Cholesterol ester hydrolase mediated conjugation of haloethanols

Jan 15, 1990 - 2,2,2-trichloroethanol, and 2-bromoethanol were obtained from. Aldrich (Milwaukee, WI). Fatty acids and fatty acid ethyl esters were pu...
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Chem. Res. Toxicol. 1990,3, 311-317

311

Cholesterol Ester Hydrolase Mediated Conjugation of Haloethanols with Fatty Acids Hari K. Bhat and G. A. S. Ansari* Divisions of Biochemistry and Chemical Pathology, University of Texas Medical Branch, Galveston, Texas 77550 Received January 15,1990 The formation of fatty acid conjugates of haloethanols was studied under in vitro conditions by using purified bovine pancreatic cholesterol ester hydrolase (EC 3.1.1.13). The enzymatic formation of 2-chloroethyl and 2-bromoethyl esters of oleic, linoleic, linolenic, and arachidonic acids was c o n f i i e d by proton nuclear magnetic resonance spectroscopy and chemical ionization mass spectrometry. 2-Bromoethanol was a better substrate than 2-chloroethanol for fatty acid esterification using cholesterol ester hydrolase. Among the chloroethanols, 2-chloroethanol was a better substrate than 2,2-dichloroethanol and 2,2,2-trichloroethanol. The saturated fatty acids (palmitic and stearic) showed a small amount of ester formation when cholesterol ester hydrolase was used. The kinetics of haloethanol and oleic acid incorporation into haloethyl oleate catalyzed by cholesterol ester hydrolase were determined. In vitro experiments were also conducted to study the conjugation of haloethanols with fatty acids using rat liver microsomes. The saturated fatty acid (palmitic) was more reactive compared to unsaturated fatty acid (oleic) when haloethanols were used. The results using rat liver microsomes were in contrast to those obtained when cholesterol ester hydrolase was used. The synthesis, purification, and characterization of 2-chloroethyl and 2-bromoethyl esters of oleic, linoleic, linolenic, and arachidonic acids are also described.

Introduction Haloethanols (HEs)' are highly toxic, yet widely used industrial chemicals (I). HEs are common contaminants in food products and medical and surgical supplies (2,3) because ethylene oxide, one of the precursors of HEs, is frequently used for the fumigation of food products and sterilization of surgical supplies (4).HEs are also formed in biological systems from other precursors such as vinyl chloride, a widely used toxic industrial chemical, which results in the formation of 2-chloroethanol (2-CE) (5). Metabolism of several compounds such as 1,2-dichloroethane, 2,2-dichloroethyl ether, and 2-chloroethyl nitrosoureas (a family of anticancer drugs) also results in the formation of 2-chloroethanol (6-12). 2-CE causes a significant reduction in the activities of drugmetabolizing enzymes and also results in the decrease of the activity of glucose-6-phosphatase (13).2-Bromoethanol (2-BE) has been shown to induce lipid peroxidation in tissue slices (14). Acute oral administration of 2-CE results in a significant decrease in the mitochondrial elongation of fatty acids (15). HEs are weak uncouplers of oxidative phosphorylation, stimulate state 4 respiration, and suppress state 3 respiration (16,17).Johnson (1967) has shown that 2-CE, when given orally to rats, resulted in rapid depletion of liver glutathione and formation of S-(carboxymethyl)glutathione (18). Haloalcohols have been shown to be effective antifertility agents in a number of species (19). Recent studies suggest that HEs are metabolized to ethylene oxide and also conjugate with fatty acids to form haloethyl esters of fatty acids (HEEs) (20-23).High levels of HEEs have been found in foods (24).Recently, HEEs have been isolated and identified from livers of rats treated with 2-CE and 2-BE (!25,26).2-CE has been shown to be released from HEEs in model digestive systems, which is *To whom correspondence should be addressed. 0893-228x/90/2703-0311$02.50/0

the focus of toxicological concern associated with HEE residues in foods (27). Since the conjugation of haloethanols with fatty acids results in the formation of more lipophilic products, the possibility that the toxicity associated with HEs may be influenced partly by the long retained lipophilic HEEs should not be ruled out. In order to understand the role of enzymes in the formation of fatty acid esters of haloethanols, we studied the involvement of cholesterol ester hydrolase and also that of the rat liver microsomes. Recently, Lange (1982) has shown that ethanol can be nonoxidatively metabolized to fatty acid ethyl esters using cholesterol ester hydrolase (28).Leighty and Fentiman (1981)have shown the conjugation of trichloroethanol with palmitic acid using rat liver microsomal preparations (29). In the present study, we have shown the enzymatic formation of 2-CE and 2-BE esters of different fatty acids under in vitro conditions.

Experimental Procedures Materials. Purified bovine pancreatic cholesterol ester hydrolase (sterol-ester acyl hydrolase, EC 3.1.1.13) was purchased from Sigma (St. Louis, MO). 2-Chl0roethan01,2,2-dichl0roeth~n01, 2,2,2-trichloroethanol,and 2-bromoethanol were obtained from Aldrich (Milwaukee,WI). Fatty acids and fatty acid ethyl esters were purchased from Sigma. lJ4C-Labeled fatty acids were purchased from New England Nuclear (Boston, MA). [4-"C]Cholesterol (specific activity 60.0 mCi/mmol; chemical purity 99%) and [4-14C]cholesteryloleate (specific activity 58.0 mCi/ mmol; chemical purity 98%) from New England Nuclear were used. Silica gel coated gass platea (20 x 20 cm,adsorbent thickness 250 and 500 Mm) were purchased from Analtech (Newark, DE). All other reagents used were of the highest purity grade comAbbreviations: HE, haloethanol; 2-CE, 2-chloroethanol;2-BE, 2bromoethanol; HEE, haloethyl eaters of fatty acids; HPLC, high-performance liquid chromatography;TLC,thin-layer chromatography;'H NMR, proton nuclear magnetic resonance apectrwopy; CIMS, chemical ionization mass spectrometry;CEH, cholesterol ester hydrolase. 0 1990 American Chemical Society

Bhat and Ansari

312 Chem. Res. Toxicol., Vol. 3, No. 4, 1990

(I) X = Br (11) x = CI

(111) X = Br (IV) X = C I

(V) X = B r (VI) X = C I

The reaction was carried out in 20-mL screw cap glass vials flushed with nitrogen and heated at 50 OC for 36 h in the dark. The reaction mixture was extracted with three 20-mL portions of n-hexane, and the solvent was removed under reduced pressure. The synthesized esters were further purified by column chromatography [glass column, 39 cm (height) X 2.5 cm (diameter), packed with aluminum oxide, 21 cm (height) (J. T. Baker, Philipsburg, NJ)]. Chloroform/hexane (1:9 v/v) was used as an eluting solvent, at a flow rate of 2 mL/min. Fivemilliliter fractions were collected and analyzed by thin-layer chromatography (TLC) on silica gel, with hexane/diethyl ether/methanol/acetic acid (90:20:52v/v) as the mobile phase. Pure haloethyl esters of fatty acids started to elute from fraction nine. Fractions WO, which showed single spots on TLC, were pooled, evaporated to dryness under reduced pressure, and analyzed by 'H NMR and CIMS. All the fatty acid esters formed were liquids at room temperature. 'H NMR values and mass spectral data for different compounds are given below. 2-Bromoethyl oleate (I): 'H NMR 6 5.34 (t, 2 H, CH-CH, J = 5.5 Hz), 4.38 (t, 2 H, OCH,, J = 6.0 Hz), 3.50 (t,2 H, CH2Br, J = 6.0 Hz), 2.34 (t, 2 H, CH2C00, J = 7.5 Hz), 2.0 (bm, 4 H, C8-H and Cll-H), 1.63 (t, 2 H, C3-H, J = 7.3 Hz), 1.55 (water impurity), 1.3 (20 H, other methylene protons), and 0.88 (t, 3 H, Cle-H, J = 6.8 Hz); CIMS 406/408 (M 18,1:1), 389/391 (M 1, l:l),281 (M - CH2CH2Br),264 (M - BrCH2CH20H),222 [M - CH2=C'+(OH)OCH2CH2Br], 166/168 [CH2=C'+(OH)OCH2CH2Br,1:0.8], and 124/126 (BrCH2CH20H,1:0.7). 2-Chloroethyl oleate (11): 'H NMR 6 5.34 (t,2 H, CH-CH, J = 5.5 Hz), 4.32 (t, 2 H, OCHZ, J = 5.6 Hz), 3.67 (t, 2 H, CH2C1, J = 5.7 Hz), 2.35 (t,2 H, CHZCOO, J = 7.5 Hz), 2.0 (bd,4 H, C8-H and Cll-H), 1.63 (bm, 2 H, C3-H),1.56 (water impurity), 1.3 (20 H, other methylene protons),and 0.88 (t,3 H, ClrH, J = 6.8 Hz); CIMS 362/364 (M + 18, 3:1), 345/347 (M + 1, 3:1), 281 (M CH2CH2Cl), 265 (M 1 - ClCH2CH20H), 264 (M C1CH2CH20H),222 [M - CH2=C'+(OH)OCH2CH2Cl],and 122/124 [CH2=Co+(OH)OCHzCH2C1,3:1]. 2-Bromoethyl linoleate (111): 'H NMR 6 5.34 (m, 4 H, CH=CHCH&H=CH), 4.38 (t, 2 H, OCH2, J = 6.0 Hz), 3.50 (t, 2 H, CH2Br,J = 6.0 Hz), 2.77 (t, 2 H, Cll-H, J = 5.7 Hz), 2.34 (t,2 H, CH2CO0,J = 7.5 Hz), 2.0 (m, 4 H, C8-Hand C14-H), 1.62 (bm, 2 H,C3-H),1.31 (14 H, other methylene protons), and 0.88 (t, 3 H, CIS-H, J = 6.8 Hz); CIMS 404/406 (M + 18, Ll), 387/389 (M + 1, l:l),307 (M - Br), 279 (M - CH2CH2Br),263 (M + 1BrCH2CH20H),262 (M - BrCH2CH20H),and 220 [M - CH2= C'+(OH)OCH2CH2Br]. 2-Chloroethyl linoleate (IV): 'H NMR 6 5.34 (m, 4 H, CH=CHCH,CH=CH), 4.32 (t, 2 H, OCH2, J = 5.6 Hz), 3.67 (t, 2 H, CH2C1, J = 5.7 Hz), 2.77 (t, 2 H, Cii-H, J = 5.3 Hz), 2.34 (t, 2 H, CH2CO0,J = 7.3 Hz), 2.0 (m, 4 H, C8-Hand C14-H), 1.64 (bm, 2 H, C3-H),1.31 (14 H, other methylene protons), and 0.89 (t, 3 H, ClgH, J = 6.8 Hz); CIMS 360/362 (M 18,31), 342/344 (M, 3 ~ 1263 ) ~ (M + 1- ClCH2CH2OH),262 (M - ClCH2CH2OH), and 220 [M - CH2=C*+(OH)OCH&H2Cl]. 2-Bromoethyl linolenate (V): 'H NMR 6 5.39 (m, 6 H, CH=CHCH&H=CHCH2CH=CH), 4.41 (t,2 H, OCHZ, J 6.2 Hz),3.53 (t, 2 H, CH2Br,J = 5.8 Hz),2.84 (t,4 H, Cll-H and C14-H, J = 5.5 Hz), 2.37 (t, 2 H, CH2CO0, J = 7.3 Hz), 2.11 (bm, 4 H, C8-Hand C17-H), 1.64 (m, 2 H, C3-H),1.35 (8 H, other methylene protons), and 1.01 (t, 3 H, C18-H,J = 7.3 Hz); CIMS 402/404 (M + 18, l:l), 384/386 (M, l:l),and 261 (M + 1- BrCH2CH20H). 2-Chloroethyl linolenate (VI): 'H NMR 6 5.35 (m, 6 H, CH=CHCH&H=CHCH&H=CH), 4.32 (t, 2 H, OCHZ, J = 5.8 Hz),3.66 (t, 2 H, CH2C1, J = 5.8 Hz),2.80 (t,4 H, C11-H and CII-H, J = 5.8 Hz), 2.34 (t, 2 H, CH2CO0, J = 7.3 Hz), 2.06 (m, 4 H, C8-H and C17-H) 1.63 (m, 2 H, C3-H),1.31 (8 H, other methylene protons), and 0.97 (t, 3 H, Cls-H, J = 7.3 Hz); CIMS 358/360 (M + 18,3:1), 340/342 (M, 3:l) and 261 (M 1 - ClCH2CH20H). 2-Bromoethyl arachidonate (VII): 'H NMR 6 5.37 (m, 8 H, CH=CHCH2CH=CHCH2CH=CHCHZCH=CH), 4.38 (t, 2 H, OCH,, J = 6.2 Hz), 3.50 (t, 2 H, CH2Br,J = 6.2 Hz), 2.81 (m, 6 H, C7-H, Cio-H, and Ci,-H), 2.36 (t, 2 H, CH2CO0, J = 7.7 Hz), 2.11 (bm, 4 H, C4-Hand Cle-H), 1.72 (m, 2 H, C3-H), 1.30 (6 H, other methylene protons), and 0.88 (t,3 H, C,-H, J = 6.9 Hz); CIMS 428/430 (M + 18, 1:l)and 411/413 (M + 1, 1:l). 2-Chloroethyl arachidonate (VIII): 'H NMR 6 5.37 (bm, 8 H, CH=CHCH&H=CHCH&H=CHCH&H=CH), 4.32 (t,

+

(vll) X = Br (vlll) x = CI

Figure 1. Structural formulas of 2-bromoethyl and 2-chloroethyl esters of different fatty acids (I, 2-bromoethyl oleate; 11, 2chloroethyl oleate; 111, 2-bromoethyl linoleate; IV, 2-chloroethyl linoleate, V, 2-bromoethyl linolenatq VI, 2-chloroethyl linolenatq VII, 2-bromoethyl arachidonate; and VIII, 2-chloroethyl arachidonate). When X = H in the above structures, then the ethyl esters of corresponding fatty acids are represented. mercially available, and all solvents used were of high-performance liquid chromatography (HPLC) grade. Instrumentation. High-resolutionproton nuclear magnetic resonance ('H NMR) spectra were acquired by using a JEOL GX 270 WB Fourier transform NMR spectrometer (6.3 TI. The samples were dissolved in deuteriochloroform, and tetramethylsilane was used as an internal reference. A nominal 0.1-Hz exponential line broadening apodization was applied prior to Fourier transformations. COSY data were accumulated as a 1024 X 266 data matrix and zemfilled to yield a 512 X 512 transformed matrix for plotting. The data were apodized with a shifted sine bell window and are shown in the absolute value mode, with high-resolution spectra plotted in place of projections. The samples were run unspinning, variable temperature controlled at 30 "C with 128 acquisitions per block. Chemical ionization mass spectrometry (CIMS) analyses were carried out with a Nermag R10-1OC mass spectrometer equipped with a PDP 11/73 data system. The mass spectrometer was operated under positive chemical ionization mode with ammonia as the reagent gas (gas pressure lo-' Torr). The sample was dissolved in chloroform (usually 1-2 pL) and placed on a tungsten filament probe tip. The solvent was allowed to evaporate, and the probe waa inserted into the ion source, set at 1.1A. The probe temperature was programmed from 0 to 500 mA at 20 mA/s. Reversed-phase HPLC was performed on a Beckman Model 334 liquid chromatograph connected with an Ultrasphere ODS column (5 particle size,25 cm X 4.6 mm i.d.; Altech knsociates; Deerfield, IL),a Beckman 165 variable-wavelength detector set at 210 nm, and a Perkin-Elmer 023 recorder. Methanol/water (232 v/v) at a flow rate of 1 mL/min was used as the mobile phase. Synthesis, Purification, a n d Characterization of Haloethyl Esters of Fatty Acids. 2-Chloroethyl and 2-bromoethyl eaters of palmitic and stearic acids were synthesized as described 26, 30). 2-Chloroethyl and 2-bromoethyl esters of earlier (25, unsaturated fatty acids, oleic, linoleic, linolenic, and arachidonic acids (Figure l), were synthesized by the following procedure. To 2 g of unsaturated fatty acid were added 10 mL of 2-chloroethanol or 2-bromoethanoland 4 drops of concentrated hydrochloric acid.

+

+

+

+

Cholesterol Ester Hydrolase 2 H, OCH2, J = 5.8 Hz), 3.66 (t, 2 H, CH2C1, J = 5.5 Hz), 2.80 (m, 6 H, CTH, Clo-H, and Cla-H), 2.36 (t, 2 H, CH2C00, J = 7.7 Hz), 2.11 (m, 4 H, C4-H and CIB-H),1.72 (m, 2 H, C3-H), 1.30 (6 H, other methylene protons), and 0.88 (t,3 H, Cm-H,J = 6.9 Hz); CIMS 384/386 (M + 18,3:1), 367/369 (M+ 1,31), and 287 (M + 1 - ClCH2CH2OH). In Vitro Experiments Using Cholesterol Ester Hydrolase. In vitro experiments were conducted by adding 40 pL of cholesterol ester hydrolase (180 pg) to 1.0 mL of 50 mM phosphate buffer (pH 7.35) containing 2 pmol of l-W-fatty acid (275 dpm/nmol) and 750 pmol of 2-chloroethanol or 2-bromoethanol and incubated a t 37 "C for 2 h. Control experiments were performed by omitting the enzyme while similar incubations were also carried out with ethanol (750 pmol), which served as a positive control (28). The reaction was stopped by the addition of chloroform (4 mL), and the lipids were extracted twice with 4-mL portions of chloroform. The organic phase was concentrated under reduced pressure, and fatty acid esters were separated from fatty acids by silica gel TLC as described above. The haloethyl esters were further purified by reversed-phase HPLC. The fractions having the retention times corresponding to that of the standard haloethyl esters were collected and analyzed by lH NMR and CIMS. Quantitation of the esters was achieved by scraping the haloethyl ester region from TLC plates and counting the 14C activity on a Searle Mark I11 liquid scintillation counter (with an automatic quench correction) after the addition of 0.5 mL of methanol/water (23:2 v/v), 0.5 mL of chloroform, and 20 mL of Tru Count (Tru Lab Supply Co., Libbertyville, IL). Background activity in the ester region was determined by dissolving the 14C-fattyacid in 1.0 mL of phosphate buffer (50 mM, pH 7.35) and extracting immediately with 4 mL of chloroform. The extracted fatty acid was applied on the TLC plate, and 14Cactivity in the ester region was counted. In Vitro Experiments Using Rat Liver Microsomes. Rat liver microsomes were prepared essentially by the method of De Duve (1971; 32)in 50 mM potassium phosphate buffer (pH 7.35). The incubation mixture consisted of 2 mL of microsomal preparation (1mg of protein/mL) in 50 mM phosphate buffer (pH 7.35), 2 pmol of fatty acid (275 dpm/nmol), and 750 pmol of 2-chloroethanol or 2-bromoethanol. Incubation was done a t 37 "C for 2 h. Control experimentswere performed by omitting the microsomes. Protein was determined by the method of Bradford using bovine serum albumin as a standard (32).The reaction was stopped by the addition of chloroform, the lipids were extracted, and haloethyl esters were quantitated as described above. Cholesterol ester hydrolase activity of the microsomes was determined by the method of Kritchevsky and Kothari (33). Hydrolytic activity was determined by using a micellar substrate containing 0.61 pmol of [4-14c]cholesteryloleate (2000 dpm/nmol), 1.5 pmol of sodium taurocholate, 1.52 pmol of phosphatidylcholine, and 1 mg of microsomal protein in 1 mL of 50 mM phosphate buffer (pH 7.0). The incubation was carried out a t 37 "C for 2 h. The reaction was stopped by the addition of chloroform, and lipids were extracted and separated on silica gel TLC using hexane/diethyl ether/acetic acid (7551 v/v) as the mobile phase. The amount of cholesterol liberated was quantitated by liquid scintillation counting of 14Cactivity. Synthetic activity was assayed by using an emulsion containing 2.06 pmol of [4-14C]cholesterol (400 dpm/nmol) 6.2 pmol of oleic acid, 4.13 pmol of sodium taurocholate, 13.33 pmol of ammonium chloride, and 1 mg of microsomal protein in 1.0 mL of 50 mM phosphate buffer (pH 6.0.), at 37 "C for 2 h. The reaction mixture was processed as described for the hydrolytic activity, and the amount of cholesterol ester formed was determined by scintillation counting. The hydrolytic and synthetic activity of purified cholesterol ester hydrolase was similarly determined by using 180pg of the enzyme instead of rat liver microsomes. Statistical Significance. Significancewas determined by two-tailed Student's t test.

Results Table I shows the formation of 2-bromoethyl, 2-chloroethyl, and ethyl esters of different fatty acids using cholesterol ester hydrolase (CEH). Among the two haloethanols, 2-BE was a better substrate for fatty acid es-

Chem. Res. Toxicol., Vol. 3, No. 4,1990 313 Table I. Formation of Ethyl, 2-Bromoethyl, and 2-Chloroethyl Esters of Different Fatty Acidso Using Bovine Pancreatic CEHb 2-bromoethyl 2-chloroethyl ethyl 9.71 f 1.01 36.64 f 1.11 palmitic 11.49 f 1.32 (1.07 f 0.04) (1.48 f 0.16) (1.17 f 0.10) 5.11 f 1.19 27.86 f 4.32 stearic 13.91 f 2.01 (2.55 f 0.19) (0.51 f 0.04) (0.78 f 0.06) oleic 387.26 f 29.35 283.99 f 9.97 54.13 f 2.78 (2.59 f 0.13) (0.76 f 0.16) (0.27 f 0.01) linoleic 208.55 f 20.97 197.56 f 15.39 48.35 f 2.55 (3.43 f 0.21) (3.51 f 0.15) (2.02 f 0.62) linolenic 49.13 f 6.45 148.38 f 9.08 18.46 f 0.47 (2.95 f 0.02) (1.30 f 0.19) (0.45 f 0.02) arachidonic 81.51 f 4.26 232.18 f 11.11 17.61 f 2.05 (4.75 f 0.69) (1.56 f 0.40) (0.67 f 0.06) OIn units of nmol of ester formed/(h-mg). bThe data are presented as an average of three experiments plus or minus the standard deviation. The data in parentheses indicate control values f SD. Control experiments were done by omitting the enzyme.

+

a

b

c

d

s

f

9

h

Figure 2. An autoradiogram of the enzymatic formation of 2-bromoethyl esters of different fatty acids using cholesterol ester hydrolase (a, b = oleic; c, d = linoleic; e, f = linolenic; and g, h = arachidonic acid; a, c, e, and g = enzymatic, and b, d, f, and h = control). After incubation of 2-bromoethanol with different individual fatty acids, the lipids were extracted with chloroform, and fatty acids were separated from fatty acid esters by TLC. The TLC plate was then exposed to an Industrex AA (Kodac) X-ray film. After exposure (2 weeks, room temperature) the film was developed by standard photographic techniques.

terifkation than 2-CE. When the different fatty acids were compared to each other, oleic acid showed the highest activity. Among the 2-chloroethyl esters, the ability of CEH to form fatty acid esters decreased from oleic acid to arachidonic acid. The saturated fatty acids (palmitic and stearic) showed the least reactivity to form 2-chloroethyl esters. Among the 2-bromoethyl esters, the activity decreased from oleic to linolenic acids, but arachidonic acid showed a slightly higher reactivity compared to linolenic acid. There was either slight or no formation of haloethyl esters when CEH was omitted from the incubation mixture (Figure 2). Among the chloroethanols, 2-chloroethanol was a better substrate for oleic acid esterification than either 2,2-dichloroethanolor 2,2,2-trichloroethanol (Figure 3) The enzymatic formation of 2-bromoethyl and 2chloroethyl esters of oleic, linoleic, linolenic, and arachidonic acids was confirmed by 'H NMR and CIMS. lH NMR of enzymatically formed esters matched that of the synthetic standards. The representative lH COSY spectra for enzymatically formed 2-bromoethyl arachidonate and that of the synthetic standard are shown in Figure 4. The two spectra match with each other, the only difference being the presence of a peak at 1.56 ppm in the enzymatic sample, which may be because of moisture present in the biological sample. The mass spectra of enzymatically

314 Chem. Res. Toxicol., Vol. 3, No. 4, 1990

Bhat and Ansari

125

100

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,

1

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L 75 0

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0 MCEO

DCEO

TCEO

b

Figure 3. Percent mono-, di-, and trichloroethyl oleate formed by using cholesterol ester hydrolase. The values given are a mean of three experiments plus or minus the standard deviation and are plotted after subtracting the control values from the experimental values. The absolute values of the nmol of ester formed after 2-h incubation are monochloroethyl oleate (MCEO),16.25; dichloroethyl oleate (DCEO), 5.34; and trichloroethyl oleate (TCEO), 1.12.

formed 2-bromoethyl and 2-chloroethylesters of different unsaturated fatty acids were similar to that of the standard HEEs. Figure 5 shows representative mass spectra of 2-bromoethyl oleate and 2-chloroethyl oleate. The kinetics of haloethanol and oleic acid incorporation into haloethyl oleate catalyzed by CEH were determined. With the 2-bromoethanol or 2-chloroethanol concentration constant at 750 mM and the oleic acid concentration varied from 0.25 to 20 mM, a calculated K', of 2.20 mM and V'of 14.38 nmol/(min.mg of protein) with respect to bromoethanol and a K L of 4.76 mM and a Vh,- of 6.18 nmol/ (min-mgof protein) with respect to chloroethanol were obtained. These results were compared with ethanol and oleic acid incorporation into ethyl oleate, which showed a K6,of 1.29 mM and V'- of 169 nmol/(min.mg of protein) with respect to ethanol. When oleic acid concentration was kept constant at 5 mM and the concentration of 2-bromoethanol or 2-chloroethanol varied from 50 mM to 1 M, a Kh, of 401.6 mM and V i - of 8.95 nmol/ (min-mgof protein) with respect to 2-bromoethanol and a K6,of 148 mM and V&- of 3.37 nmol/(min.mg of protein) with respect to chloroethanol were obtained (Figure 6). Under similar conditions a K6, of 629 mM and Vh,= of 128 nmol/(min.mg of protein) with respect to ethanol were obtained. The formation of ethyl, 2-bromoethyl, and 2-chloroethyl esters of palmitic, stearic, and oleic acids using rat liver microsomes is shown in Table 11. The saturated fatty acid (palmitic)was more reactive compared to unsaturated fatty acid (oleic) when haloethanols were used (p C 0.0011 when 2-CE was used and p C 0.0018 when 2-BE was used). The difference was still significant when ethanol was used as a substrate (p C 0.0071,although the actual values were closer to each other.

Discussion Fatty acid conjugatesof several xenobiotic alcohols have been reported (34-36). Most of these xenobiotic alcohols, used as such or produced as a result of the metabolism of

Figure 4. Two-dimensional (COSY)'H NMR spectra (A, standard; B, enzymatic) of 2-bromoethyl arachidonate. Table 11. Formation of Ethyl, 2-Bromoethyl, and 2-Chloroethyl Esters of Different Fatty Acids Using Rat Liver Microsomesa ethyl 2-bromoethyl 2-chloroethyl palmitic 38.03 f 2.29 35.37 1.94 28.69 2.37 (1.17 0.10) (1.07 0.04) (1.48 0.16) 10.70 0.72 stearic 15.98 f 1.38 15.05 2.77 (0.78 f 0.06) (2.55 f 0.19) (0.51 0.04) oleic 31.14 1.25 18.33 0.97 16.27 f 1.05 (2.59 0.13) (0.76 0.16) (0.27 f 0.01)

*

*

**

* *

* *

OIn units of nmol of ester formed/(h.mg of microsomal protein). The data are presented as an average of three experiments plus or minus the standard deviation. The data in parentheses indicate control values f SD. Microsomes were omitted from the control experiments.

the parent compound, have been shown to conjugate with fatty acids under in vitro conditions using rat liver microsomes. However, in recent years many examples of xenobiotics acylated by fatty acids in vivo have been re-

Chem. Res. Toxicol., Vol. 3, No.4, 1990 315

Cholesterol Ester Hydrolase

-

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Figure 6. Double-reciprocalplot of cholesterol ester hydrolase catalyzed esterification of [W]oleicacid with 2-bromoethanol(A) and 2-chloroethanol (0).Oleic acid concentration was kept constant at 5 mM, and the concentration of 2-bromoethanol or 2-chloroethanol was varied from 50 mM to 1 M. Each point represents a mean of three separate experiments. Results are expressed as nmol of ester formed/(min-mg of protein). Incubation and separation procedures for haloethyl oleates are described in the text.

ported in the literature (25,37,38). The chloroethyl esters of capric, myristic, palmitic, and linoleic acids have been identified at levels up to 1400 ppm in many food samples, with 2-chloroethyl linoleate being the most abundant ester in all food samples tested (22). Trichloroethanol, a major metabolite of the known carcinogen trichloroethylene, has been shown to conjugate with palmitic acid under in vitro conditions using a coenzyme A fortified microsomal system (29).

The role of specific enzyme(s) in the formation of haloethanol conjugates of fatty acids has not been addressed,

so far. Lange and his co-workers in their work on the nonoxidative metabolism of ethanol have shown the formation of ethyl esters of different fatty acids, and they have suggested the role of CEH in the formation of such esters (28,40-42). CEH is a pH-dependent enzyme that can catalyze either esterification or hydrolysis of cholesterol esters and has been shown to be present in most of the tissues (43-47). Studies on the enzyme isolated from liver tissue have shown the presence of CEH activity in the lysosomal (48, 49), microsomal (50), and cytosolic (51) fractions. CEH plays an important role in the metabolism of exogenous cholesterol esters from lipoproteins (52) and in the utilization of endogenously formed sterol esters (53). Decreased activity of lysosomal CEH leads to the accumulation of cholesterol esters, which results in various human diseases (54, 55). We have studied the formation of fatty acid conjugates of haloethanols using purified CEH from bovine pancreas under in vitro conditions. It has been shown that CEHs from liver, intestine, and aorta show a high degree of similarity to that of the pancreatic enzyme (56-57). Therefore, results obtained with pancreatic enzyme may also explain the formation of such esters under in vivo conditions in other tissues. Our studies indicate that, among the fatty acid esters formed by using CEH, oleic acid shows the maximum reactivity in terms of its ability to conjugate with ethanol or haloethanols. The saturated fatty acids (palmitic and stearic) were less reactive under these conditions. Probably, unsaturation in the fatty acid moiety may have some role to play in the enzyme activity. However, as the unsaturation increases from oleic acid to linolenic acid (having the same number of carbon atoms), the ability of CEH to form fatty acid conjugates decreases. The same pattern was observed for the synthesis of cholesterol esters using CEH (44).Arachidonic acid shows a relatively better ability to form conjugates of 2-bromoethanol and ethanol

316 Chem. Res. Toxicol., Vol. 3, No. 4, 1990

compared to linolenic acid. In the case of 2-chloroethanol there is not a significant difference between linolenic and arachidonic acids (p > 0.65). The observed difference between linolenic and arachidonic acids in terms of their ability to form esters may be because of the differences in the carbon number and the degree of unsaturation. Studies with rat liver microsomes show a higher reactivity for palmitic acid (saturated) compared to oleic acid (unsaturated) when haloethanols were used for in vitro incubations. These results are in contrast to when CEH is used. The CEH hydrolytic activity of rat liver microsomes [ 1.85 nmol of cholesterol formed/(h.mg of protein)] was far less than that of the commercial CEH [253.74 nmol of cholesterol formed/(h.mg of protein)]. The synthetic activity of microsomes [2.05 nmol of cholesterol ester formed/(h*mgof protein)] was also less than that of the commercial preparation of CEH [10.23 nmol of cholesterol ester formed/(h*mgof protein)]. The lower specific activity of microsomal preparation is expected due to the low concentrations of the CEH activity in the crude sample. In addition, several other factors present in the microsomal preparation could be responsible for its lower specific activity. However, compared to the pure enzyme preparation, which showed far less synthetic activity than the hydrolytic activity, the hydrolytic and synthetic activities of the rat liver microsomal preparation were almost identical. Although we have used two different sources of the enzyme, the enzymatic properties of CEH from the rat pancreas and rat liver microsomes are similar (57), and comparable to those of bovine pancreatic CEH. The observed results using rat liver microsomes are different from the results obtained by using bovine CEH. The reactivity of oleic acid using microsomal preparation may be because of the CEH activity present in the microsomes. However, the reactivity of saturated fatty acids cannot be explained only on the basis of CEH activity of the microsomes. If the rat CEH responds to haloethanols in a manner similar to that of bovine CEH, then there may be more than one enzyme responsible for the conjugation of fatty acids with haloethanols. Further studies are needed to address the problem of differential reactivities of saturated and unsaturated fatty acids to form esters of xenobiotics.

Acknowledgment. This research was supported by Grant ES04815 awarded by the National Institute of Environmental Health Sciences. NMR and mass spectrometers were purchased through the grant funded by the M. D. Anderson Foundation, Houston, TX. Registry NO.CEH, 9026-00-0; 2-CE, 107-07-3;2-BE, 540-51-2; 2-chloroethyl linoleate, 25525-76-2; 2-bromoethyl linolenate, 127183-06-6;2-chloroethyl linolenate, 25525-73-9; 2-bromoethyl arachidonate, 127183-07-7; 2-chloroethyl arachidonate, 12718308-8; 2-bromoethyl linoleate, 127183-05-5; 2-bromoethyl oleate, 97006-98-9; oleic acid, 112-80-1; linoleic acid, 60-33-3; linolenic acid, 463-40-1; arachidonic qcid, 506-32-1; ethyl palmitate, 62097-7; ethyl stearate, 111-61-5; ethyl oleate, 111-62-6; 2-bromoethyl palmitate, 101885-48-7; 2-bromoethyl stearate, 102464-65-3; 2chloroethyl palmitate, 929-16-8; 2-chloroethyl stearate, 1119-75-1; ethyl linoleate, 544-35-4; ethyl linolenate, 1191-41-9; ethyl arachidonate, 1808-26-0; 2-chloroethyl oleate, 51479-39-1; 2,2-dichloroethyl oleate, 127207-02-7; 2,2,2-trichloroethyl oleate, 117973-23-6; 2,2-dichloroethaml, 598-38-9.

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