Acylation of cholesterol by microsomal enzymes from mouse preputial gland tumors

Acylation of cholesterol by microsomal enzymes from mouse preputial gland tumors

ARCHIVES OF BIOCHEMISTRY Acylation AND BIOPHYSICS of Cholesterol Mouse M. R. GRIGOR; Medical Division, 371375 (1972) 164, by Microsomal Pr...

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of Cholesterol Mouse

M. R. GRIGOR; Medical


371375 (1972)


by Microsomal





Oak Ridge Associated




Tumors’ AND



Oak Ridge, Tennessee 3’1880

Received December 9, 1971; accepted February 25, 1972 Cholesterol:acylCoA acyltransferase activity, assayed in subcellular fractions from mouse preputial gland tumors, was found to be microsomal. CoA and ATP were necessary for the esterification of cholesterol and palmitic acid, and the reaction was stimulated by Mg” and bovine serum albumin. The pH optimum was close to 8. The microsomal enzyme esterified t,he membrane cholesterol &s well a8 the cholesterol added to the incubation mixtures. Both cholesterol pools were esterified linearly with time, indicating that they equilibrate very rapidly to form a single pool accessible to the enzyme. The enzyme preferred palmitic acid or pentadecanoic acid as substrate over oleic and 1inoIeic acids. The cholesterol ester fraction isolated from preputial gland tumors had oleic acid as the most abundant acid, but also contained considerable amounts of a mixture of acids between Cl4 and C16.

The mouse preputial gland tumor is rich in st’erol esters, both in total quantity and in the variety of sterols esterified (1, 2). In this report, we describe some of the properties of the cholesterol: fatty acylCoA acyltransferase of preputial gland tumors and compare them with those previously reported for the same enzyme in rat liver (3-6), rat epidermis (7), and pigeon aorta (8). MATERIALS



of Enzymes

Preputial gland t.umors (ESR-586), removed 3-5 weeks after transplant,ation, were homogenized in 0.25 M sucrose as previously described (9). An International model PR-1 refrigerated centrifuge, a Servall model RC-2 centrifuge, and a Beckman model L3-50 centrifuge were used to obtain the following fractions: 600g X IO-min supernatant, 1 This investigation was supported in part by the American Cancer Society, Grant No. E-596B, and the U. S. Atomic Energy Commission. * On leave from the New Zealand Medical Research Council. 3 Oak Ridge Associated Universities Student Trainee from Northwestern University, Evanston, IL.

15,OOOgX IO-min sediment (mitochondrial fraction), 15,000g X lo-min supernatant, 109,OOOgX 1-hr sediment (microsomal fraction), and 100,OOOg X I-hr supernatant (soluble fraction). Each of the sediments was washed twice in 0.25 M sucrose. Prot.ein of each fraction was determined by the method of Lowry et al. (10). Succinate oxidase and cytochrome c:NADH reductase were assayed according to Eibl et al. (11) and Dallner (12), respectively. The cholesterol content of the microsomal fraction was determined by extracting the lipids (13), and measuring the cholesterol by photodensitometry (14) after tlc4 on Silica Gel G. The tic plates were developed with diethyl ether, sprayed with sulphuric acid, charred at [email protected]“C!, and the char quantitated using a Photovolt densitometer. In one experiment, the microsomes were labeled with [7-3H]cholesterol. The labeled cholesterol, dispersed in Tween 20, was injected directly int,o 4-week-old tumors (25 &i/tumor); after 14 hr, the tumor-bearing animals were killed, the tumors removed, and the microsomes prepared in the conventional

4 The abbreviations

@ 1972 by Academic



and washed


used are thin-layer

tography-tic; coenzyme A-CoA; phosphate-ATP; bovine serum gas-liquid chromatography-GLC. 371




adenosine trialbumin-BSA;




Incubations Incubations were carried out at 37°C using A Labline shaker. For each incubation, 0.5 ml of the enzyme preparation (frozen once), the substrates, and cofactors were made to a final volume of 3.0 ml in 0.1 M Tris buffer at pH 7.2 unless otherwise stated. The fatty acids or cholesterol were dissolved in ethanol, and 10 ~1 of the solution was added to the incubation mixtures. Unlabeled fatty acids and cholesterol were purchased from Analabs, Inc.; the labeled fatty acids ([l-‘4C]palmitic acid, (9,10-3H]palmitic acid, [1-%]oleic acid, and [l-r%] linoleic acid) and cholesterol([7-3H]-cholesterol and [4-%]cholesterol) were purchased from New England Nuclear or Amersham/Searle. CoA and ATP were obtained from P-L Biochemicals and the BSA was obtained from Sigma Chemical Co. The reactions were assayed by measuring the incorporation of radioactivity from the labelled substrates into cholesterol esters. All reactions were terminated by extracting the lipids (13). The radioactive lipid products were separated by tic on layers of Silica Gel G developed with hexane : diethyl ether:acetic acid (80:20:1 v/v) and visualizejl with iodine. Lipid classes were located according to a standard mixture cochromatographed with the radioactive sample. The radioactivity was assayed according to areas corresponding to each lipid class or by zonal profile analysis (15) in a Packard Tri-Carb liquid scintillation counter. We determined the fatty acid specificity of the enzyme for several fatty acid substrates by separating the sterol ester fraction according to the degree of unsaturation on layers of Silica Gel G containing 10% silver nitrate, which were developed in a solvent system of benzene: hexane (1:I v/v) (4)

Sterol Ester Analysis of Preputial Tumor Wax



The six fractions were saponified according to the method of Nikkari and Haaht,i (17). The nonsaponifiable fractions were analyzed by GLC in an Aerograph HiFi model 550 equipped with a flame ionization detector containing a column (5 ft X $$ in.) packed with 2.5% SE-30 on Beropak 30 (Varian Aerograph, Inc.). The column temperature was programmed from 150 to 250°C for the analysis of mixt,ures of fatty alcohols and sterols, and maintained isothermally at 250°C for the analysis of mixtures containing only sterols. The relative mass response of fatty alcohols and sterols, based on the responses of hexadecanol and cholesterol, was determined to be 2.5. The fatty acids isolated from the saponification mixtures were converted to methyl esters by heating in a sealed tube with 7% HCl in methanol at 100°C for 30 min. The methyl esters were analyzed in a Victoreen model 4000 chromatograph with dual flame ionization detectors. The unit was equipped with a column (6 ft X x in.) packed with 10%: EGSS-X on Gas Chrom P (Applied Science Labs, Inc.); column temperatures were programmed from 160 to 200” at 2”/min. RESULTS



Products Synthesized from Labeled Substrates Figure 1 illustrates the zonal profile scans of the labeled products formed from [1-*4C]palmitic acid and unlabeled cholesterol and


The sterol ester-wax ester fraction was isolated from the total lipids of 6-week-old tumors using silicic acid chromatography; 775 mg of lipids were applied to a 20-g column of silicic acid (Unisil, 100-200 mesh, Clarkson Chemical Co.) which had been washed with hexane, and the sterol and wax esters (500 mg) were eluted with 2% diethyl ether in hexane. Fifty milligrams of this ester fraction were then applied to a magnesium oxide column (adsorptive catalytic grade, 200 mesh, Matheson, Coleman, and Bell) and eluted first with 109 ml 0.570 acetone in hexane and then with 200 ml of 1 .O% acetone in hexane (16). Twenty-milliliter fractions were collected and combined into six fractions on the basis of their chromatographic behavior on layers of alumina developed with hexane:acetone (99: 1 v/v).



5 AB

10 C



FIG. 1. Zonal profile scan (5 mm) of incubation products. The tic plate was developed in hexane: diethyl ether:acetic acid (80:20:1 v/v). Radioactivity was incorporated from [l-14C]paImitic acid (0.25 pCi) and [7-3H]cholesterol (0.5 &i). The letters designate the following lipid classes: A, phospholipids; B, cholesterol; C, free fatty acids; D, triacylglycerols; and E, cholesterol esters. Samples were incubated in 3 ml of 0.1 M Tris buffer at pH 7.2 with t,he subst,rate for 60 min in a s stem that contained 10 PM CoA, 10 mM ATP, 4 mM MgCl2,5 mg BSA, 1.2 mg microsomal protein, 50 nmoles (18 pM) palmitic acid, and 50 nmoles (18 MM) cholesterol. (e), from palmitic acid; (A), from cholesterol.



from unlabeled palmitic acid and [7-3H]cholester01 when the substrates were incubated with the 100,OOOgsediment, and all cofactors (CoA, ATP, Mgz+, and BSA). Radioactivity from the [l-14C]palmitic acid was distributed in the phosphoglycerides, the triacylglycerols and the cholesterol esters, as well as in the fatty acid fraction. Most of the r4C-activity of the phosphoglyceride fraction was associated with phosphatidylcholine (61%). When [3H]cholesterol was the substrate, only the unreacted cholesterol and the cholesterol ester fraction contained 3H-activity. With identical substrate concentrations, much more radioactivity was found in the cholester01 ester fraction formed from the labeled fatty acid than from the labeled cholesterol. These data indicate that the membrane cholesterol caused considerable isotope dilution of the added labeled cholesterol. ~Subcellular Location The cholesterol esterifying activity of preputial gland tumors is primarily confined to the 100,OOOgparticulate fraction (microsomes) (Table I). The activity of the 15,000g particulate fraction (mitochondrial fraction) can be explained in terms of microsomal contamination, since, when cytochrome c: NADH reductase was measured in similar preparations as a marker enzyme for the microsomes, the specific activity of the mitochondrial fraction was almost 50 % of that found in the microsomal fraction. The cholesterol esterifying activity of rat liver is also microsomal (6). TABLE



Co-factor Requirewmts and pH Optimum The formation of cholesterol esters in the preputial gland tumor microsomes from fatty acids and cholesterol required CoA and ATP as cofactors and was stimulated by the addition of magnesium ions and bovine serum albumin. These results are similar to those reported for the esterification of cholesterol in liver (4, 5) and epidermis (7) of rats, and are consistent with the presence of a cholesterol: acylCoA acyltransferase. There was a broad pH optimum near 8 Effect of Substrate Concentration The rate of cholesterol ester synthesis increased as the concentration of palmitic acid was increased (up to 52 pbr), whereas at higher concentrations of acid (102 PM) the rate decreased (Fig. 2). The effect of varying the amount of added cholesterol was less marked. Since considerable synthesis of cholester01 esters was still observed without adding cholesterol, it would appear that cholesterol of the microsomal membranes must be accessible to the enzyme. Nature of Substrate When the acyltransferasc reaction was assayed with [3H]cholesterol, the quantity of



6OOg supernatant 15,000g sediment 15,000g supernatant 100,OOOgsediment 100,000g supernatant

@moles ester/mg protein/60 min) 0.9 3.7 0.8 9.9 0.3

a Co-factor and substrate concentrations were identical to those listed in the legend of Fig. 1; 0.25 pCi[S, lo-3Hlpalmitic acid was used as labeled substrate.

FIG. 2. Effect of exogenous substrate concentration on cholesterol ester from cholesterol (50 nmoles, 17 m) and varying fatty acid concentrations (0) and from palmitic acid (55 nmoles, 18 PM) and varying cholesterol concentrations (A). Other conditions were identical to those described in the legend of Fig. 1. [l-W]Palmitic acid (0.25 rCi) was used as the radioactive substrate. Each value represents the mean of duplicate incubations.








[1-W]Palmitate [7-*H]Cholesterol Assuming no isotope dilution Allowing for isotope dilution total microsomal cholesterol

(nmoles/mg microsomal protein/60 min) 3.9


0.8 3.2

a Fifty-five nmoles of each added to incubation mixtures. Other conditions were identical to those listed in the legend of Fig. 1.

cholesterol ester synthesized [assuming no equilibration with the endogenous (membrane) cholesterol] was only 20 % of that observed when [l-14C]palmitic acid was the substrate. Based on the cholesterol content of the microsomes (1509160 nmoles per incubation), the cholesterol ester synthesized from the labeled cholesterol was calculated to be similar to that synthesized from [lJ4C]palmitic acid (Table II). Thus, these data indicate that the membrane cholesterol and the added cholesterol act as a single substrate pool with respect to the rsterifying enzyme and that the equilibration between the two sources is rapid. Because the cholesterol content of the preputial gland tumor microsomes (at around 300 nmoles per mg protein) is several times greater than that previously reported for rat liver microsomes (18, 19), the isotope dilution effect observed in our studies is more pronounced than that found with the liver enzymes (6). The failure to observe acylation of exogenous cholesterol in aortic subcellular particles (8) may also be partly due to a very large isotope dilution by the endogenous cholesterol pool, since the cholesterol content of aortic microsomcs (20) is similar to that of the preputial tumor microsomes. To try and measure the rate of cquilibration between the membrane cholesterol and the added cholesterol, we performed an experiment using microsomal membranes labeled with cholesterol in viva. The mem-


FIG. 3. Effect of time on cholesterol ester synthesis from membrane cholesterol, labeled with [7JH]cholesterol (O), and from added [4-W]cholesterol (50 nmoles) (A). Other conditions were identical to those described in the legend of Fig. 1. Each value represents the mean of duplicate incubations.

branes labeled with [7-3H]cholesterol were incubated with added [4-14C]cholesterol, unlabeled palmitic acid, CoA, ATP, Mg”+, and BSA for different periods of time, and the extent of esterification of the membrane cholesterol and the added cholesterol was determined. The rate of esterification of both the cholesterol sources was linear up to 3 hr (Fig. 3). These data confirm that the cholesterol of microsomal membranes does act as substrate for the csterifying enzyme, and suggest that t’he equilibration between the membrane cholesterol and the added cholesterol is completed during the first minutes of the reaction. Dcykin and Goodman (21) have stated that there is also rapid equilibration of membrane cholesterol and exogenous cholesterol in rat’ liver parbiculate systems. Fatty Acid Specificity

The major fatty acids of a sterol ester fraction, containing 84 % cholesterol esters, obtained from 7-week preputial gland tumors were oleic (22.7 %) and a poorly resolved mixture of acids between Cl4 to Cl6 (32.3 %). Palmitic acid comprised 8.4% and linoleic acid only 2.7 % of the total fatty acids. The specificity of the esterifying enzyme system for fatty acids of different degrees of unsaturation was investigated in






Saturated Monounsaturated Diunsaturated



16:0, l&l, 18:2

mo, 1s: 1, 18:2

53.5” 19.3 16.7

63.5 15.3 12.4

5 Percent of total cholesterol ester activity; means of duplicate analyses. [7-3H]Cholesterol (55 nmoles) and 67 nmoles of each fatty acid were used as substrate. Incubations were run 2 hr. Other conditions were the same as those listed in the legend of Fig. 1.

an experiment where [3H]cholesterol was incubated with equimolar amounts of oleic acid and linoleic acid, plus either palmitic acid or pentadecanoic acid (67 nmoles each, 22 PM) for 2 hr. The radioactivity associated with the saturated, monounsaturated, and diunsaturated cholesterol esters as determined by silver nitrate-tic is shown in Table III. In both cases over half of the cholesterol ester activity cochromatographed with the saturated cholesterol esters, showing that the enzyme system was selective for saturated fatty acids. The higher amount of activity found in the saturated cholesterol esters in the incubations containing pentadecanoic acid suggests that, of the two saturated fatty acids tested, the pentadecanoic acid is the preferred substrate. Our results showing a preference for palmitic acid or pentadecanoic acid in vitro contrast with the observations reported for the esterifying enzyme from rat liver where oleic acid is the preferred substrate (4, 5).

1. KANDUTSCH, A. A., .~ND RUSSELL, A. E. (1960) J. Biol. Chem. 236,2256. 2. SNYDER, F., MALONE, B., AND BLANK, M. L. (1970) J. Biol. Chem. 246,179O. 3. MUKHERJEE, S., KUNITAKE, G., .IND ALFINSLATER, R. B. (1958) J. Biol. Chem. 230,91. 4. GOODMAN, D. S., DEYKIN, D., IND SHIRATORI, T. (1964) J. Biol. Chem. 239,1335. 5. SWELL, L., LAW, M. D., SND TREIDWELL, C. R. (1964) Arch. Biochem. Biophys. 104, 128. G. STOKKE, K. T., AND NORUM, K. R. (1970) Biochim. Biophys. Acta 210,kZ. ’ 7. FREINKEL, R. K., AND Aso, K. (1971) Biochim. Biophys. Acta 239, 98. 8. ST. CL.~IR, R. W., LOFL~ND, H. B., AND CLARKSON, T. B. (1970) Circ. Res. 27,213. 9. SNYDER, F., MALONE, B., AND WYKLE, R. L. (1969) Biochem. Biophys. Res. Commun. 34, 40. LO. LOWRY, 0. H., ROSEBROUGH, N. J., FARR, A. L., AND RANDALL, R. J. (1951) J. Biol. Chem. 193, 265. 11. EIBL, H., HILL, E. E., AND LANDS, W. E. M. (1969) Eur. J. Biochem. 9,250. 12. DALLNER, G. (1963) Acta Pathol. Microbial. Scand., Suppl. 166. 13. BLIGH, E. G., AND DYER, W. J. (1959) Can. J. Biochem. Physiol. 37,911. 14. PRIVETT, 0. S., BLANK, M. L., CODDING, D. W., AND NICKELL, F. C. (1965) J. Amer. Oil Chem. Sot. 42, 381. 15. SNYDER, F., AND KIMBLE, H. (1965) Anal. Biothem. 11, 510. 16. NICOLAIDES, N. (1970) J. Chromatogr. Sci. 8, 717. T., AND HAAHTI, E. (1964) Acta 17. NIKKARI, Chem. Stand. 18, 671. 18. PASCAUD, A., AULIAC, P., AND PASCAUD, M. (1968) Biochim. Biophys. Acta 160,326. 19. COLBEAU, A., NACHBAUR, J., .IND VIGNAIS, P. M. (1971) Biochzm. Biophys. Acta 249, 462. 20. PORTMAN, 0. W., ALEXANDER, M., -*ND OSUGA, T. (1969) Biochim. Biophys. Acta 167,435. 21. DEYKIN, D., AND GOODMAN, D. S. (1962) Biothem. Biophys. Res. Commun. 8,411.