Induction of abnormal fatty acid metabolism and essential fatty acid deficiency in rats by dietary DDT

Induction of abnormal fatty acid metabolism and essential fatty acid deficiency in rats by dietary DDT

ARCHIVES OF Induction BIOCHEMISTRY AND BIOPHYSICS 175, 262-269 (1976) of Abnormal Fatty Acid Metabolism and Essential Deficiency in Rats by Die...

723KB Sizes 12 Downloads 72 Views







175, 262-269 (1976)

of Abnormal Fatty Acid Metabolism and Essential Deficiency in Rats by Dietary DDT




The Hormel Institute,



Fatty Acid


University of Minnesota, Austin, Minnesota 55912 Received December 3, 1975

Rats maintained on lab chow diet or on an essential fatty acid (EFA)-deficient diet were dosed with 5 or 500 ppm, o,p’-DDT, 5 ppmp,p’-DDT, or 100 ppm Kelthane for 30,60, or 90 days. After 30 days all groups exhibited reduced body weights and increased liver weights, lipid contents of livers were elevated only in the group receiving 500 ppm o,p’DDT. No significant difference was found between the incorporations of tritium from T,O into liver lipids from groups fed 5 ppm o,p’-DDT, p,p’-DDT, 100 ppm Kelthane and control groups. The distribution of o,p’-DDT among subcellular fractions paralleled the distribution of lipid content. The rates of desaturation and chain elongation of palmitate and desaturation of 20:3w6 by liver microsomes were decreased significantly in the groups fed o,p’-DDT. The fatty acid composition of phospholipids from liver microsomes revealed significantly diminished contents of ~6 metabolites, increased w9 acids, and increased triene/tetraene ratios in the animals fed DDT in lab chow although the intake of linoleate was adequate (2%). Dermatitis of EFA deficiency was likewise observed in the DDT-fed rats. The desaturation and chain elongation of palmitate by normal microsomes in vitro were significantly inhibited by low concentrations ofo,p’-DDT added to the incubation medium.

Chlorinated hydrocarbons have permeated the environment to such an extent that it is unlikely one can avoid ingesting measurable quantities of these substances. A review of the literature revealed that from 1 to 7 ppm of DDT’ and its metabolites are found in most items included in a staple diet (l-3). The known physiological and metabolic effects of chronic oral doses of chlorinated hydrocarbons have been reviewed by Conney and Burns (4). Highly membranous organelles are strongly affected by DDT and its metabolites. Large doses (l-300 mg/kg body wt/ day) ofo,p’-DDT [l-(O-chlorophenyl)-l-(p’ Abbreviations used: EFA, essential fatty acids; o,p’-DDT, l-(O-chlorophenyl)-l-(p-chlorophenyl)2,2,2-trichloroethane; o,p’-DDD, l-(O-chlorophenyl)l- (p-chlorophenyl)-l,l-dichloroethane; p,p’-DDT, 2,2 - bis- (p - chlorophenyl) - l,l - dichloroethane; pp’ DDT, 2,2-bis-(p-chlorophenyl)-l,l-dichloroethylene; tic, thin-layer chromatography; glc, gas-liquid chromatography; BSA, bovine serum albumin; DMSO, dimethyl sulfoxide.

chlorophenyl)-2,2,2-trichloroethanel and o,p’-DDD [l-(O-chlorophenyl)-l-(p-chlorophenyl)- l,l-dichloroethanel induced increased rates of detoxification of barbiturates (58), antipyrene (2,3-dimethyl-lphenyl-3-pyrazolin&one) (9), and phenylbutazene (Cbutyl-1,2-diphenyl-3,Bpyrozolidine-dione) (10) by mixed-function oxygenases of rat liver microsomes. Micromolar concentrations ofp,p’-DDT 12,2-bis-(pchlorophenyl)-l,l-dichloroethanel or p,p’DDE [2,2-bis-(p-chlorophenyl)-l,l-dichlorethylene] in the media had deleterious effects on photosynthesis in marine diatoms (11) and on isolated chloroplasts (12). Koch reported in vitro inhibition of cation-dependent ATPase in mitochondrial preparations of several tissues (13-W. Gross effects of chronic oral doses of chlorinated hydrocarbons have been observed on lipid metabolism. Compounds related to DDT, particularly the Arochlor series of chlorinated biphenyls, fed to rats at low rates (20-100 ppm) caused fatty liv262

Copyright 0 1976 by Academic Press, Inc. All rights of reproduction in any form reserved.



ers (16). When 30 mg/kg/day of dieldrin (1,2,3,4,10,10-hexachloro-6,7-epoxy1,4,4a,5,6,7,8,8a-octahydro-1,4-endo-exo5,gdimethano naphthalene) was fed, histopathologies appeared and lipid accumulated, mostly as triglyceride (17). Incorporation of labeled acetate into fatty acids was significantly reduced in the same rats. Geyer reported similar findings from liver biopsies of human females fed 382 mg of o,p’-DDT over 105 days (18). The biosynthesis of fatty acids by liver is mediated by membrane-bound enzyme systems whether in mitochondria or microsomes. The de nouo biosynthesis of saturated fatty acids from acetate has been shown to be adversely affected (17), but no evidence was found for aberrations in the microsomal fraction. From the abnormally large amounts of lipid material produced in livers of animals fed chlorinated biphenyls (17, 18), it seemed likely that aberrations in lipid metabolism may occur. The elucidation of these changes should explain macroscopic effects (16) as well as symptomology. MATERIALS


Chemicals. The carboxyl-labeled acids, 16:0, 20:3w6, and 18:306 were obtained from New England Nuclear Corp., Boston, Mass. The radio and chemical purity of these compounds was found to be >99% by tic and glc. Nonlabeled aids were obtained from the Lipids Preparation Laboratory of The Hormel Institute, Austin, Minn., or Nu-Chek Prep, Inc., Elysian, Minn. Cofactors were purchased from Sigma Chemical Co., St. Louis, MO., and Cal-Biothem, LaJolla, Calif. The technical grade o,p’-DDT andp,p’-DDT were provided by Montrose Chemical Corp. of California, Newark, N. J. and Allied Chemical Corp., Jackson, Miss., respectively. Technical Kelthane (dicofol) [l,l-bis(4-chloropheny1)2,2,2trichloroethanoll was provided by Rohm and Haas Co., Kansas City, MO. Pure pesticides were obtained for glc and tic standards from Supelco, Inc., Bellefonte, Pa., Applied Science Labs, Inc., State College, Pa., and PolyScience Corp., Niles, Ill. All other chemicals were reagent grade and solvents were chemically purified and distilled. The petroleum ether used had a boiling point range from 30 to 60°C.

Dietary treatments. Male weanling the Sprague-Dawley strain weighing maintained on Purina Lab Chow or cient in essential fatty acids (EFA). contained 4.7-5.1% extractable lipids

albino rats of 30-40 g were on a diet detiThe lab chow of which 182




was 35.18, 18:303 was 3.9%, and 18306 was 1.3% of total fatty acids. The EFA-deficient diet contained 68% sucrose, 2% hydrogenated coconut oil, 30% vitamin-free casein, and all known essential vitamins and minerals (19). Tests diets were prepared containing 5 ppm o,p’DDT, p,p’-DDT or 100 ppm Kelthane added as acetone solutions to the chow diet, and the solvent was removed by vacuum. Groups of rats were fed the test diets for 30, 60, or 90 days, with appropriate control groups. Preparation of microsomes. Rats were weighed and anesthetized with ether, and their livers were excised into cold 250 mM sucrose:5 mM MgCl, (20, 21). The chilled livers were weighed, minced, and placed into fresh cold sucrose:MgCl,. All subsequent steps were carried out in ice-cold conditions. The tissue was liquified in a blendor, strained through 20-mesh plastic window screen, and placed in a cell-disruption pressure bomb (22). The vessel was sealed and the contents were sustained under 1000 psi N, with constant agitation for 30 min. The homogenate was released 6-om the bomb into 50-ml centrifuge tubes and centrifuged at 17,300g for 30 min and the supernatant was recentrifuged in a Spinco 30 rotor at 105,OOOg for 2 h at 0-5°C. The pellet was suspended in sucrose:MgC12 solution and the suspension was stored under N, at -20°C in screw-cap tubes. Protein was determined by the method of Lowry et al. (23). Microsomes stored in this manner maintained high activity for at least 60 days. Analysis of lipid classes. Total lipids were extracted according to Bligh and Dyer (24) and lipid classes were separated on activated ITLC-SA papers (Gelman Instrument Co., Ann Arbor, Mich.) using petroleum ether:diethyl ether:glacial acetic acid @O:lO:ll as the developing solvent. Spots were made visible with 2’,7’-dichlorofluorescein and uv light. Spots representing lipid classes were cut out and placed in scintillation vials for determination of radioactivity (20) or transesterified with 14% (w/v) BFs:methanol containing 20% benzene for 1 h at 100°C. The methyl esters were extracted with petroleum ether and separated by glc on a ‘/a-in. x 6 ft 20% EGS + 2% H,PO, on Gas Chrom P (100/120) columns and FID detectors. The columns were operated at 190°C and a flow rate of 60 ml He/min. In this laboratory the methyl ester profile derived from the phospholipids is used as an indicator of EFA deficiency and a 20:3o9/20&6 ratio of >0.4 (25) indicates EFA deficiency. Tabulations of profiles include the summation of members of a fatty acid family minus the major metabolic precursor, i.e., ~6 acids - 18206. In this way the content of the precursors themselves do not obscure the variations of the metabolic products. Analysis of chlorinated hydrocarbons. Pesticide residues were extracted from intact tissue and from




subcellular fractions by partitioning between acetonitrile and water followed by chromatography using florisil columns according to Coffin et al. (26) as modified by Kadoum (27) and Stanley et al. (28). Clorinated hydrocarbons were made visible with AgNO,-a- 2-phenoxyethanol chromogenic spray (29) under uv light, eluted with 7% benzene in petroleum ether, and quantified by glc on 2 mm i.d. x 6 ft glass columns packed with 10% DC200 (12,500 cs) silicone oil or 4% SE-30 + 6% QF-1 on Anakrom Q (80/90, Analabs, Inc., New Haven, Conn.) operated at 200°C. Carrier gas was 5% methane in argon at 40 ml/min. The B3Ni electron capture detector was operated at 50-ps pulse intervals and at 240°C. Samples were run on both columns and the resulting data were averaged. Nucleic acid content was determined by the method of Schneider (30). Enzymatic methods. The fatty acyl desaturase activities in the 105,OOOg microsomal pellet were determined using the procedure of Paulsrud et al. (20) except that the substrates used, 16:0 and 20:306, were Na+ salts complexed with essentially fatty acid free BSA fraction V (Sigma Chemical Co., St. Louis). Reaction products were extracted, transesterified and extracted as previously described. Separation of substrate and product was done on 10% AgNO,-Silica Gel H 0.3-mm tic plates. Petroleum ether:diethyl ether (100:5) was used to separate methyl-16:O and methyl-16:lo7 and petroleum ether:diethyl ether:glacial acetic acid (70:30:1) was used to separate methyl-20:306 and methyl-20:4o6. Spots were made visible and their radioactivity was determined as previously described. The chain elongation assay followed the method of Mohrhauer et al. (31) except Na+-BSA complex was used instead of the NH,+ salt as substrate. The incubation was terminated, and total lipids were extracted and transesterified by methods previously described. Methyl esters were separated by reversed-phase chromatography on ITLC-SA papers previously activated and developed in 5% DC200 silicone oil (10 centistoke) in petroleum ether. The developing solvent was acetonitrile:glacial acetic acid:water (70:30:10). Spots were made visible on solvent-free papers by spraying with 1% a-cyclodextrin in methanol, drying, rehumidification and then exposure to I, vapors. The in vitro inhibition experiments used microsomes obtained from rats fed an EFA-deficient diet. DDT or Kelthane was added in 1 ~1 of DMSO to the microsomes plus cofactor mixture and preincubated 15 min with constant agitation at 37°C prior to addition of the carboxyl-labeled substrate. Controls were similarly treated with 1 ~1 of DMSO, other conditions and methods being the same. Triolein (0.5 mr& in DMSO was added to one set of tubes to determine whether the inhibitions observed could be induced by lipophilic compounds in general. Rate of lipogenesis. The tissue slice method of

AND HOLMAN Eliott (321 was used to measure the incorporation of 3Hp0 into the lipid pool. The incubation mixture contained 80 mCi of 3H20 (100 mCi/g) in 37.5 ml isotonic salt solution and 15 g of freshly excised liver slices. The mixture was agitated for 2 h at 37°C. The reaction was terminated by decanting the supernatant fluid, replacing it with several volumes of icecold CHCl,:MeOH (2:l) and immediately homogenizing the sample. The homogenate was repeatedly washed on a filter with 0.88% KC1 until no activity was detected in the wash water. The lipid classes were separated and collected and radioactivity was determined by previously described methods. RESULTS


In vivo effects on body and liver weights.

Table I shows the body and liver weights of rats fed 5 or 500 ppm o,p’-DDT, 5 ppmp,p’DDT, or 100 ppm Kelthane, and the lipid extracted per gram of fresh liver. Rats fed chronic oral sub-LD, doses of o,p’- or p,p’DDT in a complete commercial chow diet for 30 days had body weights significantly smaller, and liver weights larger than the control animals. There was no significant difference in body and liver weights between animals fed o,p’-DDT or p,p’-DDT. Animals fed 500 ppm o,p’-DDT chow diet for 30 days were smaller, had heavier livers, and 80% more liver lipids than the animals fed 5 ppm. Rats fed the same diets for 60 days showed no significant differences between any of the treatments. All animals fed the higher dosage had statistically significant liver enlargement. All diets containing DDT caused livers to have a greater proportion of lipids, especially the animals fed 500 ppm o,p’-DDT. Rate of lipogenesis in vivo as affected by chronic oral doses of DDT. The accumula-

tion of triglycerides in livers exposed to persistent low levels of chlorinated pesticides observed in the literature and in the preceding section suggests either an increase in the anabolic or decrease in the catabolic metabolism of triglycerides. Increased triglycerides were observed in rats fed either 5 or 500 ppm o,p’-DDT for 30 or 60 days. Increased biosynthesis of triglycerides should be evidenced by a disproportionate share of labeled fatty acid synthesized de novo. Tissue slices incubated with 3H,0 should reveal any change in de novo synthesis of fatty acid and the incorporation of labeled fattv” acids into triglscer--




are shown in Table II. The inclusion of DDT in the chow diet containing adequate amounts of EFA elevated the proportions of 16:1, l&l, and 20:3o9 in a fashion parallel to the elevations known to be caused by EFA deficiency and again demonstrated by negative controls in this experiment. Conversely, the inclusion of DDT in the chow diet suppressed the proportions of all EFA components in the microsomal phospholipids, as is true in the EFA-deficient

ides. We observed no differences in the de of fatty acids (35.1% 2 0.1, 35.5% + 1.8, and 33.0% 2 0.8 of incorporated label) or triglycerides (27.2% 2 3.4, 27.6% + 5.2, and 30.1% ? 3.0 of incorporated label) in livers from rats fed 5 ppm o,p’-DDT,p,p’-DDT in a chow diet or chow alone, respectively. This suggests that the accumulation of triglyceride may be due to differences in catabolism, or that change in de novo synthesis of fatty acids, if present, was below the limits of detection by the method. novo biosynthesis



of EFA deficiency by dietary

Fat-free Fat-free + DDT n

16:l 18:l 20:3w9 18:206 20:206 20:3w6 20&6 Total 06 metabolites 20:3co9/20&6

Body weight (g)

Lab chow + DDT

5 1.5 9.3

5 4.0" 22.6 19.6 2.5 0.8 1.4 8.8

4 6.6 21.7 4.4 1.2 1.4 7.6

17.7 2.0 1.6 22.5

5 5.0 24.4 18.0 3.4 ndb tr” 10.2














Liver weight (g)


SkhCIl- a


231.0 f 10.2 9.2 + 0.7 142 f 30 days 13.6 + 1.6 131 f 60 days 372.1 +- 25.0 196.0 -c 7.1” 14.6 k 0.2" 162 c 30 days 178 + 60 days 368.0 2 14.1 14.1 f 1.4 148 zt 30 days 200.3 + 8.0" 15.1 k 0.2" 367.1 + 11.9 14.7 + 1.3 172 k (5 ppm) 60 days 255 zt 30 days 182.9 k 26.7” 16.8 k 1.V o,p’-DDT 21.0 + 2.6' 279 f (500 ppm) 60 days 376.4 + 27.9 228.2 k 6.8 10.1 + OX Kelthane 30 days 148 2 14.5 + 1.5 154 * (100 ppm) 60 days 377.0 + 26.7 n 0 = no dermatitis, + = moderate dermatitis, ++ = severe dermatitis. b Mean 2 SD of seven rats except the 5 ppm p,p’-DDT which had only four, c Statistically significant from controls at <0.05 level by Student’s t test distribution

Control (lab-chow) o,p’-DDT (5 ppm) p,p’-DDT

Lab chow

u Percentage of total fatty acids from phospholipids. * Not detectable. c Trace, less than 0.01%. ’ Total w6 acids minus 18:206.




DDT. When 5 ppm o,p’-DDT was added to the complete chow diet, the rats became EFA deficient after 90 days, as indicated in two experiments by triene/tetraene ratios of 1.6 and 1.8, and by scaly skin on the feet and tail. The same amount of DDT in the EFA-deficient diet did not significantly worsen either criterion of deficiency because increasing degrees of EFA deficiency are more difficult to induce. The additon of 100 ppm Kelthane caused no dermatitis. The fatty acid profiles of the phospholipids from liver microsomes from the four groups of rats were determined, because phospholipids reveal most clearly effects upon metabolism of polyunsaturated acids and because they are essential and major lipid components of the membranous organelles which accomplish both chain elongation and desaturation of essential fatty acids. The results of these analyses




16 14 14 18 15 16’ 18c 2F 20 19

0 0 + + + + + ++ 0 0







state. The suppression of the 20:206 and 20:3w6 by DDT in chow-fed animals was even greater than in animals fed the fatfree diet. Using the criterion of total ~6 metabolites (all 06 acids except linoleic), the EFA deficiency of the rats fed lab chow + DDT was more severe than that of the rats fed EFA-free diet. The feeding of DDT with EFA-free diet likewise intensified the deficiency by this criterion. Using the triene/tetraene ratio as criterion (29, the EFA deficiency of rats fed lab chow + DDT was comparable to that induced by EFAfree diet. The EFA deficiency induced by DDT added to a diet containing approximately 2% EFA is unmistakeable and moderately severe by all these criteria.

diet. In liver microsomes from rats fed DDT for 30-90 days, the elongation activity was reduced to one-half that of controls. One hundred parts per million Kelthane in chow diet induced 20-30% reductions in desaturase activity and 30-40% reductions in chain elongation activities in both the 60 and 90 day groups. The desaturation of 20:306 to 20:406 in rats fed 5 ppm o,p’-DDT for 90 days was 44% less than controls whereas the elongation of l&306 to 20:306 was not affected (Table III). These results indicate that saturated fatty acids and PUFA are elongated by different enzyme systems. Strong evidence of this exists in the literature

Effect on microsomal desaturase and chain elongation activities. Fatty acid bio-

The addition of o,p’-DDT in 1 ~1 DMSO to microsomes extracted from chow fed rats caused maximum reduction of palmitate desaturase and chain elongation at the lower concentrations of pesticides (Fig. 1). Maximum inhibition of palmitoleate formation occurred at 0.25 mM o,p’-DDT, and above this concentration activities increased toward normal levels. Maximum inhibition of palmitate elongation occurred at 0.42 mM (~50% of control). The mechanism by which low levels of inhibitor have greater effect than higher concentration has been previously reported (35) but is unexplained. The data suggest that the inhibition may be via physical modification of the microsomal membrane rather than steric or substrate-product inhibition.


synthesis has been reported to be adversely affected by chlorinated hydrocarbons in the diet, whereas microsomal enzymes were not affected (17). The data presented here document the inhibitory effects of DDT on the microsomal biosynthesis of fatty acids in uiuo and in vitro. Desaturase activity (16:O + 16:l) decreased in all groups given dietary pesticide (Table III). Although the diminished desaturase activities in the test groups were significantly different from controls, the activities observed afier 30, 60, or 90 days were not significantly different from each other. Chain elongation activity for conversion of palmitate to stearate was also reduced by the presence of 5 ppm DDT in the chow








Treatment Lab-chow


Kelthane WO

(5 pm)


30 60 90 30 60 90 30 60 90

days days days days days days days days days


+ 16:l

1.18 1.08 1.06 0.88 0.61 0.60

+ 2 2 -+ 2 +

0.W 0.08 0.30 0.11’ 0.03” 0.11”


+ 20:4w6

1.54 f 0.48

0.87 ” 0.32c -

0.70 rf: o.o(F 0.78 + 0.14c

(1 Nanomoles of product formed per minute b Mean 2 SD of five rats. c p < 0.05 by Student’s t test (34).

16:0 +


2.89 2.61 2.63 1.47 1.28 1.21

0.31 0.40 0.42 O.lF 0.09 0.03c

k k f 2 2 2

18~306 -+ 20:306

1.70 k 0.2oc 1.81 re_ [email protected] per milligram

of microsomal



1.54 2 0.48

1.70 k 0.64

1.68 k 0.48 20 min.






had increased two- to threefold in all fractions. The 105,OOOg supernatant contained an unexpected quantity of both lipid material and DDT residue. That o,p’-DDT residues do partition into lipid-rich organelles is indicated by the similarity of partitioning ratios. The residues found in the three subcellular fractions (17,300g pellet:105,000g supernatant:105,OOOg pellet) were approximately 1:2:1. This is closer to the lipid distribution, 2.2:2.3:1.0, than it is to the nucleic acids, 5.5:0:4.0, or to protein, 3:3:1. The hydroxyl group on the methine carbon atom of Kelthane increases its solubility in water. This explains the low amounts of pesticide residues observed in Kelthane fed rats, even considering the larger dose level used. The Kelthane present in the lipids did partition between the subcellular compartments in the proportions 1:2:1, the same as for DDT. The degrees of enzymic inhibition observed for DDT and Kelthane in vivo (Table III) do not correlate with the quantities of residues detected.

id&es in liver. A cause and effect relation-

ship between dietary DDT and the inhibition of anabolic fatty acid pathways is possible only if there is pesticide present in the microsomal membrane and if it is found in the subcellular fractions in the same proportions as all the microsomal lipids. No pesticide residues were detected in any fraction of the livers of rats fed commercial chow or EFA-deficient diets (Table IV). The content of pesticide residues in the commercial chow was less than the detectable limit. In animals fed 5 ppm o,p’DDT for 30 days, residues were detectable in all three fractions of liver organelles. After 60 days on the test diet, the residues

CONCLUSIONS 1. The effect of concentration of added o,p’DDT upon the chain elongation and desaturation of palmitic acid by liver microsomes from normal rats. Standard deviations of values for 12 to 16 measurements fell within the size of the circles marking the points. FIG.

The in vitro inhibition by DDT of desaturation and elongation of fatty acids by lipid-rich microsomal preparations suggest that the mode of action may be via incorporation of the chlorinated hydrocar-









of ol,g’--DT per gram of



5 ppm DDT in chow diet 5 ppm DDT in chow diet Control-chow diet only 5 ppm DDT, in EFA de% cient diet EFA deficient diet Extract of 100 g chow diet

105,OOog supernatant

105,oOOg pellet







30 days







60 days







60 days







60 days







60 days









No residue detected




bon into the membranes, altering their properties. The affinity of DDT for the lipid pool supports this conjecture. Fatty acid biosynthesis in rat liver microsomes is probably reduced significantly by DDT at all steps of desaturation and chain elongation of saturated fatty acids. The suppression of the desaturation of essential fatty acids, demonstrated here only at the step 20:306 + 20:406, tends to induce a functional EFA deficiency despite normally adequate intake of linoleic acid. Despite the normal intake of linoleic acid, the paucity of this acid in the microsomal phospholipids suggests that DDT also may inhibit the transfer of linoleic acid into phospholipids. The moderately severe EFA deficiency developed in these studies was induced by levels of pesticides which can and do occur in human foodstuffs. This suggests that chlorinated hydrocarbons chronically ingested at low levels may be another contributory cause of degenerative diseases associated with abnormal fatty acid composition of tissue lipids. They may contribute to diminished concentrations of PUFA in lipid components involved in normal transport of lipids and in normal metabolism of fatty acids. The generally accepted low acute toxicity of DDT and similar compounds may be misleading. The metabolic consequence of the low levels of DDT demonstrated here require a longer period for expression than is usually involved in acute experiments. That is, if EFA deficiency is one major consequence of DDT poisoning, chronic poisoning is more hazardous than is acute poisoning because sudden EFA deficiency has not been induced by even the most drastic nutritional conditions. The moderate inhibition of desaturation and elongation of EFA by DDT for a short time would be of little immediate consequence and would escape notice, but even a very low dosage of DDT for a long time would cause major changes in the pattern of polyunsaturated acids in tissues leading to EFA deficiency. Chronic intake of DDT induced EFA deficiency in rats as judged by dermal symptoms and classical abnormalties in pattern


of fatty acid composition of tissue lipids. Low levels of DDT intake may therefore be a stress, contributing with other factors to the abnormal lipid metabolism generally associated with the atherosclerosis syndrome. ACKNOWLEDGMENTS This work was supported in part by Public Health Service Grant AM 04524 from the National Institutes of Health; Public Health Service Grant HL 08214 from the Program Projects Branch, Extramural Programs; National Heart Institute; and by The Hormel Foundation. The authors are grateful to Dr. Robert Koch for valuable consultations. REFERENCES 1. WHO/FAO (1967) in Evaluation of Some Pesticide Residues in Food, Addendum, p. 63-66. 2. The Effects of Pesticides on Fish and Wildlife (1965) U.S. Dept. of the Interior, Fish and Wildlife Service, Circular 226. 3. BURNETT, R. (1971) Science 174, 606-608. 4. CONNEY, A. H., AND BURNS, J. J. (1972) Science 178, 576-586. 5. AZARNOFF, D. L., GRADY, H. J., AND SVOBODA, D. J. (1966) Biochem. Pharmacol. 15, 1985 1993. 6. STRAW, J. A., WATERS, I. W., AND FREGLY, N. J. (1965)Proc. Sot. Exp. Biol. Med. 118,391-394. I. HART, L. G., AND FOUTS, J. R. (1965) NaunynSchmiedebergs Arch. Pharmokol. Exp. Pathol. 249, 486-497. 8. CONNEY, A. H., WELCH, R. M., KUNTZMAN, R., AND BURNS, J. J. (1967) Clin. Pharmacol. Ther. 8, 2-10. 9. KOLOMODIN, B., AZARNOFF, D. L., AND SJOQUIST, F. (1969) Clin. Pharmacol. Ther. 10, 638-642. 10. POLAND, A., SMITH, D., KUNTZMAN, R., JACOBSON, M., AND CONNEY, A. H. (1970) Clin. Pharmacol. Ther. 11, 724-732. 11. MOSSER, J. L., FISHER, N. S., TENG, T-C., AND WURSTER, C. F. (1972) Science 175, 191-192. 12. BOWES, G. W., AND GEE, R. W. (1971) Bioenergetics 2, 47-60. 13. KOCH, R. B. (1969/70) Chem. BioZ. Interactions 1, 199-209. 14. DESIAH, D., CUTCOMP, L. K., AND KOCH, R. B. (1973) Life Sci. 13, 1693-1703. 15. DESIAH, D., CUTCOMP, L. K., AND KOCH, R. B. (1972) Biochem. Pharmacol. 21, 857-865. 16. KIMBROUGH, R. D. (1972)Arch. Environ. Health 25, 354-364. 17. BHATIA, S. C., AND VEN RITASIJBRAMANIAN, J. (1972) J. Agr. Food Chem. 20, 993-996. 18. GEYER, G. (1962) Acta Endocrinol. 40, 332-348.




IY. HOLMAN, R. T. (1971) Progress in the Chemistry of Fats and Other Lipids, Vol. 9, p. 308, Pergamon Press, Oxford. 20. PAIJLSRUD, J. R., STEWART, S. T., GRAFF, G., AND HOLMAN, R. T. (1970)Lipid.s 5,611-616. 21. CHANG, H. C., JANKE, J., PIJ~CX-I, F., AND HOLMAN, R. T. (1973) Biochim. Biophys. Actu 306, 21-25. 22. SHORT, C. R., MAINES, M. D., AND DAVIES, L. E. (1972)Proc. Sot. Exp. Biol. Med. 140,58-65. 23. LOWRY, 0. H., ROSEBROUGH, N. J., FARR, A. L., AND RANDALL, R. J. (1951) J. Biol. Chem. 193, 265-275. 24. BLIGH, E. G., AND DYER, W. J. (1959) Canad. J. Biochem. Physiol. 37, 911-917. 25. HOLMAN, R. T. (1960) J. Nutr. 70, 405-410. 26. COFFIN, D. E., AND SAVIARY, G. (1964) J. Assoc. Off. Agric. Chem. 47, 875-881. 27. KADOIJM, A. M. (1968) Bull. Environ. Toxicol. 3, 65-70. 28. STANLEY, R. L., AND LEFAVOURE, H. J. (1965) J. Assoc. Off. Agric. Chem. 48, 666-667. 29. KOVACS, M. F. (1963) J. Amer. Oil. Chem. Sot.





46, 884-893. 30. SCHNEIDER, W. C. (1955) [99] in Methods in Enzymology (Colowick, S. P., and Kaplan, N. O., eds.), pp. 680-685, Vol. III, Academic Press, New York. 31. MOHRHAUER, H., CHRISTIANSEN, K., GAN, M., DEUBIG, M., AND HOLMAN, R. T. (1967) J. Biol. Chem. 242, 4507-4514. 32, ELLIOTT, K. A. C. (1955) Ill in Methods in Enzymology (Colowick, S. P., and Kaplan, N. O., eds.) Vol. I, pp. 3-9, Academic Press, New York. 33. LEE, C. J., AND SPRECHER, H. (1971) Biochim. Biophys. Actu 248, 180-185. H. (1974) Biochim. Biophys. Actu 34. SPRECHER, 360, 113-123. 35. BRENNER, R. R. (1974) Mol. Cell Biochem. 3,4151. 36. KOCH, R. B. (1975), personal communication. 37. SNEDECOR, G. W., AND C~CHRANE, W. G. (1967) Statistical Methods 6th ed., pp. 59-61, Iowa State Univ. Press.