Accumulation, Metabolism, Effects Organochlorine ...Accumulation, Metabolism, and Effects...

Vol. 46, No. 1MICROBIOLOGICAL REVIEWS, Mar. 1982, p. 95-1270146-0749/82/010095-33$02.00/0

Accumulation, Metabolism, and Effects of OrganochlorineInsecticides on Microorganisms

RUP LAL* AND D. M. SAXENADepartment of Zoology, University of Delhi, Delhi-110007, India

INTRODUCTION ............................................................... 95

ENTRY OF ORGANOCHLORINE INSECTICIDES INTO MICROBIALENVIRONMENTS .......................................................... 97

ACCUMULATION ............................................ .......... 97

Accumulation of Various Organochlorine Insecticides in Microorganisms ... .......... 97Mechanism of Accumulation................................................... 100

METABOLISM ................................................................. 101

Major Reactions of Organochlorine Insecticide Metabolism .... ..................... 101Reductive dechlorination..................................................... 101Dehydrochlorination ......................................................... 103

Oidation .................................................................. 103

Isomerization ............................................................... 105

Metabolism of Individual Insecticides and Pathways of Metabolism................... 105

1,1,1-Trichloro-2-2-bis(p-chlorophenyl)ethane ................................... 105

(i) Metabolism............................................................ 105(ii) Pathways of metabolism.................................................. 106

1,2,3,4,5,6-Hexachlorocyclohexane ............................................. 107

(i) Metabolism ............................................................ 107(ii) Pathways of metabolism................................................. 108

Aldrin and dieldrin .......................................................... 109ti) Metabolism ............................................................ 109

(ii) Pathways of metabolism................................................. 110

Heptachlor ................................................................. 110

(i) Metabolism ............................................................ 110(ii) Pathways of metabolism................................................. 111

Mirex and Kepone........................................................... 111

Endosulfan .................................................................

Chlorobenzilate and chloropropylate ....... .................................... 112MIscellaneous organochlorine insect.ides......h................................. 112Enzymes Involved in Metabolism of Organochlorine Insecticies ..................... 113

EFFECTS ................................................. 114

Effects on Populations of Microons.114

Cytological and Biochemical Effects..... 118

Cell membrane................................................. 118

Amino acids and proteins................................................. 119

Nucleic acids................................................. 119

Enzymes and catabolic pathways .............................................. 119

Photosynthesis ................................................. 120

Cell morphology................................................. 121

CONCLUSIONS ................................................. 121

LITERATURE CITED.................................................. 122

INTRODUCTIONOrganochlorine insecticides (see Table 1 for

chemical names of organochlorine insecticidesand their metabolites) mainly include DDT,DDD, Kelthane, chlorobenzilate, chloropropy-late, methoxychlor, aldrin, dieldrin, heptachlor,lindane, endosulfan, isodrin, isobenzan, endrin,chlordane, toxaphene, mirex, and Kepone.These are synthetic chemicals, and some ofthem have been extensively used for controllingthe diseases of humans and domestic animalsthat are carried by insects and mites and alsoagainst insect pests which cause great damage to

95

agricultural crops. Due to their efficiency asinsecticides, these compounds were considereda boon to the fields of agriculture and medicalentomology. However, the use of organochlo-rine insecticides has been minimized or termi-nated in technologically advanced countries dueto their persistence in nature, susceptibility tobiomagnification, and toxicity to higher animals.But the ever-increasing world population andpoor health conditions, especially in developingcountries, may outweigh the disadvantagescaused by the extensive use of these insecti-cides.

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TABLE 1. Chemical names of the insecticides and their metabolitesCommon name Chemical name

Aldrin

BHBHCa-BHCP-BHCB-BHC

BPChlordaneChlordane expoxideChlordeneChlorobenzilateChloropropylateDBHDBPDDADDCNDDDo,p'-DDDDDEDDMDDMSDDMUDDNSDDNUDDOHDDTo,p'-DDTDicofolDieldnn

DPMEndosulfan

Endrin

HeptachlorHeptachlor epoxideIsobenzanKelthaneKepone

LindaneMethoxychlorMirexa-PCCH1-PCCHPCPAa-TCCH1-TCCH-y-TCCH&TCCHToxaphene

1,2,3,4,10,10-Hexachloro-1,4,4a,5,8,8a-hexahydro-1 ,4-endo,exo-5,8-dimethanonaphthalene

BenzhydrolA mixture of isomers of 1,2,3,4,5,6-hexachlorocyclohexanea isomer of 1,2,3,4,5,6-hexachlorocyclohexanep isomer of 1,2,3,4,5-6-hexachlorocyclohexane& isomer of 1,2,3,4,5,6-hexachlorocyclohexane y-BHC isomer of 1,2,3,4,5,6-hexachlorocyclohexane

Benzophenone1 ,2,4,5,6,7,8,8-Octachloro-3a,4,7,7a-tetrahydro-4,7-methanoindane4,5,6,7,8,8-Hexachloro-2,3-epoxy-3a,4,7,7a-tetrahydro-4,7-methanoindene4,5,6,7,8,8-Hexachloro-3a,4,7,7a-tetrahydro-4,7-methanoindeneEthyl 4,4'-dichlorobenzilateIsopropyl 4,4'-dichlorobenzilate4,4'-Dichlorobenzhydrol4,4'-DichlorobenzophenoneBis(p-chlorophenyl)acetic acidBis(p-chlorophenyl)acetonitrile2,2'-Bis(p-chlorophenyl)-1,1-dichloroethane2-(o-Chlorophenyl)-2-(p-chlorophenyl)-1,1-dichloroethane2,2'-Bis(p-chlorophenyl)-1,1-dichloroethyleneBis(p-chlorophenyl)methane2,2-Bis(p-chlorophenyl)-1-chloroethane1-Chloro-2,2-bis(p-chlorophenyl)ethylene2,2-Bis(p-chlorophenyl)ethane2,2'-Bis(p-chlorophenyl)ethylene2,2'-Bis(p-chlorophenyl)ethanol1,1 ,1-Trichloro-2-2-bis(p-chlorophenyl)ethane2-(o-Chlorophenyl)-2-(p-chlorophenyl)-1,1,1-trichloroethane2,2-Bis(p-chlorophenyl)-1,1,1-trichloroethanol1,2,3,4,10,10-Hexachloro-6,6-epoxy-1,4,4a,5,6,7,8,8a-octahydro-1,4-endo,exo-5,8-dimethanonaphthalene

Diphenyl methane6,7,8,9,10,10-Hexachloro-1,5,5a,6,9,9a-hexahydro-6,9-methano-2,2,4,3-

benzodiexathiepin-3-oxide1,2,3,4,10,10-Hexachloro-6,6-epoxy-1,4,4a,5,6,7,8,8a-octahydro-1,4-endo, endo-5,8-dimethanonaphthalene

1 ,4,5,6,7,8,8-Heptachloro-3a,5,7,7a-tetrahydro-4,7-methanoindene1 ,4,5,6,7,8,8-Heptachloro-2,2-epoxy-3a,4,7,7a-tetrahydro-4,7-endomethanoindene1,3,4,5,6,7,8,8-Octachloro-1,3,3a,4,7,7a-hexahydro-4,7-methanoisobenzofuran2,2'-Bis(p-chlorophenyl)-1,1,1-trichloroethanol1,la,3,3a,4,5,5a,6-decachlorooctahydro-1,3,4-methano-2H-cyclobuta(co)pentalene-2-one

-y isomer of 1,2,3,4,5,6-hexachlorocyclohexane1,1 ,1-Trichloro-2,2-bis(p-nethoxyphenyl)ethaneDodecachlorooctahydro-1,3,4-methan-2H-cyclobuta(cd)pentalenea-2,3,4,5,6-Pentachlorocyclohex-1-enep-2,3,4,5,6-Pentachlorocyclohex-1-enep-Chlorophenylacetic acida-3,4,5,6-Tetrachlorocyclohex-1-ene-3 ,3,5,6-Tetrachlorocyclohex-1-ene

-y-3,4,5,6-Tetrachlorocyclohex-1-ene8-3 ,4,5,6-Tetrachlorocyclohex-1-eneChlorinated camphene containing 67 to 69o chlorine

Organochlorine insecticides are more toxic toinsects and less toxic to nontarget organisms,but these chemicals can damage a wide varietyof beneficial as well as harmful organisms due totheir persistence in the environment. Therefore,organochlorine insecticides have important eco-

logical effects in addition to those usually intend-ed. Among these, the interaction of organochlo-rine insecticides with microorganisms isimportant, since microorganisms are involved inmany basic ecological processes, such as bio-geochemical cycles, decomposition processes,

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ORGANOCHLORINE INSECTICIDES AND MICROORGANISMS 97

energy transfer through trophic levels, and nu-merous microbe-microbe, microbe-plant, andmicrobe-animal interactions.The literature on accumulation and metabo-

lism of DDT in microorganisms has been re-viewed recently (51, 102, 192, 239). Informa-tion on accumulation and metabolism of otherorganochlorine insecticides is rather scanty.Further, certain important aspects, such asmechanisms of accumulation, pathways of me-tabolism, and the role of enzymes in metabolismof organochlorine insecticides, need further clar-ification and elucidation.The effects of organochlorine insecticides on

populations of microorganisms have receivedconsiderable attention (33, 192, 239, 249). Thecytological and biochemical effects of these in-secticides are important in order to understandtheir mode of action. Thus, the compilation ofdata pertaining to cytological and biochemicaleffects of organochlorine insecticides is needed.The present review aims to bring together theavailable information on the accumulation, me-tabolism, and effects of the organochlorine in-secticides in relation to microorganisms.

ENTRY OF ORGANOCHLORINEINSECTICIIDES INTO MICROBIAL

ENVIRONMENTSGenerally, organochlorine insecticides are ap-

plied to soil and aquatic environments to kill thetarget organisms. A small amount of these insec-ticides remains in the air during application ordue to volatilization of these compounds fromthe soil surfaces. However, it is the soil andaquatic environments which are commonly stud-ied in relation to the effects of insecticide usageon microorganisms.The organochlorine insecticides in the soil

originate from direct application to soil andrunofffrom leaves and stems of plants on whichthey are sprayed (49, 94). The insecticides whichare taken up by the target organisms and by theother nontarget biota also reach the soil aftertheir death and decay. Smaller quantities ofinsecticides may enter the soil via the excreta ofanimals which have accumulated the insecti-cides either due to their feeding habits or bysome other means. Occasionally, disposal ofinsecticides and their containers may producelocalized regions of high insecticide concentra-tion in the soil.

Organochlorine insecticides enter the aquaticenvironment by intentional application for thepurpose of controlling insects. The variousmodes of entry of organochlorine insecticidesinto water environments are through spraying ofinsecticides and disposal of unused insecticidesand insecticide containers containing excessiveresidues. Drift from aerial or ground applica-

tions of insecticides and movement via air andwater and by soil erosion are often the majorindirect sources of entry of insecticides (48, 59,239). Air drift of insecticides is not restricted toareas adjacent to the site of application, sincedust particles carrying insecticides may be trans-ported in the atmosphere to great distances fromthe site of application (45, 52).

Organochlorine insecticides have relativelylow solubility in water. Therefore, the amountreaching the aquatic environment via runoff isgenerally low. However, exceptionally high ac-cumulation of insecticides may result from acci-dental spillage or careless disposal of surplusamounts in streams or rivers.

ACCUMULATION

Accumulation of Various OrganochlorineInsecticides in Microorganisms

Accumulation can be defined as the transfer ofan insecticide from the environment into anorganism. Many terms, such as bioconcentra-tion, biomagnification, and ecological magnifi-cation have been used to express the concept ofaccumulation (14). Nevertheless, discussion ofaccumulation of organochlorine insecticides inmicroorganisms is being restricted here to onlytwo terms, bioconcentration and biomagnifica-tion. Kneip and Laur (118) while stressing theneed for clear distinction between these twoterms, defined them as follows:

Bioconcentration is the ability of an organism toconcentrate a substance from the aquaticenvironment. This has been quantitatively ex-pressed in terms ofpercent accumulation andthe concentration factor. Percent accumula-tion is the amount of substance accumulatedas a percentage of the amount initially addedto the medium. The concentration factor isthe ratio of the concentration of substance inbiological materials expressed in parts permillion (dry weight) to its concentration inwater in parts per million.

Biomagnification is the occurrence of a sub-stance at increasingly higher concentrationswith increasing trophic levels in the foodchain.The role of microorganisms in the accumula-

tion of organochlorine insecticides is not fullyunderstood. Some microorganisms have beenreported to accumulate very high concentrationsof insecticides (Table 2). For instance, threespecies each of fungi, streptomycetes, and bac-teria differed markedly in ability to bioconcen-trate DDT and dieldrin from distilled water (39).After 4 h of incubation, the fungi accumulated 60to 83% of the DDT and 75% of the dieldrin.

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Insecticide

DDT

-y-BHC

Aldrin

Dieldrin

TABLE 2. Accumulation of organochlorine insecticides by microorganismsInsecticide in % removed from medium or

Microorganism medium concn factor

Agrobacterium tumefaciens 0.1-1 ppm 100%OStreptomycetes 0.1-1 ppm 60-83%Rhodotorula gracilis 2 ppm 97%Torulopsis utilis 2 ppm 94%Dunaliella tertiolecta 80 ppb 97.8%Cyclotella nana 80 ppb 96.6%Thalassiosira fluviatilis 80 ppb 94.4%Amphidinium carteri 80 ppb 93.2%Rumen protozoans 40 ppm 95%Stylonychia notophora 1 ppm 0.5%Tetrahymena pyriformis 1 ppm 0.5%Blepharisma intermedium 1 ppm 90%O

Aerobacter aerogenes 1 ppb 1,140-3,400Bacillus subtilis 1 ppb 1,140-3,400Microcystis sp. 1,000 ppb 230Anabaena sp. 1,000 ppb 268Scenedesmus sp. 1,000 ppb 134Oedogonium sp. 1,000 ppb 207Amphidinium carteri Ambient 80,000Syracosphaera sp. Ambient 25,000Anacystis nidulans 1 ppm 849Scenedesmus obliquus 1 ppm 626Euglena gracilis 1 ppm 99Skeletonema costatum 0.7 ppb 38,400Cyclotella nana 0.7 ppb 58,100Amphidinium carteri 0.7 ppb 9,600Tetraselmis chuii 0.7 ppb 6,300Ankistrodesmus amalloides 0.72 ppb 616Cylindrotheca closterium 100 ppb 190Paramecium bursaria 1 ppm 264Paramecium multimicronucleatum 1 ppm 964Stylonychia notophora 1 ppm 295Tetrahymena pyriformis 1 PPm 224

Achromobacter delicatusBacillus megateriumBacillus subtilisCaulobacter vibrioidesChromobacterium violaceumEnterobacter aerogenesEscherichia coliChlorella sp.

Zalerion maritimum

StreptomycetesAgrobacteriumBacillus sp.Serratia sp.Cyclotella sp.Microcystis sp.Anabaena sp.Scenedesmus sp.Oedogonium sp.Ankistrodesmus amalloidesZalerion maritimum

Chlordane Scenedesmus quadricaudaCaulobacter vibrioides

Methoxychlor Aerobacter aerogenesBacillus subtilis

rpr

0.1 ppm0.1 ppm0.1 ppm0.1 ppm0.1 ppm0.1 ppm0.1 PPm10.6 ppb

10-100 ppm

0.1-1 ppm0.1-1 ppm0.1-1 ppm0.1-1 ppm

0.1 ppb1 ppm1 ppm1 ppm1 ppm0.72 ppb

10-100 ppm

0.1-100 ppb0.1 ppb

1 ppb1 ppb

20070100300701020

2,301-6,561

2,000

83%90%o16%16%

80-90%o130-270130-270130-270130-270

3202,000

6,000-15,00055,900

1,140-3,4001,140-3,400

Reference

3939232326262626127133132Lal, Ph.D.

thesis1041042332332332335151828282

2022022022021781138282133133

8383838383832087

211

39393939

246233233233233178211

7883

104104

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Agrobacterium tumefaciens accumulated 100%of the DDT and 90% of the dieldrin (the resultsfrom other microorganisms were inconclusive,probably due to losses through cell washing).Yeast cells have also been reported to accumu-late high concentrations ofDDT, with accumula-tion ranging from 94 to 97% (23). A ciliateprotozoan, Blepharisma intermedium, accumu-lated more than 90To of the DDT from mediuminitially containing 1 ppm (R. Lal, Ph.D. thesis,University of Delhi, Delhi, India, 1980); thisorganism was also found to retain high DDTconcentrations subsequently, presumably due tolow rate of metabolism and excretion of theinsecticide.

Rice and Sikka (202) used different species ofalgae to study the differential response of thesespecies to insecticide accumulation. Althoughall species tested accumulated DDT rapidly fromvarious media initially containing 1 ppb, thediatoms Skeletonema costatum and Cyclotellanana accumulated DDT to the highest level. Theaccumulation of DDT was linearly related to itsconcentration in the medium. However, an in-crease in cell density decreased the capacity ofindividual cells to accumulate DDT, and DDTconcentration was also inversely related to cellsize. Rice and Sikka (203) also measured dieldrinuptake in different algae, using a constant diel-drin concentration. The uptake of dieldrinshowed a linear relationship between dieldrinaccumulation and cell density in Amphidinium,but the uptake was not linear in Skeletonema,Tetraselmis, Cyclotella, Isochrysis, and Olistho-discus.A blue-green alga (cyanobacterium) isolated

from a stream, Synechococcus elongatus, wasfound to contain DDT (252). The culture of thisspecies was raised under laboratory conditionsand exposed to 99 ppb DDT. The organismsrapidly removed a high percentage of DDT fromthe medium, and the accumulated levels weresubsequently retained. Ware et al. (238) alsofound that the alga Cladophora accumulatedDDT and retained it without any apparent excre-tion; they suggested that this organism couldserve better than Oscillatoria as an indicator ofDDT contamination in surface irrigation water.Without any apparent adverse effect, axenic

cultures of three ciliate protozoans, Stylonychianotophora (133), Tetrahymena pyriformis (132),and B. intermedium (Lal, Ph.D. thesis), accumu-lated DDT from medium initially containing 1ppm. S. notophora and T. pyriformis accumulat-ed about 0.5% of the DDT, whereas in B.intermedium, DDT accumulation continued un-til the end of the experiment, when the organismhad accumulated 90%o of the DDT. Anotherflagellate protozoan, Euglena gracilis, also ac-cumulated DDT without eliminating it (56). As

pointed out earlier, such species can be used asindicators of DDT contamination in water.The second direct method for expressing the

bioconcentration of organochlorine insecticidesis the use of a concentration factor. Concentra-tion factors of different insecticides in microor-ganisms have been listed in Table 2. It is evidentthat concentration factors vary from species tospecies and are extremely high in certain cases.Since aquatic microorganisms in freshwater andmarine environments serve as important nutri-ent sources for a broad spectrunm of aquaticfilter-feeding organisms, the accumulation oforganochlorine insecticides in microorganismsat the primary trophic level and transfer oforganochlorine insecticides through successivetrophic levels can constitute a hazardous link inthe food chain to fish and higher vertebrates.The direct evidence for such a biomagnificationof insecticide was provided by Butler (34). Heobserved that DDT and its metabolites appliedto estuarine waters were absorbed by planktonalmost immediately. The biomagnification ofresidues in the food web progressed from anestimated 1.0 ppb DDT and related metabolitesin water to 70 ppb in plankton to 15 ppm in fishand up to 800 ppm in porpoise blubber.The widespread distribution of organochlorine

insecticides has become a classic example ofbioconcentration and biomagnification. Al-though the most suitable way to study the bio-concentration potential of a compound shouldbe by direct exposure of organisms to the partic-ular compound, the experimental approach forsuch studies still lacks international standardiza-tion. From the environmental point of view, thisbecomes mote important when the acute toxici-ty of a compound is low and the physiologicaleffects go unnoticed until the chronic effectsbecome evident. It is for these reasons that theuse of an indirect method which involves themeasurement of the partition coefficient of thecompound under consideration has been sug-gested.The partition coefficient is the ratio of the

equilibrium concentrations of a chemical in anonpolar solvent and a polar solvent. The parti-tion coefficients of chemicals derived from asystem of nonpolar (octanol) and polar (water)solvents have been reported, and in a number ofcases correlations between partition coefficientsand bioconcentration factors have been estab-lished (161). Traditionally, the partition coeffi-cient of an organic compound between octanoland water phases is calculated by adding thecompound to a system containing octanol andwater and determining the distribution of thecompound between octanol and water (235).However, several workers have suggested theuse of more sophisticated systems, such as

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liquid chromatography and thin-layer chroma-tography, for this purpose (37, 166). Recently,Renberg and Sundstrom (201) have reported theuse of reversed-phase thin-layer chromatogra-phy for the prediction of bioconcentration po-tential. It has also been shown, by Neely et al.(176), that log partition coefficient correlateswell with log bioconcentration factor for a vari-ety of diverse organic chemicals.Nonpolar compounds, such as organochlorine

insecticides, are accumulated in an organismaccording to their partitioning between the medi-um and the lipid contents ofthe cell (51). Kenaga(114) also suggested that such compounds parti-tion in octanol and water in a ratio similar to thepartition coefficients between the lipid of aquaticorganisms and the medium containing the organ-isms. Recently, an attempt has been made tovalidate such a system by using the ciliate proto-zoan T. pyriformis (132). Concentration factorsin this ciliate were in the following order: p,p'-DDD > p,p'-DDE > o,p'-DDT > DDMU >p,p'-DDT. The values of partition coefficients inoctanol-water for these compounds also ap-peared in the same order, but such a relationshipcould not be established during hexane-waterpartitioning. Thus, the octanol-water partition-ing system can perhaps be used for determiningthe relative bioconcentration potentials of or-ganochlorine insecticides. However, more workis needed on the kinetics of partitioning oforganochlorine compounds in different systems,using standard methods, before any valid con-clusion can be drawn.

Mechanism of AccumulationWhereas most studies on the accumulation of

insecticide have been designed to show howmuch of the insecticide is accumulated by themicroorganism, they have said little about themechanism of accumulation. In general, the rateof accumulation of organochlorine insecticidesin microorganisms is very rapid, and in certaincases it takes only a few seconds to accumulatean organochlorine insecticide to the highest con-centration. However, the time required to accu-mulate to maximum concentration differs fromorganism to organism, and among the microor-ganisms bacteria show an extraordinary abilityto rapidly accumulate organochlorine insecti-cides. For instance, Aerobacter aerogenes andBacillus subtilis required only 30 s to accumulate80 to 90% of the DDT residue accumulated in 24h (39), whereas the algae Tetraselmis chuii andAnkistrodesmus amalloides required 1 to 3 h toaccumulate DDT to maximum concentrations(178, 202). However, in the alga Chlorella pyren-oidosa (216) and the yeast Torulopsis utilis (23),accumulation of DDT was very rapid: these

species required only 15 s and 3 min, respective-ly, to accumulate DDT to maximum concentra-tions. Such differences are also not uncommonamong yeasts. Saccharomyces cerevisiae re-quired 30 s (236) and Rhodotorula gracilis re-quired 3 min (23) to accumulate DDT to maxi-mum concentrations. Protozoans are relativelyless efficient than bacteria and algae in thisrespect. The ciliate protozoan S. notophora re-quired 1 h (133), T. pyriformis required 4 h (132),and Crithidia fasciculata required 6 h (J. E.French, Diss. Abstr. Int. B 37:96-97, 1976) toaccumulate DDT to maximum concentrations.Other organochlorine insecticides are accu-

mulated less readily than DDT. In C. pyrenoi-dosa, y-BHC was accumulated less readily thanDDT (217). The rate of accumulation of dieldrinwas still slower in C. pyrenoidosa (216) andScenedesmus obliquus (200), and maximumamounts were accumulated between 6 and 36 h.However, A. amalloides and Nitzschia sp. accu-mulated dieldrin more readily: equilibrium wasestablished between 30 and 60 s (35, 178, 246).Flavobacterium harrisonii, B. subtilis, and C.pyrenoidosa accumulated methoxychlor rapidly,reaching equilibrium within 30 min, whereasAspergillus sp. accumulated methoxychlorslowly and required 16 h (185). Aspergillus sp.and Aeromonas proteolytica (Aeromonas hydro-phila subsp. proteolytica) also required longerperiods-24 and 48 h, respectively-to accumu-late chlordane to maximum concentrations (J. P.Nakas, Diss. Abstr. Int. B 38:77, 1977). Toxa-phene, like DDT, was accumulated rapidly in F.harrisonii, B. subtilis, and C. pyrenoidosa (186).Hansen (87) studied the mechanism of accu-

mulation of-y-BHC in Chlorella. He showed thatthe rate of uptake of y-BHC per cell in Chlorellasp. was very high on day 1, declined until day 3,and increased again until day 6. The initialaccumulation of y-BHC was assumed to be dueto adsorption. The rate of adsorption was rapidin the beginning, for the whole culture had manyyoung cells, which had more capacity to adsorbthe insecticide. The increase of accumulation inthe sublethal range from day 3 to day 6 wasattributed to change in the metabolic activity ofthe cells.

In addition to the physiological state of micro-organisms, density and size of the microorga-nisms also play an important role. Rice andSikka (202) showed that accumulation of DDTwas inversely proportional to the number ofcells in the medium. They also emphasized thatthe rapid accumulation of organochlorine insec-ticides in microorganisms is mainly due to ad-sorption and that the extent of accumulation ofan insecticide is decided by the surface area ofthe organisms. The amount of surface per unitmass or volume increases as the size of the

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particle decreases. Consequently, if adsorptionis a significant process in accumulation, smallparticles or organisms might be expected toaccumulate higher amounts of insecticides.There is no relevant report on organochlorineinsecticides to support this hypothesis, but men-tion may be made of a report on the distributionof polychlorinated biphenyls in Chesapeake Bay(173). The suspended sediment (particle size,<0.5 ,um) contained 0.92 ppm polychlorinatedbiphenyls on a dry weight basis, and bottomsediment (particle size, <100 ,um) contained0.28 ppm.The possibility that organochlorine insecti-

cides are incorporated into microorganisms byactive or passive transport also must be consid-ered. In various studies it was found that notonly live microbial cells but also autoclaved cellsshow similar uptakes of organochlorine insecti-cides (35, 80, 186, 203), which appears to indi-cate that an actual metabolic factor is not in-volved in the accumulation process. Forinstance, Johnson and Kennedy (104) found thatthe accumulation of DDT and methoxychlor byautoclaved cells of A. aerogenes was greaterthan that of living cells of the bacterium. Theysuggested that molecular polarity and lipid solu-bility influence the accumulation of insecticidein A. aerogenes cells. A similar conclusion wasdrawn in studies with A. aerogenes, B. subtilis,and S. cerevisiae, where the accumulation ca-pacities of the organisms increased after boiling(106, 239). The alga C. nana is an exceptionalcase: dead cells adsorbed less dieldrin than didliving cells (246).

Liyr and Ritter (141) examined the penetrationof H-BHC isomers into S. cerevisiae andshowed that the rate of uptake increased in theorder <<y < a, which was different from theirorder of solubility in organic solvents and alsofrom their order of toxicity to S. cerevisiae, thisbeing 8 < y < I < a. When the cells in culturewere exposed to 30 M concentrations of individ-ual isomers in the medium, the intracellularconcentrations of the a, p, fy, and 8 isomersreached constant levels corresponding to up-takes of 60, 18, 40, and 24%, respectively, of thecompound in the medium. Thus the most bioac-tive isomer (8) was taken up marginally morereadily than the slightly active p isomer, where-as the least toxic isomer (a) was absorbed morereadily. For this reason, the stereochemical dif-ferences between isomers were considered tohave overriding influence on BHC activity.

It is evident from the literature that for someorganochlorine insecticides, e.g., DDT, consid-erable information has been obtained regardingtheir accumulation in microorganisms. Howev-er, little information is available on the extent towhich accumulation is influenced by other fac-

tors, e.g., the physical and chemical characteris-tics of the insecticide, environmental conditions,and the nature of the organism. This makes itdifficult to draw any conclusion of ecologicalsignificance. Since such studies have been con-ducted under different experimental conditions,no comparison can be made. It is understand-able that many environmental factors whichinfluence the accumulation cannot be consid-ered together for obvious reasons of practicaldifficulty, but attempts should be made to con-sider some of the major factors, such as pH,temperature, limited nutrients, competition, etc.

It is also clear from the foregoing account thatalthough there have been many reports on theaccumulation of organochlorine insecticides inmicroorganisms, we know little about the mech-anisms of accumulation. Investigation on thetransfer of insecticide through cell membranes,the distribution of insecticides in the cell, andthe structural differences among insecticidesthat influence accumulation should be carriedout for better understanding of the mechanism ofaccumulation.

METABOLISMMicroorganisms play an important role in the

metabolism of organochlorine insecticides.However, the persistence of a number of or-ganochlorine insecticides in soil and water forvery long periods has been reported. This maybe due either to the resistance of the insecticideto microbial degradation or to the formation of acomplex with some component of the environ-ment which is largely resistant to microbialattack (3). This section deals with the metabolismof individual insecticides (see Table 3) and thepathways of metabolism of insecticides in micro-organisms. Because microorganisms degrade or-ganochlorine insecticides by reductive dechlori-nation, dehydrochlorination, oxidation, andisomerization of the parent molecule, these reac-tions are discussed briefly.

Major Reactions of Organochlorine InsecticideMetabolism

Reductive dechlorination. Reductive dechlori-nation of organochlorine insecticides is an im-portant microbial reaction. The reaction pro-ceeds by replacing a chlorine atom on anonaromatic carbon with a hydrogen atom. Con-version of DDT to DDD is a classic example ofreductive dechlorination and has been shown tooccur in yeast (109), Proteus vulgaris (10), Ser-ratia marcescens (219); soil actinomycetes, in-cluding Nocardia erythropolis and five speciesof Streptomyces (40); A. aerogenes (158); plant-pathogenic and saprophytic bacteria (103); andin soil samples under anaerobic conditions (84).

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TABLE 3. Metabolism of organochlorine insecticides in microorganismsInsecticide Microorganism Metabolite(s) Reference(s)

Bacteria from ruminantsMicroorganisms from intestines of

ffies and mammalsMicroorganisms from stable fly

excretaMicroorganisms isolated from

cheeseNocardia sp.Streptomyces aureofaciensStreptomyces annamoneusStreptomyces viridochromogenesKlebsiella pneumoneaeAerobacter aerogenesEscherichia coli

Pseudomonas sp.Bacillus sp.Micrococcus sp.Arthrobacter sp.Hydrogenomonas

Microorganisms from sewage sludge

Cylindrotheca closteriumSkeletonema costatumCyclotella nanaIsochrysis galbanaAmphidinium carteriThalassiosira fluviatilisDunaliella sp.Nitzschia sp.Saccharomyces cerevisiaeTrichoderma virideMucor alternans

Acanthamoeba castellanjiStylonychia notophoraBlepharisma intermediumTetrahymena pyriformis

'y-BHC Bacillus coliClostridium sporogenesClostridium sp.Clostridium rectumClostridium sphenoidesPseudomonas putida

Escherichia coliCitrobacter freundiiMixed microbial culture

Chlorella vulgarisChlamydomonas reinhardii

Pseudomonas sp.Micrococcus sp.Bacillus sp.Unicellular algaeDunaliella sp.Aspergillus nigerAspergillus flavusPenicillium notatumPenicillium chrysogenum

DDDDDD

DDD, DDE

DDE

DDDDDDDDDDDDDDDDDDDDD, DDMU, DDMS,DDNU, DDOH, DDA, DBH

DDDDDDDDDDDDDDD, DDE, DDMS, DDMU,DBH, DDM, DDA, DPM,PCPA

DDCNDDCN, DDDDDEDDEDDEDDEDDEDDEDDD, DDE, DDOH, DDNSDDEDDDDDD, DDE, DDNS, DDASolvent-soluble and water-

soluble metabolitesDDE, DDD, DBPDDE, Kelthane, DDMUo,p'-DDT, DDEDDMU, DDE, o,p'-DDT

Benzene, monochlorobenzeneBenzene, monochlorobenzene,y-TCCHy-TCCHy-TCCH-y-PCCH, y-TCCH, CO2ly-TCCH, a-BHCy-PCCH, a-BHCy-PCCH, y-TCCH-y-PCCH; a-, ,B-, and &BHC;

ly-TCCHy-PCCHy-PCCH

trans-Aldrindioltrans-Aldrindioltrans-AldrindiolDieldrinDieldrinDieldrinDieldrinDieldrinDieldrin

127, 16510, 30, 159; Feil et

al.a219

136

40404040241241134, 135

1881881881885, 66, 67, 191, 213,

214

210111426, 2022022022022626, 189168109153, 157, 1887

194133Lal, Ph.D. thesis132

66143, 14485, 98, 179, 1809213, 691502379861

222222

188188188115189123123123123

DDT

Aldrin

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TABLE 3-ContinuedInsecticide Microorganism Metabolite(s) Reference(s)

Trichoderma viride trans-Aldrindiol, photodieldrin 188, 223

Dieldrin Pseudomonas sp. trans-Aldrindiol 24, 149Bacillus sp. trans-Aldrindiol 152Aerobacter aerogenes trans-Aldrindiol 153Pseudomonas marginalis Photodieldrin, trans-aldrindiol 234Pseudomonas fluorescens Photodieldrin, trans-aldrindiol 234Pseudomonas syringae Photodieldrin, trans-aldrindiol 234Pseudomonas morsporunorum Photodieldrin, trans-aldrindiol 234Pseudomonas glycinea Photodieldrin, trans-aldrindiol 234Pseudomonas sp. CO2 97Arthrobacter sp. CO2 97Bacillus sp. CO2 97Mycobacterium CO2 97Mycococcus CO2 97Nocardia sp. CO2 97Dunaliella Photodieldrin, trans-aldrindiol 189Neurospora sp. Photodieldrin, trans-aldrindiol 234Trichoderma koningii CO2 16Soil microorganisms trans-Aldrindiol, photodieldrin 188, 223

Heptachlor Soil microorganisms Heptachlor epoxide, 1- 38hydroxy-2,3-epoxychlordene

Soil microorganisms Heptachlor epoxide, 163chlordene, chlordeneepoxide, 1-hydroxychlordene, 1-hydroxy-2,3-epoxychlordene

Endrin Pseudomonas sp. Ketoendrin, ketone, aldehyde 155, 188Micrococcus sp. Ketoendrin 188Arthrobacter Ketoendrin 188Trichoderma viride Ketoendrin 188

a V. J. Feil, E. J. Thacker, R. G. Zaylskie, C. H. Lamoureure, and E. Styrvoky, Abstr. Meet. Am. Chem.Soc. Div. Pestic. Chem. 162nd, abstr. no. 47, 1972.

The strains of Hydrogenomonas which wereused to degrade DDT analogs under anaerobicconditions also exhibited the ability to dechlori-nate DDT reductively to DDM (67).

dechlorinat ionc/

DDT DDD

Degradation of the insecticide y-BHC to ly-TCCH in Clostridium rectum (179) and Pseudo-monas putida (69, 151) also proceeds by reduc-tive dechlorination. Reductive dechlorination ofheptachlor in bacteria and actinomycetes resultsin the formation of chlordene (163). Matsumuraet al. (154) have reported another type of reduc-tive reaction, which involves the reversal ofepoxidation from dieldrin to aldrin.

Dehydrochlorination. Dehydrochlorination in-volves the simultaneous removal of hydrogenand chlorine from organochlorine insecticides.Typically, the reaction takes place between thesaturated chlorinated carbon and the adjacenthydrogen on the neighboring carbon. The forma-

tion of DDE from DDT and the formation of -y-PCCH from y-BHC are the most familiar exam-ples of this reaction.

CQ~ O v dehydrochlonnation- to~~~~~~~~~~n-Cd2C-DDT DDE

Oxidation. The oxidative reactions which areimportant in degrading organochlorine insecti-cides in higher organisms are less common inmicroorganisms, probably due to the lack of adefined mixed-function oxidase system in micro-organisms (149). The oxidation of aldrin to itsepoxide, dieldrin, in microorganisms was report-ed by Korte et al. (123). Other examples ofoxidation reactions are the formation of hepta-chlor epoxide from heptachlor (138) and theformation of Kelthane from DDT (153).

ci c ~ L- cc- \Vj -ci+oC=O

OH DBP

4,4'-dichlorobenzilic acid

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104 LAL AND SAXENA

O, P'- DOT

Cl I

}cloIoCHCI2

0, ;- DOD

'$2~'-HC3lCC12

tCl / ODDT ODE

uMetblte reductive pathway(dechlornotion)

CHCI2 CHCI

DDD oDMU

CH2CI4 DOMS

CH24 DDNU

CH3I DONSH

~ ~~O~H t O

SHD

H H t

I~~~~~~~~~

H OH 0

0DD OSH DSP1 ring openingzC 3 COO U<CH_HH OH

PCPA I(p-chloropheni) ethanol

cl- J-C-CHOOH

FIG. 1. Proposed pathways of metabolism of DDT in microorganisms.

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The opening of aromatic ring of DDT in Hy-drogenomonas sp. (67) and of -y-BHC in P.putida (13) also involves oxidation. Decarboxyl-ation reactions during the oxidation of 4,4'-dichlorobenzilic acid, a hydrolysis product ofboth chlorobenzilate and chloropropylate, toDBP have also been reported to occur in theyeast R. gracilis (167).

Isomerization. Isomerization reactions inmicroorganisms during the metabolism of or-ganochlorine insecticides are the conversions ofy-BHC to a-BHC (13), dieldrin to photodieldrin(149, 155), and 8-ketoendrin to endrin (158). Ineach of these cases the isomer differs from theparent compound due to the difference in theposition of the chlorine atom in the benzenering.

cl ci cl cl y-isomerization

CI y-BHC ca-BHCMetabolism of Individual Insecticides and

Pathways of Metabolism1,1,1-Trichloro-2-2-bis(p-chlorophenyl)ethane.

(i) Metabolism. DDT is one of the most exten-sively studied insecticides. The primary meta-bolic mechanism is the reductive dechlorinationofDDT to DDD (Fig. 1). A number of studies onDDT metabolism in microorganisms have shownthat under anaerobic conditions reductive de-chlorination ofDDT predominates, whereas un-der aerobic conditions dehydrochlorination isthe dominant reaction. Indeed, the rapid degra-dation of DDT residues under anaerobic con-ditions has prompted many investigators toexamine flooding as one of the means for decon-taminating soil with high DDT residues (84,112).A. aerogenes was incubated with DDT in

parallel experiments conducted aerobically andanaerobically (158, 242). In these experiments itwas shown that DDT was degraded to DDDunder both conditions, but anaerobic conditionsfavored higher yields of DDD. However, John-son et al. (103), using pure cultures of microor-ganisms, found that no species, including A.aerogenes, produced DDD under aerobic condi-tions, but most produced DDD under anaerobicconditions. Langlois (134) studied the metabo-lism of DDT by Escherichia coli grown in vari-ous broths or skim milk and reported over 50%dechlorination of DDT to DDD in the brothsafter 2 days and 90%o conversion by 7 days.Negligible dechlorination occurred in skim milk.Later, Langlois et al. (135) concluded that caseinin milk forms a complex with DDT which isresistant to bacterial degradation. They studiedseven bacteria, incubated in tryptic soy brothunder both aerobic and anaerobic conditions, for

DDT-degrading ability. Aerobically, Pseudomo-nas fluorescens and Staphylococcus aureus didnot degrade DDT, whereas Bacillus cereus, Ba-cillus coagulans, and B. subtilis degraded DDT.For E. coli and Enterobacter aerogenes, DDDwas the major or the only product of aerobicdegradation, but trace amounts of DDMU,DDMS, DDNU, DDOH, DDA, and DBP weredetected under anaerobic conditions. However,the three bacilli produced trace levels of thesame metabolites under aerobic conditions. Inthe above experiments aerobic incubation mighthave become anaerobic during the period ofexperimentation, since the liquid media were inscrew cap flasks.

Fries (71) reviewed the literature on DDTmetabolism in microorganisms under aerobicand anaerobic conditions. He reported that in-vestigators (103, 158) who had shaken theirculture flasks containing microorganisms to en-sure aerobic conditions did not report DDDformation, whereas those (103, 116, 153, 159,188) who reported aerobic production of DDDapparently did not shake their cultures. Thelatter experiments may have involved a periodof unintentional anaerobic degradation, whichcould account for the production of much of thereported product. Zoro et al. (256) examinedthese aspects carefully, and, working withmixed cultures of microorganisms, observedthat DDT was degraded to DDD rapidly underanaerobic conditions. They further suggestedthat under anaerobic conditions, DDT forms acomplex with reduced iron-porphyrin which ismore susceptible to degradation, whereas such acomplex is not formed under aerobic conditions.A species of Hydrogenomonas isolated from

sewage sludge by Focht and Alexander (67) hasbeen extensively studied for its ability to de-grade DDT and its analogs (5, 66, 67, 191). Themetabolites of DDT produced by this bacteriumare DDD, DDMS, DDMU, DBH, DDM, andDDA. This bacterium, though unable to utilizeDDT as a sole carbon source, has been reportedto cleave one of the rings of DDM, a product ofDDT metabolism, to yield p-chlorophenylace-tate under aerobic conditions (67). Deo andAlexander (57) isolated a strain of Arthrobacterfrom lake water by using an inorganic salt solu-tion as an enrichment medium to study themetabolism of p-chlorophenylacetate earlier ob-tained during DDM metabolism in Hydrogeno-monas (67). The p-chlorophenylacetate was con-verted to two products by resting cells of theArthrobacter strain. One of the products was 4-chloro-3-hydroxyphenylacetate.

Synergistic degradation of DDM, a metaboliteof DDT, by Hydrogenomonas and Fusarium sp.was reported by Focht (65). Hydrogenomonassp., capable of metabolizing DDM to p-chloro-

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106 LAL AND SAXENA

phenylacetate, was unable to metabolize DDMto CO2 and H20 when used alone. However,when used in combination with Fusarium sp., itmetabolized DDM to H20 and CO2. This indi-cates that pure cultures of microorganismswhich have been shown to be incapable ofmetabolizing insecticides may metabolize themin combination with other organisms. It will beof special interest to know the biochemicalevents which occur during such interactions.A new compound, DDCN, has been reported

as a main product of the degradation of DDT inanaerobic sewage sludge (2, 101). In the experi-ments conducted by Albone et al. (2), radiola-beled DDT and nonlabeled DDT at 37°C wereincubated in a vessel containing anaerobic sew-age sludge. Only DDCN was produced fromDDT. Jensen et al. (101) incubated DDT withsewage sludge enriched with nitrogen at 20°C for8 days; in this case it was noticed that, inaddition to DDCN, a small amount of DDD wasalso present. In both the cases microbial degra-dation was implicated.

Fungi also seem to play a significant role inthe degradation of DDT. In this group of micro-organisms DDD is the major metabolic productof DDT. Matsumura and Boush (153) observedthat DDT is extensively metabolized by differentstrains of Trichoderma viride isolated from soil.Out of 18 isolates tested for DDT metabolism, 8produced DDD and dicofol, 3 produced DDDonly, and 1 produced DDE. Later, Matsumuraet al. (157) identified a "dicofol-like" compoundin place of dicofol which was subsequently con-firmed to be DDNS by Matsumura et al. (157).In addition, two strains were also able to metab-olize DDT to DDD, DDA, and DDNS (188). Inshake cultures with Mucor alternans, [14C]DDTwas partially metabolized into a hexane-solubleand two water-soluble metabolites (7). Thesemetabolites were not identified when examinedby thin-layer chromatography and autoradiogra-phy, and none of the compounds had Rf valuesidentical to those ofDDD, DDE, DDMU, DDA,DBP, dicofol, and DDNS.

Algae, as compared with bacteria, are lessefficient in the metabolism of DDT. The mostcommon product of DDT metabolism in algae isDDE (in contrast with DDD, which is the majormetabolic product of DDT in bacteria). Anotherdifference is the low ability of algae to metabo-lize DDT, for the amounts of DDE producedfrom DDT ranged from 3 to 10% (26). In additionto DDE, trace levels of polar metabolites in S.costatum (202) and DDD in A. amalloides (178)have also been reported. In another study, Patilet al. (189) reported DDD, DDE, and DDNS asthe metabolites of DDT in Dunaliella sp.Information on the metabolism of DDT in

protozoans is not sufficient to make any state-

ment regarding the role played by them in thedegradation of DDT. Rumen protozoans con-verted DDT to DDD and DDE (127). The soilamoeba Acanthamoeba castellanii metabolizedDDT to DDE, DDD, and DBP (194). Freshwaterciliates showed significant differences in theirability to metabolize DDT. S. notophora (133)and T. pyriformis (132) were quite active inmetabolizing DDT, and the amounts of metabo-lites were always higher than the amount ofDDT accumulated in the organisms. S. noto-phora metabolized DDT to Kelthane, DDMU,and p,p'-DDE, whereas DDT was converted toDDMU, DDE, and o,p'-DDT in T. pyriformis.In contrast, B. intermedium accumulated veryhigh amounts ofDDT and metabolized it to tracelevels ofDDE and o,p'-DDT (Lal, Ph.D. thesis).It is interesting to note here that DDD, which isa common metabolic product ofDDT in most ofthe microorganisms, was not detected in thesefreshwater ciliates.Commercial DDT generally contains about

15% o,p'-DDT, which is converted biologicallyto corresponding o,p'-DDD (158) and o,p'-DDE(90). Conversion of o,p'-DDT to p,p'-DDT inrats has been reported (117). However, A. aero-genes was unable to convert o,p'-DDT to p,p'-DDT under aerobic and anaerobic conditions(158). Rumen microorganisms are known to con-vert o,p'-DDT to o,p'-DDD (72).

(ii) Pathways of metabolism. The pathways ofDDT metabolism in microorganisms are notfully understood. The major step ofDDT metab-olism in microorganisms is reductive dechlorina-tion of DDT to DDD. Engst and Kujawa (62)reported that DDD was formed from DDTthrough DDE in Fusarium oxysporum, but laterstudies by Plimmer et al. (193) revealed thatDDD is directly formed from DDT and that noother intermediate is involved. DDD is furtherdegraded through dechlorination, dehydrochlo-rination, and decarboxylation to DBP or to amore reduced form, DDM. Ring cleavage ofDDM can occur to form PCPA, which in turn isshown to degrade to p-chloroglycoaldehyde. Asecond pathway of DDT metabolism involvesdehydrochlorination of DDT to DDE. Forma-tion of Kelthane also follows an independentroute and is formed due to oxidation of DDT.

This account of DDT metabolism reflects ageneralized scheme of DDT metabolism inmicroorganisms and is by no means a com-plete documentation .of the types of variousreaction sequences that can occur in individ-ual species. Wedemeyer (242) studied the path-ways of metabolism of DDT in A. aerogenes,which were as follows: DDT -) DDD -- DDMU-+ DDMS DDNU -* DDOH -- DDA -. DBPand DDT DDE. These pathways were eluci-dated by incubating proposed intermediates with

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organisms and examining the products formed.A major criticism of the scheme was that prod-ucts beyond DDNU could not be detected afteraerobic or anaerobic incubation of any of thepreceding metabolite with A. aerogenes. Fur-ther, DDA was only observed after anaerobicbreakdown of DDNU in the presence of anexogenous energy source, and DBP was onlydetected by anaerobic breakdown of DDAalone, and not if any of the earlier breakdownproducts were used. Further studies by Wede-meyer (243) with A. aerogenes revealed thatDDA is converted to DBP via DPM and DBH.The low yield of DBH and DBP compared withthat of DPM suggests either that DBP is rapidlyconverted into some other unknown metaboliteor that DBH and DBP are only minor productsand the majority of DDA is converted via someother pathways into some other products, whichremain undetected.DDT is also degraded to water-soluble metab-

olites in rumen microorganisms (72), bacteriaisolated from seawater (107), algae (168, 202),and fungi (7, 107). Fries et al. (72) suggested thatthe formation of water-soluble metabolites in-volves an entirely independent and separatemetabolic pathway which is different from thosealready reported for solvent-soluble metabo-lites.A strain of Pseudomonas known earlier as

Hydrogenomonas (67) has been extensivelystudied for its ability to metabolize DDT and itsanalogs. Focht and Alexander (67) demonstratedthe metabolism of DDM, itself a metabolite ofDDT in the sequence that involves ring cleavageto PCPA. Further studies by Subba-Rao andAlexander (221) revealed that Pseudomonasalso metabolized DDM to DBH, DBP, benzhy-drol, benzophenone, p-chlorophenyl ethanol,and p-chlorophenyl glycolaldehyde in the se-quences shown in Fig. 1. It may be mentionedhere that no such nonchlorinated moleculeshave been detected from natural environments.This is not unexpected, because, once formed,they would most likely be degraded rapidly.The ciliate protozoan T. pyriformis has recent-

ly been used to study the pathways of metabo-lism of DDT (132). T. pyriformis metabolizedDDT to DDE, o,p'-DDT, and DDMU. Incuba-tion of the species with different metabolites-viz., DDD, DDE, o,p'-DDT, and DDMU-re-vealed that none of the compounds was metabo-lized further except for DDD, which was con-verted to DDMU. This indicates that inTetrahymena, o-p'-DDT and p,p'-DDE areformed directly from p,p'-DDT and no otherintermediate is involved. However, DDD, anintermediate during the formation of DDMU inmost microorganisms, was not detected whenTetrahymena was incubated with DDT. This

probably indicates either a divergent metabolicpathway ofDDT metabolism to DDMU in Tetra-hymena or that DDD is converted readily toDDMU so that its levels always remain belowdetectable limits.The exact pathway for DDCN formation from

DDT in microorganisms is not known. Howev-er, Albone et al. (2) and Jensen et al. (101)suggested that DDCN is formed directly fromDDT and that no other intermediate is involved.

1,2,3,4,5,6-Hexachlorocyclohexane. (i) Metab-olism. In general, BHC is less persistent thanDDT, aldrin, dieldrin, and heptachlor. The rapiddisappearance of -y-BHC from soil and aquaticenvironments has been attributed to its suscepti-bility to degradation by microorganisms (28,144, 198, 255). Like DDT metabolism, microbialdegradation of -y-BHC (Fig. 2) has been ob-served to occur more rapidly under anaerobicconditions (120, 145, 212, 254). Jagnow et al. (98)studied the degradation of y-BHC and its iso-mers in anaerobic and facultatively anaerobicbacteria. The amounts of -y-BHC or its isomerswere determined by gas-liquid chromatography.The results of their experiments revealed thatnot only the strict anaerobic bacteria but alsosome of the facultatively anaerobic bacteria(grown aerobically and subsequently anaerobi-cally) belonging to the Enterobacteriaceae andBacillaceae metabolized these compounds ac-tively.

Although microorganisms degrade -y-BHC andits isomers, the extent of degradation is depen-dent on the spacial arrangement of chlorineatoms in the benzene ring. As observed byMacRae et al. (144), the four isomers of BHC(a-, 1B-, y-, and b-BHC) were decomposed inflooded rice soils at different rates (-y-BHC > a-BHC > 1-BHC = b-BHC). A similar pattern ofdegradation was observed with a mixed cultureof anaerobic microorganisms isolated from soiland also with axenic cultures of Citrobacterfreundii (98), with the exception of Citrobacterbutyricum, which degraded a-BHC to nearly thesame extent as y-BHC. Despite a reasonablerate of anaerobic degradation, a-BHC is one ofthe most frequently observed environmentalcontaminants. This may be due to its release intothe environment by the application of technical-grade BHC, which contains large amounts of a-BHC, and also to the conversion of -y-BHC to a-BHC by microorganisms (13, 91).The major metabolites of y-BHC in microor-

ganisms are -y-PCCH and -y-TCCH. The formeris formed by dehydrochlorination of -y-BHC, andthe latter is formed by dechlorination of y-BHC.Francis et al. (69) detected ly-PCCH from -y-BHCin E. coli. However, Pseudomonas putida me-tabolized y-BHC to -y-PCCH, y-TCCH, and CO2(13, 151). Mixed microbial cultures with P. aeru-

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108 LAL AND SAXENA

cl

Cl ?-PCCH

dehYdrochlornation

dechlormnotic

,'L'

Cl CI

Isomrer,atlon

-HC(%,I PCl -- Ho ~ICl

dchydro- y hydroxylationchlorination Chloro- Chlorophenot

benzene Ir ing openinq

FIG. 2. Proposed pathways of metabolism of y-BHC in microorganisms.

ginosa as the predominant species converted a-,3-, 'y-, and B-BHC to -y-PCCH and y-TCCH (61).Several other microbes have been reported tometabolize y-BHC to y-PCCH, y-TCCH, a-TCCH, chlorinated benzenes, and phenols (128,227, 228).MacRae et al. (143) and Sethunathan et al.

(210) reported an unknown metabolite of y-BHCin Clostridium which was later identified as y-TCCH (226). Cell suspensions of Clostridiumsphenoides have been reported to convert a- and-y-isomers of BHC to a- and y-TCCH, respec-tively (91). Both of these intermediates werefurther metabolized to unknown compounds. InC. rectum, -y-TCCH was the major metabolicproduct of -y-BHC, and the ability of the bacteri-um to metabolize -y-BHC was inhibited by theaddition of oxygen (179).

Dechlorination of-y-BHC during the formationof y-TCCH releases chlorine ions. This has beendemonstrated in experiments with microbial cul-tures, using y-[36C1]BHC. In mixed cultures ofbacterial flora, Haider and Jagnow (86) showedthat about 90%o of -y-BHC was converted tochloride and chlorine-free metabolites. In anoth-

er experiment, different microorganisms, includ-ing Clostridium butyricum, Clostridium pasteur-ianum, and C. freundii, were reported todechlorinate y-BHC completely, thus liberatingchloride and chlorine-free metabolites (98).Complete degradation of y-BHC, which in-

volves ring cleavage to produce C02, was re-ported to occur in submerged soil by MacRae etal. (144). Later studies by Sethunathan et al.(210) revealed that Clostridium sp. is the speciesin soil which is mainly responsible for the con-version of y-BHC to CO2. Further studies byMatsumura and Benezet (150) with P. putidarevealed that an exogenous substrate (reducedflavin adenine dinucleotide) promotes the com-plete degradation of y-BHC, which is convertedfirst to y-TCCH and then to CO2.

Little information is available on y-BHC me-tabolism in algae and fungi, and protozoansappear not to have been studied. Sweeney (222)found that Chlorella vulgaris and Chlamydomo-nas reinhardi metabolized -y-BHC to y-PCCH.In fungi, -y-BHC was metabolized to chlorinatedbenzenes and phenols (63, 128).

(ii) Pathways of metabolism. The pathways of

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-y-BHC metabolism have been studied mainly inrats and houseffies, and the steps of degradationare known to involve dechlorination, dehydro-genation, and dehydrochlorination (29, 41, 224).As is evident from the literature, microorga-nisms have the capacity to degrade -y-BHC ex-tensively, but there have been few studies on thepathways of microbial metabolism of y-BHC.Matsumura et al. (151) reported the existence oftwo pathways of -y-BHC metabolism in P. pu-tida. The first pathway was the dehydrochlori-nation of -y-BHC to -y-PCCH and was not depen-dent on any exogenous substrate. The secondpathway was the dechlorination of -y-BHC to -y-TCCH and was nicotinamide adenine dinucleo-tide dependent. The latter pathway was alsoconfirmed by Jagnow et al. (98) in C. freundiiand C. rectum; further, they observed that exog-enous substrate acts as an electron donor for -y-BHC degradation. Studies by Ohisa et al. (180)on the mechanism offormation of y-TCCH fromy-BHC revealed that the exogenous substrate(dithiothreitol) donates two electrons, whichcause the release of chlorine ions and y-TCCHfrom y-BHC. Thus, the first step of y-BHCdechlorination by C. rectum is C6H6C16 (y-BHC)+ 2e -- C6H6Cl4 (-y-TCCH) + 2Cl.The metabolism of -y-TCCH to monochloro-

benzene remained unclear for many years, al-though monochlorobenzene as a metabolite of y-BHC was detected as early as 1955, by Allan (6),in Clostridium sporogenes and Bacillus coli(Escherichia colt) isolated from cattle dips. Re-cently, Ohisa et al. (180) have investigated thesteps involved during the formation of mono-chlorobenzene from y-BHC in C. rectum. Intheir experiment they observed that y-BHC wasconverted to y-TCCH first and that then y-TCCH was attacked by a pair of electrons fromthe exogenous substrate, which would producedichlorocyclodienes. However, dichlorocyclo-dienes were not detected, for they are quiteunstable and probably spontaneously dehydro-chlorinated to monochlorobenzene. The schemeof formation of y-TCCH and monochloroben-zene is shown in Fig. 2.The third pathway of metabolism of y-BHC in

microorganisms involves its isomerization to a-BHC, as observed in P. putida (157) and E. coli(237). According to Vonk and Quijns (237), thisisomerization requires the conversion of config-uration of one of the carbon atoms of -y-BHC.Since the position of chlorine atoms in -y-BHC iseeeaaa (e, equatorial; a, axial), whereas that ina-BHC is eeeeaa, it may be speculated that anintermediate containing a double bond isformed. Formation of a-BHC from the potentialintermediates y-TCCH and y-PCCH requires theaddition of two chlorine atoms to y-TCCH andHCl to y-PCCH, which does not seem very

probable. Another potential intermediate couldbe hexachlorocychohexene, but this metabolitehas not been detected in microorganisms. Inter-estingly, the formation of a-BHC from -y-BHCwas always accompanied by the formation of -y-TCCH in P. putida (151) and E. coli (237), andthe formation of a-BHC was also stimulated byexogenous substrates (151). Since y-TCCH doesnot seem to be a probable precursor of a-BHC,an enzyme system similar to that operating in -y-TCCH formation might be responsible for theformation of both of these compounds or theymight have a common precursor (151).

Aldrin and dieldrin. (i) Metabolism. Proposedpathways of metabolism of aldrin and dieldrinare shown in Fig. 3. Aldrin is less persistent thanDDT and BHC and volatilizes rapidly from soil(89). This had placed aldrin among the effectiveorganochlorine insecticides for soil insect con-trol. However, dieldrin residues were detectedfor several years wherever aldrin was applied.In contrast to aldrin, dieldrin is a highly persis-tent and nonvolatile insecticide.For several years it was assumed that dieldrin

residues appeared as a result of microbial epoxi-dation of aldrin in soil in areas where dieldrinitself had never been used. The indirect evi-dence for the conversion of aldrin to dieldrin byepoxidation was provided by Lichtenstein andSchulz (138). They found that aldrin was rapidlyconverted to dieldrin in nonsterile soil with littlecoversion in sterile soil. Aldrin also disappearedmore quickly in moist soil than in dry soil,because microorganisms are more active in

Cl2 CI

rcCl aIdrin

1101

C12 CI

Ct s ln

trans- aidrindi

isomeriza- Htion Ct

. Cl ketooldrin0

DH phd

iol photodigldrin

FIG. 3 Proposed pathways of metabolism of aldrinand dieldrin in microorganisms.

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110 LAL AND SAXENA

moist soil. Subsequently, Lichtenstein et al.(139) showed that the conversion of aldrin todieldrin in soil was inhibited by synergists atrelatively high rates and that this inhibitionparalleled a decrease in microorganism popula-tions.The first direct evidence that microorganisms

metabolized aldrin to dieldrin was provided byKorte et al. (123). They found that cultures ofAspergillus niger, Aspergillus flavus, Penicilli-um notatum, and Pencillium chrysogenum me-tabolized aldrin to dieldrin and to four otherunidentified metabolites. In the light of thesefindings, Tu et al. (231) carried out an extensivestudy to prove the role of soil microorganisms inconversion of aldrin to dieldrin. They screened92 pure cultures of microorganisms for theiraldrin-degrading activity. A majority of themicroorganisms showed the ability to convertaldrin to dieldrin. Among the fungi, Tricho-derma sp. was most active, followed by Fusar-ium and Penicillium. Except for Bacillus sp.,bacteria were of less importance. The ability ofmicroorganisms to metabolize aldrin to dieldrinwas linear in relation to time in some cases,whereas in other cases a period of adaptationappeared to be necessary, after which came amarked increase in the production of dieldrin.

Little information is available on the metabo-lism of aldrin in algae. Khan et al. (115) reportedthat algae metabolized aldrin to dieldrin. Inaddition to dieldrin, trans-aldrindiol was formedby Dunaliella.

Dieldrin is more persistent than aldrin in soil.The degradation of this compound has beenmainly attributed to microorganisms (138). Of577 isolates of soil microorganisms, only 10 werefound to degrade dieldrin, namely, 6 Pseudomo-nas, 2 Bacillus, and 2 Trichoderma isolates.trans-Aldrindiol was the major metabolite, andsix other metabolites were water-soluble. T.viride isolated from an Ohio orchard converteddieldrin to trans-aldrindiol and to solvent-solu-ble and water-soluble metabolites (154).Matsumura and Boush (153) isolated a strain

of Pseudomonas from soil near a dieldrin-pro-ducing factory in order to study the difference inmetabolic activity of this strain. They found thisPseudomonas sp. to be capable of producing the6-keto derivative, and alcohol, an aldehyde, andan acid. Later, Matsumura et al. (156) identifiedphotodieldrin as the metabolic product of diel-drin in this isolate. In addition to photodieldrinand trans-aldrindiol, Vockel and Korte (234)also reported trace amounts of ketoaldrin inPseudomonas marginalis, P. fluorescens, Pseu-domonas syringae, Pseudomonas mors-prun-orum, Pseudomonas glycinea, and Neurosporasp.An interesting aspect of dieldrin metabolism is

its ring cleavage to CO2 in Trichoderma, report-ed by Bixby et al. in 1971 (16). A year later,Jagnow and Haider (97) reported the formationof CO2 in several isolates of bacteria and fungi.Out of 175 isolates of bacteria and fungi, 91could degrade dieldrin to CO2. In addition, manywater-soluble and solvent-soluble metabolites ofdieldrin have also been reported (97, 124).

Patil et al. (189) reported metabolism of dieldrinin several species of algae, including Dunaliellasp. The metabolites were photodieldrin, trans-aldrindiol, and several unidentified compounds.They emphasized that the role of light during themetabolism of dieldrin in algae must be consid-ered, because light is capable of affecting boththe rates and the routes of metabolism of diel-drin.

(ui) Pathways of metabolism. Little informationis available on the pathways of metabolism ofaldrin and dieldrin. Microbial epoxidation ofaldrin produces dieldrin. The reverse of epoxi-dation involves the reduction of dieldrin to al-drin. Dieldrin is further hydrolyzed by microor-ganisms to trans-aldrindiol. Photodieldrin is theresult of an isomeric change in dieldrin. Theformation of these compounds from dieldrin isdirect and no other intermediate seems to beinvolved. The conversion of dieldrin to water-soluble and solvent-soluble metabolites and toCO2 has been reported, but the pathways ofmetabolism of dieldrin to these compounds havenot been established.

Heptachlor. (i) Metabolsm. Heptachlor, likealdrin, is oxidized to its epoxide in soil (11, 73,174, 248) (Fig. 4). In addition to heptachlorepoxide, Weisgerber et al. (245) reported hy-droxydihydroheptachloro -1 - methoxychlordeneand 1-hydroxychlordene as the degradationproducts of heptachlor in soil. The first evidencethat heptachlor is degraded by soil microorga-nisms came from the experiments of Langlois etal. (135), when less of the epoxide was recov-ered from sterilized than from nonsterilized soiltreated with heptachlor. Heptachlor is also con-verted chemically to 1-hydroxychlordene in soil,and microbial attack on this product results inthe production of 1-hydroxy-2,3-epoxychlor-dene (38). Mixed cultures of microorganismsisolated from cutting fluid of the Texaco Co. alsometabolized heptachlor to chlordene, 1-hy-droxy-2,3-epoxychlordene, and heptachlor ep-oxide (22).

Miles et al. (163) have shown that heptachloris metabolized by soil microorganisms to manydifferent products by many independent meta-bolic pathways. In their studies they used 92isolates of soil bacteria and fungi. Heptachlorepoxide, chlordene, chlordene epoxide, 1-hy-droxychlordene, and 1-hydroxy-2,3-epoxychlor-dene were the products of microbial degradation

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ORGANOCHLORINE INSECTICIDES AND MICROORGANISMS

Heptechlor epoxide

FIG. 4. Pathways of metabolism of heptachlor in microorganisms (from Miles et al. [163]).

of heptachlor. Further studies by Miles et al.(164) on mixed microbial cultures isolated fromsandy loam revealed that heptachlor and itsepoxide were readily converted to chlordeneand 1-hydroxychlordene, respectively. Thesetwo metabolites were not degraded further bymixed cultures of microorganisms. This could bethe reason for the occurrence of high levels of 1-hydroxychlordene and low levels of heptachlorepoxide in heptachlor-treated soils.

(U) Pathways of metabolsm. Oxidation of hep-tachlor by microorganisms converts heptachlorto its epoxide. Heptachlor is also chemicallyhydrolyzed to 1-hydroxychlordene, which, uponepoxidation by microbes, produces 1-hydroxy-2,3-epoxychlordene. Microbial dechlorination ofheptachlor produces chlordene, which under-goes microbial epoxidation to form the corre-sponding chlordene epoxide.Mirex and Kepone. Mirex is a polycyclic or-

ganochlorine insecticide and is used in bait gran-ules to control fire ants (36). It is highly resistantto microbial degradation and thus can persist insoil for long periods (105). However, Andradeand Wheeler (8) reported that in sewage sludgeunder anaerobic conditions in the dark, about80%o of mirex was degraded into unknownmetabolites.Kepone is similar to mirex in structure and,

like mirex, is used in bait granules to control fireants (36). The persistent nature of Kepone hasled to many severe environmental problems.Kepone and its derivatives have been detectedin soil (21) and aquatic environments (181) and inworkers exposed to high concentrations of Ke-pone during plant operation (17). However, littleis known about the agents responsible for the

transformation of Kepone and its products. Re-cently, Orndorff and Colwell (182) have ob-served microbial metabolism of Kepone in Pseu-domonas aeruginosa and in a mixed cultureobtained from sewage sludge lagoon water. Purecultures of P. aeruginosa and mixed microbialculture were found to dechlorinate Kepone tomonohydro-Kepone and dihydro-Kepone underaerobic conditions. Although the concentrationsof total Kepone transformation products weresimilar for both sets of microorganisms, themixed culture produced less dihydro-Keponeand more monohydro-Kepone than did purecultures of P. aeruginosa. Similar quantitativedifferences between mixed and pure cultureswere also observed in microorganisms isolatedfrom James River sediments (181).

Endosulfan. Pathways of endosulfan metabo-lism are shown in Fig. 5. Soil bacteria and fungidegraded endosulfan to endosulfate and endo-sulfandiol (146). Endosulfate was the major me-tabolite in fungi, and endosulfandiol was themajor metabolite in bacteria. Degradation ofendosulfan in these microorganisms was depen-dent on pH. Microorganisms were active inendosulfan degradation below pH 7, but abovepH 7, chemical degradation was predominant.Other metabolic products identified were en-doether, endohydroxyether, chlorendic acid,and endolacetate. El-Zorgani and Omer (60), intheir study on a- and ,B-endosulfan degradationin A. niger, reported endosulfandiol as the majormetabolic product, in contrast to the findings ofMartens (146), who reported endosulfate as themajor metabolite.

Recently, Miles and Moy (162) studied thepathways of endosulfan metabolism in mixed

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112 LAL AND SAXENA

SsOH \0

ci NN

co

- - endosulfan

N

c,

endg,sullansutfate

N NH

Ci

Cl

3- endosulfan

I

i* i N NmC"zH ")-C1N2ON'ci~~~~~~~cCl C Cl

c, endosulfan etherendosulfan diol

H~ ON

C CIc,

cndosulfan oC-hydroxyether

H 0

CN,Nci Ici

l a tendosulftan (actone

FIG. 5. Pathways of metabolism of endosulfan in microorganisms (from Miles and Moy [162]).

cultures of soil microorganisms. They reportedthe interconversion of a- and P-endosulfan.Both a- and P-endosulfan were further con-verted primarily to endosulfandiol. The endo-sulfandiol was then converted chiefly to thea-hydroxyether with a minor pathway to endo-sulfanether. When incubated separately,endosulfanhydroxyether was converted almostcompletely to endosulfan lactones.

Chlorobenzilate and chloropropylate. Chloro-benzilate and chloropropylate are similar toDDT in some of their physical and chemicalproperties. However, very little is known abouttheir fate and behavior in the environment.Although considerably less persistent thanDDT, their narrow spectrum of insecticidal ac-tivity limits their usefulness as successors toDDT. Both chlorobenzilate and chloropropylatewere metabolized to 4,4'-dichlorobenzilic acid inR. gracilis (167). 4,4'-Dichlorobenzilic acid wasfurther converted to DBP. The rates of chloro-propylate metabolism and chlorobenzilate me-tabolism were greatly increased when citratewas added to the microbial growth medium. Thepresence of citrate in the medium stimulated thedecarboxylation of 4,4'-dichlorobenzilic acid,

whereas 2-ketoglutaric acid inhibited the proc-ess. On the basis of the chemical similarity ofcitric acid to 4,4'-dichlorobenzilic acid, Miyaza-ki et al. (167) suggested that these two substratescould be handled by the same decarboxylation-dehydrogenati6n system. Further in vitro stud-ies are needed to confirm such a possibility.

Miscellaneous organochlorine insecticides. Patilet al. (188) reported that endrin was converted toketoendrin in 20 microbial isolates, including T.viride, Pseudomonas sp., Micrococcus sp.,Arthrobacter sp., and Bacillus sp. In anotherstudy, Matsumura et al. (155) reported at leastseven metabolites of endrin in addition to ke-toendrin, including ketones and aldehydes withfive and six chlorine atoms. A. aerogenes, P.aeruginosa, and three unidentified microorga-nisms degraded endrin, but the metabolites werenot identified (J. A. Dazzio, Diss. Abstr. B28:2893, 1968). In algae, endrin was metabolizedto ketoendrin and to an unidentified metabolite(189).Mendel et al. (158) studied the metabolism of

methoxychlor in A. aerogenes under anaerobicconditions. They reported 1,1-dichloro-2,2-bis(p-methoxyphenyl)ethylene and 1,1-dichloro-

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2,2-bis(p-methoxyphenyl)ethane as the metabol-ic products of methoxychlor. The soil fungi A.flavus, A.- niger, and P. notatum degraded iso-benzan to hydrophilic metabolites (125).

Recently, Clark and Matsumura (43) reportedthe metabolism of 14C- and 'Cl-labeled toxa-phene (a complex insecticidal mixture of polych-lorobornans) in aquatic sediments, enriched bac-terial cultures, and washed-cell cultures of P.putida. Oxidation of toxaphene occurred underaerobic conditions and was proposed as themajor metabolic pathway for toxaphene metabo-lism.

Enzymes Involved in Metabolism ofOrganochlorine Insecticides

Most synthetic insecticides do not share anoverall structural resemblance to natural prod-ucts, but synthetic and natural chemicals havemany substituent and bond arrangements incommon. It is probable, therefore, that most ofthe synthetic insecticides are degraded by en-zymes evolved in response to the presence ofnatural substrates. The enzymes induced in re-sponse to certain synthetic chemicals other thanorganochlorine insecticides may also degradeorganochlorine insecticides. Thus, enzymes in-volved in the metabolism of organochlorine in-secticides may be constitutive or may requireinduction by insecticides (111) or by compoundsother than insecticides (3, 4). For instance, C.rectum stored in the absence of y-BHC in liquidmedium and subsequently incubated with -yBHCwas able to degrade y-BHC without a lag (179),indicating that this species has constitutive en-zymes for -y-BHC metabolism. Tu et al. (231)isolated some strains of soil microorganismswhich required the induction of enzymes todegrade aldrin. They observed that the activityof microorganisms in metabolizing aldrin to diel-drin was linear in relation to time in some cases,whereas in other cases a period of adaptationappeared to be necessary, after which came amarked increase in the production of dieldrin. Inthe latter case, microorganisms required a timelag to form an enzyme system which was notalready present when the organisms were grownin the absence of the added substance.

Indirect evidences for the involvement of mi-crobial enzymes in the metabolism of DDT indifferent strains of T. viride have been reported(157, 188). Evidence for the involvement of asingle enzyme at each step during metabolism ofDDT by A. aerogenes was obtained by Wede-meyer (242, 243). Wedemeyer (243) also foundthat cyanide, fluoride, iodoacetate, and malon-ate inhibited the formation of DDMS fromDDMU, but did not affect the decarboxylationof DDA to DBP. Since cyanide is a specificinhibitor of cytochrome c oxidase, Wedemeyer

(243) suggested the involvement of cytochromesas cofactors for DDT metabolism.A comprehensive study on enzymatic degra-

dation of DDT by F. oxysporum has been car-ried out by Engst and Kujawa (62, 127a). Theyreported that F. oxysporum degraded DDT toDDD, DDE, DDMU, DDA, DDOH, and DBP invivo. Fractionation of a cellular homogenate andsubsequent incubation of each fraction withDDT yielded one fraction which contained allenzymes responsible for DDT metabolism. Ion-exchange chromatography of this fraction ondiethylaminoethyl cellulose yielded five enzy-matic fractions. Fraction 1 contained enzymeswhich metabolized DDT to DDMU, DDMU toDDOH and DBP, and DDA to DDM and DBP.Fractions 2 and 3 were not active, for only traceamounts of these metabolites were produced.Fractions 4 and 5 contained enzymes whichmetabolized DDT to DDE, DDA to DDOH, andDDM to DBP. Further studies on separation ofenzymes by gel filtration and sedimentation gavenegative results. However, their results revealedthat several enzymes with similar molecularweights are responsible for DDT metabolism.French and Hoopingarner (70) studied the

ability of cellular fractions of E. coli to metabo-lize DDT. Reductive dechlorination of DDT toDDD occurred in the 'membrane fraction, andaddition of the soluble fraction enhanced themetabolic rate of DDD formation. Addition ofreduced flavin adenine dinucleotide to the mem-brane fraction also enhanced the rate of DDDformation. It was concluded from these resultsthat reductive dechlorination is a membrane-associated process and that the rate offormationof DDD from DDT is stimulated by a factor(s)present in the cytoplasm.

In the light of these findings, studies werecarried out on the enzymatic metabolism ofDDT in T. pyriformis (Lal, Ph.D. thesis). Differ-ent fractions were obtained from a cellular ho-mogenate of T. pyriformis, and each fractionwas incubated with DDT under anaerobic condi-tions at 30°C. DDT was dechlorinated to DDDby total homogenate and nuclear and microsom-al fractions. The microsomal fraction was mostactive: about 45% of the DDT was converted toDDD. However, dechlorination of DDT did notoccur in the postmicrosomal fraction and infractions boiled before incubation. This con-firms the finding of French and Hoopingarner(70) that enzymes for DDT metabolism are asso-ciated with membranes.

Recently, Heritage and MacRae (92) reportedfor the first time the enzymatic dechlorination ofy-BHC to y-TCCH in C. sphenoides. A cell-freepreparation of C. sphenoides degraded ly-BHCto -y-TCCH in the presence of glutathione. Theenzyme responsible for dechlorination was asso-

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114 LAL AND SAXENA

ciated with the membrane fraction, for this frac-tion showed more rapid metabolism comparedwith the soluble fraction. The time course ofmetabolism of ly-BHC revealed that the enzymeinvolved is fairly stable. However, Heritage andMacRae made no attempt to isolate and charac-terize the enzyme.

EFFECTS

Effects on Populations of MicroorganismsIt is evident from the data reviewed that

microorganisms are important agents in deter-mining the accumulation and metabolism of or-ganochlorine insecticides in the environment.However, they may themselves be subject toorganochlorine insecticide toxicity. The natureof the action of an insecticide on microorga-nisms depends upon the nature of the insecti-cide, the concentration of the insecticide, thetypes of microorganisms, and the environmentin which the microorganisms are growing. Thissection will deal with the effects of organochlo-rine insecticides on microbial growth. The termmicrobial growth will be restricted only to themultiplication of a population, which is de-creased, increased, or not affected at all by theaddition of the insecticide.

Several workers have studied the toxicities oforganochlorine insecticides to microorganisms.Although it is difficult to reach any conclu-sion, some trends are evident. Albone et al. (1)found that technical DDT at 100 ppm in agarconsistently reduced counts of both aerobic andanaerobic bacteria cultured from mud. Earlierstudies by Kokke (121) revealed that DDT inagar at 0.1 ppm reduced the population of bacte-rial isolates from nursery soil. Ko and Lock-wood (119) also demonstrated an inhibitory ef-fect of 10 ppm DDT on several microorganisms.In contrast, massive doses of DDT (112 kg/hectare) did not alter numbers of soil microflora(207).

In loam, DDT and dieldrin reduced the num-bers of bacteria whereas the numbers of fungiwere increased, indicating that bacteria are moresusceptible to the effects of these insecticidesthan are filamentous microorganisms (220). Col-lins and Langlois (47) reported that the effects ofinsecticides on microorganisms were also de-pendent on the nature of the culture medium.They found that DDT and heptachlor at 50 and100 ppm were capable of affecting the growth ofP. fluorescens and S. aureus on agar plates, butno effect was observed on the growth of testorganisms in skim milk containing DDT, diel-drin, or heptachlor.

Martin (147) showed that annual applicationsof aldrin, dieldrin, chlordane, DDT, and toxa-phene at 5 to 20 lb/acre (ca. 6 to 23 kg/hectare) to

Romana sandy loam had no appreciable effecton the numbers of soil bacteria or fungi. Evenvery high concentrations (2,000 ppm) of dieldrinand aldrin had no significant effect on popula-tions of bacteria or fungi (230). A related cyclo-diene insecticide, endrin, when applied to soil atmore than three times the concentration expect-ed from endrin-treated tree seed, exerted noappreciable effect on the numbers of soil mi-crobes, and it was concluded that a very highconcentration of endrin in soil would be neces-sary to alter soil microbial activities (18).

Richardson and Miller (204), using plate cul-ture techniques and Rhizoctonia solani, demon-strated that y-BHC, which has a relatively highwater solubility (6 to 7 ppm), was most toxic atsupersaturation (25 ppm). Heptachlor, chlor-dane, and aldrin were highly toxic even at lowconcentrations. They suggested that the differ-ence in toxicity of these insecticides is due totheir difference in vapor pressure and solubilityin water.

Aldrin, dieldrin, chlordane, endrin, and hepta-chlor at 10,000 ppm had a selective effect onbacterial growth in that they inhibited a widerange of gram-positive bacteria without affectinggram-negative bacteria (9). This selective effectwas attributed to blockage in the electron trans-port chain and to some factor(s) associated withthe chemical composition of the cell wall ormembrane.Kepone, like most of the organochlorine in-

secticides, is highly persistent in the environ-ment and has been detected in both soil andaquatic environments (105, 181), but its effectson microorganisms are yet to be studied. Re-cently, Orndorff and Colwell (182) reported theresults of a field survey on the toxicity ofKepone carried out over a period of 2 years.They found that only gram-negative bacteria,predominantly Pseudomonas spp., Vibrio spp.,and Aeromonas hydrophila, were susceptible to<1 mg of Kepone per liter.There is evidence of considerable diversity in

susceptibility among aquatic microorganisms tothe effects of organochlorine insecticides (Table4). It can also be noted from Table 4 that in manyof the studies, levels of organochlorine insecti-cides far exceeding their water solubility havebeen used.The toxicity of DDT to the diatom S. costa-

tum increased with increasing concentrations,and it was concluded that low levels of DDT innatural waters might have deleterious effects onphytoplankton populations (253). A similar con-clusion was drawn by Sodergren (216), whoreported that the growth of Chlorella sp. wasaffected by less than 0.3 ppb DDT. The results ofother studies are somewhat at variance withthese observations. DDT at 0.6 ppm was not

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TABLE 4. Effects of organochlorine insecticides on microbial populationsMicroorganism Concn Effect Reference(s)

Bacteria, actinomycetes,fungi

Bacteria, fungi, protozoa

Soil bacteria

Pseudomonas fluorescensEscherichia coliStaphylococcus aureusAspergillus flavus

Aspergillus parasiticus

PhytoplanktonsChlorella sp.

Chlorella sp.Chlorella pyrenoidosa

Chlorella ellipsoideaAnacystis nidulansScenedesmus obliquusScenedesmus quadricauda

Chlamydomonas rei0thardiiChlamydomonas sp.Dunaliella sp.Cyclotella sp.

Skeletonema costatum

Synechococcus elongatusAnkistrodesmus falcatus

Euglena gracilisEuglena gracilisSoil protozoansSoil amoebae and flagel-

latesParamecium bursariaParamecium multimicronu-

cleatumEuplotes vannus

Crithidia fasciculata

Stylonychia notophora

Blepharisma intermedium

Tetrahymena pyriformis

Bacteria, fungi

Ammonifying bacteriaRhizobium sp.Azotobacter sp.Bacillus subtilis

Nitrifying bacteria

10 lb/acre (ca. 11 kg/hectare)

400 lb/acre (ca. 454kg/hectare)

100 ppm (technicalDDT)

50-100 ppm50-100 ppm50-100 ppm2 ppm

2 ppm

0-6 ppm1 ppm

5 ppm0.3 ppb10-100 ppm

100 ppm1 ppm1 ppm0.1-1000 ppb0.1 ppm

20 ppm100 ppm

1 ppm1 ppm

100 ppb

20 ppm1-25 ppm

1 ppm100 ppm200 ppm10 ppm

1 ppm1 ppm

10 ppm

425 ppm

50-100 ppm

50-100 ppm

50-100 ppm

400 lb/acre (ca. 453kg/hectare)1,000-1,500 ppm1,000 ppm1,000 ppm1,000 ppm

100 ppm

None

None

Reduction in populationdensity

NoneNoneNoneSlight increase in popula-

tion densitySlight increase in popula-

tion densityNoneReduction in population

density

NoneClumping of cellsNoneNoneNoneNoneNoneSlight reduction in popula-

tion densityNoneNoneNoneReduction in population

densityReduction in population

densityNoneReduction in population

densityNoneNoneNoneReduction in population

densityNoneNone






None

NoneNoneNoneReduction in population


density

VOL. 46, 1982

Insecticide

DDT 19

215

1

47474731

31

232Ellis andGould-inga

25021542, 23344828214028

16944160170

160

16953

8242, 195106, 215145

8282

190

Frenchb

131

130

129

80, 81

58, 251585858

215

BHC (techni-cal grade)


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TABLE 4-ContinuedMicroorganism

Aspergillus niger

Chlorella pyrenoidosa

Ankistrodesmus braunji

Anacystis nidulans

Rhizobium japonicum

Marine phytoplanktonsProtococcus sp.

Anabaenopsis racibroskii

Anabaenopsis aphanizo-menoides



Soil bacteria, fungi

Soil bacteriaRhizobium sp.

Microcystis aeruginosa

Euglena gracilis

Soil bacteriaDictyostelium discoideum


Cyclotella nana

Scenedesmus quadricauda


Cyclotella nana

Chlorella pyrenoidosa


Cylindrospermum sp.

Scenedesmus obliquusChlorella variegataNitzschia palea

Concn

0.5 ppm

0.001-10 ppm

0.1-100 ppm

0.1-100 ppm

50-100 ppm

5 ppm1 ppm

10 ppm

10 ppm

5-10 ppm

50-100 ppm

100 lb/acre (ca. 113kg/hectare)10 kgthectare1,000-2,000 ppm

5 ppm

50-100 ppm

10 kg/hectare50 ppm

5 ppm

100 ppb

0.1-1 ppm

5 ppm

100 ppm

200 ppm

0.1-100 ppb

0.1-50 ppb

100 ppb

100 ppb

2 ppm

2 ppm

2 ppm2 ppm2 ppm

Effect






NoneIncrease in population den-

sityReduction in population




density

None

NoneReduction in population



density

NoneReduction in population




density



None

Increase in population den-sity

Increase in population den-sity


None



NoneNoneNone

a S. W. Ellis and K. H. Goulding, Br. Phycol. J. 8:208, 1973.b J. E. French, Diss. Abstr. Int. B 37:96-97, 1976.

Insecticide

y-BHC

Aldrin

Dieldrin

Endrin

Reference(s)

31

75, 77

122

122

55, 229

232232

54

54

99, 197

55

64, 138,187

88108

233

195

8832

233

160

32

233

160

50, 68,240

79

79

79

126

183

183

183183183

Chlordane Soil bacteria, fungi

Scenedesmus quadricauda

Chlamydomonas sp.

Chlamydomonas sp.

Methoxychlor

Toxaphene


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lethal to marine phytoplankton (232). Even high-er concentrations (20 ppm) did not affect thealgal population (233). Christie (42) found thatDDT at 10 and 100 ppm did not affect the growthof C. pyrenoidosa cultured axenically. The lackof effect of DDT was attributed to the lowsolubility ofDDT in water and rapid degradationof this compound by the organisms. However,these should not be taken as the only factorswhich make Chlorella more resistant to DDT ascompared with other algae which are more sus-ceptible. Some other intrinsic factors, such asthe lipid contents and physiological status of theorganism, also play an important role in decidingthe toxicity of the insecticide.

Species-specific susceptibility to organochlo-rine insecticide may have a pronounced effect onnatural microbial populations. Although effectson total viable counts may be minimal, thesusceptibility of one type or group of organismsmay result in an appreciable change in thebehavior of the population. For instance, Men-zel et al. (160) observed that different species ofmarine phytoplankton varied considerably intheir responses to the organochlorine insecti-cides DDT, dieldrin, and endrin. DDT at 100 ppbinhibited cell division in S. costatum (a coastaldiatom), but had no effect on Coccolithus hux-leyi (an open-ocean alga). In contrast, endrinhad little effect on cell division in S. costatum.Dunaliella tertiolecta (an estuarine species) wasapparently insusceptible to all of the compoundsup to 1,000 ppb, whereas cell division wasinhibited in C. nana (an open-ocean alga) atconcentrations of 0.1 to 1 ppb. Menzel et al.suggested that the resistance of the estuarinespecies, compared with the more susceptiblecoastal and open-ocean forms, may reflect theirgreater adaptability to insecticides. Consistentwith these observations, Bowes (26) reportedthat with the exception of S. costatum, whosegrowth was inhibited at 80 ppb DDT, sevenother species of marine phytoplankton were notaffected at all.

In a related study, Mosser et al. (170) con-firmed that D. tertiolecta was unaffected by1,000 ppb DDT, as reported by Menzel et al.(160), and that other species, including C. rein-hardii and E. gracilis, were also relatively resis-tant (170). Mosser et al. (171) further investigat-ed the effects of DDT on mixed cultures ofmarine algae containing the susceptible diatomThalassiosira pseudonana (earlier known as C.nana) and the resistant alga D. tertiolecta inequal proportion. It was found that growth of D.tertiolecta was not inhibited at any DDT concen-tration tested. However, T. pseudonana, whichgrew faster and soon outnumbered D. tertiolectain control cultures, was affected by 10 ppb DDTto the extent that its competitive success was

significantly diminished, even though T. pseu-donana was unaffected by 10 ppb DDT in pureculture. The levels ofDDT used by Mosser et al.(171) do occur in natural waters. Thus, it isreasonable to assume that DDT can induce suchspecies-specific changes in natural environ-ments.

In natural environments, microorganisms maybe exposed to several chemicals simultaneously.The interaction of chemicals with insecticidesmay influence microbial tolerance of insecti-cides. The possibilities of such alterations in theeffects on microorganisms were investigated byMosser et al. (172). A mixture of organochlorinecompounds (DDE and polychlorinated biphe-nyls) was added to axenic cultures of T. pseu-donana. The results indicated that polychlori-nated biphenyls and DDE in combination weremore toxic to the organisms than they wereindividually, indicating that polychlorinated bi-phenyls and DDE act synergistically, thus in-creasing the toxicity. However, addition of 50ppb polychlorinated biphenyls restored thegrowth of organisms which had been affected by500 ppb DDT. In the blue-green alga Nostocmuscorum, the toxicity of BHC was reducedwhen concentrations of nutrients, such asK2HPO4, Ca(NO3)2, and CaCl2, were increasedbeyond normal levels (110). In this case, BHCinteraction with nutrients to form a complexwith less toxicity was suggested as the reason.

Several reports exist on the effects of insecti-cides on microorganisms. However, less isknown about the possible adverse effects of theirmetabolites. In Anacystis nidulans, DDD andDDE inhibited growth markedly (25). In thiscase DDD was more toxic, followed by DDEand DDT. The growth of an alga, Exuvilla bal-tica, was inhibited by both DDT and DDE, butDDE was more toxic than DDT (1%). Of threeinsecticides and their metabolites tested for theireffects on blue-green algae, aldrin was the leasttoxic to algae, and isolates of Agmentallumquadruplicatum were more tolerant than A. ni-dulans (12). Algal growth was depressed by 950ppb dieldrin and by its metabolite photodieldrin.Endrin was more toxic to the algae than was itsmetabolite ketoendrin.

Little is known about the effects of DDT onprotozoans in both soil and aquatic environ-ments. Available information suggests that pro-tozoans are more resistant to DDT than areother microorganisms. Gregory et al. (82) ob-served that 1 ppm DDT did not have any toxiceffect on E. gracilis, Paramecium bursaria, andParamecium multimicronucleatum, thoughthese species concentrated DDT to 99- to 964-fold. Even concentration ofDDT up to 100 ppmdid not alter the growth of E. gracilis (195). Onthe contrary, the populations of several protozo-

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ans, including amoebae and flagellates, werereduced upon treatments with 5 and 50 ppmDDT (145). DDT and DDE separately caused a 3to 10% decrease in cell numbers of the ciliateprotozoan Euplotes vannus at 10 ppm (191),whereas a flagellate protozoan, C. fasciculata,was resistant to DDT and a higher concentration(425 ppm) of DDT was required to suppress thegrowth appreciably (French, Diss. Abstr. Int. B37:96-97, 1976). In three ciliate protozoans, S.notophora (131), B. intermedium (130), and T.pyriformis (129), lower concentrations of DDT,up to 1 ppm, affected neither morphology norcell division. However, higher concentrations,such as 50 and 100 ppm, inhibited cell divisionappreciably in all the three ciliates.

Determination of the effects of organochlorineinsecticides on microbial growth is commonlydone by means of viable counts, which does notestablish whether the size of the cell as a wholeis affected or not. To obtain definitive informa-tion about the effect ofDDT on the cell size of T.pyriformis in addition to viable counts, surfacearea and volume were also determined (129).DDT at 50 and 100 ppm brought about a declinein cell number as well as a reduction in surfacearea and volume.

Reports on the effects of BHC on protozoansare somewhat confusing. BHC at concentrationsbetween 0.001 to 0.5 ppm decreased the cellnumber of T. pyriformis significantly (76). Thisspecies was shown to be resistant to DDT atthese concentrations (129). Daubner et al. (55)reported stimulation of growth in T. pyriformistreated with 0.0018 ppm -y-BHC, and higherconcentrations, between 50 and 100 ppm, wererequired to inhibit cell division significantly. Incontrast, Jeanne-Levain (99) reported that 5ppm ly-BHC inhibited the multiplication of T.pyriformis and that 10 ppm of y-BHC inhibitedcell division completely. Although such discrep-ancies may be explained in experiments con-ducted under natural conditions, it is disturbingthat they should occur in pure cultures, illustrat-ing the inherent variability of microbial reactionsto insecticides.The effects of BHC on algae have not been

extensively studied. BHC at 50 or 100 ppb didnot cause any detectable effect on algal growthor progeny size in a mixed culture containing asusceptible (T. pseudonana) and a resistant (D.tertiolecta) species of marine phytoplankton(15).The literature reviewed above deals with the

primary effects of organochlorine insecticides onmicroorganisms. The secondary effects of or-ganochlorine insecticides have received littleattention, but they are also important. Amongthe most commonly reported secondary effectsof organochlorine insecticides in aquatic envi-

ronments is the formation of phytoplanktonblooms. This is mainly due to the indirect effectof insecticides on algae, because more sensitivezooplankton species either feed on algae orcompete for food with algae. For instance, DDTtreatments offorests for defoliation control or ofstreams for Simulium control were followed bythe formation of phytoplankton blooms due tothe decimation of aquatic arthropods that feedupon them (95). Phytoplankton activity in a NewJersey salt marsh treated with DDT at 1 lb/acre(ca. 1.1 kg/hectare) increased nearly fourfoldbecause the larvae of mosquitos (Aedes sollici-tana) were eliminated (206), and in Minneapolisponds only Volvox responded to DDT treatment(1 lb/acre), which increased to seven times thenormal levels (106).A stream in the Isle of Man contaminated by

BHC discharged from sheep-dips developed anovergrowth of Cladophora, accompanied bysome Spirogyra, because of the destruction ofthe herbivorous arthropods, such as mayflynymphs and Gammarus amphipods (96). Appli-cation of ly-BHC at S kg/hectare in Philippinerice fields resulted in a bloom of blue-green algaedue to the elimination of ostracods (199).

Cytological and Biochemical EffectsOur present knowledge of cytological and

biochemical effects of organochlorine in micro-organisms is too fragmentary to provide anysubstantial evidence regarding their modes ofaction. In higher organisms, such as mammals,where the site of action has been clearly identi-fied, organochlorine insecticides interfere withthe ionic permeability of nerve cell membranesand so produce an unstable state in which spuri-ous nerve impulses induce uncontrolled activityin the whole organism. However, in microorga-nisms these insecticides affect one or more sites,depending upon the type of microorganism. Themajor targets which are susceptible to DDT arecell membranes, enzymes, and nucleic acids. Inalgae, organochlorine insecticides seem to inter-fere primarily with photosynthetic activity,which may bring many other biochemicalchanges in cellular metabolism. As mentionedearlier, our present knowledge of the effects oforganochlorine insecticides at the cellular levelmay not be sufficient to point out the site(s) ofaction, but it does form the basis of developingworking concepts to better understand themechanism(s) of action.

Cell membrane. Independent of whether theinsecticide is adsorbed or absorbed, it seemsreasonable to believe that interaction of organo-chlorine insecticides with cell membranes isimportant in determining the primary target oftheir action. Lethal action of DDT in B. subtiliswas both dose and time dependent, and it was

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related to the binding ofDDT to membranes (93;G. F. Hicks, Diss. Abstr. Int. B 36:4313, 1976).Lipid analysis revealed that DDT treatment inthis species altered the lipid composition of themembranes. Recent studies on the qualitativeand quantitative alterations in the lipids of cellmembranes in E. coli showed that DDT, aldrin,and dieldrin altered the ratio of polar phospho-lipid head groups and the composition of fattyacids (205). These changes might be expected toalter the structure and function of membranes,because polar phospholipids are the main lipidconstituents of cell membranes.Amino acids and proteins. Interference of or-

ganochlorine insecticides with the structure andchemistry of membranes can affect the perme-ability of the cell, which can in turn have seriouseffects on cellular activity. For example, DDT atconcentrations of 1 to 5 ppm decreased theuptake of amino acids in Chlamydomonas niva-lis, A. amalloides, and Scenedesmus basiliensis(53). However, transport of certain amino acids,such as tyrosine, valine, leucine, isoleucine, andhistidine, remained unaltered or was slightlyhigher than in the controls. Higher concentra-tions, from 10 to 25 ppm, still reduced thetransport of amino acids in C. nivalis and A.amalloides, whereas in S. basiliensis the trans-port of two amino acids, serine and arginine,was increased. This further indicates that DDTtreatment alters the structural integrity of themembrane in such a way that it becomes morepermeable to certain amino acids while inhibit-ing the transport of many other amino acids.

In C. fasciculata, DDT treatment even at 425ppm increased the uptake of [3H]leucine fromthe medium (French, Diss. Abstr. Int. B 37:96-97, 1976). However, in the ciliates S. notophora(208), B. intermedium (R. Lal and D. M. Saxena,Acta Protozool., in press), and T. pyriformis(209), the uptake of [3H]lysine from mediumcontaining 100 ppm DDT was inhibited signifi-cantly, thereby inhibiting the synthesis of pro-teins. The inhibition of the incorporation of[3H]lysine was reversible, for the ciliates recov-ered from the effect when transferred to toxi-cant-free medium. These ciliates are capable ofquickly metabolizing and eliminating DDT whentransferred to toxicant-free medium, which maybe the reason for their rapid recovery (132).

Nucleic acids. Organochlorine insecticides, inaddition to their interference with the structureand function of membranes and with proteinsynthesis, also affect the synthesis of nucleicacids. In C. fasciculata, DDT (425 ppm) inhibit-ed the uptake of [3H]thymidine and [3H]uridine,thereby inhibiting the synthesis of deoxyribonu-cleic acid and ribonucleic acid, respectively(French, Diss. Abstr. Int. B 37:96-97, 1976).This inhibition in the synthesis of nucleic acids

was attributed to interaction of DDT with com-plex regulatory processes associated with thetransport of precursors necessary for nucleicacid synthesis. In T. pyriformis, DDT at 50 and100 ppm inhibited the uptake of [3H]thymidineand [3H]uridine added directly to the medium inwhich the organisms were grown (209). Howev-er, in S. notophora (131, 133) and B. interme-dium (Lal and Saxena, in press.), [3H]thymi-dine-[3H]uridine was provided throughprelabeled Tetrahymena because the organismswere unable to take up the isotope directly. Theorganisms were able to incorporate the isotopethrough Tetrahymena, but failed to synthesizedeoxyribonucleic acid or ribonucleic acid whentreated with DDT. This indicates that the inhibi-tion of nucleic acid synthesis may not be due toalterations in cell permeability alone, but rathermay be due to the action of DDT on the internalenvironment of the cell so that the organism isnot able to initiate the synthesis of nucleic acideven though the precursor is available.

It is not yet possible to distinguish whether theinhibitory effect is due to the binding of theorganochlorine insecticides to cytoplasmic ornuclear sites. G. G. Polikarpov, V. G. Tsytsu-gina, and A. V. Tokoreva (Bioferachel Matr.Vscs. Simp. 1:278, 1975) have reported that inProcentrum micans, radiolabeled DDT entersthe nuclear membrane, suggesting its direct in-terference with the genetic machinery. Alterna-tively, DDT may alter the internal milieu of thenucleus, thus disturbing the whole process ofnucleic acid synthesis.

In Dunaliella bioculata and Amphidinium car-teri, y-BHC inhibited cell division and the syn-thesis of deoxyribonucleic acid and ribonucleicacid (100). The synthesis of deoxyribonucleicacid was strongly inhibited during the first cellcycle, followed by complete inhibition in thesecond cell cycle. The synthesis of ribonucleicacid was strongly inhibited during the first cellcycle as compared with the second cell cycle.The latter effect can be attributed to the adapta-tion of the organisms to the toxicant and subse-quent dilution of -y-BHC in the daughter cells.Enzymes and catabolic pathways. Aldrin at

1,000 and 2,000 ppm inhibited the metabolism ofpentose, hexose, and tricarboxylic acid cycleintermediates in Rhizobium sp., presumablythrough its interference with oxidative enzymes(108). A comprehensive study on enzyme sys-tems in T. pyriformis revealed that BHC at 0.001ppm did not affect the activity of 8-aminolevulin-ate dehydrogenase, hexokinase, and pyruvatekinase, whereas it stimulated the activity ofglutamic dehydrogenase, isocitrate dehydroge-nase, and malate dehydrogenase (74). Increasingthe concentration of BHC from 0.001 to 0.1 ppmincreased the activity of almost all of the en-

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zymes except alanine aminotransferase, the ac-tivity of which was depressed. Further increasesin concentration to 1 ppm inhibited the activityof almost all of the enzymes.Nelson and Williams (177) observed that or-

ganochlorine insecticides (chlordane, hepta-chlor, heptachlorepoxide, aldrin, dieldrin, en-drin, and mirex) inhibited cell division in S.cerevisiae. The inhibition of growth was as-sumed to be due to insecticide interference withoxidative metabolism. This was supported byfindings that chlordane inhibited cell division ofS. cerevisiae on all nonfermentable substrates(glycerol, lactate, and ethanol) provided as ener-gy sources, but not when fermentable sugars(glucose, galactose, and fructose) were providedas energy sources. This further indicates thatchlordane does not inhibit the transport of sub-strates into the cell and further strengthens theinterpretation that the primary site of chlordaneaction is oxidative metabolism. Whether chlor-dane inhibits cell division by direct action on thecomponents of oxidative phosphorylation or bygenetically altering the synthesis of these com-ponents remains to be determined.Chlordane is also known to inhibit the activi-

ties of succinate dehydrogenase and reducednicotinamide adenine dinucleotide dehydroge-nase in B. subtilis and reduced nicotinamideadenine'dinucleotide dehydrogenase in E. coli(225, 247). In the marine bacterium A. proteoly-tica, chlordane inhibited endopeptidase activity(175; Nakas, Diss. Abstr. Int. B 38:77, 1977).Further, in this species chlordane was associat-ed with membranes, suggesting that specificalterations in the enzyme activity are due to theassociation of chlordane with membranes.

Photosynthesis. Various detrimental effects onautotrophic microorganisms engaged in photo-synthesis have been reported for DDT and otherorganochlorine insecticides (77, 213, 232). Thishas prompted further research on the effect ofthese insecticides on photosynthesis, and thusseveral reports have appeared in the literatureindicating the inhibition of photosynthesis inalgae by organochlorine insecticides (26, 27, 33,40, 46, 170, 171, 253). This is particularly signifi-cant because algae play an important role inmarine food webs and in the biosphere's oxygenbalance. This section will examine the effects oforganochlorine insecticides on photosynthesis inmicroorganisms.

Photosynthesis consists of light and dark reac-tions. Adenosine 5'-triphosphate (ATP) and re-duced nicotinamide adenine dinucleotide phos-phate are synthesized in the light reaction, andCO2 is given off. In the dark reaction CO2 isreduced and incorporated into various organiccompounds using ATP and reduced nicotin-amide adenine dinucleotide phosphate. A toxic

compound can interfere with photosynthesis dueto its influence on (i) the development andstructural integrity of chloroplasts, (ii) the pho-tochemical pathways involved in the conversionof radiant energy to chemical energy (light reac-tion), or (iii) the biosynthetic pathway involvedin the production of carbohydrates (dark reac-tion).The effects of DDT on the morphology of the

Nitzschia delicatissima chloroplast indicatedthat even the lowest concentration (9.4 ppb)distorted the chloroplast, and at 1,000 ppb themorphology of the chloroplast was completelydestroyed (142). Kopecek et al. (122) reportedthat 10 ppm -y-BHC reduced the chlorophyllcontents in Ankistrodesmus braunii and A. nidu-lans, whereas concentrations below 10 ppmstimulated photosynthesis. BHC at 10 ppm in0.33% acetone also decreased the chlorophyllcontent in C. pyrenoidosa (75, 184). The finestructures of the chloroplasts showed some de-gree of degeneration in C. pyrenoidosa (184) andD. bioculata (197).

Clegg and Koevening (44) determined the ef-fects of four organochlorine insecticides (DDT,aldrin, chlordane, and dieldrin) on the lightreaction of photosynthesis by measuring theproduction of ATP. Three species of algae,Chlorella ellipsoidea, Chlamydomonas sp., andE. gracilis, were exposed to 100 ppm of eachinsecticide separately for 3 days. ATP was thenextracted and assayed by measuring lumines-cence in a luciferin-luciferase solution. Cleggand Koevening showed that all of the insecti-cides significantly reduced the amounts of ATPdetected in algal extracts, but did not significant-ly alter the population densities. They suggestedthat the insecticides interfere with photophos-phorylation in the light reaction of photosynthe-sis in these algae.The assimilating power, reduced nicotinamide

adenine dinucleotide phosphate, and ATP gener-ated in the photochemical reaction are utilizedfor fixation and reduction of CO2. As organo-chlorine insecticides interfere with the produc-tion of ATP due to their effect on photophos-phorylation in the light reaction, this willcertainly affect CO2 fixation. Thus, 14C uptakeas a measure of photosynthesis in algae treatedwith insecticides has been used by many investi-gators. Wurster (253) found that fixation of14CO2 in four species of marine algae declined asthe concentration of DDT increased up to 100ppb. Menzel et al. (160) observed that DDTaffected 14CO2 uptake differently in differentmarine algae, and they suggested that this wasdue to the differential penetration of the insecti-cide through the cell walls, and membranes indifferent organisms.A detailed study on the effect of DDT on

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photosynthesis was carried out by Lee et al.(137) with Selanstrum capricormutum. They alsoused 14C fixation as a measure of photosynthesisand further traced the path of radiolabeled car-bon in the alga. Their studies revealed that DDTconcentrations between 3.6 and 36 ppb wereinhibitory to photosynthetic CO2 fixation, andlonger exposure to DDT further increased theinhibition. The incorporation of 14C from 14CO2indicated that DDT stimulated the incorporationof 14C into glycolic acid, a major compound ofphotorespiration, and caused the concomitantsuppression of flow of 14C into aspartic acid, amajor component of the C4-dicarboxylic acidpathway. This shift from an efficient pathwayinto a nonefficient pathway by DDT was inter-preted to be caused by interruption of cyclicphotophosphorylation.Based on the current status of knowledge on

the effects of organochlorine insecticides onphotosynthesis, it is difficult to postulate theexact site of interference of an insecticide withthe photosynthetic system. However, it seemsto center on inhibition of ATP and reducednicotinamide adenine dinucleotide productionduring the light reaction. It is hoped that thecurrently available information will serve as amodel that can be subjected to rigorous andsophisticated experimentation.

Cell morphology. Apart from biochemicalchanges, organochlorine insecticides have beenreported to alter cellular morphology. S. noto-phora treated with 100 ppm DDT revealed alarge number of abnormalities in the nuclearmorphology, such as deep incisions, loose chro-matin, and fragmented macronuclei (131). In C.fasciculata, DDT at 425 ppm changed the archi-tecture of the plasma membrane and producedmitochondrial swelling (French, Diss. Abstr.Int. B 37:96-97, 1976).

In D. bioculata, y-BHC at 10 ppm altered thenumber of cellular organelles and caused degen-eration of the nuclear apparatus (99). Furtherdetailed study at the ultrastructural level in D.bioculata and A. carteri revealed that y-BHCtreatment enlarged the cells, decreased the num-ber of microtubules, and enlarged the Golgiapparatus (100). -y-BHC (10 ppm) treatment inAcetabularia altered the structure of plastidsmarkedly (20). In the basal part of the plastidsthe lamellae were extended with one or severalcarbohydrate grains, whereas in the apical partsmall chloroplasts with numerous thalakoids andwith small polysaccharide granules were ob-served.A detailed study on the effects of hexachloro-

benzene (chemically related to BHC) and ace-tone (used as solvent) on the growth and ultra-structure of C. pyrenoidosa revealed thatacetone alone at 0.33% did not produce any

change in the ultrastructure of the organism(184). However, 3.33% acetone resulted in thedegeneration of cellular structure (184). Hexa-chlorobenzene at 10 ppm in 0.33% acetone dam-aged the cell membrane and cell organelles,leaving only starch grains, the pyrenoids, andsome endomembranes. These effects were at-tributed primarily to the damage caused to thecell membrane, which allowed the leakage ofcellular material.

CONCLUSIONSThe literature on accumulation, metabolism,

and effects of organochlorine insecticides onmicroorganisms has been reviewed. Microorga-nisms in soil and water accumulate these insecti-cides fairly readily and bioconcentrate them tomany times their concentrations in the surround-ing environment. However, no attempt has beenmade to study the mechanism of transport ofthese insecticides through the cell membrane,though microorganisms can prove very conve-nient experimental material for this purpose.

It is clear from the data that bacteria and fungiplay a significant role in the metabolism oforganochlorine insecticides. The contribution ofalgae and protozoans is uncertain and has re-ceived relatively little attention. It is also evi-dent from the literature that most of these stud-ies have been carried out with pure culturesunder laboratory conditions, and results havebeen interpreted to indicate what may happenunder natural conditions. However, in naturethe metabolism of insecticides is influenced byenvironmental factors (e.g., temperature, oxy-gen concentration, limited nutrients, competi-tion). It is therefore possible that the samemicroorganisms might metabolize an insecticidedifferently according to the environmental con-ditions.Our knowledge of the enzymatic metabolism

of organochlorine insecticides in microorga-nisms is incomplete and contains many inconsis-tencies. Only a few reports are available on DDTand BHC which indicate the involvement ofenzymes in their degradation. Greater emphasisshould be placed on the isolation and character-ization of the enzyme involved in insecticidedegradation. Whether these enzymes are consti-tutive or induced is again an unsettled question.

In previous years, some reviewers of theeffects of organochlorine insecticides on popula-tions of microorganisms have claimed, justifi-ably, that adverse effects were mainly the resultof high rates of application of insecticides. How-ever, with the considerable advances that havebeen made in pesticide technology, it is becom-ing clear that even lower doses of insecticides atnormal field rates may have adverse effects.The elucidation of the effects of metabolites of

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different organochlorine insecticides is also im-portant, because these metabolites may be moretoxic, may be more persistent, or may havedifferent mobilities than the parent compoundsin microbial environments.Our present state of knowledge of cytological

and biochemical effects of organochlorine insec-ticides is too fragmentary to provide any sub-stantial evidence regarding their mode of action.The available data indicate that organochlorineinsecticides interfere with cell membrane perme-ability, photosynthesis, oxidative metabolism,and synthesis of nucleic acids and proteins. Ithas been recently demonstrated that organo-chlorine insecticides alter the architecture of theplasma membrane and change its lipid composi-tion. On the basis of these results it is temptingto suggest that organochlorine insecticides mayinterfere primarily with the plasma membrane,thus changing cell permeability, which, in turn,can alter cellular physiology.

ACKNOWLEDGMENTS

We are thankful to Ravi Kant, J. P. Khurana, S. K. Gupta,and Vipin Rastogi for their advice and critical comments onthe manuscript. Our sincere thanks are also due to RanjanaMathur and B. V. Prasad Reddy for their help during thepreparation of the manuscript.

This project was funded in part by grants from the Depart-ment of Science and Technology, Government of India.

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2. Albone, E. S., G. En , N. C. Evam, and M. M.Rhead. 1972. Formation of bis-(p-chlorophenyl) acetoni-trile (p,p'-DDCN) from p,p'-DDT in anaerobic sewagesludge. Nature (London) 240:420-421.

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9. Andus, L. J. 1960. Microbiological breakdown of herbi-cides in soil, p. 1-19. In E. K. Woodford and G. R. Sagar(ed.), Herbicides and soil. Blackwell Scientific Publica-tions, Ltd., Oxford.

10. Barker, P. S., F. 0. Morrlson, and R. S. Whbtaker. 1965.Conversion of DDT to DDD by Proteus vulgaris. Abacterium isolated from the intestinal flora of a mouse.Nature (London) 205:621-622.

11. Barthel, E. F., R. T. Murphy, W. G. Mitchell, and C.

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