Metabolic Transformation of DDT, Dieldrin, Aldrin, and Endrin by Marine Microorganisms Krishna C. Patil, Fumio Matsumura,' and G. Mallory Boush Department of Entomology, University of Wisconsin, Madison, WI 53706
To study the transformation process of stable chlorinated hydrocarbon pesticides in marine environments, samples of seawater, bottom sediments from both ocean and estuarines, surface films, algae, and marine plankton were collected and treated with radiolabeled insecticides at the collection sites and incubated for 30 days in the laboratory. Also, various microorganisms were isolated from these samples and their metabolic activities on the insecticides were monitored along with some laboratory cultures of unicellular algae. Transformation of DDT and cyclodiene insecticides took place in samples with biological materials such as surface films, plankton, and algae, but not in waters from open ocean. A number of marine microorganisms in pure culture also metabolized the pesticides. The patterns of metabolic activities by the microorganisms were similar to those observed in the fieldcollected samples.
D
espite the enormous concern with the persistence of DDT and other cyclodienes, very little is known about the biodegradation processes of these insecticides in the oceanic environments. There are some reports about the degradation of these insecticides by soil and nonaquatic microorganisms-e.g., Barker et al., 1965; Chacko et al., 1966; Matsumura and Boush, 1967; Matsumura et al., 1968; Tu et al., 1968). We have reported earlier the metabolism of DDT by soil, fresh water, and lake bottom sediments (Lake Michigan) (Patil et al., 1971), and cyclodiene insecticides by various microorganisms (Patil et al., 1970; Matsumura et al., 1971). We have extended our studies for the metabolism of DDT and other chlorinated hydrocarbon insecticides under oceanic conditions and report the results of our investigations, Experimental
Metabolism Under Oceanic Conditions. For studying the metabolism under oceanic conditions, the insecticides were added to several samples of water (1000 ml) directly at the site of collection in Oahu, Hawaii, during the summer of 1970. Descriptions of the collection sites and conditions are summarized in Table I. Temperature and pH of the samples at the time of collection were in the range of 26-27°C and 7.6-7.7, throughout. The water samples were incubated with 0.1 pmole of labeled insecticides which were added with 100 p1 of 9 5 Z ethanol and kept at 23°C in the laboratory for 30 days. Sediments and other visible particulate matter were removed from the water samples through precipitation by gravity and subsequent decantation. All samples of sediments were collected with water from the collection site (2 parts of water to 1 part sediment) in a glass bottle filled up to the top to avoid aeration. Surface films were collected by use of a specially designed collection apparatus (Harvey, 1966). Plankton samples were collected simultaneously by using DDT
~~
' To whom correspondence should be addressed.
Clarke-Bumpus apparatus with No. 20 mesh net (G. M. Manufacturing Co., New York, N.Y.). In all cases the insecticides were incubated with plain seawater (from open sea) under identical experimental conditions as controls to compensate chemical and photochemical decomposition of the pesticidal substrate, if any. Metabolism with Pure Microbial Cultures. Algal cultures tested were laboratory colonies of a Dunaliella sp. and Agmenellum quadraplicatum (Strain PR-6) (Batterton, 1970) and field-collected algae containing water samples. All the insecticide-treated algal cultures were maintained under constant illumination with fluorescent lamps. The control tubes with plain media were also kept under illumination to correct photolytic degradation. To study the microbial metabolism of these insecticides in pure cultures, a few samples of water, surface films, and bottom mud with water were used for isolation of pure cultures from Hawaii and the ship canal area, Houston, Tex. The water samples from Houston contained varying degrees of surface slicks of crude oil. The isolation and culture techniques used were similar to those described before (Matsumura and Boush, 1967). All microbial cultures are kept in either continuous culture or stored in liquid nitrogen to ascertain the reproducibility of the results. For studying the degradation abilities of these microorganisms, 10 pl of 10-3MC14-labeledDDT solution in ethanol was added to yeast extract mannitol cultural media (Fred and Waksman, 1928) in a screw-capped 20-ml test tube with a microsyringe. They were kept for 30 days in an incubator at 3OoC without illumination. The extraction and clean-up procedures were similar to that described by Matsumura and Boush (1967). During the incubation period the tubes were capped to prevent volatilization of water and the insecticide. Also each insecticide was incubated with plain sterile cultural media for the same duration under the identical conditions as a control. Identification of Metabolites. For extraction of the water samples (1 liter), an equal amount of chloroform and methanol mixture (9: 1) was used. Each sample was extracted twice, and the chloroform phase combined and passed over sodium sulfate, The residue was taken up in a small amount of chloroform, and a small portion was spotted, using a glass micropipette, on an activated silica gel thin-layer plate. Throughout the experiments the chromatograms were developed until 15 cm from the origin with ether-hexane (1 :1) as a mobile phase for separation of dieldrin, endrin, and aldrin and their metabolites and n-pentane-ethyl acetate (15 :1) for DDT and their metabolites. Different solvent systems were, however, used for comparing R, values of the metabolites with the known reference compounds. Detection of resolved and radioactive compounds on the chromatogram was accomplished by radioautography using No-Screen Medical X-ray Safety-Film (Eastman Kodak Co., Rochester, N.Y.). An exposure period of 30 days was given in all cases except for the cochromatographic comparison tests of condensed samples against reference compounds, where 10 days exposure was sufficientbecause of their high radioactivities. The radioactive regions on the chromatogram correspondVolume 6, Number 7, July 1972 629
Table I. Some Active Microbial Isolates in Degrading DDT from Hawaii and Houston, Tex. Solvent-H20 DDT and its transformation productsd formed Active" partition (total radioactivity in the solvent phase, %) isolate, TDE DDT Aqueous Solvent Origin DDOH DDNS DDE no. Source4 (total no. of isolates)* 96.6 3 . 5 2 . 0 56.9 28.3 1699 3.4 5.9 ... Pearl Harbor (12), 3.0 ... 44.1 ... 55.7 1702 44.3 8.6 ... bottom sediments (silt) 69.6 3.1 0.6 33.3 6.6 30.4 26.0 ... 1704 20.4 ... ... 14.3 2.4 1708 74.1 3.7 ... ... 9.8 3.4 52.6 32.5 ... 1709 47.4 6.9 5.0 83.6 94.7 ... ... 5.3 ... 1710 6.1 Pearl City (16), 20.9 4.9 ... 15.0 79.1 3.1 15.6 24.3 1719 near-shore sediments 3.0 54.8 2.4 70.4 ... 5.0 1.4 1720 29.6 75.7 9.7 ... ... 8.1 36.0 24.3 9.1 1724 ... 2.0 2.3 54.2 ... 63.0 1725 2.5 37.0 Fish pond (4), semistagnant 0.82 54.4 17.4 82.6 15.7 1.8 5.2 1727 0.82 water 3.3 36.5 72,l 2.2 2.0 1735 27.9 7.6 8.6 Lanai pineapple field 2.0 1736 4.8 66.0 3.1 96.9 7.8 4.8 1.4 (drainage to ocean) 2.7 73.8 94.6 1.4 2.1 1739 5.3 5.6 1.4 Hana (12), shoreline soil, 0.5 51.0 12.3 87.7 4.7 4.8 16.2 2.1 1741 Maui Island 1.3 0.7 59.6 1742 3.6 96.4 5.7 17.8 2.5 0.9 70.5 90.0 1744 10.0 7.8 2.9 3.9 0.7 18.0 0.9 ... 0.4 14.4 1745 37.7 2.8 62.3 1.8 26.1 0.5 0.6 19.9 0.4 1746 1.5 73.9 Algal culture (1) 3.2 2.4 81.0 1750 99.5 6.1 1.7 1.1 0.5 Surface film (9), 21.6 ... 2.0 6.0 7.6 46.5 1754 10.8 89.2 collected in Kahee 4.9 ... 1.9 0.6 2.6 34.7 1755 53.9 46.1 Lagoon, Oahu ... ... 4.3 17.4 25.3 ... 74.7 174-T 3.6 Houston Ship Canal (24)) 3.8 16.8 ... 14.3 ... 3.4 181-T 38.3 61.7 approximately 100 meters ... 3.2 1.1 33.0 5.9 21.1 67.0 1.7 189-T from San Jacinto Monument 2.2 ... ... 21 . o 3.6 13.3 1.9 191-T 79.0 ... 3.9 28.1 2.6 36.9 0.7 63.1 1.6 195-T ... 2.5 23.9 2.1 1.7 5.0 200-T 35.9 64.1 1.4 57.3 2.4 8.3 201-T 14.6 89.9 4.6 10.2 3.0 19.7 18.4 2.6 11.9 0.5 1756 88.1 Kaneohe Bay (17), 65.9 0.7 1.0 8.3 1.3 3.6 80.7 19.3 1761 Oahu, surface and ... ... 8.0 22.6 12.0 19.3 68.6 1764 33.4 near-surface water 18.4 38.0 ... ... 7.0 13.7 80.6 1768 19.4 20.4 47.0 ... ... 10.0 8.1 85.7 14.3 1772 ... 9.0 16.3 55.3 9.0 ... 89.9 1773 10.1 .
I
.
-
Others
... ... ...
... ... ... 16.2 3.8 12.9 2.0 3.8 11.9 10.0 6.6 9.1 8.7 3.4 1.1 1.4 4.1 5.3 9.1
...
... ... ... ... 0.7 ... 1.2 ... 6.4 3.5
No microorganisms were isolated from the water samples from deep sea and near-surface area from open sea.
* Altogether 95 isolates have been tested for DDT metabolism.
All microbial cultures here being identified by the culture numbers are kept in this laboratory for futurereference. d Chemical names used in this paper are (see also Figure 1 ) : DDT: l,I,l-trichloro-2,2-bis(pchlorophenyI)ethane; TDE:l,l-dichloro-2,2-bis(pchloropheny1)ethane; DDNS : 2,2-bis(p-chlorophenyl)ethane;DDOH : 2,2-bis(p-chlorophenyl)ethanol; and DDE:I, I-dichloro-2,2-bis(p-chlorophenyl)ethylene. c
ing to darkened areas on the film were marked and scraped into 20-ml scintillation vials for direct measurement of resolved radioactivity using a liquid scintillation spectrometer. The chromogenic agent for detection of the nonradioactive reference compounds was prepared by dissolving 1 gram of silver nitrate in 5 ml of distilled water, adding 10 ml of 2phenoxyethanol, and making up the volume to 200 ml with acetone. A small drop of 3 0 z hydrogen peroxide was added to the mixture as a preservative. The silver nitrate reagent was stored in a dark brown glass bottle. For comparison of the resolved metabolites of DDT, the chloroform phases of the most active cultures showing similar patterns of degradation were combined. The combined samples were spotted on thin-layer plates and each band scraped and eluted with ether. The ether extract was evaporated to 0.5 ml and respotted again in different solvent systems of acetone and hexane (1 :4); pentane and ethyl acetate 630 Environmental Science & Technology
(15 : l ) ; hexane-ethanol-acetic acid (17 :2 :1) and CC14 as mobile phases. The metabolites of cyclodienes were chromatographically matched against known candidate compounds by using thin-layer chromatographic methods (Matsumura et al., 1968,1971). Results and Discussion About 100 microbial isolates from Hawaii and Houston, Tex., were screened to investigate the role of these microorganisms in degrading DDT. The results of the general survey on the degradation activities of the microorganisms are presented in Table I. Of the total cultures tested, 35 appeared to be active in degrading DDT. The percentage distribution of radioactivity among DDT and its solvent-extractable metabolites are also presented in Table I. It is apparent from the table that TDE is the predominant metabolite representing more than 50
Table 11. Degradation of Cl4-DDTby Field-Collected Samples of Marine Water, Sediments, and Associated Biological Materials5 Partition Metabolites Sample no. Source Water Solvent Origin DDOH DDNS TDE DDT DDE HWOll-T Algae culture 11.5 88.5 32.8 6.1 1.4 16.3 12.6 5.2 (Dunaliella sp.) Plankton, 70.6 29.4 2.8 ... ... 4.3 25.3 ... HW022-T net collection HW024-T Sediment, 65.0 35.0 2.0 0.5 ... 25.8 3.0 0.6 Kahee Lagoon Fishpond, Oceanic 2 6 . 3 73.7 16.6 ... 9.3 31.0 10.0 , , . HW040-T Institute (mullet culture, open) HW044-T Algae, collection 1.5 98.5 5.0 0.8 0.7 52.1 37.3 ... from a fish pond (stagnant)
Others 1.4
... 3.1
6.7
3.3
5 Altogether 17 samples have been examined. Only active samples have been listed here. All insecticides were added to the samples at the collection site, and the system was maintained in the laboratory for 30 days thereafter.
of the radioactivity from the solvent phase. In addition to this, many of them also form DDNS and DDOH as minor metabolites (Figure 1). The isolates from the soil from Lanai, fish pond, and surface film form additional unknown metabolites. The isolates from Houston, Tex., on the whole, showed relatively low metabolic activities. Table I1 represents the water, sediments, surface film, and plankton samples with direct exposure to CX4-DDT.Seventeen samples consisting of water samples from the open sea and estuaries (6), algal cultures (4), surface films (2), one each of plankton and fish pond samples, and three samples of sediment were exposed to DDT for one month. The results of such direct tests indicated generally poor metabolic activity of the water samples. None of the water samples from the open sea, estuarines (such as Kaneohe Bay, Pearl Harbor) and seashores showed any metabolic activities. It is possible, however, that the experimental conditions adopted herein do not favor either chemical or photochemical degradation, and that the insecticides could still degrade under natural conditions. On the other hand, the algal samples, surface films, sediments, fish pond, and plankton samples were found to be relatively efficient in metabolizing DDT. The major metabolite is again TDE. Also, the two other minor metabolites, DDNS and DDOH,
a s p + -
-
C
I
OY
CI-c-CI
CI-c-CI
DDT
TDE
tl
O
C
are formed. For the identification of DDT metabolites from microbial isolates and other samples, matching tests were carried out using different solvent systems. The minor metabolites were identified as DDNS and DDOH by use of the reference compounds. It is of interest to note that both algal cultures produced a metabolite which chromatographically matched with DDOH as the major metabolite. To study their role in degrading the cyclodiene insecticides, the above 17 samples were also exposed directly to radioactive dieldrin, aldrin, and endrin, respectively, and incubated for one month. Table I11 summarizes the results of these test findings. The metabolites of dieldrin, aldrin, and endrin were identified by matching tests with the reference compounds and then tentatively identified (Figure 2). Photodieldrin is the main metabolite from dieldrin, and a small amount of diol and other unidentified metabolites were found in some cases. Dieldrin and trans-aldrindiol are the metabolic products of aldrin and ketoendrin from endrin. It must be mentioned here that both photodieldrin and ketoendrin are known to form as a result of either photochemical reactions (Rosen et al., 1966; Robinson et al., 1966; Zabik et al., 1971), or by microbial actions (Matsumura et al., 1970,1971).Although no photolytic reaction could be observed in any of the control tubes which were illuminated in the absence of microorganisms, there is a
l
A
ALDRIN
A
dH
DDNS
DDOH
C=O OH
Figure 1. Metabolic pathway of DDT under oceanic conditions
\
0 DIELDRIN
0 PHOTO DIEURIN
bH ALDRIN DOL
Figure 2. Metabolic pathway of aldrin and dieldrin under oceanic conditions Volume 6, Number 7, July 1972 631
Table 111. Degradation of C14-Dieldrin,Aldrin, and Endrin by Field-Collected Samples of Marine Water, Sediments, and Associated Biological Materials. Major metabolites*, Sample no. Source Insecticide formed HWOl 1-DL Algae (Dunaliella SP.)
HW026-DL
HW041-DL
HW045-DL
Dieldrin Photodieldrin (8 5 % and unknown metabolite (3.273 Surface film Dieldrin Photodieldrin (1.1 %) and 2 unknown metabolites (2.6%:) Fish pond, Oceanic Dieldrin Diol (2.3 %) Inst. (mullet and unculture, open) known metabolite (4.1%) Algae, collection Dieldrin Photodieldrin (0.5 %) and from a fish pond (stagnant) unknown metabolite (0.5%) Algae (Dunaliella Aldrin Dieldrin SP.) (23.2 %) and diol (5 2%) Surface film Aldrin Diol (8.1 %) Fish pond Endrin Unknown metabolite (35.5 %) Algae, collection Endrin Ketoendrin from a fish pond (24.4 %> (stagnant)
On the other hand, most of the strong degradation activity was found to be associated with biological samples such as algae, plankton, surface films, and microorganisms. The patterns of degradation of insecticides, in general, closely resemble those found in the terrestrial and aquatic (freshwater) flora and fauna (Matsumura and Boush, 1971), except that algal cultures appear to convert DDT exclusively to a DDOHlike compound. Apart from the pure cultures of microorganisms and algae, there is a question of the closeness of the laboratory incubation conditions to the actual field situations. This is particularly true for the surface films which contain delicate microscopic organisms. These organisms are expected to die within 24 hr after collection. This is the reason why the insecticides were given at the site within 1 hr after collection. Nevertheless, change in microflora and fauna during the incubation process is inevitable in all field samples, In the absence of truly adequate methods to simulate field conditions, the approach of comparing the degradation patterns in field-incubated samples vs. those in pure cultures of organisms isolated from the corresponding field samples at least should provide a general information. Acknowledgment We thank John Batterton for his help in testing algal cultures.
Literature Cited Barker, P. S., Morrison. F. 0.. Whitekar. R. S.. Nature (Lon.don), 205, 621-2 (1965). ’ Batterton, J., PhD thesis, “A Study of Halo Tolerance in Blue HW028-A Green Algae,” University of Texas, 1970. HW042-E Chacko, C. I., Lockwood, J. L., Zabik, M., Science, 154,893-5 (1966). Fred, E: B., Waksman, S. A., “Laboratory Manual of General Microbiology with Special Reference to the Microorganisms HW046-E of the Soil,” 145 pp, McGraw-Hill, New York, N.Y. (1928). Harvey, G. W., Limnol. Oceanogr., 11,608-13 (1966). Matsumura, F., Boush, G. M., “Metabolism of Insecticides by Microorganisms” in “Soil Biochemistry,” A. D. McSame samples as the ones tested for DDT. Only active samples have Laren and J. Skujins, Eds., Vol 2, p 320, Marcel Dekker been listed here. b Chemical names used here are (see also Figure 2): dieldrin: 1,2,3,4,Inc., New York, N.Y. (1971). 10,10-hexachloro-1,4,4a,5,6,7,~,Sa-octahydro-6,7 - epoxy - 1.4-endo - 5,sMatsumura. F.. Boush. G. M.. Science. 156. 959-61 (1967). exodimethanonaphthalene; aldrin: 1,2,3,4,10,10-hexachloro-1,4,4a,Matsumura; F:, Boush, G. M., Tai, ’A., ’Nature (London), 5,8,8a-hexahydro-1,4-endo-5,8-exodinlethanonaphthalene;photodiel219. 965-7 (1968). drin: a structural isomer of dieldrin, resulting from 2-9 intramolecular bridge formation; trans-aldrindiol: 6,7-trans-dihydroxydihydroaldrin; Matsumura, F., Patil, K. C., Boush, G. M., Science, 170, endrin: 1,2,3,4,10,10-hexachloro-1,4,4a,5,6,7,8,8a-octahydro-6,7-epoxy- 1206-7 (1970). 1,4-endo-5,8-endodimethanonaphthalene;and ketoendrin: a structural Matsumura, F., Khanvilkar, V. G., Patil, K. C., Boush, isomer of endrin, resulting from 2-7 intramolecular bridge and ketone G. M., J . Agr. Food Chem., 19,27-31 (1971). (6-position) formation. Patil, K. C., Matsumura, F., Boush, G. M., Appl. Microbiol., 19, 879-81 (1970). Patil, K. C., Matsumura, F., Boush, G. M., Nature, 230,325-6 possibility that a photosensitizing substance is present among (1971). the microbial products and that the reaction is of photoRobinson, J., Richardson, A., Bush, B., Elgar, E., Bull. Environ. Contam. Toxicol., 1, 127-32 (1966) chemical-biochemical nature. Rosen, J. D., Sutherland, D. J., Lipton, G. R., ibid., 1966, The most significant observation in the entire study appears pp 133-41. to be that these insecticides are not metabolized in plain seaTu, C. M., Miles, J. R. W., Harris, C. R., Life Sci., 7, 311-22 water. The insecticides were not degraded even in relatively (1968). polluted estuarine waters such as the ones from Kaneohe Bay Zabik, M. J., Schuetz, R. D., Burton, W. L., Pape, B. E., J. Agr. Food Chem., 19, 308-13 (1971). and Pearl Harbor, Hawaii. The only water sample which showed any degree of degradation was the one from fish ponds which contained algal populations. Sea bottom sediReceived for review September 1 , 1971. Accepted January 1I , ments did not show a strong degradation activity, as the 1972. Supported in part by research grant EP-00812 from the sample from Kahee Lagoon (Table 11) being the only sediment Emironmental Protection Agency and by the Brittingham sample which showed a low level of degradation. Others tested Fund of the Uniuersity of Wisconsin Trust. Supported by the were the sandy sediments from various depths of sea shelf College of Agricultural and Life Sciences and Marine Studies Program, University of Wisconsin. and silts and mud of estuarine and bay sediments.
HWOl I-A
632 Environmental Science & Technology
