Microbial Transformation of Pesticides

Microbial Transformation of Pesticides

JEAN-MARC BOLLAG Laboratory of Soil Microbiology, Department of Agronomy, The Pennsylvania State Uniuersity, University Park, Pennsylvania I. Introduction ...................................... 11. Mechanisms of Pesticide Transformation .............. A. Pesticide as a Nutrient Source ................... B. Cometabolism ................................ C. Conjugate Formation .......................... D. Microbial Accumulation of Pesticides ............ 111. Enzymatic Reactions in Pesticide Metabolism .......... A. Oxidative Reactions ............................ B. Reductions ................................... C . Hydrolysis ................................... D. Dehalogenation Mechanisms .................... E. Synthetic Reactions ........................... IV. Chemical Structure and Microbial Transformation Relationship ...................................... A. Microbial Transformation of Pesticidal Groups .... B. Effect of Various Substitutions on Biodegradability . . C. Molecular Recalcitrance and Pesticide Transformation V. Conclusions ..................................... References .......................................

I.

75 77 78 78 80 80 81 81

97 99 103 109 114 115 119 121 122 124

Introduction

The fate of applied xenobiotic compounds, such as pesticides, in the environment is of great importance, since disappearance, persistence, or partial transformation of such a compound determines its usefulness or its potential hazardous effect. There may be chemical and physical factors that influence the fate of a pesticide, but the least predictable transformation is usually caused by microorganisms. Because these compounds have become an integral part of our economy, there can be no question that considerable effort must be expended in order to gain an understanding of the mechanisms of pesticide transformations. There is a need to know the actual biochemical reactions involved in pesticide metabolism, since this can give a basis for the understanding of their short or long persistence in a natural environment and can also contribute to the clarification of the relationship between chemical structure and susceptibility to probable microbial transformations. For this purpose it is necessary to investigate the metabolic reactions and the enzyme systems and to isolate and identify the resulting products in laboratory experiments. The information derived from such studies can serve as a signpost for subsequent investigations in a natural ecosystem which 75

76

JEAN-MARC BOLLAG

is usually too complex as a primary research medium for conclusive results. The soil, for instance, which can be considered as the most complex microbial habitat and an area where pesticides or their derivatives are deposited either by direct application or by decaying foliage, possesses many characteristics for an efficient chemical or physical attack on a molecule and for its removal by adsorption, leaching, volatilization or photodecomposition. On the other hand, the soil simultaneously offers, in many instances, the necessary prerequisites for the proliferation of a vast variety of living organisms. Consequently, it is very difficult to discriminate in a soil environment between microbial, chemical, and physical factors contributing to the removal or transformation of a pesticidal molecule. It was also stated that breakdown products of biological and physicochemical activity in soil are often similar because the reagents-oxygen, water, and nucleophiles-are the same in each instance (Crosby and Li, 1969). Photolysis, for instance, can cause the hydrolysis of esters and amides, dealkylation of amines, and other effects, and the same reaction can be initiated by enzymatic activity resulting in identical “products.” Biological transformation of xenobiotic compounds, for example, pesticides in a soil, freshwater, or estuarine ecosystem, appears to be caused primarily by bacteria, actinomycetes, and fungi. It has to be emphasized that much less attention in research has been devoted to the possible interference of the microfauna and algae, and this subject is only partially covered in this review. Initial research on microbial pesticide metabolism was characterized solely by isolating organisms capable of using a compound as their only source of both carbon and energy, but later it was realized that other mechanisms, such as cometabolic transformation, conjugation reactions, or the mere accumulation of a pesticide within a microbe, are important factors of microbial interference. The complete degradation of a pesticidal molecule to its inorganic parts or its fragmentation into components that can be further used in an oxidative cycle, like the Krebs cycle, removes its potential toxicity completely from the environment. The mechanisms which cause only partial change or temporary removal do not eliminate the potential hazard of an applied chemical or its transformation product in nature. There are several review articles that include aspects on the microbial degradation of pesticides ( Menzie, 1969; Alexander, 1969; Helling et a,?., 1971) and numerous reviews that cover microbial attacks on pesticidal classes or related topics, some of which are referred to in this article. In this review it was attempted to assess the probable transformation capabilities of pesticides by microorganisms and to characterize

MICROBIAL TRANSFORMATION OF PESTICIDES

77

the specific enzymatic reactions that are part of the metabolism for the major known groups of pesticides. II.

Mechanisms of Pesticide Transformation

If a pesticide is exposed to a microbial species, there are four major possibilities for its transformation or inactivation by the organism: (1) the pesticide can serve as a substrate for growth and energy; ( 2 ) the xenobiotic compound can undergo “cometabolism,” i.e., microorganisms transform it, but cannot derive energy for growth from it; ( 3 ) the entire pesticidal molecule or an intermediate of it can be conjugated with naturally occurring compounds; and (4) the pesticide is incorporated and accumulates within the organism. It is self-evident that in many cases the transformation of a pesticide does not occur by only one type of mechanism during its exposure to one organism or to a whole microflora under natural conditions. In addition, a specific compound can be metabolized by various pathways in the environment; consequently, different products can result from the same initial material. All microbial transformations are caused by enzymes, and since all the applied pesticides are foreign materials, possessing a molecular configuration that may not occur in nature, it is understandable that many of the enzymes catalyzing a certain reaction are induced. This often causes an initial lag period until metabolic activity can be determined. Although many enzymes are induced, the transformations that they catalyze are usually reactions also encountered in the metabolism of natural substances. However, it is difficult to predict which molecular change can be expected by a specific microbe, since each group of organisms, even various strains of one genus, can alter a selected molecule differently. For example, the insecticide carbaryll was hydroxylated by different species of Penicillium on the ring, at the side chain, or not at all, respectively ( Bollag and Liu, 1972a); Fusarium muniliforme dealkylated the ethyl group of the herbicide atrazine, while F . roseum showed a stronger activity in the removal of the isopropyl substitution (Kaufman and Blake, 1970); and DDT was metabolized to TDE and a dicofol-like compound by Trichoderma uiride, while variants of the same species produced DDA or DDE ( Matsumura and Boush, 1968). Nevertheless, a certain mode of biological attack can be anticipated on the basis of the molecular structure of the pesticide, and the knowledge acquired should help to foresee such a transformation (Section 111). Cosinion and chemical designation of pesticides referred to in this text are listed in Table IV.

78

JEAN-MARC BOLLAG

AS A. PESTICIDE

A

NUTRIENTSOURCE

From a practical point of view the complete microbial breakdown of an organic molecule to its inorganic components is the desired mechanism if one is interested in avoiding the persistence of a potentially hazardous compound in the environment. As elaborated in laboratory experiments, numerous organic pesticides can serve as the sole carbon or energy source for growth and proliferation of certain microorganisms. If a pesticide can be used in such a way, it is degraded and fragmented to compounds that can be channeled into known oxidative cycles such as the Krebs cycle, and thus the organism can derive all the necessary energy . In order to determine whether a pesticide can serve as the only carbon source needed for growth, the general experimental approach proceeds by an enrichment culture technique. After isolation of the surviving microorganisms, the pesticide is added to a basal salts medium and further observations are usually performed with a pure culture. However, it was also shown that in certain cases a pesticide can be decomposed only in the presence of two different microbial species (Gunner and Zuckerman, 1968). It appears to be an obvious conclusion that in a natural ecosystem like the soil, with an abundance of various microbes, there is an even greater possibility for the single or combined transformation and complete use of a specific pesticide by the microbial population.

B. COMETABOLISM The phenomenon that a microorganism can transform a chemical without deriving energy to support its growth is a relatively recent observation, and its detection and significance is related to the modern use of xenobiotic compounds in various environments. Foster (1962) used the expression “co-oxidation,” Jensen ( 1963 ) suggested the term “cometabolism,” and Ruiz-Herrera and Starkey ( 1969) designated this process as “co-dissimilation,” but although all designations try to express the same thought, it appears that cometabolism is the most general term, and therefore it will be used in this review. The potential importance of cometabolism for the transformation of pesticides was first pointed out by Alexander ( 1967), and the findings of many investigations can now be explained by this process (Table I ) (Horvath, 1972). Cometabolism generally does not result in extensive degradation of a pesticidal molecule, but it can cause a reduction, elimination or probably increase of toxicity in the environment. However, it was also demonstrated that different microorganisms can degrade a certain pesticide

79

MICROBIAL TRANSFORMATION OF PESTICIDES

TABLE I PESTICIDES SUBJECTTO COMETABOLISM A N D ACCUMULATED PRODUCTS Substrate Chlorobenzilate

Product

4,4’-l)ichlorobenzophenone Chloroneb 2,5-Dichloro-4methoxyphenol DDT p,p-Dichlorodiphenylmethane 3,t5-I>ichlorocatechol 3,5-Dichloro-%hy(metabolite of droxymuconic 2,4-D, 2,4,5-T, semialdehyde and 2,3,6-TBA) p,p’-Diehlorodiphen- p-Chlorophenyl aceylmethane (metahtic acid olite of D D T ) 3-Nitrophenol Nitro hydroquinone 2,4,.i-T 2,3,6-TBA

Organism

R hodotorula gracilis Fusarium sp. Aerobacter aerogenes Achromobacter sp.

Reference Miyasaki et a1 (1970) Wiese and Vargas (1973) Wedemeyer (1967) Horvath (1970b)

Hydrogemonas sp.

Focht and AIexander (1971)

Flavobacterium sp.

Raymond and Alexander ( 1971) Horvath (1970a) Horvath (1971)

3,5-L>ichlorocatechol Brevibacterium sp. 3,S-l)ichlorocatechol Brevibacterium sp.

considerably by subsequent cometabolic attack; for instance, the herbicides 2,4,5-trichlorophenoxyacetateand 2,3,6-trichlorobenzoate are converted by a cometabolic oxidation to 3,5-dichlorocatechol by a Breuibacterium sp. (Horvath, 1970a,1971), whereas an Achromobacter sp. was capable of cooxidizing the resulting 3,5-dichlorocatechol to 3,5-dichloro2-hydroxymuconic semialdehyde ( Horvath, 1970b). It was also speculated that cometabolism could account for complete mineralization of a chemical if a carbon and energy source were supplied to mixed microbial populations in the form of a biodegradable analog of the chemical under investigation ( Horvath, 1972). The process of cometabolism is effected by bacteria as well as actinomycetes and fungi, and therefore it can be assumed that its occurrence is widespread in natural ecosystems. Many observations of microbial transformations that could not be understood, since the microbes did not derive any energy or nutritional use from it, are now interpretable by this mode of metabolism. The process of cometabolism, especially as a factor in the microbial transformation of pesticides, requires still further intensive study for a clear understanding of its actual cause, its transformation capabilities as related to the structure of chemicals, and the extent to which such a transformation forms a more or less persistent compound. Although it is difficult to demonstrate unequivocally that microbial cometabolism also occurs under natural conditions,

80

JEAN-MARC BOLLAG

there is little doubt that it takes place, and therefore the ecological importance of this biological reaction has to be fully explored. C. CONJUCATE FORMATION

Whereas the biotransformations described previously include the attack of the original molecule, conjugation reactions are syntheses by which a pesticide, or any of its metabolites, is combined with naturally occurring compounds, generally amino acids or carbohydrates. The formation of a conjugate usually makes the molecule more polar, and therefore more water- and less lipid-soluble. Conjugations of pesticides and other xenobiotic chemicals are common and frequent reactions in all higher organisms, but they have not been found to a similar extent in microorganisms. It is not likely that this observation is an experimental oversight, but it could present a metabolic characteristic of microbes, especially bacteria. Nevertheless, this mode of transformation requires further exploration, since a pesticide, or an intermediate of it, is only bound to another molecule. A conjugated compound can easily be cleaved again, and the released chemicals can subsequently exert a toxic influence. The herbicide amitrole, for instance, was coupled with alanine by Escherichia coli and subsequently incorporated into cellular protein ( Williams et al., 1965). This conjugation reaction occurred presumably since the conjugated metabolite showed structural similarity to histidine and functioned as an analog of histidine. Various examples of conjugate formation are described under Section II1,E.

D. MICROBIALACCUMULATION OF PESTICIDES The possibility that pesticides are incorporated into microorganisms by an active or passive accumulation mechanism provokes special concern, since the microbial interference means-as in a conjugation reaction-only temporary removal of a toxic compound. Most observations of pesticide accumulation within the cells were registered with chlorinated hydrocarbons like DDT, dieldrin, aldrin, and heptachlor. Mycelia of actinomycetes and fungi added to soil containing dieldrin, DDT, and pentachloronitrobenzene accumulated these compounds to levels above ambient concentrations (KO and Lockwood, 1968). This observation also was confirmed with specific bacteria, actinomycetes, and fungi in culture solutions containing DDT and dieldrin (Chacko and Lockwood, 1967). In various studies it was found that not only live bacterial cells, but also autoclaved cells, show a similar uptake of pesticides, which appears to indicate that an actual metabolic factor is not involved in the accumulation process. Johnson and Kennedy (1973) found that the accumulation

MICROBIAL TRANSFORMATION OF PESTICIDES

81

rate of DDT and methoxychlor by autoclaved cells was greater than that for the living bacteria; for instance, after autoclaving the cells of Aerobacter aerogenes, the uptake of methoxychlor was double the amount absorbed by living cells. They suggested that the molecular polarity and lipid solubility influences the retention of the organochlorine insecticides by the bacterial cells. Experiments with yeast, Sacchuromyces cerevisiae, also showed that the adsorption capacity for lindane and dieldrin increased after boiling of the organism, and that the two insecticides could be removed by washing with fresh water (Voerman and Tammes, 1969). Adsorption and concentration of the insecticide aldrin was determined for floc-forming bacteria which were isolated from Lake Erie, and it was suggested that the adsorption capacity of flocculent bacteria might even be evaluated for removal of pesticides in an aqueous environment ( Leshniowsky et al., 1970). Since aquatic microorganisms and plankton in freshwater and marine environments are an important nutrient source for a broad spectrum of aquatic filter-feeding organisms, their accumulation of pesticides can constitute a hazardous link in the food chain to fish and higher vertebrates. Therefore, the findings of extensive biomagnification by these organisms has to provoke considerable concern. A marine diatom, Cylindrotheca closterium, adsorbed and concentrated DDT up to approximately 200-fold from its culture medium containing 0.1 ppm of the insecticide ( Keil and Priester, 1969). Likewise, it was found that cultures of the blue-green alga Anacystis nidulans, the green alga Scenedesmus obliquus, the flagellate Euglena gracilis, and the two ciliates Paramecium bursaria and P . multimicronucleatum concentrated DDT and parathion after exposure for 7 days at a rate of 100 to 964 and 50 to 116 times, respectively (Gregory et al., 1969). Ill.

Enzymatic Reactions in Pesticide Metabolism

Following is an attempt to classify the enzymatic reactions into groups that cover the majority of biotransformations which pesticides undergo. However, it is clear that all attempts to categorize natural processes have their shortcomings; therefore, this should be considered as a trial to assort the essential enzymatic characteristics for easier evaluation.

A. OXLDATIVE REACTIONS 1 . Hydroxylation Introduction of a hydroxyl group to a pesticide is a frequent primary transformation of a molecule resulting in the formation of a compound which can be biologically more reactive, often more polar, and conse-

82

JEAN-MARC BOLLAG

quently more soluble in water. Enzymes that catalyze this reaction have been variously termed “hydroxylases,” “monooxygenases,” or mixed-function oxidases. For plants and animals, the insertion of a hydroxyl group into a compound often provides a center at which conjugation can occur, but this is rarely the purpose of microbial hydroxylations. Hydroxylation can occur with aliphatic as well as aromatic compounds, and often it constitutes only a step in a more complex reaction; for instance, dealkylation reactions proceed via a hydroxylated intermediate, which is, however, in many cases an unstable compound. All studies in which microbial hydroxylation of pesticides was investigated in more detail claim that the reaction takes place only in the presence of oxygen and the reduced form of nicotinamide adenine dinucleotide phosphate ( NADPH ) , or nicotinamide adenine dinucleotide (NADH), indicating that the process is catalyzed by a mixed-function oxidase, whereby the molecular oxygen is apparently incorporated without intermediate water formation. Like most other processes of xenobiotic compounds, more detailed studies on the mechanism of hydroxylation have been performed with mammalian liver microsomal systems ( DaIy, 1971). Hydroxylation of aromatic pesticides is an important step as a tool for introducing polar groups into the molecule as well as a prerequisite for further degradation by ring cleavage. Most observations on hydroxylations at various positions and with different microorganisms were made with phenoxyalkanoate pesticides which probably constitute the group of herbicides whose microbial degradation was most thoroughly studied. While 2,4-D was hydroxylated to the 6-hydroxy derivative by a Pseudomoms sp. (Evans et al., 1971b), the fungus Aspergillus niger produced essentially 2,4-dichloro-5-hydroxyphenoxyacetic acid and, to a lesser extent, 2,5-dichloro-4-hydroxy0-CH,COOH

HO

/ Pseudomonas sp.

Cl

Asperffillus niger OH

83

MICROBIAL TRANSFORMATION OF PESTICIDES

phenoxyacetic acid ( Faulkner and Woodcock, 1964; 1965). The latter product indicates a shift of a chlorine atom coupled with the replacement of a hydroxyl group which will be considered further as a NIH-shift under epoxidation reactions. The herbicide MCPA was transformed similarly to 2,4-D, resulting in the production of the 6-hydroxy (Evans et al., 1963) and the 5-hydroxy derivative (Faulkner and Woodcock, 1964) by a Pseudomonm sp. and A. niger, respectively. Hydroxylation appears to be a common mode of attack by A. niger on phenoxyacetic acids. From 4-chlorophenoxyacetic acid it was possible to isolate 4-chloro-2-hydroxy- and 4-chloro-3-hydroxyphenoxyaceticacid, and the exposure of phenoxyacetic and 2-chlorophenoxyacetic acid to A. niger resulted in the formation of all possible hydroxy derivatives ( Faulkner and Woodcock, 1961; Clifford and Woodcock, 1964). Chlorinated phenoxyalkanoic acids undergo cleavage of the ether linkage by the metabolic activity of various bacterial species resulting in the corresponding phenol. Bollag et al. (1968a) isolated a soluble enzyme preparation from a soil Arthrobacter sp. which converted 2,4-dichlorophenol and 4-chlorophenol to 3,5-dichlorocatechol and 4-ch1orocatecho1, respectively. The enzyme involved appears to be a mixed-function oxidase, since both oxygen and NADPH were required for the hydroxylation reaction. Oxidation of an aromatic amine group can be initiated by N-hydroxylation, which appears to be a major pathway for the oxidation of chlorinated anilines by Fusarium oxysporum. p-Chloroaniline, which constitutes an intermediate of several herbicides, was hydroxylated to pchlorophenylhydroxylamine, which could accumulate temporarily in the growth medium of the fungus up to 76%of the amount theoretically possible ( Kaufman et al., 1973). p-Chlorophenylhydroxylamine was subsequently metabolized to p-chloronitrosobenzene and p-chloronitrobenzene.

+-+Q-Q NH,

NHOH

NO

NO*

c1

c1

c1

c1

The insecticide carbaryl was oxidized by hydroxylation at different positions; from the growth medium of the fungus Gliocladium roseum, it was possible to isolate and identify 1-naphthyl N-hydroxymethyl carbamate as well as 4-hydroxy- and 5-hydroxy-1-naphthyl methylcarbamate, which indicated side chain and ring hydroxylation, respectively ( Liu and Bollag, 1971a).

84

JEAN-MARC BOLLAG

O H I1 I 0 - C -N-CH3

I

O H II I 0- C-N-

O H II I 0-C-N-CH,

&J 4-d CH,OH

/

OH

I:':

0- C-N-CH,

@ OH

Soil fungi were tested in relation to their ability to hydroxylate carbaryl, and it was found that hydroxylation in the side chain or in the positions on the aromatic ring varied qualitatively as well as quantitatively with various fungal species (Bollag and Liu, 1972a). An isolated Mucor species, for example, accumulated essentially ring-hydroxylated products, whereas Aspergillus terreus hydroxylated mostly on the side chain. Considerable differences could be detected even within one genus, where different species hydroxylated at different positions or showed no activity at all. Wallnofer et al. (1972), investigating the metabolism of the systemic fungicide 2,5-dimethyl-3-furancarboxylicacid anilide, found that the pesticide ( 60 pmoles/liter ) was hydroxylated by Rhizopus japonicus to 2-hydroxymethyl-5-methyl- and 2-methyl-5-hydroxymethyl-3-furancarboxylic acid anilide at a ratio of 23 and 12 pmoles/liter, respectively. The formed metabolites were not further degraded by the fungus.

nNH;)-J CH,OH

CH3

2. Dealkylation Numerous pesticides, such as phenylureas, acylanilides, carbamates, s-triazines possess alkyl moieties which very often present active groups

85

MICROBIAL TRANSFORMATION OF PESTICIDES

producing a desired toxic influence. Therefore, dealkylation reactions are of great importance since they are a first step in the detoxication of pesticides and alkyl groups of side chains are frequently the first target of microbial attack. Most of our knowledge on the mechanism of dealkylation originates from studies performed with the microsomal fraction of liver (Gram, 1971), and only a few reports exist on the dealkylating activity of pesticides by microbial enzymes. Usually it is assumed that a dealkylation reaction results in the dealkylated product and an aldehyde: R-X-CHZ-R'

-+

R-X

+

R'-CHO

X can represent an N or 0 atom, and the dealkylation can produce an amine or an alcohol, respectively. Both N- and O-dealkylation are catalyzed by a mixed-function oxidase requiring a reduced nicotinamide nucleotide as a hydrogen donator. a. N-Dealkylation. The mechanism of n-dealkylation is not yet clear. The question arises especially around the possible formation of N-oxide as an intermediate, and studies with microsomal liver systems are not conclusive (Gram, 1971). However, in several cases it was possible to isolate an N-hydroxylated intermediate, which in turn can be metabolized to the dealkylated product. For example, carbaryl is transformed by the fungus Aspergillus terreus to l-naphthyl N-hydroxymethyl carbamate and subsequently to l-naphthyl carbamate ( Liu and Bollag, 1971b). o n

O H II I 0 -C-N--CH,OH

0 II O-C-NH,

--& -fJ$

11 I 0 - C -N-CH,

The N-hydroxymethyl intermediate is also chemically degraded to l-naphthyl carbamate, .but the study gave evidence that, through the additional biological activity, the formation of the dealkylated product was increased considerably. In many other cases of N-dealkylation it was suspected that an N-hydroxyalkyl intermediate might be formed, but these compounds are often chemically unstable and decompose further to the dealkylated product. Hydroxylation of the methyl group in the side chain of carbaryl was also observed with many other soil fungi (Liu and Bollag, 1971a; Bollag and Liu, 1972a), and it can be assumed that further transformation of the 1-naphthyl N-hydroxymethyl carbamate results in the dealkylated product. A carbamate-related amide, the herbicide diphenamid, was stepwise dealkylated by Trichoderma viride and Aspergillus candidus to N-methyl

86

JEAN-MARC BOLLAG

2,2-diphenylacetamide and, subsequently, to 2,2-diphenylacetamide ( Kesner and Ries, 1967) :

Po -qo II

6

HC-C-N,

,CH,

CH,

CH,

II

6

HC-C-N’

H ‘

~

Po HC-C-N, II

0

,H

H

An interesting observation was that diphenamid, the applied herbicide, is less toxic to the target plants than the two dealkylated metabolites. N-Dealkylation has been demonstrated to be a major reaction in the metabolism of dimethyl-substituted phenylureas ( Geissbuhler, 1969). In liquid cultures of mixed soil bacteria, Geissbiihler et al. (1963) isolated and identified 3-(4-chlorophenoxy )phenyl-l-methylurea and 3- ( 4-chlorophenoxy ) phenyhrea from chloroxuron:

/ Similar patterns of dealkylation were also reported for other phenylureas: diuron is stepwise dealkylated in soil to 3-( 3,4-dichlorophenyl) -1methylurea and 3,4-dichlorophenylurea ( Dalton et al., 1966); monolinuron and linuran were dealkylated by an Aspergillus niger sp. (Borner, 1967); metobromuron was converted by the fungus Talaromyces wortmanii to 1-(p-bromophenyl ) -3-methoxyurea and 1-(p-bromophenyl ) 3-methylurea, indicating a dealkylation and dealkoxylation reaction, and subsequently to p-bromophenylurea (Tweedy et al., 1970a); and Rhizoctonia solani metabolized chlorbromuron to the demethylated product ( Weinberger and Bollag, 1972). Wallnofer et al. ( 1973) also found that Rhizopus japonica was active in demethylation of phenylurea herbicides; however, buturon did not lose the N-methyl, but the N-butyryl ( l-methyl-2-propynyl) group, resulting in the formation of 3- (p-chlorophenyl ) -1-methylurea. s-Triazine herbicides are metabolized by soil microorganisms, and the removal of the alkyl side chains appears to be the primary mode of attack. In pure culture studies, simazine was dealkylated by Aspergillus and the ring porfumigatus to 2-chloro-4-ethylamino-6-amino-s-triazine,

MICROBIAL TRANSFORMATION OF PESTICIDES

87

tion of the molecule remained intact (Kearney et al., 1965); attempts to isolate a cell-free preparation from the mycelium were not successful. The N-dealkylation of atrazine to either 2-chloro-4-amino-6-ethylaminos-triazine or 2-chloro-4-an~ino-6-isopropyl-amino-s-triazinewas shown by 12 different soil fungi ( Kaufman and Blake, 1970) : c1 I

All the fungi investigated were able to dealkylate the herbicide by either alkylamino group, but the removal of the ethyl side chain or the isopropyl group by various fungal species was quantitatively different. Aspergillus fumigatus, for example, removed essentially the ethyl moiety, whereas Rhizopus stolonifer metabolized the isopropyl group more readily. In these experiments there was no evidence that the ring of the s-triazine molecule was cleaved. It is noteworthy that dealkylation of s-triazines does not necessarily mean reduction in their herbicidal or phytotoxic activity (Knuesli et al., 1969). Additional pesticides which undergo dealkylation as an initial degradation reaction include : the dipyridyl herbicide paraquat, which was apparently demethylated by an unidentified bacterium ( Funderburk and Bozarth, 1967); trifluralin, which was dealkylated by removal of a propyl group by A . niger (Funderburk et al., 1967); and dinitramine, which is degraded to the dealkylated chemical by a cell extract from A. fumigatus in the presence of NADPH and ferrous ions (Laanio et al., 1973). b. O-Dealkylation. The removal of a methyl or another alkyl group from an oxygen atom functioning as a linkage to the other molecular moiety can be considered as an ether cleavage or an O-dealkylation reaction. Although the enzymatic mechanism for cleavage of the oxygen from the hydrocarbon appears to be very similar in all investigated reactions, in this review O-dealkylation as the removal of an alkyl group was distinguished from cleavage of the ether linkage as the separation of another hydrocarbon from oxygen.

88

JEAN-MARC B O U A G

Axelrod (1956) first demonstrated the enzymatic cleavage of methoxylated compounds by microsomal preparations from rat liver; both oxygen and NADPH were necessary for the conversion of anisole to phenol and formaldehyde. Methoxylated aromatic compounds are demethylated by soil fungi; 0-,m-, and p-methoxybenzoic acid were converted to the corresponding hydroxybenzoic acids and veratric acid was demethylated to vanillic acid by species of Hormodendrum, Haplographium, and Penicillium ( Henderson, 1957). Cell-free preparations from Pseudomonas fluorescens, capable of converting vanillate to protocatechuate and formaldehyde, were first obtained by Cartwright and Smith (1967), and it was established that oxygen, reduced nicotinamide nucleotides, and reduced glutathione are required for the demethylation reaction:

Q COOH

COOH .+

HCHO

OCH,

OH

OH

However, it was not possible to clarify whether the protocatechuate was directly formed from vanillate or indirectly via p-hydroxybenzoate. The 0-dealkylases, induced growth of species of P. flu0rescen.s and Nocardia on 4-alkoxybenzoates, specifically attack the ether linkage and are indifferent to the nature of the alkyl group which, itself, determines reaction rates (Cartwright et al., 1971). Enzymes of an Arthrobacter sp. also converted vanillate to protocatechuate and m-methoxybenzoate to m-hydroxybenzoate with the simultaneous formation of formaldehyde (Raymond and Alexander, 1972). The 0-demethylase system from P . testosteroni was shown to be composed of at least two protein fractions (Ribbons, 1971), and an enzyme extract from P. puticla was further resolved and three protein components were purified ( Bernhardt et al., 1971). Stenersen (1969) found that the insecticide bromophos was oxidized by a double-dealkylation to the bisdemethylated bromophos by the fungi Alternaria tenius and Trichoderma lignorum: CHJO,,/s c1 HO S cH3>odBr HO o

- x d'-

0-Dealkylation was also found with anisole, whose chlorinated derivatives were occasionally found as intermediates during the breakdown of chlorinated phenoxyacetic acids; replacement cultures of Aspergillus niger oxidized anisole to phenol (Bocks et al., 1964). Mycelia of Rhizoctonia solani and some other fungi converted the ( Hock and Sissoil fungicide chloroneb to 2,5-dichloro-4-methoxyphenol

89

MICROBIAL TRANSFORMATION OF PESTICIDES

ler, 1969), and in some cases a further dealkylation to 2,5-dichlorohydroquinone was observed ( Wiese and Vargas, 1973):

@

c1

-.p"

OCH,

OCH,

>cl@cl OH

Boothroyd et al. (1961) found that different fungi demethylate the fungicide griseofulvin at different methoxy groups attached to the molecule. Botrytis allii formed the 2'-demethylgriseofulvin, Cercospora rnelonis produced the 6-demethylgriseofulvin, and Microsporum canis generated 4-demethylgriseofulvin.

3. Cbauage of Ether Linkage As outlined under 0-dealkylation reactions, which can also be considered as a cleavage of an ether linkage, it appears that the ether cleavage enzymes investigated are quite versatile in the range of substrates they can oxidize. This can probably be explained by the natural abundance of methoxylated compounds in connection with lignin and other organic material in soil. In view of this, it is not very surprising that many pesticides containing ether linkages, like 2,4-D, related phenoxyacetates, and dicamba, are relatively easily metabolized. The cleavage of an ether linkage is considered to be a reaction caused by a mixed function oxidase insofar as all detailed studies revealed that reduced pyridine nucleotides and molecular oxygen are required. A large number of microorganisms capable of metabolizing phenoxyalkanoic herbicides produce the corresponding phenols as intermediates (Loos, 1969). This transformation could also be shown with cell-free extracts from an Arthrobacter sp. (Bollag et al., 1967; Loos et al., 1967a). In order to clarify the mechanism of ether cleavage, Helling et al. (1968), using phenoxy-'*O acetic acid, demonstrated that the cell-free extract of a MCPA-grown Arthrobacter sp. catalyzed, in the presence of oxygen, the cleavage between the aliphatic side chain and the ether-oxygen atom as indicated by the complete retention of lXOin the phenol molecule. Two mechanisms are apparently involved which cause the cleavage of the ether linkage: (1) a reductive reaction observed with higher phenoxyacetic acids ( MacRae and Alexander, 1963) and ( 2 ) an oxidative reaction which was established with 2,4-D and MCPA (Tiedje and Alexander, 1969; Gamar and Gaunt, 1971). MacRae and Alexander (1963) reported that a Flavobacterium sp. caused the cleavage of the ether linkage of ,-linked (omega-linked) 2,4-dichlorophenoxyalkyl carboxylic acids from the propionic to the

90

JEAN-MARC BOLLAC

61'

undecanoic homolog. The cleavage apparently resulted in the production of a phenol and the fatty acid corresponding to the aliphatic moiety: O-(CH,),-COOH

+

\

CH,-(CH,),-COOH

\

c1

c1

A soluble enzyme preparation from an ATthTobacteT sp. catalyzed the cleavage of the ether linkage of 2,4-D resulting in the formation of 2,4-dichlorophenol and alanine (Tiedje and Alexander, 1969). S'ince acetate and glycolate were not attacked by the cell-free extract, but glyoxylate was rapidly metabolized, it was suggested that glyoxylate was the initial product, and its further transformation resulted in the production of alanine. A similar investigation was performed by Gamar and Gaunt (1971) using MCPA as the substrate and a crude extract prepared from a Pseudomonas sp. grown on a basal medium with MCPA as sole source of carbon. MCPA was oxidized by the cell-free preparation in the presence of a reduced nicotinamide nucleotide (NADH or NADPH) to 2-methyl-4-chlorophenol and glyoxylic acid: 0-CH,COOH

Q.'" c1

4

2

H

3

+

CHO-CCOOH

c1

4. Oxidation of Aromatic Ring Numerous pesticides are cyclic compounds, and consequently their complete biodegradation can be achieved only after cleavage of the ring. Many microorganisms have the ability to oxidize aromatic substances and can use the resulting aliphatic compounds as substrates in the intermediary metabolism. Intensive studies contributed to a quite clear understanding of the general catabolic pathways of aromatic molecules, and many microbial enzymes involved in this reaction sequence could be isolated and characterized (Dagley, 1971; Stanier and Ornston, 1973). It is generally accepted that dihydroxylation is a prerequisite for enzymatic cleavage of the benzene ring. Ring fission can be brought about by dioxygenases through three different pathways depending upon the distribution of the hydroxyl groups; catechol, for example, which possesses hydroxyls on adjacent carbons, can be oxidized by ortho or meta

MICROBIAL TRANSFORMATION OF PESTICIDES

91

cleavage forming cis,cis-muconic acid or a-hydroxymuconic semialdehyde, respectively :

ortho cleavage

metu cleavage

A third pathway can occur with para-dihydric phenols; the ring of gentisic acid, for example, can be cleaved between the hydroxyl and the adjacent carboxyl group, resulting in the formation of maleylpyruvic acid:

qcmy OH

OH

B g - C O O H

OH

Aromatic pesticides-and also other xenobiotic compounds-are usually distinguished by multiple and variable substitutions on the ring or different ring formations, causing a molecular structure which often cannot be easily attacked. Therefore, each pesticide possessing a cyclic structure has to be investigated independently to determine whether ring fission can be provoked by microbial activity. Evans (1969) and Chapman ( 1972) summarized known bacterial pathways of various phenolic compounds, but detailed knowledge on the microbial ring cleavage of aromatic pesticides is quite scarce, and only fragmentary knowledge exists on the fate of products after ring cleavage. However, one group of pesticides, the chlorinated phenoxyalkanoic herbicides, were intensively studied in relation to ring cleavage, and the fate of the chlorinated catechols was followed and elaborated with enzyme preparations from different bacteria. Cell-free extracts from an Arthrobacter sp. metabolized 3,5-dichlorocatechol and 4-chlorocatechol, degradation products of 2,4-D and 4-~hlorophenoxyacetate,to ring fission products which retained the halogens (Bollag et al., 1968a). By dilution of the cell-free extract it was possible to achieve the accumulation of the muconic acids. Tiedje et al. (1969) identified cis,cis3-chloro- and cis,cis-2,4-dichloromuconic acid from 4-chloro-and 3,5-dichlorocatechol,respectively :

?H c1

c1

92

JEAN-MARC BOLLAG

The formation of the cis&-muconic acids implies an ortho-fission mechanism. Extracts of the bacterium also converted catechol, 3- and 4-methylcatechol to the corresponding muconic acids. Evans et al. (1971b) also described the conversion of 3,5-dichlorocatechol, suspecting that they had found cis&-muconic acid in the culture medium of a Pseudomonas sp., but another Pseudomonus strain apparently produced a-chloromuconate, and therefore, they concluded that, in the latter case, dechlorination at the p-position has taken place at some stage before ring cleavage. The intermediate muconic acid of MCPA was tentatively identified as &,cis-y-chloro-a-methylmuconate as shown in the reaction sequence of a cell-free system from a Pseudomonas sp. (Gaunt and Evans, 1971); the ring-fission enzyme required Fe2+or Fe3+ and reduced glutathione €or activity, as do many other catechol oxygenases. A meta-cleaving oxygenase, a catechol 1,6-oxygenase, from an Achromobacter sp. was active on methylated and chlorinated catechols ( Horvath, 1970b). 3-Methylcatechol, 4-ch1orocatecho1, and 3,5-dichlorocatechol were oxidized to 2-hydroxy-3-methylmuconic semialdehyde, 4-chloro-2-hydroxymuconic semialdehyde, and 3,5-dichloro-2-hydroxymuconic semialdehyde, respectively:

-

OHC H W

T CHS

&OH c1

~

oF : c1

HOOC c1

c1

c1

It is of interest to emphasize that halogen or alkyl substitutions of the aromatic compound were not released prior to the ring cleavage, and therefore, the fate of the resulting ring-fission products have to be followed further if one is concerned with their toxic impact. An important practical question is related to the microbial degradation of DDT which is considered to be of the most persistent pesticides in the environment. Whereas several microbes were found to be capable of dehalogenating DDT to DDD and also DDE (see Section III,D), there is still no clear knowledge as to the extent to which the cleavage of the aromatic rings of DDT or its metabolites takes place. Focht and Alexander (1971) isolated a Hydrogemonas sp. from sewage effluent capable of cleaving the ring of DDT analogs. Cell suspensions of the bacterium, which were grown on diphenylmethane, did not metabolize DDT and p,p’-dichlorobenzophenone, but they did transform the corre-

MICROBIAL TRANSFORMA'MON OF PESTICIDES

93

sponding monochloro and nonchlorinated compounds. They concluded that the presence of the para-chlorine substitution on the phenyl rings and the substitution of carbonyl or trichloromethyl group on the carbon atom binding the two phenyl groups inhibited the metabolism of DDT or analogs by the Hydrogemonas sp. However, one of the benzene rings of diphenylmethane, p,p'-dichlorodiphenylmethane, and 1,l-diphenyl2,2,2-trichloroethane was cleaved as indicated with the formation of phenylacetic, p-chlorophenylacetic, and 2-phenyl-3,3,3-trichloropropionic acid, respectively:

Q

c1 I

-0 6-

CH,-COOH I

c1

Q HC-CQ,

CH,(CCI$COOH

I

-0

-

In a subsequent investigation it was found that high protein concentrations in bacterial extracts of a Hydrogemom sp. caused the transformation of DDT, DDD, and other products under anaerobic conditions. If whole cells were added and aerobic conditions were provided, subsequently, the formation of p-chlorophenylacetic acid could be determined (Pfaender and Alexander, 1972). This observation infers that one microbe is capable of attacking DDT and causing its degradation to a single chlorinated benzene compound. Gunner and Zuckerman ( 1968) described the microbial degradation of the pyrimidyl ring of the insecticide diazinon in the presence of two microorganisms. When a Streptomyces sp. or an Arthrobacter sp. were incubated individually with diazinon, the pyrimidyl ring was not attacked, but the two microbes together cleaved and metabolized the ring structure. The ring-fission mechanism of several important groups of pesticides by microorganisms, if it actually occurs, is still very obscure and requires a lot of experimental work for its clarification. There are only indicative

94

JEAN-MARC BOLLAG

data that the rings of s-triazine herbicides ( Kaufman and Kearney, 1970), the substituted anilines which are intermediates from numerous pesticides (Chisaka and Kearney, 1970), and 1-naphthol (Bollag and Liu, 1972b), resulting from the bicyclic ring of certain methylcarbamate insecticides, are cleaved and further metabolized. Most information leading to the assumption that ring cleavage takes place results from experiments with radiolabeled pesticides whereby it was possible to trap "CO,. In the case of s-triazines, for example, evolution of IfC02has been reported from microbial systems treated with 14C-ring-labeleds-triazines, but in nearly all cases, only up to 4% of the applied herbicide evolved as ' C O , ( Kaufman and Kearney, 1970).

5. p-Oxidation This reaction was found especially in the oxidation of long-chain phenoxyalkanoate herbicides. p-oxidation of an aliphatic side chain proceeds by the stepwise removal of two-carbon fragments from a fatty acid, and the shortened acid can then be further oxidized. Bacteria (Taylor and Wain, 1962), actinomycetes (Webley et al., 1957), and fungi (Byrde and Woodcock, 1957) can metabolize w-phenoxyalkanoic acids by p-oxidation. Oxidation of 2,4-dichlorophenoxyalkanoicacids, for instance, with an even number of carbon atoms in the side chain results in the formation of 2,4-dichlorophenoxyacetate whereas acids with an odd number of carbon atoms are converted to 2,4-dichlorophenol (Loos, 1969) : CH2-CHz-CH,-CHa+CH2-COOH

@ I

;

-

c1

qcl

J

CHa-CHa+CH,I 0

COOH

C1

CH,-COOH I 0

ecl CHz-CHaI

C1

c1

\

CH,-fCH,-COOH

CHa- CH,- COOH I

I

__t

\

c1

-

6-" c1

MICROBIAL TRANSFORMATION OF PESTICIDES

95

The mechanism of p-oxidation of phenoxyalkanoate herbicides was first established with pure cultures, but subsequently, the same kind of metabolism was also shown to occur in natural soil (Gutenmann et al., 1964). 6. Epoxidation

Addition of an oxygen atom to a double bond represents an epoxidation reaction, which is now recognized as a common process for the metabolism of xenobiotic compounds. In recent years it became clear that this is a widespread process in the metabolism of aromatic substances. Arene oxides are intermediates which are transformed enzymatically or nonenzymatically to dihydrodiols and ( pre ) mercapturic acids and cause considerable concern, since, as metabolic intermediates, they are capable of initiating tissue necrosis and carcinogenesis (Daly et al., 1972). Arene oxide, lP-naphthalene oxide, has been identified in a biological system as the obligatory intermediate in the formation of naphthols from naphthalene (Jerina et al., 1969). Monooxygenases of bacteria and fungi can introduce a hydroxyl group to an aromatic compound accompanied by migration of an original substituent. This phenomenon, termed NIH-shift, proceeds via the intermediary formation of an arene oxide. Guroff et al. (1966) found that halogen substituents may be displaced from carbon-4 to carbon-3 when phenylalanine is hydroxylated by Pseudomonas:

c1

OH

Although it was shown that various microorganisms cause the NIH-shift, its importance in the microbial degradation of pesticides has yet to be clearly established. However, one example is well known: Faulkner and Woodcock (1964, 1965) found that Aspergillus niger metabolized the herbicide 2,4-D to 2,4-dichloro-5-hydroxy-and 2,5-dichloro-4-hydroxyphenoxyacetic acid:

b"

OCH,-COOH

\

C1

OCH,-COOH

OCH,-COOH

HOf q\C 1

c1

+.

c1QC1\ OH

96

JEAN-MARC BOLLAG

The latter product generated by the fungal activity shows the shift of a chlorine atom which was coupled with the introduction of a hydroxyl group in its place. When the herbicide MCPA, in which the chlorine at carbon-2 is substituted by a methyl group, was exposed to the same fungus, hydroxylation occurred, but the NIH-shift was not observed ( Faulkner and Woodcock, 1965). An enzyme system from Pseudomonm oleovorans was shown to catalyze the epoxidation of alkenes; 1,7-octadiene was converted to both 7,8-epoxy-l-octene and 1,2-7,8-diepoxyoctane, whereas l-octene was oxidized to both 7-octenel-01 and 1,2-epoxyoctane (May and Abbott, 1973). It appeared that the enzymatic epoxidation reaction is mechanistically similar to the reaction causing the methyl group hydroxylation of alkanes and fatty acids. Both enzyme systems are composed of three protein components ( reductase, rubredoxin, and hydroxylase) and require molecular oxygen and NADH (NADPH cannot be substituted) for activity. Several cyclodiene insecticides, such as aldrin, isodrin, and heptachlor, can -undergo epoxidation by various microorganisms yielding products with increased toxicity in the environment. Korte et al. (1962) demonstrated that the fungi Aspergillus niger, A. fEavus, Penicillium notatum, and P . chrysogenum converted aldrin to dieldrin:

“pJJ c1

c1

o* l c -

dl

c1 Cl

c1 I

Ninety-two different strains of bacteria, actinomycetes, and fungi isolated from soil were tested for their ability to transform aldrin, and most of them could epoxidize the original pesticide dieldrin (Tu et al., 1968). The most active fungal isolate, a Fusarium sp., oxidized about 9% of the added aldrin to dieldrin during the 6-week incubation period. A similar epoxidation of a chlorinated hydrocarbon containing an isolated double bond was observed with the insecticide heptachlor, which was oxidized to heptachlor epoxide (Miles et al., 1969). Thirty-five of 47 fungi, and 26 of 45 bacteria and actinomycetes isolated from soil produced the epoxide; the greatest activity was shown by a Nocardia sp. which epoxidized 6%of the applied heptachlor. Another epoxidation reaction occurred after an initial chemical hydrolysis of heptachlor to l-hydroxychlordene. The chemical intermediate product was converted to l-hydroxy-2,3-epoxychlordeneby 43 of 47 fungi, but only 4 of the 45 bacteria and actinomycetes tested showed activity.

hlICROBIAL TRANSFORMATION OF PESTICIDES

97

7. Sulfoxidation This reaction consists of the oxidation of divalent sulfur to the sulfoxide and, sometimes, to the sulfone: > S + >SO + >SO2, but evidence from detailed studies was received only from experiments with microsomal enzymes from plant and animal systems. Sulfoxidation of pesticides in soil was attributed to biological transformation with aldicarb ( Coppedge et al., 1967), phorate (Getzin and Chapman, 1960), and the s-triazine prometryne (Plimmer and Kearney, 1969), but no specific microorganisms were isolated capable of sulfoxidizing the various pesticides. Ahmed and Casida ( 1958), investigating the metabolism of organophosphorus insecticides, determined that the green alga Chlorella pyrenoidosa and the yeast Torulopsis utilis oxidized phorate to its respective sulfoxide, but slowly converted this product to the phosphorothiolate sulfoxide with little formation of the salfide or sulfone.

B. REDUC~IONS Several groups of pesticides are subject to reduction, but this reaction is usually less common than oxidation in the transformation of xenobiotic compounds. The reduction of the nitro group to amine has been found during the metabolism of various pesticides by the activity of different bacteria and fungi. It is anticipated that the reduction takes place in stages involving the intermediate formation of a nitroso and hydroxyamino group : R-NO,

R-N=O

i

--t

R-NHOH

-+

R-NHZ

In experiments with Escherichia coli, Saz and Slie (1954) found that various organic nitro compounds are reduced in the presence of cysteine to the corresponding arylamines. 2,4-Dinitrophenol, which is used in fungicidal preparations, was reduced by Fusarium oxysporum in a liquid basal medium to 2-amino-4-nitrophenol and 4-amino-2-nitrophenol ( Madhosingh, 1961) . Formation of the 4-amino-2-nitrophenol compound appeared to be favored in acid cultures, whereas the 2-amino isomer dominated at a higher pH value. The degradative pathway of the herbicide DNOC by a pseudomonad isolated from a garden soil was followed by isolation of intermediates from growing cultures, cell suspensions, and cell-free extracts, and the reactions sequence was determined as : 3,5-dinitro-o-cresol+ 3-amino-5nitro-o-cresol + 3-methyl-5-nitrocatechol -+ 3-methyl-5-aminocatechol + 2,3,5-trihydroxytoluene (Tewfik and Evans, 1966). Hamdi and Tewfik (1970) also determined that DNOC is reduced to 3-aminod-nitro-ocresol by Rhizobium leguminosarum.

98

JEAN-MARC BOLLAG

The fungicide pentachloronitrobenzene ( PCNB ) is easily reduced in culture solution by various actinomycetes and fungi to the corresponding aniline (Chacko et al., 1966; Nakanishi and Oku, 1969), and the same transformation was observed in soil (KO and Farley, 1969).

-

cl@ c1

cl@cl

' c1

c1

c1

c1

c1

Lichtenstein and Schulz (1964) found that the organophosphorus insecticide parathion was metabolized in soil either by hydrolysis or by reduction to its amino, form, apparently depending 0x1 populations of soil microorganisms. Since Neuberg and Welde (1914) showed that in the presence of yeast nitrobenzene was reduced to aniline, Lichtenstein and Schulz also tested the effect of yeast on the metabolism of parathion and found it responsible for the reduction of the insecticide to aminoparathion; bacteria apparently did not participate in this reduction. Likewise, the reduction of a nitro to an amino group occurred during the metabolism of the organosphosphorus insecticide Sumithion ( Fenitrothion) by Bacillus subtilis (Miyamoto et al., 1966); experiments with washed cell suspensions of B . subtilis indicated that the nitro group of various phosphorothioates was reduced under aerobic as well as under anaerobic conditions. Various chlorinated hydrocarbons are transformed by reductive dehalogenation processes, but these reactions will be considered in Section II1,D. Frequently, it is also observed that reduction is a process which can produce a center for conjugation, and such a pathway was found in various metal-containing agricultural pesticides. In the case of arsene-containing compounds, it was found that a strain of Methanobacterium reduced arsenate ( 5 + ) to dimethylarsine ( 3 - ) under anaerobic conditions, whereas reduction and methylation reductions occurred intermittently (Fig. 1); in these experiments, methylcobalamin served as methyl donor of choice ( McBride and Wolfe, 1971). The sequence of reactions involved the reduction of arsenate (5+, arsenic valency ) to arsenite (3+ ). This intermediate was methylated to methylarsonic acid, subsequently reduced and methylated to dimethylarsinic acid ( I f ) , and finally further reduced to form the dimethylarsine ( 3 - ) . This reaction could also be shown with cell extract from the Methanobacterium, but adenosine triphosphate and hydrogen had to be added.

99

MICROBIAL TRANSFORhlATION OF PESTICIDES

-

OH I HO-As5+OH

7%

As3+-OH I1 0

II

0 Arsenate

HO-AS3"OH I1

0

Arsenite

Methylarsonic acid

J

?H3 As3--CH,

y

3

HO--As'+-CH, II

I

n

0

Dimethy larsine

Dimethylarsinic acid

FIG.1. Metabolic transformation of arsenate by a Methanobacterium sp.

C. HYDROLYSIS Hydrolysis is a reaction type that can be initiated enzymatically or chemically, and therefore, it may be difficult in some cases to determine the true origin. Generally, it can be assumed that hydrolysis converts a lipophilic compound into a hydrophilic, water-soluble substance. In the microbial breakdown of pesticides, hydrolytic enzymes include amidases, esterases, nitrilases, and phosphatases which yield an acid on one side and an alcohol or amine on the other: I

-C-C-N-

Ester

I l l -C-C-0-C-

Nitrile

-C-C=N

I

0

I

I

I

-

R

Amide

I

I I

-

0 I II -C-C-OH

I

I I

0 II

-C-C-OH

+

H-N-

+

HO-C-

I I I

0

I II -C-C-NH, I

--C-C-OH

I

I

0 It

f

NH,

Carbamates can be considered simultaneously as esters and amides and represent special cases insofar as the intermediate carbamic and carbonic acids are unstable compounds and degrade spontaneously with the liberation of CO,. However, it is evident that, independent of an esterase or an amidase reaction, the end products are the same. Hydrolysis appears to be the major reaction for acylanilide herbicides causing the cleavage of the C-N bond and the release of the side chain. Two species of Penicillium and one species of Pululluria isolated

100

JEAN-MARC BOLLAG

from soil were capable of hydrolyzing the herbicide karsil (Sharabi and Bordeleau, 1969), whereas 3,4-dichloroaniline and 2-methylvaleric acid were identified as intermediates. A cell-free extract was prepared from a Penicillium sp., and the specificity of the partially purified acylamidase was tested on various anilides and structurally related compounds. Activity was enhanced with increasing chain length, up to fourcarbon compounds. Substitution of the N-acyl group or the phenyl ring also influenced the enzyme activity. A phenylurea, diuron, and a phenylcarbamate herbicide, CIPC, were not attacked by the enzyme preparation. An acylamidase which hydrolyzed propanil to 3,4-dichloroaniline and propionic acid was also isolated from the mycelium of Fusarium solani (Lanzilotta and Pramer, 1970). Other acylanilides such as dicryl and karsil as well as phenylureas (monuron and fenuron) appeared to be unaffected by this enzyme system. Extensive studies were performed on the enzymatic hydrolysis of the phenylcarbamate chlorpropham ( CIPC ) with partially purified cell-free preparations obtained from a strain of Pseudomonas striata isolated from soil enrichment cultures (Kearney, 1965). An enzyme, purified by ammonium sulfate precipitation and column chromatography by gradient elution on DEAE-celluIose, catalyzes the hydrolysis of CIPC to 3-chloroaniline, carbon dioxide, and isopropyl alcohol:

However, it is not clear whether the enzymatic cleavage proceeds by hydrolysis of the ether linkage or of the amide bond or both, since the initial products produced by both reactions would be unstable. It is interesting to observe that this enzyme exhibits a broad substrate specificity, since a large number of structurally related phenylcarbamates and acylanilides were hydrolyzed, but ureas and methylcarbamates were not metabolized. The metabolism of urea herbicides was studied and ehcidated especially with Bacillus sphaericus, which was isolated from soil treated with monolinuron. Whole cells or cell-free extracts of B. sphaericus ( Wallnofer, 1969; Wallnofer and Bader, 1970) degraded various N’-methoxyphenylurea compounds by releasing CO, from the ureido portion of the molecule and leaving the corresponding aniline moieties as well as an unidentified product. The cell-free extract hydrolyzed the N’-methoxyphenylurea compounds monolinuron, linuron, chlorbromuron, and metobromuron, but the N’,N’-

MICROBIAL TRANSFORMATION OF PESTICIDES

101

dimethylphenylureas monuron, diuron, buturon, and fluometuron were not attacked (Engelhardt et al., 1971). However, there was no clear explanation as to why the decomposition of urea appears to be specific for the methoxy-substituted phenylureas. Thirteen acylanilides, which are used partially as herbicides or fungicides, were hydrolyzed by the cell-free extract at a rate at least 10 times higher than that of the methoxy-substituted phenylureas. In all these studies cell-free extract activity was found only if the enzyme preparation was induced after growth on the herbicide of choice, namely, linuron. Engelhart et al. (1971, 1972) also clarified the hydrolytic pathway of linuron. After incubation with extracts of B. sphaericus, it was possible to identify 3,4-dichloroaniline and CO, as well as N,O-dimethylhydroxylamine by characterization of its dinitrophenyl derivative. This leads to the general conclusion that phenylamide compounds are hydrolyzed to the corresponding anilines and acids, but the acid moiety formed during the composition is dissociated rapidly to the alkylalkoxyamine and CO,. The proposed reaction sequence for linuron is as follows:

c=o I

C1

I

Cl

L in u r 0 n

3,4-Dichloroaniline

N , 0-Dimethylhydroxylamine

Another mechanism of phenylurea degradation was proposed in the review of Geissbiihler (1969) from experiments with soil samples, where stepwise dealkylation of dimethyl-substituted phenylureas would precede hydrolysis. N-Lauroyl-1-valine, an amino acid derivative used as a pesticide with a preventive effect against rice blast, was metabolized by Pseudomonas aeruginosa. Since one product was identified as lauric acid, it was suggested that cleavage of the N-acyl linkage occurred, resulting in the formation of lauric acid and valine, but the latter compound was not detected because it might have been metabolized rapidly after its release ( Shida et al., 1973). Nitrile hydrolysis was shown when sterile and nonsterile soil treated

102

JEAN-MARC BOLLAG

with ioxynil were compared. No breakdown of the herbicide was detectable in the sterile soil, but in the nonsterile soil ioxynil was converted to 3,5-diiodo-4-hydroxybenzoicacid, with 3,5-diiodo-4-hydroxybenzamide as an intermediate product: CN

I

CONH,

COOH I

OH

OH

I

1

OH

Hydrolytic degradation also constitutes one of the major reactions in the metabolism of organophosphorus insecticides which contain either the P=O (phosphate) or the P=S ( phosphorothioate) groupings. It must be stressed that phosphorus esters are also easily susceptible to catalytic cleavage induced by nitrogenous compounds like amino acids and by heavy metal ions. Consequently, it has to be expected that nonenzymatic hydrolysis of phosphorus occurs easily in soil and other habitats. Mounter et al. (1955) demonstrated the presence in freeze-dried bacterial cells of a phosphatase which hydrolyzed dialkylfluorophosphates with the release of fluoride ions, and Mounter and Tuck (1956) showed that Escherichia coli and PropionihacteTium pentasaceum hydrolyze paraoxon, TEPP, and the diethyl, diisopropyl, and di-n-butyl fluorophosphates. Ahmed and Casida (1958) concluded that Pseudomonas fluorescens hydrolyzed Phorate ( Thimet ) without a subsequent oxidation reaction since, after incubation with the microbes, the residual organophosphate recovered partitioned completely into hexane from an acetone-water mixture. The soil fungus Trichoderma &ride and a Pseudomonas sp. degraded malathion, presumably by hydrolysis of the ester groups, to various carboxylic acid derivatives ( Matsumura and Boush, 1966). The insecticide trichlorfon ( Dipterex) was metabolized in a culture medium by AspeTgillus niger, Penicillium notatum, and a Fusarium sp. to hydrolytic products; one was identified as O-methyl-2,2,2-trichloro1-hydroxyethylphosphonic acid, and a second metabolite was tentatively acid ( Zayed et al., identified as 2,2,2-trichloro-l-hydroxyethylphosphonic 1965) : 0 OH C1 It I I CH,O-P-CH-C-C1 I I OCH, C1

0 +

II

OH I

C1 I

HO-P-CH-C-CI I I OCH, C1

-

0 OH CI I1 I I HO-P-CH-C-C1 I

OH

I

c1

Conversion of carbaryl to 1-naphthol also appears to be a hydrolytic reaction generated by various soil microorganisms, but the simultaneous chemical hydrolysis of the insecticide makes it difficult to establish to what extent the reaction is biological or chemical ( Bollag and Liu, 1971).

hIICROBIAL TRANSFORMATION OF PESTICIDES

103

The partial conversion of dieldrin by a hydrolytic reaction was shown in laboratory experiments. Matsumura and Boush ( 1967) isolated several species of Pseudomonas and Bacillus from soil samples, and they suggested, on the basis of an identical R, value with an authentic compound, that 6,7-trans-dihydroxydihydroaIdrin( aldrin diol ) might be a major product, whereas Wedemeyer (1968) came to a similar conclusion with the bacterium Aerobacter amogenes after various chromatographic analyses. c1 c1 I

c1

Cl

Although there are numerous reports with plants and animals which describe enzymatic hydrolysis of chlorotriazines to the hydroxy derivative, there is only one report claiming that atrazine is transformed by a microorganism, by the fungus Fusarium roseum, to its corresponding hydroxy analog (Couch et al., 1965).

MECHANISMS D. DEHALOGENATION Halogenated aliphatic and aromatic pesticides are widely used, and therefore, they are of increasing importance in the environment. The major metabolic problem which they pose relates to the question of the stability of the carbon-bound halogen. The transformation of the organic halogen to an inorganic form can usually be considered as a detoxication reaction, but if the metabolism does not involve the release of the halogen, the resulting intermediate may cause concern in the various ecosystems. The possible biological attack on halogenated compounds varies widely, and it is clear that the carbon-halogen bond and the number of halogen substitutions, as well as other structural features of the molecule, determine the metabolic fate of such a compound; therefore, one might expect different enzymatic mechanisms. The microbial enzymes which catalyze the removal of the halogen from the organic molecule were divided into three groups (Table 11): ( 1 ) hydrolytic dehalogenation, in which a hydroxyl group replaces the halogen atom; ( 2 ) reductive dehalogenation, where halogens are exchanged with hydrogen; and ( 3 ) dehydrohalogenation, in which both hydrogen and chlorine are removed from the molecule with the resultant formation of a double bond. The dehalogenating enzymes involved in these reactions are not clearly characterized, and therefore the division into various dehalogenating

TABLE I1 DEHALOQENATION MECHANISMS OF PESTICIDES OR THEIRMETABOLIC INTERMEDIATES BY MICROORQANISMS Schematic reaction 1. Hydrolytic dehalogenation

RCHz(halogen) RCHzOH

-+

Example

Refer en ces

Fluoroacetate + glycolic acid 3-Bromopropanol 3-hydroxypropionic acid 2-Hydroxyphenoxyacetic acid 4 Zchlorophenoxyacetic acid 3-Chlorobenzoic acid 3-hydroxybenzoic acid

Goldman (1965) Tonomura et al. (1965) Castro and Bartnicki (1965) Faulkner and Woodcock (1961)

2-Fluorobenzoic acid + catechol

Goldman et al. (1967)

DDT + D D D (TDE)

Kallman and Andrews (1963), Mendel and Walton (1966), Chacko el al. (1966), Wedemeyer (1966, 1967), Johnson et al. (1967), Plimmer et al. (1968), Braunberg and Beck (1968), Matsumwa and Boush (1968), French and Hoopingarner (1970) Miles et al. (1969)

-+

Halogen

on

-+

2. Reductive dehalogenation RC(ha1ogen)s -+ RCH(ha1ogen)2

Heptachlor -+ chlordene 3. Dehydrohalogenation RCHJ3(halogen)a -+ RCH =C(halogen)z

DDT-t DDE

Lindane -+ y-pentachlorocyclohexene -pChloro-a-methylmuconic acid + r-carboxymethylene-a-methyl-Aaflbutenolide

Johnston et al. (1972)

Stenersen (1965), Mendel and Walton (1966), Guenzi and Beard (1967), Matsumura and Boush (1968), Chacko et al. (1966), Johnson et al. (1967), Langlois (1967), Braunberg and Beck (1968) Yule et al. (1967) Gaunt and Evans (1971)

105

MICROBIAL TRANSFORMATION OF PESTICIDES

TABLE I1 (Continued) Schematic reaction

Example

References

cis, cis-3-Chloromuconic acid + (4-carboxymetliylene but-%enolide) -+ maleylactic acid eis,cis-2,4-Dichloromuconic acid -+ (2-chloro-4-carboxymethylene but-2enolide) --+ chloromaleylacetic acid pyruvic acid Dalapon 3-Chloropropionic acid ---f acrylic acid Ethylene dibromide -+ ethylene

Bollag et al. (1968b), Tiedje et al. (1969), Evans et al. (1971a),

---$ -+

Bollag et al. (1968b), Tiedje et al. (1969), Evans et al. (1971b) Kearney et al. (1964) Bollag and Alexander (1971) Castro and Belser (1968)

mechanisms should be considered only as an attempt to categorize the observed microbial removal of the halogens.

1. Hydrolytic Dehalogenution In this reaction the halogen is replaced by a hydroxyl group, but the specific enzymatic mechanism involved was not elaborated in the case of microbial pesticide metabolism. The introduced hydroxyl group can be generated from water as demonstrated by experiments in lSO enriched water (Goldman and Milne, 1966), or the oxygen of the hydroxyl group can originate from the reduction of molecular oxygen by the catalytic activity of a NADPH-dependent hydroxylase ( Kaufman et aE., 1962). Aliphatic compounds containing C atoms bearing only one halogen will be transformed to alcohols. The halidohydrolases catalyze relatively simple reactions in which a halogen at the 2-position of a short-chain fatty acid is replaced by a hydroxyl group. An enzyme of this kind was found in a soil Pseudomonas sp, ( Goldman, 1965) which catalyzes chloro-, fluoro-, and iodoacetate: Hal. CH2COO-

+ HO-

+

HOCH2COO-

+ Hal.-

Fluoroacetate, on which the bacteria were grown, was the preferred substrate, and chloride and iodide are released from their substrates at only 15 and 0.53, respectively, of the rate of fluoride release. Castro and Bartnicki (1965) found that a pseudomonad grown in a medium containing 3-bromopropanol also replaced the halide with a hydroxyl group, and they isolated and identified 3-hydroxypropionic acid.

106

JEAN-MARC BOLLAG

Dehalogenation of aromatic compounds by microorganisms usually occurs after ring cleavage at it is described later under “dehydrohalogenation” in the metabolism of various chlorinated phenoxyacetic acids. However, it was also found that a halogen can be directly replaced on a benzene ring by a hydroxyl group. Faulkner and Woodcock (1961) observed that 2-chlorophenoxyacetic acid is converted to 2-hydroxyphenoxyacetic acid by Aspergillus niger, and Johnston et al. (1972) determined that a Pseudomonas sp. transformed 3-chlorobenzoic acid to 3-hydroxybenzoic acid. There are no investigations reported on the microbial enzymes performing this reaction, and therefore, it is not possible to make comparisons with the microsomal hydroxylating system of rat liver which is capable of converting both 4-chloro- and 4-fluoroaniline to 4-hydroxyaniline ( Daly et al., 1968). Goldman et al. (1967) concluded that dicarboxylation and defluorination occurs simultaneously when 2-fluorobenzoate is converted to catechol by a pseudomonad. Further support for this conclusion was obtained when the reaction was carried out in an atmosphere containing 50% laOzand 50%‘*Ox, since the catechol produced contained either 2 atoms of le0or 2 atoms of l8O, indicating a one-step reaction ( Milne et al., 1968).

2. Reductive Dehalogenation This mechanism of halogen removal has been demonstrated with numerous microorganisms in the conversion of DDT to DDD (TDE) : c1

c1

dl

Ci DDT

DDD (TDE)

The process was first described with microorganisms in studies with yeast (Kallman and Andrews, 1963), and subsequentIy with Escherichia coli (Mendel and Walton, 1966), Proteus vulgaris (Barker et al., 1965), Serratia marcescens ( Stenersen, 1965), soil actinomycetes, namely Nocardia erythropolis and five species of Streptomyces (Chacko et al., 1966), Aerobacter aerogenes ( Mendel et al., 1967), plant pathogenic and saprophytic bacteria (Johnson et al., 1967), and in soil samples under anaerobic conditions (Guenzi and Beard, 1967). Most of the studies provide evidence that anaerobic conditions favor reductive dechlorination over competing reactions. Wedemeyer (1966) isolated a cell-free system from Aerobacter

MICROBIAL TRANSFORMATION OF PESTICIDES

107

aerogenes which catalyzed, anaerobically, the reduction of DDT to DDD. Since the addition of 0.001 M cyanide or carbon monoxide com-

pletely inhibited the conversion, Wedemeyer suggested that reduced cytochrome oxidase is probably responsible for the reductive dechlorination. French and Hoopingarner ( 1970) obtained membrane fractions from Escherichia coli which also produced DDD under anaerobic conditions if flavine adenine dinucleotide (FAD) was added. The cytoplasmic factor, alone or in the presence of boiled membrane fraction, was completely inactive. Plimmer et al. (1968) demonstrated conclusively with deuterated DDT that DDD is the result of a direct reductive dechlorination which does not involve the formation of DDE as an intermediary metabolite. Retention of the deuterium atom in DDD excluded the possibility of dehydrohalogenation and subsequent reduction. One pathway of degradation of the insecticide heptachlor is caused by bacteria and actinomycetes and apparently proceeds by reductive dechlorination resulting in the formation of chlordene as an intermediate ( Miles et al., 1969). 3. Dehydrohalogenution

The simultaneous removal of a hydrogen and halogen was found especially with chlorinated hydrocarbon insecticides. The formation of DDE from DDT is perhaps the most familiar reaction in this group of pesticides :

DDT

DDE

In comparative studies of soil samples kept under anaerobic and aerobic conditions, it was found that DDT is rapidly converted to DDD in the absence of oxygen, whereas the transformation to DDE occurred aerobically and at a slow rate (Guenzi and Beard, 1968). In several cases the formation of TDE was accompanied, even anaerobically, by DDE, but on a much smaller scale (Stenersen, 1965; Mendel and Walton, 1966; Guenzi and Beard, 1967). Matsumura and Boush (1968) found that the majority of variants of the soil fungus Trichoderm viride produced TDE, while some variants produced only DDE; this indicates that different enzyme systems causing the degradation of DDT exist even among variants of the same species.

108

JEAN-MARC BOLLAG

Soil microorganisms appear to be responsible for the breakdown of lindane, and the isolation of 7-pentachlorocyclohexeneimplies a dehydrochlorination reaction (Yule et al., 1967). In the metabolism of chlorophenoxyacetates by soil pseudomonads it was shown that, after ring fission of 4-chloro-substitued phenoxyacetates, the formed chloromuconic acid derivative was lactonized by dehydrochlorination. For example, an enzyme preparation from a Pseudomonm sp. catalyzed the transformation of y-chloro-a-methylmuconic acid, an intermediate of MCPA, to 7-carboxymethylene-a-methyl-a"e-butenolide (Gaunt and Evans, 1971) :

Cl

The lactonizing enzyme, functioning simukaneously as a dehydrochlorinase, required Mn'+ or Mg2+ as cofactors and was stimulated by Fez+ and Co". Therefore, Gaunt and Evans (1971) concluded that it bears no resemblance to the DDT-dehydrohalogenating enzyme investigated by Lipke and Kearns (1959)) which had no cofactor requirement and also differed in other tests. The observation that a chloride is removed from a benzene ring oply after ring cleavage and the formation of a muconic acid was also established in the bacterial metabolism of 2,4-D and 4-chlorophenoxyacetic acid (Bollag et al., 196813; Tiedje et al., 1969; Evans et aZ., 1971a,b). Castro and Belser ( 1968) established that soil-water cultures dehydrohalogenated nematocidal soil fumigants. Ethylene dibromide was converted almost quantitatively to ethylene in sterilized soil which was inoculated with soil suspensions: BrCH2CH2Br .+ CHz=CHz

+ 2Br'

and meso- and dl-2,3-dibromobutane were transformed by soil-water suspensions to bromine and butene: B r q B r CH, meso

-

H&./--..CH,

109

MICROBIAL TRANSFORMATION OF PESTICIDES

Kearney et al. (1964) reported the isolation and partial purification of an enzyme from an Arthrobacter sp. that removed chlorine from dalapon resulting in the formation of pyruvic acid, and this reaction should probably also be categorized under dehydrohalogenation. It was not possible to isolate intermediates from this system and to make a clear conclusion concerning the mechanism involved, but it was proposed that 2-chloroacrylate and 2-chloro-2-hydroxypropionate are unstable intermediary products : c1 I

CH,-C-COOH c I1

-I

C&=C-COOH dl

+

I

c1

-

0 ll CH,-C-COOH

The partially purified enzyme had its greatest activity on dalapon with less activity on 2-chloropropionate, dichloroacetate, and 2,2-dichlorobutyrate. No activity was detected on any p-chloro-substituted aliphatic acid. This observation differs from the isolated enzyme system of Micrococcus denitrificans which dehalogenates chlorinated aliphatic acids only with the halogen in the P-position (Bollag and Alexander, 1971). 3- and 4-Carbon acids-even unsaturated compounds like acrylic or crotonic acid-were dechlorinated only if the halide was in the p-position, but the dehalogenating enzyme system failed on the chlorinated acetic acids and on all other aliphatic acids with halogens solely on the @-carbon. There was more evidence that the enzyme preparation from M . denitrificans dechlorinated 2-chloropropionic acid via acrylic acid and not via 3-hydroxypropionic acid, indicating that the halogen is removed by dehydrohalogenation.

E. SYNTHETICREACTIONS In this category of enzyme reactions, the formation of a conjugate or condensate during pesticide metabolism is considered. A conjugation reaction implies the coupling of a pesticide, or an intermediate thereof, to an endogenous substrate resulting in the formation of, for example, methylated or acetylated compounds, amino acid conjugates, or glycosides, while the formation of a condensate implies the enzymatic condensation of a pesticide or an intermediate thereof. Williams (1971) stated that synthetic reactions require a source of energy that is usually supplied via adenosine triphosphate ( ATP ) . Several methylation reactions have been observed in the microbial metabolism of pesticides. A few examples are known in which O-methylation of chlorinated phenolic compounds took place (Fig. 2), and the

qcl

110

JEAN-MARC BOLLAG

cl$

\

c1

c1

c1

___)

\

c1

c1

cl+

c1

\

c1

c1

FIG.2. 0-methylation of 2,4-dichlorophenol, pentachlorophenol, and 2,5-dichloro4-methoxyphenol by various microorganisms.

methylation of heavy metals which are used as pesticides deserves special attention. Loos et al. ( 1967b) isolated 2,4-dichloroanisole during the metabolism of 2,4-D in the growth medium of an Arthrobacter sp., and they suspected that it might have been produced by an enzymatic 0-methylation of 2,4-dichlorophenol. Cserjesi and Johnson ( 1972) found that pentachlorophenol, a substance of fungicidal and various other pesticidal activities, can be methylated by the fungus Trichoderma viride in a growth medium, and the resulting product was identified by melting point determination and infrared spectroscopy as pentachloroanisole. The fungicide chloroneb was demethylated by numerous microorganisms to 2,5-dichloro-4-methoxyphenol,but some of the same microbes, especially Trichoderma viride and Mucor ramunnianus, could also reverse the reaction and methylate the dealkylated fungicide ( Wiese and Vargas, 1973). Both reactions could be shown to be independent if the two compounds were amended to a liquid basal medium. In addition, it was observed that some fungi could both methylate and demethylate 2,5-dichloro-4-methoxyphenol to produce chloroneb and 2,s-dichlorohydroquinone, respectively. A noteworthy observation relates to the finding that pentachlorothioanisole was a product during the metabolism of pentachloronitrobenzene by various species of Fusariurn oxysporurn ( Nakanishi and Oku, 1969). No enzymatic studies were reported on the methylation of the chlorinated phenol pesticides, and it can only be an assumption that the enzymes are comparable to the 0-methyltransferases which use S-adenosylmethionine as a methyl donor (Axelrod, 1971). Mercury fungicides that can cause serious poisoning effects have come under critical examination. Jernelov (1969) showed in a review that metallic mercury is oxidized chemically, but the subsequent conversion

MICROBIAL TRANSFORMATION OF PESTICIDES

111

of divalent inorganic mercury to methylmercury and dimethylmercury is caused by microorganisms : Hg

Chemical

Hgt2

Biologiral

CH3Hgf2

Biological

CHsHgCHS

Evidence for microbial methylation of mercury was presented by Wood et al. (1968)) who demonstrated that extracts of a Methanobacterium strain transferred the methyl group from methylcobalamin ( Co3+) to Hg2+.Yamada and Tonomura (1972) reported the methylation reaction of mercury in a pure culture of Clostridium cochlearium which was isolated from soil. McBride and Wolfe (1971) showed that under anaerobic conditions the same organisms also synthesize dimethylarsine from a variety of arsenic derivatives; adenosine triphosphate and hydrogen were found to be essential for this reaction with cell-free extract. In these studies it was also established that selenium and tellurium are readily methylated by a Methanobacterium sp. However, it should be pointed out that the transfer of methyl groups from Co3+to Hg2+may also occur as a nonenzymatic process (Imura et al., 1971; Bertilisson and Neujahr, 1971)) but it is enhanced by anaerobic conditions and by increasing numbers of bacteria capable of synthesizing alkylcobalamins ( Lezius and Barker, 1965; Wood and Wolfe, 1966). The formation of an arsenic gas compound generated by a fungus was already observed in the last century (Gosio, 1893), but only forty years later it was possible to correctly identify the volatile compound as trimethylarsine ( Challenger et aZ., 1933). Challenger described extensively in his review (1945) the ability of Scopulariopsis breuicaulis to methylate organic and inorganic forms of arsenic and other metalloids. Challenger et al. (1954) partially established the mechanism of methylation of arsenic. Arsenic-metabolizing microorganisms were isolated from soil and sewage and tentatively identified as Candida humicola, Gliocladium roseum, and a Penicillium species; these fungi formed trimethylarsine gas from monomethylarsonic acid and dimethylarsinic acid (COXand Alexander, 1973). An acetylation reaction in the metabolism of phenylurea herbicides was observed by Tweedy et al. (1970a,b). In many mammalian species acetylation is a common conjugation reaction especially for foreign aromatic amines ( Weber, 1971), but there is !ittle knowledge involving microorganisms using this process in pesticide transformation. The fungi Talaromyces wortmanii and Fusarium oxysporum metabolize metobromuron by demethylation and demethoxylation with the apparent, subseiuent acetylation of the aniline intermediate. p-Bromoaniline was not found as an intermediate, but it can be assumed that acetylation of the aniline is a fast process; consequently, it does not accumulate in

112

JEAN-MARC BOLLAG

the culture medium. If p-bromoaniline was used as a substrate, it was completely acetylated to p-bromoacetanilide by the two fungi tested as well as by a Bacillus sp. and Chlorella vulgaris (Tweedy et al., 1970a).

Likewise, p-chloroaniline was converted to p-chloroacetanilide in the growth medium of Fusarium oxysporum, but only approximately 3%of the parent aniline was acetylated (Kaufman et al., 1973). One report states that formylation of aniline in soil was detected as a transformation process ( Kearney and Plimmer, 1972); 3,4-dichloroformylanilide was identified as a product of 3,4-dichloroaniline, but the possible participation of microorganisms in this reaction was not examined. Conjugation reactions of sulfhydryl-containing compounds, with natural metabolites such as amino acids, has been shown to take place in vitro. Kaars Sijpesteijn et al. (1962), studying the transformation of dithiocarbamate fungicides, showed that when cell suspensions of various microorganisms were incubated with the sodium salt of dimethyldithiocarbamate, the compound was converted to 7 - ( dimethylthiocarbamoythio ) -a-aminobutyric acid and the corresponding keto acid:

(CH,),: N * C . S (CH,), II

.CO .COOH

S

The studies were performed with washed cell suspensions of Saccharomyces cerevisiae, Hansenula anomala, and Escherichia coli as well as mycelial pellets of Glomerella cingulata, Aspergillus niger, and Cladosporium cucumerinum; all these microorganisms produced at least one conjugate from dimethyldithiocarbamate. A sulfhydryl-oxidizing enzyme system which catalyzes the conversion of dithiocarbamates to the corresponding disuIfides was isolated from the cuIture filtrate of PiricuZaria oryzae and Polyporus versicolor (Neufeld et al., 1958). For instance, sodium diethyldithiocarbamate was oxidized by atmospheric oxygen to tetraethylthiuram disulfide: 2(CZHs)2N-C-SH

II

S

+ 3402

-+

(CZH~)~N-C-S-S-C-N(C~H~)~

II

S

I1

S

+ H2O

MICROBIAL TRANSFORMATION OF PESTICIDES

113

There is some concern related to aniline-based herbicides like phenylureas, phenylcarbamates, and acylanilides whose aniline intermediate product can be polymerized to an azo-derivative, a group of compounds with possible carcinogenic effects in animals (Weisburger and Weisburger, 1966). Bartha and Pramer (1967) reported first that soil treated with propanil, 3’,4’-dichloropropionanilide, produced as a major metabolite 3,3’,4,4’-tetrachloroazobenzene (Fig. 3, B ) . The synthesis of the azo compound was a result of microbial activity, since the condensation product was not detected in sterilized soil that received propanil or 3,4-dichloroaniline. Bartha et al. (1968) studied the ability of aniline and mono- and dichlorinated anilines to form azo compounds; aniline did not form a condensation product, but all monochloro- and some dichloroanilines were transformed to their corresponding dichloro- and tetrachloroazobenzenes. Of particular interest are reports describing the formation of asymmetric as well as symmetric azo compounds which are the result of different anilines added simultaneously to soil (Bartha, 1969; Kearney et al., 1969). Hybridization, between different substituted anilines released from propanil and solan, produced two symmetrically formed azobenzenes, 3,3’,4,4‘-tetrachloroazobenzene ( Fig. 3, B ) and 3,3‘-dichloro-4,4’-dimethylazobenzene, as well as the hybrid, 3,3‘,4-trichloro-4’methylazobenzene ( Bartha, 1969). Condensation of halogenated anilines can be further complicated as

Cl (C)

FIG.3. Formation of azobenzenes from 3,4-dichloroaniline.

114

JEAN-MARC BOLLAG

illustrated by the isolation of 1,3-bis( 3,4-dichlorophenyl)triazene ( Fig. 3, D), which apparently arises from the reaction of 3,4-dichloroaniline with nitrite to form an intermediate diazonium cation, which subsequently reacts with another molecule of free aniline to produce the triazene ( Plimmer et al., 1970). Another aniline condensation product is 4-( 3,4-dichloroanilino)-3,3’,4’-trichloroazobenzene ( Fig. 3, C ) which resulted from the addition of another 3,4-dichloroaniline molecule to the previously formed azobenzene ( Linke, 1970). A report by Daniels and Saunders (1953) described the synthesis of 4,4’-dichloroazobenzene from monochloroaniline by a peroxidase; therefore, it was assumed that an analogous enzyme system is also active in the soil. Bartha et al. (1968) demonstrated that there was a considerable similarity in the azobenzene condensation of various anilines by the selected soil and a horseradish peroxidase. It was also possible to extract peroxidase from soil which would catalyze the conversion of chloroanilines to chloroazobenzenes after addition of H,O, ( Bartha and Bordeleau, 1969). A pathway of chloroazobenzene formation was proposed by Bordeleau et al. (1972). They concluded from their studies that an initial attack of peroxidase produced a free chloroanilino radical, which was transformed to another labile intermediate, chlorophenylhydroxylamine, which condensed spontaneously with excess chloroaniline and formed chloroazobenzene. This reaction sequence was suggested as the main pathway, although another pathway could also be anticipated. IV.

Chemical Structure and Microbial Transformation Relationship

The concern over the persistence of a pesticide, or a derivative of it, and the related probable toxic hazard in the environment evoked much speculation on the ability of microbial organisms to transform such a xenobiotic compound. It is self-evident that an experimental approach contributes much to elucidate this problem, but the innumerable possibilities of chemical structures to which a microorganism can be exposed makes it unreal to test all possible transformations of each compound under the various conditions. Therefore, it is desirable to know the possible avenues of microbial attack in relation to specific molecular configuration. Acquaintance with enzymatic reactions in the metabolism of investigated pesticides or other compounds helps to anticipate certain chemical changes; on the other hand, it has been recognized that factors like cellular permeation of the chemical and its steric and electronic characteristics influence microbial activity. A new approach, combining chemical reactivity with substituent parameters and understanding of the multiconditional character of structure-activity or structure-degrad-

MICROBIAL TRANSFORMATION OF PESTICIDES

115

ability relationships using regression analysis, was initiated and developed by Hansch and other authors (Hansch, 1969, 1971; Verloop, 1972). It was shown that hydrophobic, steric, and electronic factors could be used to formulate mathematical structure-activity relationships, and the value of the equations has been tested on numerous bioactive compounds. Publications related to the application of the Hansch approach have been numerous in recent years (Verloop, 1972), but specific investigations on the relationships between microbial enzymes and transformation of a chemical structure have not yet been elaborated. From practical experience and from laboratory tests some general conclusions can be made concerning the probable microbial transformation of certain groups of pesticides and the possible enzymatic attack of certain linkages or substitutions.

A. MICROBIALTRANSFORMATION OF PESTICIDAL GROUPS There have been relatively few systematic studies designed to obtain data relating the chemical structure of pesticides to their probable microbial transformation. Most investigations in the past focused only on those microorganisms that could be isolated and were able to use the pesticidal molecules as a source of carbon, nitrogen, phosphorus, sulfur, or energy, or as a combination of these. The phenomenon of cometabolism, for instance, was not taken into consideration. It must be emphasized that laboratory conditions, using axenic cultures, do not permit the observation of the combined activity of various microbial species, as it has to be anticipated in an ecosystem with its inherent microflora. The same is true if metabolism would occur by auxotrophic microorganisms that require for their proliferation specific conditions that are not provided in general screening experiments. Consequently, investigations on the structure-biodegradability relationship have to be critically evaluated, but indications in the laboratory often have been shown to be compatible with the applied experience in pesticide use. In addition, the knowledge acquired on the biochemical or enzymatic processes-as outlined in Section III-provides the actual reactions that occur during the transformation of individual chemical groups of pesticides. From these results it is possible to establish a relationship to the persistence of the various pesticidal groups (Table 111). Pesticides which are attacked initially by a hydrolytic mechanism, like phenylcarbamates or organophosphates, are relatively short-lived in soil, whereas pesticidal groups which are dealkylated by a primary reaction appear to be more persistent. Halogenated alkanoic acids undergo dehalogenation and persist a relatively short time, but haloge-

TABLE 111 PERSISTENCE OF PESTICIDES IN SOILA N D ENZYMATIC TRANSFORMATION REACTIONS

Example

Pesticidal group Herbicides Halogenated alkanoic acids

c1 0 I II €I -C-C & -OH I

c1

Period of persistence in soil 2 to 8 Weeks

Enzymatic transformations by microorganisms Primary reactions

Subsequent reactions

Dehalogenation

Dalapon

5'

s-Triazines

18 Months

N-Dealkylation, dehalogenation

Deamination

4-18 Weeks

poxidation, cleavage of ether linkage, ring hy droxylation

Ring hydroxylation, ring cleavage

4-15 Months

N-Dealkylation, hydrolysis

Hydrolysis, acetylation, condensation reaction

Simazine

Phenoxyalkanoic acids

O--CH,--COOH QC'

Cl

2.4-D

Phenylureas

H O I II N-C-N,

,CH,

Cl

Linuron

F*

0

Phenylcarbamates

H

O I It N-C-0-C

H,CHS

7 Weeks

Ester hydrolysis

'CH,

Hydroxylation, ring cleavage

dl CIPC

Acylanilides

'::: N-c-cH&H,

Hydrolysis

Condensation reaction, acetylation

-zB

8 $

Cl

Propanil

Benzoic acids

bC1

> 6 Weeks

0

Decarboxylation, dehalogenation

Y

NH*

c1

Amiben

(Continued)

w F a,

TABLE I11 (Continued)

Pesticidal group

Example

Period of persistence in soil

Enzymatic transformations by microorganisms Primary reactions

Insecticides Organophosphates

Subsequent reactions ~ ~

~

Ring cleavage

Diazinon

Halogenated hydrocarbons

>9 Years

Epoxidation, dehalogenation

Hydrolysis

2-8 Weeks

Side-chain hydroxylation, ring hydroxylation, hydrolysis

Hydrolysis, ring cleavage

Sulfoxidation

Hydrolysis

Heptachlor

Methylcarbamates

Carbaryl

Fungicides Thiocarbamates

yL

HsC--S -C -C=N-O-C-N-CH, I 1 I/ I H,C H O H Temik

_

_

MICROBIAL TRANSFORMATION OF PESTICIDES

119

nated hydrocarbons with a more complex molecular configuration, like heptachlor, lindane, or DDT, persist for extended time periods. Generally, it was found that certain linkages in pesticides are readily susceptible to cleavage, and the rate at which these linkages cleave depend on the characteristics of the remaining molecule. Such observations indicate some general trends in the biodegradability of pesticide groups, but it must be stressed that numerous environmental conditions can interfere in the availability of a pesticide to microbial attack.

B. EFFECTOF VARIOUSSUBSTITUTIONS ON BIODEGRADABILITY Minor alterations in the structure of pesticide molecules frequently cause a drastic change in the susceptibility of such compounds to biotransformation. Introduction of polar groups, such as OH, COOH, NO2, and others, often affords microbial systems a site of attack, while others such as halogen or alkyl substitutions make a molecule more resistant. The rate of a reaction is also strongly influenced by steric and electronic factors of other atoms in the molecule. Generally, it can be stated that the type, the number, and the position of substitutions affect the rate of microbial decomposition of organic compounds. Which of these three factors is most influential depends upon the organic compounds studied. Investigations on this topic related to pesticide degradation were concerned especially with the influence of halogens in aliphatic acid herbicides and with the effect of substitution in the benzene ring. Experiments using sewage microorganisms, for instance, confirmed that unsubstituted aliphatic acids are degraded readily, but the rate of decomposition was much slower with substituted acids as substrates ( Dias and Alexander, 1971). A single halogen substitution, particularly if on the a-carbon, makes the molecule less susceptible to attack, and dihalogenated compounds were even more resistant to biodegradation. Jensen (1959) found that strains of Trichoderma viride, Clonostachys sp., and Acrostalagmus sp. degraded monochloroacetate more rapidly than dichloroacetate, but trichloroacetate was not attacked by these fungi. Many similar studies also indicated that increasing the number of halogen substitutions increased the resistance of a molecule to biodegradation. It was also shown that the rate of decomposition depends on the specific halogen substituent, i.e. chlorine, bromine, fluorine, or iodine, but no general conclusion concerning the possible attack by various organisms could be drawn ( Hirsch and Alexander, 1960). Introduction of substituents on a benzene ring influences its degradation considerably, and since many pesticides have an aromatic ring as an essential part of the molecule, its substitution is a determining factor for resistance to biodegradation. Systematic surveys of the effect of

120

JEAN-MARC BOLLAG

chemical structure on the microbial degradation of substituted benzenes have their shortcomings, since specific test conditions have to be selected. Results from various studies-with compounds that were not necessarily pesticidal-showed a certain agreement and trend related to the influence of the position of the substituent on the aromatic ring and its chemical nature. Kameda et al. (1957) found that not one of 34 soil pseudomonads was capable of degrading meta isomers of nitro-, amino-, and methoxybenzoates, but some could use the corresponding ortho- and para-substituted molecule. Likewise, Alexander and Lustigman ( 1966) determined that, in studies with a mixed soil microflora using an ultraviolet spectrophotometric assay to follow the destruction of aromaticity, meta isomer substitutions of various groups were almost invariably degraded more slowly than the ortho- or para-substituted analogs. In experiments with different monosubstituted compounds, they showed that phenol and benzoate were degraded rapidly, aniline and anisole were attacked less readily, and benzenesulfonate and especially nitrobenzene appeared to be most resistant to microbial transformation. With aromatic compounds containing two substituents, carboxyl and phenolic hydroxyl groups favored microbial degradation of the molecule while other groups, such as chloro, nitro, and sulfonate substitutions, reduced rates of metabolism. It should be pointed out that these experiments were performed while the test compound was supplied as the sole carbon source and cometabolic transformations were not considered. The effect of the chemical structure on the persistence of chlorinated phenoxyalkanoate herbicides was summarized by Alexander ( 1965b). He concluded that the type of linkage of the aliphatic acid to the ring, and the position-in this case not the number of chlorines-determines the persistence of these pesticides. Compounds containing a chlorine in the meta position are not metabolized to a significant extent, and substances which have the ring linked to the aliphatic side chain at the alpha position are more resistant to degradation. Similar observations were made with chlorophenols which have fungicidal activity. Also, CartWright and Cain (1959) reported that organisms could easily be isolated for growth on 0- and p-nitrobenzoic acids but with difficulty on the meta-substituted derivative. On the other hand it was determined that microbial degradation of chlorinated N-phenylcarbamates in soil perfusion studies was more rapid if substitution occurred at the meta-position than with the ortho- or para- substituted compounds ( Kaufman, 1966). However, Kearney ( 1967), studying the influence of physicochemical properties of phenylcarbamates that influence hydrolysis by a microbial enzyme, found that the isopropyl ester of p-nitrophenyl carbamate is hydrolyzed considerably faster than the corresponding meta compound. In addition, it was deter-

MICROBIAL TRANSFORMATION OF PESTICIDES

121

mined that reaction rates of hydrolysis decreased with the following meta-.substituents on the ring: NO, > CH,CO > CI > CH,O > H. The size of the molecule also has an effect on enzymatic hydrolysis as indicated by the faster degradation of the isopropyl ester of phenyl as compared to the 2-naphthylcarbamic acid. J.-M. Bollag, N. M. Henninger, and B. Bollag (unpublished data, 1973) found that the fungus Rhixoctonia solani metabolized chlorinated and brominated anilines most rapidly if the halogen was substituted in the para position; mta-substituted anilines were transformed slower and a substitution in the orthoposition proved to be most resistant to fungal attack. Studies on the persistence of DDT with a Hydrogenomonas sp. revealed that especially the para-chloro substitution and substituents on the methylene-carbon governed the resistance of the DDT molecule to microbial metabolism (Focht and Alexander, 1970). Many other examples of specific investigations which indicate a change of resistance to microbial transformation by a simple substitution in a pesticidal molecule could be cited, but much more research needs to be done, hopefully also by use of the Hansch approach, to be able to generalize in clearer terms the effect of small molecular alterations in a specific chemical group. No rules that are generally applicable can yet be identified, but certain trends could be established that influence biodegradability. It can also be stated that it is not possible to find a relationship between the effects of chemical structure of pesticides on toxicity to a target organism and the effects of a molecule on its microbial degradability. This problem has to be evaluated for each pesticidal group or even each single compound; this indicates the complexity for developing from theoretical considerations and experimental data a useful and practical pesticide. AND PESTICIDE TRANSFORMATION C. MOLECULARRECALCITRANCE

Alexander (1965a) attributed two main causes to the recalcitrance of chemicals: ( a ) environmental conditions not conducive to microbial ability to change a certain molecule and ( b ) the structural configuration of a compound, which makes it either totally or partially resistant to biodegradation under all circumstances. Whereas the first parameter for a compound as nonbiodegradable is generally accepted, the second cause cited often arouses criticism if the definition is considered from a basic scientific point of view. While Alexander (1965a) states that “every biologically synthesized organic molecule doubtlessly will, under some set of circumstances, be destroyed by one or several species,” he doubts that all synthetic organic compounds, which are increasingly produced

122

JEAN-MARC BOLLAG

and discharged into the various ecosystems, can be biodegraded. There is no doubt that, for instance, certain pesticidal groups are difficult to attack by microorganisms, and they-or a derivative of them-may persist for a considerable length of time in the environment. Although in several cases it appeared that a pesticide is nontransformable biologically, this finding had to be corrected when on-going research discovered that a pesticidal molecule could be metabolized, at least partially, under certain environmental conditions and by specific organisms. Are there really synthetic compounds that cannot be altered by microorganisms after mutational or nongenetic adaptation if there is a need for it? Presently, for instance, the question of possible biological transformation of synthetic polymers with a high molecular weight is yet unresolved, and it presents a justified practical concern related to the pollution of the environment. It appears that whether there are synthetic chemicals intrinsically resistant to biological degradation raises an academic question whose answer may, with synthesis of new chemicals, always be delayed.

V.

Conclusions

Microbial and biochemical processes affecting the fate and behavior of pesticides have been investigated essentially in model systems using isolated microbial cultures or enzyme systems. This appears-with the presently available techniques-to be the only feasible approach, if one is interested in elaborating the mechanism of pesticide transformation, clarifying the actual microbial activity by isolation and identification of intermediates, and establishing the rates at which these processes occur. With respect to pollution, the transformation of a xenobiotic compound should not be a matter of conjecture, since it is important to know the fate of the original pesticide as well as the resulting transformation products. The clarification of the extent to which microorganisms interfere and transform introduced chemicals or their decomposition products should help in determining the potential hazard of their use. It is also necessary to keep in mind that under various conditions or in different ecosystems, a chemical can be transformed by different metabolic pathways or organisms and consequently, the resulting product can vary. The knowledge of enzymatic reactions in the metabolism of pesticides or their identified intermediates should contribute to understanding transformation possibilities of newly developed compounds. This general problem, which is related to the molecular configuration and the resistance to microbial attack, needs far more research for pertinent and applicable conclusions, i.e., for the development of new pesticides which

123

MICROBIAL TRANSFORMATION OF PESTICIDES

have the desired toxic activity and are simultaneously susceptible to adequate microbial metabolism. One must be aware of the difficulty in extrapolating from experiments and results obtained in vitro to the complex environment of a natural habitat. Isolated organisms which alter a pesticide in pure culture conditions are not necessarily those responsible for its transformation in uiuo. However, it appears that basic laboratory studies are a prerequisite for establishing the possibilities of microbial pesticide transformations in a natural environment. PESTICIDlC3

Common or trade name Aldicarb Aldrin Amit,role Atrazine Brornophos Buturon Carbaryl Chlorbromuron Chlorobeneilate Chloroneb Chloroxuron Chlorpropham CIPC 2,4-1) I1a1apo n DDA 1IIIII (TIIE) 1lDE IIDT Iliazinon Dicamba Ilicofol ilicryl Dieldrin Dinitramine Diphenamid Dipterex Diuron

TABLE I V MENTIONICD I N 'PHI', TEXTAND THEIR CHEMICAL

I)ESIGNATION

Chemical designation 2-Methyl,-2-(methylthio)propionaldehyde0-(methylcarbamoy1)oxime 1,2,3,4,10,l0-Hexachloro-1,4,4aJ5,8,8a-hexahydro-1,4-endo,eso-5,8diniethanonaphthalene %Amino- 1,2,4-tariazole 2-Chloro-4-(ethylamino)-6- (isopropy1amino)-s-triazine 0-(4-Bromo-2,5-dichlorophenyl) 0,O-dimethylphosphorothioate 3-(p-Chlorophenyl)- 1-methyl- 1-( 1-methyl-2-propyny1)urea 1-Naphthyl N-methylcarbamate 3- (3-Chloro-4-bromophenyl)-l-methoxy-l-methylurea Ethyl 4,4'-dichlorobenzilate 1,4-Dichloro-2,5-dimethoxybenzene 3-[4-(p-Chlorophenoxy)phenyl]-l, 1-dimethylurea See CIPC Isopropyl N-(3-chlorophenyl) carbamate 2,4-Dichlorophenoxyacetic acid 2,2-Dichloropropionic acid 2,2-Bis(p-chlorophenyl)aceticacid 2,2-Bis(p-chlorophenyI)- 1,l-dichloroetharie 2,2-Bis (p-chloropheny1)- 1,1-dichloroethene 2,2-Biu (p-chloropheny1)-1, 1,1-trichloroethane 0,O-Diethyl 0-(2-isopropy1-4-methyl-6-pyrimidinyl) phosphorothioate 3,6-L)ichloro-o-anisic acid 1,l-bis (p-~hlorophenyl)-2,2,2-trichloroethanol N-(3,4-IIichlorophenyl) methacrylamide 1,2,3,4,10,10-Hexachloro-6,7-epoxy-1,4,4a,5,6,7,8,8a-octahy~ro1,4-endo,exo-5,8-dimethanonaphthalene N3,N~-r)iethyl-2,4-dinitro-6-trifluoromethyl-m-phenylenediamine N,N-Dimethyl 2,2-diphenylacetamide See Trichlorfon 3. (3,4-lXchlorophenyl)-1,l-dimethylurea (Continued)

124

JEAN-MARC BOLLAG

TABLE I V (Continued) Common or trade name

DNOC Fenitrothion Fenuron Fluometuron Griseofulvin Heptachlor Ioxynil Isodrin Karsil Lindane Linuron Malathion MCPA Methoxychlor Metobromuron Monolinuron Monuron Par aoxon Paraquat Parathion PCNR Phorate Prometryne Propanil Simazine Solan Sumithion 2,4,5-T 2,3,6-TBA TDE Temik TEPP Thimet Trichlorfon Trifluralin

Chemical designation 3,5-Dinitro-o-cresol See Sumithion l,l-Dimethyl-3-phenylurea l,l-I~imethyl-3-(~,or,a-trifluoro-m-tolyl)urea 7-Chloro-4: 6 :2’-trimethoxy-6’-methylgris-2’-ene-3 :4’-dione 1,4,5,6,7,R,8-Heptachloro-3a,4,7,7a-tetrahydro-4,7-methanoindene 4-Hydroxy-3,5-diiodobenzonitrile 1,2,3,4,10,10-Hexachloro1,4,4a,.5,8,8a-hexahydi-ol,Cendo,endo5,s-dimethanonaphthalene N-(3,4-lXchlorophenyl) 2-methylpentanamide y- 1,2,3,4,.~,6-Hexschlorocyclohexane 3-(3,4-I~ichlorophenyl)-l-methoxy-l-methylurea 0,O- Dimethyl S-bis (carboethox y)ethyl phosphorodi thioate 4-Chloro-Zmethylphenoxyacetic acid 2,2-Ris(p-methoxyphenyl)-l, 1,l-trichloroethane 3- (p-Bromopheny1)- 1-methoxy- 1-methylurea 3-(p-Chlorophenyl)-l-melhoxy-1-methylurea 3-(p-Chlorophenyl)-l,1-dimethylurea 0,O-diethyl-0-p-nitrophenyl phosphate 1,l’-Dimethyl 4,4’-bipyridinium salt 0,O-Diethyl 0-p-nitrophenyl phosphorothioate Pentachloronitrobenzene 0,O-Diethyl S-(ethylthiomethyl) phosphorodithioate 2,4-Bis (isopropylamino)-6-methylthio-s-triazine 3‘,4’-Dichloropropionanilide ZChloro-4,6-bis(ethylarnino)-s-triazine N-(3-Chloro-4-methylphenyl)-2-methylpentanamide 0,O-Dimethyl 0-(3-methyl-4-nitrophenyl)phosphorothionate 2,4,5-Trichlorophenoxyacetic acid 2,3,6-Trichlorobenzoic acid See D D D 2-Methyl-2-(mrthylthio) propionaldehyde-l 0-(methylcarbamoyl) oxime Tetraethyl pyrophosphate See Phorate 0,O-Dimethyl 2,2,2-trichloro- 1-hydroxyethyl phosphonate ol,a,a-Trifluoro-~,6-dinitro-N,~-dipropyl-p-toluidine

REFERENCES Ahmed, M. K., and Casida, J. E. (1958). J. Econ. Entomol. 51,59. Alexander, M. (1965a). Aduan. Appl. Microbiol. 7, 35. Alexander, M. ( 196513). Soil Sci. SOC. A m r . , Proc. 29, 1. Alexander, M. (1967). In “Agriculture and the Quality of Our Environment,” Publ. No. 85, pp. 331-342. Amer. Ass. Advan. Sci., Washington, D.C. Alexander, M. (1969). In “Soil Biology,” pp. 209-240. UNESCO, Paris. Alexander, M., and Lustigman, B. K. ( 1966). J. Agr. Food Chem. 14,410. Axelrod, J. (1956). Biochem. J. 63, 634.

MICROBIAL TRANSFORMATION OF PESTICIDES

125

Axelrod, J. ( 1971). In “Handbuch der experimentellen Pharmakologie” ( B. B. Brodie and J. R. Gillette, eds.), Vol. 28, Part 11, pp. 609-619. Springer-Verlag, Berlin and New York. Barker, P. S., Morrison, F. O., and Whitaker, R. S. (1965). Nature (London) 205, 621. Bartha, R. (1969). Science 166, 1299. Bartha, R., and Bordeleau, L. M. (1969). Soil Biol. G Biochem. 1, 139. Bartha, R., and Pramer, D. (1967). Science 156, 1617. Bartha, R., Linke, H. A. B., and Pramer, D. (1968). Science 161, 582. Bernhardt, F.-H., Ruf, H. H., Staudinger, H., and Ulrich, V. ( 1971). Hoppe Seyler’s 2. Physiol. Chem. 352, 1091. Bertilisson, L., and Neujahr, H. Y. ( 1971). Biochemistry 10, 2805. Bocks, S. M., Lindsay-Smith, J. R., and Norman, R. 0. C. (1964). Nature (London) 201, 398. Bollag, J.-M., and Alexander, M. (1971). Soil Biol. G Biochem. 3, 91. Bollag, J.-M., and Liu, S.-Y. ( 1971). Soil Bio2. 6 Biochem. 3, 337. Bollag, J.-M., and Liu, S.-Y. ( 1972a). Nature (London) 236, 177. Bollag, J.-M., an,d Liu, S.-Y. (1972b). Can. J. Microbiol. 18, 1113. Bollag, J.-M., Helling, C. S., and Alexander, M. (1967). Appl. Microbiol. 15, 1393. Bollag, J.-M., Helling, C. S., and Alexander, M. (1968a). J. Agr. Food Chem. 16, 826. Bollag, J.-M., Briggs, G. G., Dawson, J. E., and Alexander, M. ( 196813). J . Agr. Food Chem. 16, 829. Boothroyd, B., Napier, E. J,, and Somerfield, G. A. (1961). Biochem. J. 80, 34. Bordeleau, L. M., Rosen, J. D., and Bartha, R. (1972). J. Agr. Food Chem. 20, 573. Mrner, H. ( 1967). Z. Pflan~enkr.(Pflanzenpathol.) Pflanzenschutz 74, 135. Braunberg, R. C., and Beck, V. (1968). J . Agr. Food Chem. 16, 451. Byrde, R. J. W., and Woodcock, D. ( 1957). Bioclzep. J. 65, 682. Cartwright, N. J., and Cain, R. B. (1959). Biochem. J. 71, 248. Cartwright, N. J., and Smith, A. R. W. (1967). Biochem J. 102, 826. Cartwright, N. J., Holdom, K. S., and Broadbent, D. A. (1971). Microbios 3, 113. Castro, C. E., and Bartnicki, E. W. (1965). Biochim. Biophys. Acta 100, 384. Castro, C. E., and Belser, N. 0. (1968). Enuiron. Sci. Techno2. 2, 779. Chacko, C . I., and Lockwood, J. L. ( 1967). Can. J. Microbial. 13, 1123. Chacko, C. I., Lockwood, J. L., and Zabik, M. (1966). Science 154, 893. Challenger, F. ( 1945). Chem. Reu. 36, 315. Challenger, F., Higginbottom, C., and Ellis, L. (1933). J. Chem. SOC. London p. 95: Challenger, F., Lisle, D., and Dransfield, P. (1954). J. Chem. SOC., London p. 1760. Chapman, P. J. (1972). I n “Degradation of Synthetic Organic Molecules in the Biosphere,” pp. 17-55. Nat. Acad. Sci. Washington, D.C. Chisaka, H., and Kearney, P. C. (1970). J . Agr. Food Chem. 18,854. Clifford, D. R., and Woodcock, D. (1964). Nature (London) 203, 763. Coppedge, J. R., Lindquist, D. A., Bull, D. L., and Dorough, H. W. ( 1967). J . Agr. Food Chem. 15,902. Couch, R. W., Gramlich, J. V., Davis, D. E., and Funderburk, H. H. ( 1965). Proc. S . Weed Conf. 18, 623.

126

JEAN-MARC BOLLAG

Cox, D. P., and Alexander, M. (1973). Bull. Enuiron. Contam. Toxicol. 9, 84.

Crosbi, D. G., and Li, M.-Y. ( 1969). In “Degradation of Herbicides” (P. C. Kearney and D. D. Kaufman, eds.), pp. 321363. Dekker, New York. Cserjesi, A. J., and Johnson, E. L. (1972). Can. J . Microbiol. 18, 45. Dagley, S . (1971). Adoan. Microbial Physiol. 6, 1. Dalton, R. L., Evans, A. W., and Rhodes, R. C. (1966). Weeds 14, 31. Daly, J. ( 1971). I n “Handbuch der experimentellen Pharmakologie” (B. B. Brodie and J. R. Gillette, eds.), Vol. 28, Part 11, pp. 285-311. Springer Verlag, Berlin and New York. Daly, J. W., Guroff, G., Udenfriend, S., and Witkop, B. (1968). Biochem. Pharmucol. 17, 31. Daly, J. W., Jerina, D. M., and Witkop, B. ( 1972). Experientia 28, 1129. Daniels, D. G. H., and Saunders, B. C. (1953). J . Chem. SOC., London p. 822. Dias, F. F., and Alexander, M. ( 1971). Appl. Microbiol. 22, 114. Engelhardt, G., Wallnofer, P. R., and Plapp, R. (1971). Appl. Microbiol. 22, 284. Engelhardt, G., Wallniifer, P. R., and Plapp, R. (1972). Appl. Microbiol. 23, 664. Evans, W. C. (1969). In “Fermentation Advances” (D. Perlman, ed.), pp. 649-687. Academic Press, New York. Evans, W. C., Gaunt, J. K., and Davies, J. I. (1963). Abstr. Commun., Int. Congr. Biochem., 5th, 1961. p. 364. Evans, W. C., Smith, B. S. W., Moss, P., and Femley, H. N. (1971a). Biochem. J . 122, 509. Evans, W. C., Smith, B. S. W., Femley, H. N., and Davies, J. I. (1971b). Biochem. J . 122, 543. Faulkner, J. K., and Woodcock, D. (1961). J. Chem. Soc., London p. 5397. Faulkner, J. K., and Woodcock, D. ( 1964). Nature (London) 203, 865. Faulkner, J. K., and Woodcock, D. (1965). J . Chem. Soc., London p. 1187. Focht, D. D., and Alexander, M. ( 1970). Science 170, 91. Focht, D. D., and Alexander, M. (1971). J . Agr. Food Chem. 19, 20. Foster, J. W. (1962). In “The Oxygenases” ( 0 .Hayaishi, ed.),pp. 241-271. Academic Press, New York. French, A. L., and Hoopingamer, R. A. (1970). J . Econ. Entomol. 63, 756. Funderburk, H. H., and Bozarth, G. A. ( 1967). J . Agr. Food Chem. 15,563. Funderburk, H. €I., Schultz, D. P., Negi, N. S., Rodriguez-Kabana, R., and CurI, E. A. ( 1967). Proc. S. Weed Conf. 20, 389. Gamar, Y., and Gaunt, J. K. ( 1971). Biochem. J . 122, 527. Gaunt, J. K., and Evans, W. C. (1971). Biochem. J . 122,533. Geissbuhler, H. (1969). In “Degradation of Herbicides” ( P . C. Kearney and D. D. Kaufman, eds.), pp. 79-111. Dekker, New York. Geissbiihler, H., Haselbach, C., Aebi, H., and Ebner, L. (1963). Weed Res. 3, 277. Getzin, L. W., and Chapman, R. K. ( 1960). J . Econ. Entomol. 53, 47. Goldman, P. (1965). J . Biol. Chem. 240, 3434. Goldman, P., and Milne, G. W. A. (1966). J. Biol. Chem. 241, 5557. Goldman, P., Milne, G. W. A., and Pignataro, M. T. (1967). Arch. Biochem. Biophys. 118, 178. Gosio, B. (1893). Arch. Ital. Biol. 18, 253. Gram, T. E. ( 1971). In “Handbuch der experimentellen Pharmakologie” (B. B. Brodie and J. R. Gillette, eds.), Vol. 28, Part 11, pp. 334-348. Springer-Verlag, Berlin and New York. Gregory, W. W., Reed, J .K., and Priester, L. E. (1969). J . Protozoal. 16, 69.

MICROBIAL TRANSFORMATION OF PESTICIDES

127

Guenzi, W. D., and Beard, W. E. ( 1967). Science 156, 116. Guenzi, W. D., and Beard, W. E. (1968). Soil Sci. SOC. Amer., Proc. 32, 522. Cunner, H. B., and Zuckerman, B. M. (1968). Nature (London) 217, 1183. Curoff, G., Kondo, K., and Daly, J. (1966). Biochem. Biophys. Res. Commun. 25, 622. Guroff, G., Daly, J. W., Jerina, D. M., Renson, J., Witkop, B., and Udenfriend, S. (1967). Science 157, 1524. Gutenmann, W. H., Loos, M. A., Alexander, M., and Lisk, D. J. (1964). Soil Sci. SOC. Amer., Proc. 28, 205. Hamdi, Y. A., and Tewfik. M. S. (1970). Soil Biol. G Biochem. 2, 163. Hansch, C. (1969). Accounts Chem. Res. 2, 232. Hansch, C. ( 1971). In “Drug Design” ( E . J. Ariens, ed.), Vol. 1, pp. 271-342. Academic Press, New York. Helling, C. S., Bollag, J.-M., and Dawson, J. E. ( 1968). J . Agr. Food Chem. 16, 538. Helling, C. S., Kearney, P. C., and Alexander, M. (1971). Adoun. Agron. 23, 147. Henderson, M. E. K. (1957). J. Gen. Microbiol. 18,686. Hirsch, P., and Alexander, M. (1960). Can. J . Microbiol. 6, 241. Hock, W. K., and Sisler, H. D. (1969). J . Agr. Food Chem. 17, 123. Horvath, R. S. ( 1970a). Bull. Enuiron. Contum. Toxicol. 5, 537. Horvath, R. S. (1970b). Biochem. 3. 119,871. Horvath, R. S. (1971). J . Agr. Food Chem. 19, 291. Horvath, R. S. (1972). Bacteriol. Reu. 36, 146. Imura, N., Sukegawa, E., Pan, S.-K., Nagao, K., Kim, J.-Y.,Kwan, T., and Ukita, T. (1971). Science 172, 1248. Jensen, H. L. (1959). Actu Agr. Scund. 9, 421. Jensen, H. L. (1963). Actu Agr. Scund. 13, 404. Jerina, D., Daly, J., Witkop, B., Zaltzman-Nirenberg, P., and Udenfriend, S. (1969). Biochemistry 9, 147. Jemelov, A. ( 1969). In “Chemical Fallout: Current Research on Persistent Pesticides” (M. W. Miller and G. G. Berg, eds.), pp. 68-74. Thomas, Springfield, Illinois. Johnson, B. T., and Kennedy, J. 0.(1973). Appl. Microbiol. 26, 66. Johnson, B. T., Goodman, R. N., and Goldberg, H. S. ( 1967). Science 157,560. Johnston, H. W., Briggs, G. G., and Alexander, M. (1972). Soil Biol. G Biochem. 4, 187. Kaars Sijpesteijn, A., Kaslander, J., and van der Kerk, G. J. M. (1962). Biochim. Biophys. Actu 62, 587. Kallman, B. J., and Andrews, A. K. ( 1963). Science 141, 1050. Kameda, Y., Toyoura, E., and Kimura, Y. (1957). Kanuzawa Duiguku Yukugukubu Kenkyu Nempo 7, 37. Kaufman, D. D. (1966). In “Pesticides and Their Effects on Soils and Water,” Symp. Pap., Spec. Publ. No. 8, pp. 85-94. Soil Sci. SOC.Amer., Inc., Madison, Wisc. Kaufman, D. D., and Blake, J. (1970). Soil. Biol. G Biochem. 2, 73. Kaufman, D. D., and Kearney, P. C. (1970). Residue Reu. 32,235. Kaufman, D. D., Plimmer, J. R., and Klingebiel, U. I. (1973). J. Agr. Food Chem. 21, 127. Kaufman, S., Bridgers, W. F., Eisenberg, F., and Friedman, S. ( 1962). Biochem. Biophys. Res. Commun. 9, 497. Kearney, P. C. (1965). J . Agr. Food Chem. 13,561. Kearney, P. C. ( 1967). J . Agr. Food Chern. 15,568. Kearney, P. C., and Plimmer, J. R. (1972). J . Agr. Food Chem. 20,584. Kearney, P. C., Kaufman, D. D., and Beall, M. L. (1964). Biochem. Biophys. Res. Commnun. 14, 29.

128

JEAN-MARC BOLLAG

Kearney, P. C., Kaufman, D. D., and Sheets, T. J. (1965). J . Agr. Food Chem. 17, 1418. Kearney, P . C., Plimmer, J. R., and Guardia, F. B. (1969). J . Agr. Food Chem. 17, 1418. Keil, J. E., and Priester, L. E. ( 1969). Bull. Enuiron. Contam. Toxicol. 4, 169. Kesner, C. D., and Ries, S. K. ( 1967 ). Science 155, 210. Knuesli, E., Berrer, D., Dupuis, G., and Esser, H. (1969). In “Degradation of Herbicides” (P. C. Kearney and D. D. Kaufman, eds.), pp. 51-78. Dekker, New York. KO, W. H., and Farley, J. 0.(1969). Phytopathology 59, 64. KO, W. H., and Lockwood, J. L. (1968). Can. J . Microbiol. 14, 1075. Korte, F., Ludwig, G., and Vogel, J. (1962). Justus Liebigs Ann. Chem. 656, 135. Laanio, T. L., Kearney, P. C., and Kaufman, D. D. ( 1973). Pestic. Biochem. Physiol. 3, 271. Langlois, B. E. (1967). J . Dairy Sci. 50, 1168. Lanzilotta, R. P., and Pramer, D. (1970). Appl. Microbiol. 19, 307. Leshniowsky, W. O., Dugan, P. R., Pfister, R. M., Frea, J. I., and Randles, C. 1. (1970). Science 169, 993. Lezius, A. G., and Barker, H. A. (1965). Biochemistry 4, 510. Lichtenstein, E. P., and Schulz, K. R. (1964). J . Econ. Entomol. 57, 618. Linke, H. A. B. ( 1970). Naturwissenschaften 57, 307. Lipke, H., and Kearns, C. W. (1959). J . Biol. Chem. 234, 2129. Liu, S.-Y., and Bollag, J.-M. (1971a). J . Agr. Food Chem. 19, 487. Liu, S.-Y., and Bollag, J.-M. (1971b). Pestic. Biochem. Physiol. 1, 366. Loos, M. A. ( 1969). In “Degradation of Herbicides” (P. C . Kearney and D. D. Kaufman, eds.), pp. 1-49. Dekker, New York. Loos, M. A., Bollag, J.-M., and Alexander, M. (1967a). J . Agr. Food Chem. 15, 858. Loos, M. A., Roberts, R. N., and Alexander, M. (1967b). Can. j . Microbiol. 13, 691. McBride, B. C., and Wolfe, R. S. (1971). Biochemistry 10, 4312. MacRae, I. C., and Alexander, M. (1963). J . Bacterial. 86, 1231. Madhosingh, C. (1961). Can. 1. Microbiol. 7, 553. Matsumura, F., and Boush, G. M. (1966). Science 153, 1278. Matsumura, F., and Boush, G. M. (1967). Science 156, 959. Matsumura, F., and Boush, G. M. (1968). J . Econ. Entomol. 61, 610. May, S. W., and Abbott, B. J. (1973). J . Biol. Chem. 248, 1725. Mendel, J. L., and Walton, M. S. (1966). Science 151, 1527. Mendel, J. L., Klein, A. K., Chen, J. T., and Walton, M, S. (1967). J . Ass. Ofic. Anal. Chem. 50, 897. Menzie, C. M. (1969). US.,Fish Wildl. Sew., Spec. Sci. Rep; Wildl. 127. Miles, J. R. W., Tu, C. M., and Harris, C. R. (1969). J. Econ. Entomol. 62, 1334. Milne, G. W. A., Goldman, P., and Holtzman, J. L. (1968). J . Biol. Chem. 243, 5374. Miyamoto, J., Kitagawa, K., and Sato, Y. (1966). Jap. J . Exp. Med. 36, 211. Miyazaki, S., Boush, M., and Matsumura, F. (1970). 1. Agr. Food Chem. 18, 87. Mounter, L. A., and Tuck, K. D. (1956). J. B i d . Chem. 221, 537.

MICROBIAL TRANSFORMATION OF PESTICIDES

129

Mounter, L. A., Baxter, R. F., and Chanutin, A. (1955). J . Biol. Chem. 215, 699. Nakanishi, T., and Oku, H. (1969). Phytopathology 59, 1761. Neuberg, C., and Welde, E. (1914). Biochem. Z. 60, 472. Neufeld, H. A,, Green, L. F., Latterell, F. M., and Weintraub, R. L. (1958). J . Biol. Chern. 232, 1093. Pfaender, F. K., and Alexander, M. (1972). J . Agr. Food Chem. 20, 842. Plimmer, J. R., and Kearney, P. C. (1969). Abstr., 158th Meet., Amer. Chem. SOC. Paper A-30. Plimmer, J. R., Kearney, P. C., and von Endt, D. W. (1968). J. Agr. Food Chem. 16, 594. Pfimmer,, J. R., Kearney, P. C., Chisaka, H., Yount, J. B., and Klingebiel, U. I. (1970). J. Agr. Food Chem. 18, 859. Raymond, D. D., and Alexander, M. (1972). Pestic. Biochem. Physiol. 2, 270. Raymond, D. G . M., and Alexander, M. (1971). Pestic. Biochem. Physiol. 1, 123. Ribbons, D. W. (1971). FEBS Lett. 12, 161. Ruiz-Herrera, J., and Starkey, R. L. (1969). J . Bacteriol. 99, 544. Saz, A. K., and She, R. B. (1954). Arch. Biochem. Biophys. 51, 5. Sharabi, N. El-D., and Bordeleau, L. M. (1969). Appl. Microbiol. 18, 369. Shida, T., Honima, Y., and Misato, T. (1973). Agr. Biol. Chem. 37, 1027. Stanier, R. Y., and Ornston, L. N. (1973). Advan. MicrobioZ. PhysioE. 9, 89. Stenersen, J. (1969). Bull. Enuiron. Contam. Toxicol. 4, 104. Stenersen, J. H. V. ( 1965). Nature (London) 207, 660. Taylor, H. F., and Wain, R. L. (1962). Proc. Roy. SOC., Ser. 156, 172. Tewfik, M. S., and Evans, W. C. (1966). Biochem. J . 99, 31p. Tiedje, J. M., and Alexander, M. (1969). J. Agr. Food Chem. 17, 1080. Tiedje, J. M., Duxbury, J. M., Alexander, M., and Dawson, J. E. (1969). J. Ag?. Food Chem. 17, 1021. Tonomura, K., Futai, F., Tanabe, O., and Yamaoka, T. (1965). Agr. Biol. Chem. 29, 124. Tu, C . M., Miles, J. R. W., and Harris, C. R. (1968). Life Sci. 7, 311. Tweedy, B. G., Loeppky, C., and Ross, J. A. (1970a). J . Agr. Food Chem. 18, 851. Tweedy, B. G., Loeppky, C., and Ross, J. A. ( 1970b). Science 168,482. Verloop, A. (1972). In “Drug Design” ( E . J. Ariens, ed.), Vol. 3, pp. 133-187. Academic Press, New York. Voerman, S . , and Tammes, P. M. L. (1969). Bull. Enoiron. Contam. Toxicol. 4 , 271. Wallnofer, P. (1969). Weed Res. 9, 333. Wallnofer, P. R., and Bader, J. (1970). App!. Microbiol. 19, 714. Wallnofer, P. R., Koniger, M., Safe, S., and Hutzinger, 0. (1972). J . Agr. Food Chem. 20, 20. Wallnofer, P. R., Safe, S., and Hutzinger, 0. (1973). Pest. Biochem. Physiol. 3, 253. Weber, W. W. (1971). In “Handbuch der experimentellen Pharmakologie” (B. B. Brodie and J. R. Gillette, eds.), Vol. 28, Part 11, pp. 564483. Springer-Verlag, Berlin and New York. Webley, D. M., Duff, R. B., and Farmer, V. C. (1957). Nature (London) 179, 1130. Wedemeyer, G. ( 1966). Science 152, 647. Wedemeyer, G . (1967). Appl. Microbiol. 15, 569.

130

JEAN-MARC BOLLAG

Wedemeyer, G. (1968). AppZ. Microbiol. IS, 661. Weinberger, M., and Bollag, J.-M. (1972). Appl. Microbiol. 24,750. Weisburger, J. H., and Weisburger, E. K. (1966). Clzern. Eng. News 44, 124. Wiese, M. V., and Vargas, J. M. ( 1973). Pestic. Biochem. Physiol. 3, 214. Williams, A. K., Cox, S. T., and Eagon, R. G . (1965). Biochem. Biophys. Kes. Commzm. 18, 250. Williams, R. T. ( 1971). In “Handbuch der experimentellen Pharmakologie” (B. B. Brodie and J. R. Gillette, eds.), Vol. 28, Part 11, pp. 226-242. Springer-Verlag, Berlin and New York. Wood, J. M., and Wolfe, R. S. (1966). 1. Bacterial. 92, 696. Wood, J. M., Kennedy, F. S., and Rosen, C. G . (1968). Nature (London) 220, 173. Yamada, M., and Tonomura, K. (1972). Hakko Kogaku Zasshi 50, 159. Yule, W. N., Chiba, M., and Morley, H. V. (1967). J. Agr. Food Chem. 15, 1000. Zaki, M. A., Taylor, H. F., and Wain, R. L. (1967). Ann. Appl. Biol. 59, 481. Zayed, S. M. A. D., Mostafa, I. Y., and Hassan, A. (1965). Arch. Mikrobiol. 51, 118.