. and Taylor, F. (1979) Biotransformation of hydrocarbons and related compounds by whole organism suspensions of methane-grown Methylosinus trichosporium OB3b. Biochem. Biophys. Res. Comm. 89, 671-677. [80] Furuhashi, K., Shintani, M. and Takagi, M. (1986) Effects of solvents on the production of epoxides by Nocardia corallina B-276. Appl. Microbiol. Biotechnol. 23, 218-223. [81] Omori, T., Jigami, Y. and Minoda, Y. (1975) Isolation, identification and substrate assimilation specificity of some aromatic hydrocarbon utilizing bacteria. Agric. Biol. Chem. 39, 1775-1779. [82] Sielicki, M., Focht, D.D. and Martin, J.P. (1978) Microbial transformations of [14C] styrene in soil and enrichment cultures. Appl. Environ. Microbiol. 35, 124-128. [83] Shirai, K. and Hisatsuka, K. (1979) Production of /3phenetyl alcohol from styrene by Pseudomonas 305-STR1-4. Agric. Biol. Chem. 43, 1399-1406. [84] Baggi, G., Boga, M.M., Catelani, D., Galli, E. and Traccain, V. (1983) Styrene catabolism by a strain of Pseudomonas fluorescens. System. Appl. Microbiol. 4, 141-147. [85] Churchman, J. and Grbic-Galic, D. (1987) Anaerobic transformation of styrene by methanogenic consortia and isolation of pure cultures. Abstracts of Annual Meeting of the American Society for Microbiology p. 287. [86] Hartmans, S., Bont, J.A.M. de, Tramper, J. and Luyben, K.Ch.A.M. (1985) Bacterial degradation of vinyl chloride. Biotechnol. Lett. 7, 383-388. [87] Vogel, T.M., Criddle, C.S. and McCarty, P.L. (1987) Transformations of halogenated aliphatic compounds. Environ. Sci. Technol. 21, 722-736. [88] Janssen, D.B., Grobben, G., Hoekstra, R., Oldenhuis, R. and Witholt, B. (1988) Degradation of trans-l,2-dichloroethene by mixed and pure cultures of methanotrophic bacteria. Appl. Microbiol. Biotechnol. 29, 392-399. [89] Little, C.D., Palumbo, A.V., Herbes, S.E., Lidstrom, M.E., Tyndall, R.L. and Gilmer, P.J. (1988) Trichloro- ethylene biodegradation by a methane-utilizing bacterium. Appl. Environ. Microbiol. 54, 951-956. [90] Nelson, M.J.K., Montgomery, S.O. and Pritchard, P.H. (1988) Trichloroetheylene metabolism by microorganisms that degrade aromatic compounds, Appl. Environ. Microbiol. 54, 1703-1708. [91] Wackett, L.P. and Gibson, D.T. (1988) Degradation of trichloroethylene by toluene dioxygenase in whole-cell studies with Pseudomonas putida F1. Appl. Environ. Microbiol. 54, 1703-1708. [92] Fathepure, B.Z. and Boyd, S.A. (1988) Reductive dechlorination of perchloroethylene and the role of methanogens. FEMS Microbiol. Lett. 49, 149-156. [93] Primrose, S.B. (1979) Ethylene and agriculture: the role of the microbe. J. Appl. Bacteriol. 46, 1-25. [94] Yang, S.F. and Hoffman, N.E. (1984) Ethylene biosynthesis and its regulation in higher plants. Ann. Rev. Plant. Physiol. 35, 155-189. [95] Primrose, S.B. (1977) Evaluation of the role of methional, 2-keto-4-methylthiobutyric acid and peroxidase in ethylene formation by Escherichia coli. J. Gen. Microbiol. 98, 519-528. [96] Billington, D.C., Golding, B.T. and Primrose, S.B. (1979) Biosynthesis of ethylene from methionine. Biochem. J. 182, 827-836. [97] Ince, J.E. and Knowles, C.J. (1986) Ethylene formation by cell-free extracts of Escherichia coli. Arch. Microbiol. 146, 151-158. [98] Fukuda, H., Fujii, T. and Ogawa, T. (1986) Preparation of a cell-free ethylene-forming system from Penicillium digitatum. Agric. Biol. Chem. 50, 977-981. [99] Fukuda, H., Fujii, T. and Ogawa, T. (1988) Production of ethylene by a growth-suppressed mutant of Penicillium digitatum. Biotechnol. Bioeng. 31,620-623. [100] Abeles, F.B., Craker, L.E., Forrence, L.E. and Leather, G.R. (1971) Fate of air pollutants: removal of ethylene, sulfur dioxide and nitrogen dioxide by soil. Science 173, 914-916. [101] Cornforth, I.S. (1975) The persistence of ethylene in aerobic soils. Plant Soil 42, 85-96. [102] Lynch, J.M. and Harper, S.H.T. (1974) Formation of ethylene by a soil fungus. J. Gen. Microbiol. 80, 187-195. [103] Smith, M.A. (1976) Ethylene production by bacteria in reduced microsites in soil and some implications to agriculture. Soil Biol. Biochem. 8, 293-298. [104] Lindberg, T., Granhall, U. and Berg, B. (1979) Ethylene formation in some coniferous forest soils. Soil. Biol. Biochem. 11, 637-643. [105] Sutherland, J.B. and Cook, R.J. (1980) Effects of chemical and heat treatments on ethylene production in soil. Soil. Biol. Biochem. 12, 357-362. [106] Hunt, P.G., Campbell, R.B. and Moreau, R.A. (1980) Factors affecting ethylene accumulation in a Norfolk sandy loam soil. Soil Science 129, 22-27. [107] Lynch, J.M. and Harper, S.H.T. (1980) Role of substrates and anoxia in the accumulation of soil ethylene. Soil. Biol. Biochem. 12, 363-367. 262 [108] Babiker, H.M. and Pepper, I.L. (1984) Microbial production of ethylene in desert soils. Soil Biol. Biochem. 16, 559-564. [109] Smith, K.A. (1977) Ineffectiveness of ethylene as a regulator of soil microbial activity. Soil Biol. Biochem. 10, 269-272. [110] Smith, K.A, and Scott Russel, R. (1969) Occurrence of ethylene, and its significance, in anaerobic soil. Nature, 222, 769-771. [111] Smith, K.A. and Dowdell, R.J. (1974) Field studies of the soil atmosphere; I. Relationships between ethylene, oxygen, soil moisture content, and temperature. J. Soil Science 25, 217-230. [112] Arshad, M. and Frankenberge, W.T. (1988) Influence of ethylene produced by soil microorganisms of etiolated pea seedlings. Appl. Environ. Microbiol. 54, 2728-2732. [113] Ginkel, C.G. van, Welten, H.G.J., Bont, J.A.M. de and Boerrigter, H.A.M. (1986) Removal of ethene to very low concentrations by immobilized Mycobacterium E3. J. Chem. Tech. Biotechnol. 36, 593-598. [114] Green, J. and Dalton, H. (1986) Steady-state kinetic analysis of soluble methane mono-oxygenase from Methylococcus capsulatus (Bath) Biochem. J. 236, 155-162. [115] Ginkel, C.G. van, Welten, H.G.J. and Bont, J.A.M. de (1987) Growth and stability of ethene-utilizing bacteria on compost at very low substrate concentrations. FEMS Microbiol. Ecol. 45, 65-69. [116] Beyer, E.M. (1984) Why do plants metabolize ethylene? in Ethylene: Biochemical, Physiological and Applied Aspects (Fuchs, Y. and Chalutz, E., eds.), pp. 65-74, Martinus Nijhoff, The Hague. [117] Sanders, I.O., Smith, A.R. and Hall, M.A. (1986) Ethylene metabolism and action. Physiol. Plant. 66, 723-726. [118] Smith, K.A. and Restall, S.W.F. (1971) The occurrence of ethylene in anaerobic soil. J. Soil Science 22, 430-443. [119] Goodlass, G. and Smith, K.A. (1977) Effects of organic amendments on evolution of ethylene and other hydrocarbons from soil. Soil Biol. Biochem. 10, 201-205. [120] Fukuda, H., Fujii, T, and Ogawa, T. (1984) Microbial production of C2-hydrocarbons, ethane, ethylene and acetylene. Agric. Biol. Chem. 48, 1363-1365. [121] Fukuda, H., Fujii, T. and Ogawa, T. (1984) Microbial production of C 3- and C4-hydrocarbons under aerobic conditions. Agric. Biol. Chem. 48, 1679-1682. [122] Fukuda, H., Kawaoka, Y., Fujii, T. and Ogawa, T. (1987) Production of a gaseous saturated-hydrocarbons mixture by Rhizopus japonicus under aerobic conditions. Agric. Biol. Chem. 51, 1529-1534. [123] Fujii, T., Ogawa, T. and Fukuda, H. (1988) Preparation of a cell-free, isobutene-forming system from Rhodutorula minuta. Appl. Environ. Microbiol. 54, 583-584. [124] Kanakidou, M., Bonsang, B., Le Roulley, J.C., Lambert, G., Martin, D. and Sennequier, G. (1988) Marine source of atmospheric acetylene. Nature 33, 51-52. [125] Clarke, P.H. (1987) Experimental enzyme evolution and the design of novel biocatalysts, in Physiological and Genetic Modulation of Product Formation (Alberghina, [126] [127] [128] [129] [130] [131] [132] [133] [134] [135] [136] [137] [138] [139] [140] L., Frontali, k. and Hamer, G., eds.), Dechema Monographs vol. 105, pp. 13-28, VCH Weinheim, F.R.G. Wu, T.T. (1978) Experimental evolution in bacteria. Crit. Rev. Microbiol. 6, 33-51. Reineke, W. (1986) Construction of bacterial strains with novel degradative capabilities for chloroaromatics. J. Basic Microbiol. 26, 551-567. Ghosal, D., You, I.-S., Chatterjee, D.K. and Chakrabarry, A.M. (1985) Microbial degradation of halogenated compounds. Science 228, 135-142. Janssen, D.B., Scheper, A., Dijkhuizen, L. and Witholt, B. (1985) Degradation of halogenated aliphatic compounds by Xanthobacter autotrophicus GJ10. Appl. Environ. Microbiol. 49, 673-677. Stucki, G., G~illi, R., Ebershold, H.-R. and Leisinger, Th. (1981) Dehalogenation of dichloromethane by cell extracts of Hyphomicrobium DM2. Arch. Microbiol. 130, 366-371. Keuning, S., Janssen, D.B. and Witholt, B. (1985) Purification and characterization of hydrolytic dehalogenase from Xanthobacter autotrophicus G J10. J. Bacteriol. 163, 635-639. Kohler-Staub, D. and Leisinger, Th. (1985) Dichloromethane dehalogenase of Hyphomicrobium sp. strain DM2. J. Bacteriol. 162, 676-681. Kohler-Staub, D., Hartmans, S., G~illi, R., Suter, F. and Leisinger, Th. (1986) Evidence for identical dichloromethane dehalogenases in different methylotrophic bacteria. J. Gen. Microbiol. 132, 2837-2843. Scholtz, R., Wackett, L.P., Egli, C., Cook, A.M. and Leisinger, Th. (1988) Dichloromethane dehalogenase with improved catalytic activity isolated from a fast-growing dichloromethane-utilizing bacterium. J. Bacteriol. 170, 5698-5704. Turner, E.M., Wright, M., Ward, T., Osborne, D.J. (1975) Production of ethylene and other volatiles and changes in cellulase and laccase activities during the life cycle of the cultivated mushroom, Agaricus bisporus. J. Gen. Microbiol. 91,167-176. Sharma, A. and Padwal-Desai, S.R. (1986) Biogenesis of some antibiotics in the presence of 2-chloroethylphosphonic acid. Appl. Environ. Microbiol. 52, 605-606. Sharma, A., Padwal-Desai, S.R. and Nadkarni, G.B. (1985) Possible implications of reciprocity between ethylene and aflatoxin biogenesis in Aspergillus flavus and Aspergillus parasiticus. Appl. Environ. Microbiol. 49, 79-82. Smith, A.M. (1973) Ethylene as a cause of soil fungistatis. Nature 246, 311-313. Sehippers, B., Boerwinkel, D.J. and Konings, H. (1978) Ethylene not responsible for inhibition of conidium germination by soil volatiles. Neth. J. PI. Path. 84, 101-107. Oremland, R.S. and Taylor, B.F. (1975) Inhibition of methanogenesis in marine sediments by acetylene and ethylene: validity of the acetylene reducton assay for anaerobic microcosms. Appl. Microbiol. 30, 707-709. 263 [141] Schink, B. (1985) Inhibition of methanogenesis by ethylene and other unsaturated hydrocarbons. FEMS Microbiol. Ecol. 31, 63-68. [142] Hyman, M.R. and Arp, D.J. (1988) Acetylene inhibition of metalloenzymes. Anal. Biochem. 173, 207-220. [143] Dilworth, M.J., Eady, R.R., Robson, R.L. and Miller, R.W. (1987) Ethane formation from acetylene as a potential test for vanadium nitrogenase in vivo. Nature 327, 167-168. [144] Knowles, R. (1982) Denitrification. Microbiol. Rev. 46, 43-70. [145] Sprott, G.D., Jarrel, K.F., Shaw, K.M. and Knowles, R. (1982) Acetylene as an inhibitor of methanogenic bacteria. J. Gen. Microbiol. 128, 2453-2462. [146] Oremland, R.S. and Taylor, B.F. (1975) Inhibition of methanogenesis in marine sediments by acetylene and ethylene: validity of the acetylene reduction assay for anaerobic microcosms. Appl. Microbiol. 30, 707-709. [147] Lidstrom, M.E. (1983) Methane consumption in Framvaren, an anoxic marine fjord, Limnol. Oceanogr. 28, 1247-1251. [148] Bont, J.A.M. de, Lee, K.K. and Bouldin, D.F. (1978) Bacterial oxidation of methane in a rice paddy. Ecol. Bull. 26, 91-96. [149] Holzapfel-Pschorn, A., Conrad, R. and Seiler, W. (1985) Production, oxidation and emission of methane in rice paddies. FEMS Microbiol. Ecol. 31, 343-351. [150] Bont, J.A.M. de and Mulder, E.G. (1976) Invalidity of the acetylene reduction assay in alkane-utilizing, nitrogen-fixing bacteria. Appl. Environ. Microbiol. 31, 640647. [151] Ottengraf, S.P.P. (1987) Biological systems for waste gas elimination. Trends Biotechnol. 5, 132-136. [152] Kirchner, K., Hauk, G. and Rehm, H.J. (1987) Exhaust gas purification using immobilised monocultures (biocatalysts). Appl. Microbiol. Biotechnol. 26, 579-587. [153] Ottengraf, S.P.P., Meesters, J.J.P., Oever, A.H.C. van den and Rozema, H.R. (1986) Biological elimination of volatile xenobiotic compounds in biofilters. Bioprocess Eng. 1, 61-69. [154] Higgins, l.J., Best, D.J. and Hammond, R.C. (1983) New findings in methane-utilizing bacteria highlight their importance in the biosphere and their commercial potential. Nature 286, 561-564. [155] Neidleman, S.L. and Geigert, J. (1983) Biological halogenation and epoxidation. Biochem. Soc. Symp. 48, 39-52. [156] May, S.W. and Abbott, B.J. (1973) Enzymatic epoxidation: Comparison between the epoxidation and hydroxylation reactions catalyzed by the ~-hydroxylation systems of Pseudomonas oleovorans. J. Biol. Chem. 248, 1725-1730. [157] May, S.W., Schwartz, R.D., Abbott, B.J. and Zaborsky, O.R. (1975) Structural effects on the reactivity of substrates and inhibitors in the epoxidation system of Pseudomonas oleovorans. Biochim. Biophys. Acta 403, 245-255. [158] Dalton, H. and Higgins, l.J. (1987) Physiology and biochemistry of methylotrophic bacteria, in Microbial Growth on C 1 Compounds (Verseveld, H.W. van and Duinen, J.A., eds.), pp. 89-94, Martinus Nijhoff Publishers, Dordrecht, The Netherlands. [159] Colby, J., Stirling, D.I. and Dalton, H. (1977) The soluble methane mono-oxygenase from Methylococcus capsulatus (Bath). Biochem. J. 165, 395-402. [160] Hou, C.T., Patel, R., Laskin, A.I. and Barnabe, N. (1980) Microbial oxidation of gaseous hydrocarbons: oxidation of lower n-alkenes and n-alkanes by resting cell suspensions of various methylotrophic bacteria, and the effect of methane metabolites. FEMS. Microbiol. Lett. 9, 267-270. [161] Habets-Crtitzen, A.Q.H., Brink, L.E.S., Ginkel, C.G. van, Bont, J.A.M. de and Tramper, J. (1984) Production of epoxides from gaseous alkenes by resting-cell suspensions and immobilized cells of alkene-utilizing bacteria. Appl. Microbiol. Biotechnol. 20, 245-250. [162] Ginkel, C.G. van, Welten, H.G.J. and Bont, J.A.M. de (1986) Epoxidation of alkenes by alkene-grown Xanthobacter spp. Appl. Microbiol. Biotechnol. 24, 334-337. [163] Furuhashi, K., Taoka, A., Uchida, S., Karube, I. and Suzuki S. (1981) Production of 1,2-epoxyalkanes from l-alkenes by Nocardia corallina B-276. Eur. J. Appl. Microbiol. Biotechnol. 12, 39 45. [164] Brink, LE.S. and Tramper, J. (1987) Production of propene oxide in an organic liquid-phase immobilized cell reactor. Enzyme Microbial Technol. 9, 612-618. [165] Hou, C.T. (1984) Propylene oxide production from propylene by immobilized whole cells of Methylosinuz' sp. CRL 31 in a gas-solid bioreactor. Appl. Microbiol. Biotechnol. 19, 1-4. [166] Hamstra, R.S., Murris, M.R. and Tramper, J. (1987) The influence of immobilization and reduced water activity on gaseous-alkene oxidation by Mvcobacterium Pyl and Xanthobacter Py2 in a gas-solid bioreactor. Biotechnol. Bioeng. 29, 884-891. [167] Meer, A.B. van der, Beenackers, A.A.C.M. and Stamhuis, E.J. (1986) Microbial production of epoxides from alkenes in continuous multiphase reactors. Chem, Eng. Sci. 41,607-616. [168] Habets-Crtitzen, A.Q.H. and Bont, J.A.M. de (1987) Effect of co-substrates on 1,2-epoxypropane formation from propene by ethenutilizing Mycobacteria. Appl. Microbiol. Biotechnol. 26, 434-438. [169] Habets-Criitzen, A.Q.H. and Bont, J.A.M. de (1985) Inactivation of alkene oxidation by epoxides in alkeneand alkane-grown bacteria. Appl. Microbiol. Biotechnol. 22, 428-433. [170] Hartmans, S., Somhorst, B.P.M., Voskuilen, G.T.H. and Bont, J.A.M. de (1988) Alkene monooxygenase from Mycobacterium parafortuYtum E3: a multicomponent enzyme, in Proceedings 2nd Netherlands Biotechnology Congress (Breteler, H., Lelyveld, P.H. van and Luyben, K.Ch.A.M. eds.), pp. 306-307, Netherlands Biotechnology Society, Delft. 264 [171] Drozd, J.W. (1987) Hydrocarbons as feedstocks for biotechnology, i n Carbon Substrates in Biotechnology (Stowell, J.D., Beardsmore, A.J., Keevil, C.W. and Woodward, J.R., eds.), IRL Press, Oxford. [172] Habets-Criitzen, A.Q.H., Carlier, S.J.N., Bont, J.A.M. de, Wistuba, D., Schurig, V., Hartmans, S. and Tramper, J. (1985) Stereospecific formation of 1,2-epoxypropane, 1,2-epoxybutane and 1-cbloro-2,3-epoxypropane by alkene-utilizing bacteria. Enzyme Microbiol. Technol. 7, 17-21. [173] Weijers, C.G.A.M., Ginkel, C.G. van and Bont, J.A.M. de (1988) Enantiomeric composition of lower epoxyalkanes produced by methane-, alkane- and alkene-utilizing bacteria. Enzyme Microbiol. Technol. 10, 214-218. [174] Archelas, A., Hartmans, S. and Tramper, J. (1988) Stereoselective epoxidation of 4-bromo-l-butene and 3- butene-l-ol with three alkene-utilizing bacteria. Biocatalysis 1,283-292. [175] Johnstone, S.L., Phillips, G.T., Robertson, B.W., Watts, P.D., Bertola, M.A., Koger, H.S. and Marx, A.F. (1987) Stereoselective synthesis of S - ( - )-/3-blockers via microbially produced epoxide intermediates, in Biocatalysis in Organic Media (Laane, C., Tramper, J. and Lilly, M.D., eds.), Elsevier, Amsterdam. [176] Weijers, C.G.A.M., Haan, A. de and Bont, J.A.M. de (1988) Chiral resolution of 2,3-epoxyalkanes by Xanthobacter Py2. Appl. Microbiol. Biotechnol. 27, 337-340. [177] Furuhashi, K. and Takagi, M. (1984) Optimization of a medium for the production of 1,2-epoxytetradecane by Nocardia corallina B-276. Appl. Microbiol. Biotechnol. 20,6 9.