Endogenous xenobiotic enzyme levels in mammalian cells

Mutation Research, 261 (1991) 29-39

29

© 1991 Elsevier Science Publishers B.V. 0165-1218/91/$03.50 ADONIS 016512189100128E MUTGEN 01688

E n d o g e n o u s x e n o b i o t i c e n z y m e levels in m a m m a l i a n cells Douglas B. McGregor a,., Ian Edwards a C. Roland Wolf and William J. Caspary c

b Lesley M. Forrester b,**

a Inveresk Research International, Ltd., Musselburgh, EH21 7UB (Scotland), b Imperial Cancer Research Fund, Hugh Robson Building, George Square, Edinburgh, EH8 9XD (Scotland) and c Cancer Genetics and Molecular Pathology Section, National Institute of Environmental Health Sciences, National Institutes of Health, P.O. Box 12233, Research Triangle Park, NC 27709 (U.S.A.)

(Received 12 September 1990) (Revision received 15 January 1991) (Accepted 4 March 1991)

Keywords: Enzyme levels, endogenous xenobiotic; Xenobiotic metabolism, enzymes involved in; Mammalian cells, response to

chemicals; Glutathione-S-transferase

Summary The response of mammalian cell lines to chemicals depends, in part, on the exogenous activation system used for the induction of a biological response. This could be attributed to differences in the expression of enzymes involved in xenobiotic metabolism. We have measured the activities of benzo[a]pyrene hydroxylase, dimethylaminoazobenzene N-demethylase, catalase, superoxide dismutase, peroxidase and glutathione-S-transferase in human lymphoblast TK6, mouse lymphoma L5178Y, Chinese hamster ovary ( C H O ) and lung (V79) and mouse C 3 H 1 0 T 1 / 2 cell lines as well as in primary hepatocytes and $9 preparations of liver from male F344 rats. Nitroreductase was also measured in some of these preparations. H u m a n lymphoblast TK6 and mouse C 3 H 1 0 T 1 / 2 cells had the capacity to metabolize dimethylaminoazobenzene and the latter cell line also metabolized benzo[a]pyrene, indicating the presence of constitutive mono-oxygenase activity. Cytochrome P450 could not be detected spectrophotometrically in the cell lines. Western blot analysis indicated that P450 from the P450IIA family is expressed in C 3 H 1 0 T 1 / 2 cells. Reactivity was also observed with an antibody to P450IA2; however, the identity of this protein remains uncertain. Superoxide dismutase, catalase and peroxidase, which protect cells against oxygen radical damage, were found in all the cell lines and in rat hepatocytes and $9. The human lymphoblast TK6 cell line, however, had the least of each of these three enzymes. Glutathione-S-transferase activity was detected at varying levels in all cell types. Nitroreductase activity was high in $9 and Chinese hamster ovary cells and lower in mouse lymphoma and Chinese hamster V79 cells.

* Present address: International Agency for Research on Cancer, 150 cours Albert-Thomas, 69372 Lyon, Cedex 08 (France). ** Present address: Division of Molecular and Developmental Biology, Mount Sinai Hospital Research Institute, 600 University Avenue, Toronto, Ont. (Canada).

Correspondence: Dr. William Caspary, Cancer Genetics and Molecular Pathology Section, NIEHS, P.O. Box 12233, Research Triangle Park, NC 27709 (U.S.A.).

30 Mammalian cells in culture can be important indicators of the mutagenic activity of chemicals. However, the mutagenic response is, among other things, dependent on the manner in which the chemical is metabolically activated and deactivated by cellular enzymes (Gelboin and Wiebel, 1971). In mammals, chemicals are substrates for metabolizing enzymes which may modify them to mutagenic or carcinogenic products (Miller, 1970; Jerina and Daly, 1974; Sims and Grover, 1974). In their adaption from in vivo to culture conditions, cells lose many specialized or differentiated functions, including the expression of enzymes involved in xenobiotic metabolism. The mechanism of this loss of gene expression is unknown (Franks and Cooper, 1972; Sato et al., 1960; Franks and Wilson, 1970; Clark and Pateman, 1978). "Immortalized" cell lines derived from different tissues a n d / o r species appear to have similar enzyme activities. For example, Lieberman and Ove (1960) monitored four human cell lines for 13 enzymatic activities and found the activities to be similar. Peppers et al. (1960) showed that the catalase activity levels of long-term cultures derived from liver, epidermis, subcutaneous connective tissue, and sarcomatous and carcinomatous tumors were essentially identical, whereas they were only 0.5% that of normal liver. Mutagenic and toxic responses are other endpoints that have been used to examine the similarity of cell lines of different origins. Caspary et al. (1988) examined the mutagenic response at the tk locus in the mouse lymphoma L5178Y and the human lymphoblast TK6 cell lines. There was no evidence of different mutagenic responses to a series of 13 chemicals in the absence of $9, suggesting that, for these two cell lines at least, the species of origin had no effect on the mutagenic responses to chemicals studied. In contrast, Singh and Gupta (1985) and Gupta (1982, 1985) demonstrated that there were differences in both mutagenic and toxic responses to certain compounds of cells derived from the human (HeLa), mouse (3T3 and L M T K - ) and hamster (Chinese hamster ovary and M3-1) to certain compounds. Other studies also showed differences in cells to the mutagenic a n d / o r toxic effects of compounds (Hoffman et al., 1984; Coppinger et al., 1984; Aaron et al., 1980).

Because many cell lines activate promutagens either poorly or not at all (Gelboin and Wiebel, 1971; Wiebel and Gelboin, 1975), the addition of exogenous tissue enzymes is often required (Bimboes and Griem, 1976; Krahn and Heidelberger, 1977; Kuroki et al., 1977; Uemeda and Saito, 1975). The importance of enzymatic supplementation was made clear when Malling (1971) demonstrated that a nonmutagen, dimethylnitrosamine, could be metabolized to a mutagen in Salmonella typhimurium TA1530 and G46 by coincubating the compound with mouse liver microsomes. In mammalian cell mutation assays, a 9000 g supernatant fraction of Aroclor 1254-induced rat liver homogenate ($9) is commonly used to produce some of the metabolic reactions which would occur in vivo. It is important to be aware of the limitations of such homogenate systems, two major ones being that they are usually deficient in the cofactors required for conjugation reactions and that they are, with respect to the target cell, extracellular. The contribution of intracellular enzymatic pathways to determine the mutagenic response is often neglected (Gelboin and Wiebel, 1971). The cytochrome P450 monooxygenases are particularly important in the metabolism of xenobiotics. This metabolism includes oxidative and reductive mechanisms, resulting in the formation of arene oxides, hydroxyl derivatives, dealkylated products, nitroso and nitro compounds, sulphoxides, sulphones, etc. These products are often conjugated to either glucuronic acid or glutathione, a process which facilitates their excretion. Both of these conjugation pathways are important in determining the toxicity profiles of xenobiotics. Glucuronides are generally of low toxicity. Glutathione conjugates can be further metabolized to biologically active intermediates as well as low toxicity mercapturic acids. The glutathione-S-transferases account for up to 10% of the soluble protein from rat liver cytosol (Jacoby et al., 1976) and 3% of that from human liver (Kamisaka et al., 1975). Consequently, there has been a particular interest in glutathione conjugation. Another series of metabolic steps of toxicological importance involves active oxygen species, such as the superoxide ion and hydrogen perox-

31 ide. These are generated in a number of reactions, including some involving cytochrome P450 (Joenje, 1983). Superoxide anions are reduced to hydrogen peroxide by superoxide dismutase (McCord and Fridovich, 1969) and peroxide is dismutared to oxygen and water by catalase (Chance et al., 1979). These enzymes protect against chromosomal damage induced by active oxygen species (Sofuni and Ishidate, 1984), so their concentrations in different cells are important for the evaluation of toxic responses. As a contribution to the study of the response mechanisms in vitro the levels of activity of certain enzymes were measured in homogenates of five different cultured cell lines [mouse lymphoma (L5178Y), Chinese hamster ovary fibroblast (CHO), Chinese hamster lung fibroblast (V79) and mouse embryo fibroblast (C3H10T1/2)] and of primary hepatocytes from male F344 rats. For comparison, measurements were also made on rat liver $9 preparations. In this paper, we report the endogenous xenobiotic enzyme levels in cell lines commonly used to measure the mutagenic and carcingenic potential of chemicals. Materials and methods

Cell cultures The cell lines were cultured as follows: (1) mouse lymphoma L5178Y clone 3.7.2 C cells were grown in suspension in Fischer's medium containing 10% horse serum and 0.1% pluronic acid; (2) CHO K1-BH4 ceils were grown in suspension in MEM containing 10% fetal bovine serum; (3) C3H10T1/2 cells were grown as monolayers in Eagle's basal medium with Earle's salts containing 10% heat-inactivated fetal bovine serum: (4) Chinese hamster V79 cells were grown as monolayers in Eagle's MEM formula No. 84-5114 (Gibco Laboratories) containing 3% fetal bovine serum. Rat hepatocyte primary cultures were established following collagenase perfusion of liver in otherwise untreated adult male F344 rats anaesthetized with sodium pentobarbitone (50 mg/kg) and killed by cervical dislocation. Hepatocytes were maintained in Williams' medium supplemented with 10% calf serum and 10 -7 M dexamethasone.

Rat liver homogenates Rat liver $9 was prepared from male Fischer F344 rats (150-300 g) which were injected once i.p. with Aroclor 1254 (diluted in corn oil to 200 mg/ml) at a dosage of 500 mg/kg, 5 days prior to homogenate preparation. Food was withheld from the animals for 16 h before they were killed. Livers were removed into ice cold 0.15 M KC1, weighed and chopped. Materials used during the preparation of homogenates were kept at 0-4°C. Cell cultures were pelleted by centrifugation at 5000 g for 5 min and resuspended in ice cold 0.15 M KCI, then pelleted again and the gross wet weight of ceils determined. Cell culture pellets and chopped rat livers were homogenized in 3 ml ice cold 0.15 M KC1 per mg of cells or rat liver. Homogenization was performed in glass homogenization vessels by eight strokes of a teflon pestle rotating at approximately 1200 rpm. Rat liver homogenate was then centrifuged at 9000 g for 10 min. Aliquots of rat liver 9000 g supernatant and the cell homogenates were stored in liquid nitrogen.

Enzyme assays, materials Common laboratory reagents, solvents, 1chloro-2,4-dinitrobenzene, L-cysteine • HC1, pnitrobenzoic acid, pyrogallol, quinine sulphate and trichloroacetic acid were obtained from BDH Ltd., Poole, Dorset, England. Benzo[a]pyrene [B(a)P] was obtained from Lancaster Synthesis Ltd., Lancaster, England. Dimethylaminoazobenzene (DAB) was a gift from the Central Toxicology Laboratory, ICI Ltd., Macclesfield, Cheshire, England. NADP, NADPH, NADH, glucose-6phosphate and xanthine oxidase (EC.1.1.3.22) were obtained from Boehringer Mannheim Ltd., Lewes, Sussex, England. Acetylacetone, paminobenzoic acid, ammonium sulphamate, cytochrome c, FMN, reduced glutathione, glutathione-S-transferase, N(1-naphthyl)ethylenediamine.2HCl and superoxide dismutase were obtained from Sigma Chemical Co. Ltd., Poole, Dorset, England. Spectrophotometric measurements were made using a Philips/Pye Unicam PU 8800 spectrophotometer and fluorescence measurements were made using a Perkin-Elmer LS-3 fluorimeter.

32 Protein concentrations were determined by the method of Lowry et al. (1951).

Enzyme assays, methods Benzo[a]pyrene hydroxylase. B(a)P hydroxylase activity was determined by a method modified from that as previously described (Wiebel and Gelboin, 1975; Wattenberg et al., 1962; Nebert and Gelboin, 1968), in which the fluorescence of phenolic products is measured, using quinine sulphate as a fluorescence standard calibrated against 3-OH-B(a)P. The assay was performed in 1 ml 0.05 M Tris-HC1 buffer, pH 7.6, containing 3 mM MgCI2, 0.5 mg NADPH and 10-200/xl of sample (giving 0.1-4 mg protein). 100 nmoles of B(a)P was added in 0.05 ml acetone and the mixture incubated at 37°C, with agitation, for between 3 and 20 min, depending upon the activity in the sample. The reaction was stopped by the addition of 1.0 ml acetone. Blanks received 1.0 ml of acetone before the addition of B(a)P. The mixture was shaken with 3.0 ml hexane and 2.0 ml of the organic layer extracted with 3 ml 1 M NaOH. The fluorescence of the extract was measured immediately at 396 nm excitation and 520 nm emission. The fluorescence was compared to that of a series of secondary standards, consisting of quinine sulphate (20-100/xg/ml) in 0.1 N HzSO 4. Quinine sulphate was calibrated against a series of 3-OH-B(a)P standards and a conversion factor obtained. Activity was quoted as pmoles 3-OH-B(a)P/mg protein/rain. N-Demethylase. N-Demethylase activity was determined by measuring formaldehyde production by a method modified from Schenkman et al. (1967), using DAB as substrate. The incubation was performed in 1.0 ml 0.1 M potassium phosphate buffer, pH 7.4, containing 6 mM MgSO 4, 0.7 mM NADP and 13 mM glucose-6-phosphate at 37°C. Sample was added in a range of 0.5-2 mg protein/ml. 20 mM DAB was added in 25/xl ethanol and the mixture was incubated with agitation for 30 min. The reaction was stopped by addition of 0.5 ml 10% trichloroacetic acid in water. Trichloroacetic acid was added to standard and blank tubes before incubation. The tubes were centrifuged at 500 g for 5 min and 1 ml

supernatant fluid transferred to 1 ml Nash reagent (150 g ammonium acetate, 2 ml acetylacetone, 3 ml acetic acid per litre, pH 6.0) and incubated for 60 min at 37°C. The absorbance of the sample at 415 nm was measured and compared with standards containing 0.3-20/xg formaldehyde. Activity units were measured in nmoles formaldehyde/rag protein/min. For high activities the Nash reagent was diluted 2-fold after addition of the supernatant fluid.

Nitroreductase. Nitroreductase activity was determined using p-nitrobenzoic acid as substrate, and assaying the formation of p-aminobenzoic acid (Bratton and Marshall, 1939; Saz and Martinez, 1956). The samples were incubated for 60 min at 37°C in sealed glass vials gassed with oxygen-free N 2 and containing 0.45 ml sample, 0.1 ml 75 mM L-cysteine, 0.1 ml 1.5 mM MnC12, 0.1 ml 1.5 mM FMN, 0.1 ml 1.0 mM NADH, 0.6 ml 20 mM Tris buffer, pH 7.5, and 0.1 ml 4.4 mM p-nitrobenzoic acid dissolved in ethanol. The substrate was replaced by either 0.1 ml water, in control incubations, or 0.1 ml paminobenzoic acid (5-150 /xg/ml), in standard incubations. After incubation, the protein was precipitated with 150 /zl 10% trichloracetic acid and sedimented by centrifugation for 5 min at 500 g. 1 ml of the protein-free supernatant fluid was then heated for 60 min at 100°C with 0.5 ml 4 N HC1 to hydrolyse arylamine conjugates. After cooling, 100/xl 0.1% NaNO 2 was added, followed after 3 rain by 100 nl 0.5% ammonium sulphamate and, after a further 2 rain, 100 /xl 0.1% N(1-naphthyl)ethylenediamine dichloride. The absorbance of the samples at 545 nm was measured using the blank as reference. Nitro reductase activity units were ng p-aminobenzoic acid/mg protein/min. Glutathione-S-transferase. Glutathione-Stransferase activity was determined by the conjugation of glutathione with 1-chloro-2,4-dinitrobenzene (Habig et al., 1974). The assay was performed at 25°C in a thermostatically-controlled 1 cm cuvette in 3 ml of 0.1 M potassium phosphate buffer, pH 6.5, containing 0.15 ml 1 mM reduced glutathione and 0.15 ml 1 mM 1-chloro-2,4-dinitrobenzene dissolved in ethanol. The rate of

33

increase in absorbance at 340 nm was determined before and after addition of the sample, from which the quantity of S-2,4-dinitrophenylglutathione was calculated (e340 = 9.6 mM/cm). Glutathione-S-transferase units were nmoles S-2,4dinitrophenylglutathione/mg protein/min.

Superoxide dismutase. Superoxide dismutase (SOD) activity was determined by the inhibition of the rate of reduction of cytochrome c in the presence of a xanthine : xanthine oxidase superoxide radical generating system (McCord and Fridovich, 1969). The assay was performed at 25°C in a thermostatically-controlled 1 cm cuvette in 3 ml of 0.05 M potassium phosphate buffer, pH 7.8, containing 0.1 M EDTA, 0.05 M xanthine and 0.01 mM ferricytochrome c. The rates of change of absorbance at 550 nm after the addition of xanthine oxidase to a concentration of 1.66 × 10 -3 Izg/ml in the presence and absence of the sample were measured. 1 unit of superoxide dismutase activity was defined as the activity required to inhibit the rate of reduction of cytochrome c by 50% under these conditions. Catalase. Catalase activity was determined by the decomposition of hydrogen peroxide measured directly by the decrease in absorbance at 240 nm (Nelson and Kiesow, 1972). The assay was performed at 25°C in a thermostatically-controlled 1 cm cuvette in 3 ml 0.05 M potassium phosphate buffer, pH 7.0, containing 10.5 mM H 2 0 2. This concentration gave an A240 of 0.500 _+0.010. 20 /~1 of sample was added to both the H 2 0 2 and the reference cuvettes and the change in A240 recorded. Results were expressed as Bergmeyer units, i.e., the amount of enzyme which liberates half of the oxygen from an H 2 0 2 solution of any concentration (ca. 10 mM) in 100 sec at 25°C (Aebi, 1974). Peroxidase. Peroxidase activity was determined by the increase in absorbance at 430 nm resulting from the oxidation of pyrogallol to purpurogallin (Chance and Machly, 1955). The assay was performed at 20°C in a thermostatically controlled 1 cm cuvette in 3 ml of 33 mM potassium phosphate buffer, pH 6.0, containing 7.8 mM H 2 0 2 and 5 mg/ml pyrogallol. 100/zl of sample

was added and the rate of increase in absorption at 430 nm is measured, from which the quantity of purpurogallin was calculated (e430 = 2.47 n M / cm).

Western blotting. Cells or $9 samples were diluted to 3 mg protein/ml, mixed (1:1) with buffer (50 mM Tris containing 2% SDS, 5% mercaptoethanol, 10% glycerol and 0.005% bromophenol blue) and heated to 100°C for 5-10 min. Proteins were separated by SDS-PAGE (9%), then transferred to nitrocellulose (Towbin et al., 1979) in 20 mM sodium phosphate containing 20% methanol for 16 h at 250 mA. Filters were washed in 50 mM Tris-HC1, pH 7.9, containing 150 mM NaCI and 0.05% Tween 20 (TBST) (2 × 10 min) and blocked in dried milk powder, 3% (w/v), in TBST. They were then exposed for 1 h to polyclonal antisera against purified rat cytochrome P450 isoenzymes, P450IA2 and P450IIA1 (Adams et al., 1985) used at a dilution of 1:500. Bound antibody was visualized using horseradish peroxidase-linked donkey anti-rabbit IgG (1:1000; 1 h) and the substrate, 4-chloro-l-naphthol. Sensitivity was increased by incubating the nitrocellulose with I125-protein A, followed by autoradiography. Results

For this study, five cell lines were chosen, two from mice (L5178Y and C3H10T1/2), two from hamsters (Chinese hamster ovary and Chinese hamster V79) and one from man (human lymphoblast TK6). All have been used extensively to examine either the mutagenic or transforming properties of chemicals. We have compared the enzymatic activity in these cell lines with the metabolic systems commonly used in mutation studies, i.e., rat hepatocytes and Aroclor 1254 induced rat $9. The results are summarized in Fig. 1.

Enzyme levels from exogenous sources of "metabolic activity The enzyme levels of the cytochrome P450 related activities, benzo[a]pyrene hydroxylase and

34

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Fig. 1. Enzyme activities in homogenates of Aroclor 1254-induced rat liver, in rat hepatocytes and in various cultured cells. Error bars: standard deviations on N independent assays. The number of experiments performed for each assay is presented on top of each bar. Units: Catalase, Bergemeyerenzyme units/mg protein; superoxide dismutase, enzyme units/mg protein; peroxidase, mg purpurogallin/mg protein/min; nitroreductase, ng p-aminobenzoicacid/rag protein/rain; DAB-N-demethylase,nmoles formaldehyde/mg protein/min; B(a)P hydroxylase, pmoles 3-hydroxy-B(a)P/mgprotein/min; glutathione-S-transferase, tzmoles S-2,4-dinitrophenylglutathione/mg protein/min.

dimethylaminoazobenzene N-demethylase, and that of glutathione-S-transferase were higher in $9 from Aroclor 1254 induced rats than in noninduced hepatocytes. For the enzymes that are involved in protecting cells from oxidative damage (peroxidase, superoxide dismutase and catalase), the levels of activity were higher in the noninduced rat hepatocytes. This indicates either that a significant proportion of the activity was associated with the fraction sedimenting at 9000 g or that these activities were reduced by Aroclor treatment.

Comparison of enzyme activities in liver homogenate and cultured cells Benzo[a]pyrene hydroxylase and N-demethylase activities were higher in $9 than in the cell lines examined. While these levels were also higher in rat hepatocytes when compared to the cells in culture, the differences were smaller. Decreases in enzyme activity in the cell lines compared to rat hepatocytes were especially pronounced for B(a)P hydroxylase, peroxidase and catalase. Glutathione-S-transferase activity was significantly lower in mouse lymphoma and hu-

35 man lymphoblast cells, compared with rat hepatocytes while the activity of Chinese hamster ovary, Chinese hamster V79 and C 3 H 1 0 T 1 / 2 ceils were roughly equivalent to that found in hepatocytes.

Cytochrome P450 enzymes The two cytochrome P450 activities, benzo[a]pyrene hydroxylase and dimethylaminoazobenzene N-demethylase, were absent from mouse lymphoma, Chinese hamster ovary, or Chinese hamster V79 cells. Interestingly, some activity was observed in C 3 H 1 0 T 1 / 2 cells, which were originally selected because of their low spontaneous transformation frequency and their susceptibility to transformation by polycyclic aromatic hydrocarbons. This evidence supports the reports of cytochrome P450 in these ceils (Gehly et al., 1979; Gehly and Heidelberger, 1982; Rudo et al., 1986). Since both benzo[a]pyrene hydroxylase and N-demethylase functions are at least predominantly cytochrome P450 mediated, it would be expected that these two would increase or decrease concomitantly. This was found, however, not to be true. In human lymphoblast TK6 cells, no benzo[a]pyrene hydroxylase activity was measurable, but N-demethylase activity was observed.

The presence of only the latter enzyme may indicate either the absence of specific isoenzyme or an alternative mechanism for N-demethylation. Benzo[a]pyrene hydroxylase activity, although detectable in C 3 H 1 0 T 1 / 2 cells, was 1/100th of that found in hepatocytes and 1/1500th of that in $9. Dimethylaminoazobenzene N-demethylase activity, on the other hand, was only two-fold higher in hepatocytes and six-fold higher in $9 than in C 3 H 1 0 T 1 / 2 cells. The detected benzo[a]pyrene hydroxylase and N-demethylase activities in the C 3 H 1 0 T 1 / 2 cells were supported by Western blots (Fig. 2) where antibodies to P450IA2 and P450IIA1 recognized proteins in these cells. The mobility of the protein reacting with the rat P450IIA1 antibody is consistent with the known molecular weight (approximately 48 kD) of P450IIA1 protein in rat and mouse and had the same molecular weight as a protein identified by this antibody in mouse liver (Fig. 2, tracks I and J). The mobility of the band recognized by the P450IA2 antibody was faster than that associated with either P450IIA1 or P450IA2 in mouse liver (Fig. 2, tracks I and J) and its exact identity remains to be established. The P450IIC6 or P450IIIAI antibodies did not

anti P45o- ]I A1

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Fig. 2. CytochromeP450 isoenzyme expression in cell lines. Western blot analysis of 14 ~g rat $9 (D) and 45/zg of each of the following cell homogenates: Chinese hamster ovary (E), V79 (F), mouse lymphoma (G), and C3HIOT1/2 (H). Mouse liver microsomal protein, 2.5/zg (I) and 5.0 tzg (J) and 0.4, 0.2 and 0.1 pmoles purified rat P450IA2(A,B,C respectively) were loaded as controls. Bands reacting with the anti-P450-1A2or anti-P450 IIA1 were visualized using x25I-protein A and the autoradiographs shown were exposed for 6 days. Proteins from whole cell homogenates(45 #g), $9 (4/zg), or standards (0.4, 0.2, 0.1 phaoles) were run on 9% SDS-polyacrylamidegels and then transferred to nitrocellulose. Bands reacting with the cytochromeP450 antibodies were then identified using the procedure described in the Materials and Methods section using I ta5 protein A. The autoradiographs shown were developed after a 6 day exposure. The antibodies used were to rat P450IA2(1MC-la) and P450IIA1(UT1) and P450IIC6 (PV-2c).

36 identify bands attributable to cytochrome P450s. All the other cell lines tested were negative with all the four antibodies. Oxidative enzymes Superoxide dismutase, catalase and peroxidase activities, all of which protect cells against oxygen radical damage, would be expected to be expressed in any cell surviving in an aerobic environment. Although the levels of activities were quite variable, they were measurable in all of the cell lines studied. Superoxide dismutase activity was lowest in the single human lymphoblast TK6 homogenate. In most of the other homogenates, the activities were 2-4-fold higher, except for the rat hepatocyte samples which, at 34.5 _+ 3.6 units/mg protein, were approximately double the activities in most of the other samples and seven times higher than the superoxide dismutase activity found in human lymphoblast TK6 cells. This last comparison was made upon a single sample result. Catalase activities were lowest and similar in the L5178Y, human lymphoblast TK6 and C3H10T1/2 cell homogenates. They were approximately 4-5-fold higher in the Chinese hamster ovary and V79 cells and approximately 70-fold higher in rat hepatocytes. The lower level of catalase activity found in $9, than in hepatocytes, is probably due to loss of catalase by sedimentation of a proportion of the peroxisomes. Peroxidase activities were similar in L5178Y, Chinese hamster ovary, Chinese hamster V79 and C3H10T1/2 cells. Mean values covered only a 1.6-fold range in these cell types, but these were 6-10-fold higher than in human lymphoblast TK6 cells. In two hepatocyte samples very different peroxidase concentrations were found. One of these, 22.3 /xg purpurogallin/mg protein/min, was very similar to the activity measured in induced rat $9, but the other was 3-fold higher. Reductive enzymes The study of reductive enzymes was limited to nitroreductase, using p-nitrobenzoic acid as the substrate. Such reactions are important for the metabolism of nitro compounds to hydroxylamines, which may be reactive and damaging products. Nitroreductase activity was lowest in

Chinese hamster V79 cells and only about 2-fold higher in L5178Y cells. The values were approximately 16-18-fold higher, respectively, in Chinese hamster ovary cells and the induced rat liver $9. Activities were not measured in the remaining cell homogenate, due to shortage of sample. Conjugating enzymes Glutathione-S-transferase activities were similar and low in the L5178Y and human lymphoblast TK6 cells, where there were only 0.07 and 0.06 units/mg protein, respectively. Activity in C3H10T1/2 cells was approximately 3-fold higher and the values were approximately 6-17fold higher in the other cell homogenates. In induced rat liver $9 the glutathione-S-transferase activity was about 29-fold higher than in the L5178Y cell homogenate. Thus, there is considerable variation in the potential for conjugation with glutathione between these cell types. Discussion

The purpose of this study was to demonstrate the activities of some enzymes with xenobiotic metabolizing capabilities in different cell lines, $9 and rat hepatocytes which are commonly used in many in vitro toxicology systems. In many instances, the capacity is present for conjugation and for peroxidation reactions. The presence of antioxidative enzymes is common and to be expected since they are necessary for cell survival in an aerobic environment. Most cell lines had no capacity for cytochrome P450-mediated reactions although its presence was found in the C23H10T1/2 cell line. In competent cell lines, the two cytochrome P450 enzyme activities, benzo[a]pyrene hydroxylase and N-demethylase, did not follow similar patterns. Thus, the ratio of N-demethylase to benzo[a]pyrene hydroxylase activities ($9 being 100% in each case) were: hepatocytes, 0.17/0.18; C3H10T1/2, 0.08/0.01; human lymphoblast TK6, 0.07/0. While these studies add to previously published information regarding enzymic activities in cell lines (Wiebel et al., 1980), they do not, on their own, provide a basis for explaining intercellular differences in response. Knowledge of cellular metabolic capacity is but a necessary compo-

37 nent of our understanding of in vitro test systems used to evaluate the role of individual components in complex processes such as carcinogenesis. The results reported here suggest that if cytochrome P450 enzymatic activity was required for the evocation of a particular type of response, then the use of $9 might be preferable to rat hepatocytes due to the higher levels of activity obtained from the $9 system. If an exogenous enzyme system is not usable (for example, in chronic, low dose in vitro studies), the use of C 3 H 1 0 T 1 / 2 cells might be selected. This study suggests that a member of the P450IIA gene family is expressed in C 3 H 1 0 T 1 / 2 cells. It also showed the reactivity of the antibody to P450IA2 with a protein in this cell line although its mobility did not correspond to that of the expected antibody-protein complex in mouse liver and its identity could not be established from the work presented here. In a recent study, cytochrome P450IA family genes were not detected in this same cell line (Pottenger and Jefcoate, 1990). Our study is not in conflict with these findings in that the P450IA1 antibody used by Pottenger and Jefcoate did not cross react with any proteins in the cell and their work did not preclude the possibility that the P450IIA1 was expressed. The consequences of the differences in enzyme activity on the biological activity of different chemicals cannot be evaluated at this stage, but it is possible that the combination of relatively high superoxide dismutase levels with low catalase levels observed in L5178Y and C 3 H 1 0 T 1 / 2 cells could result in enhanced intracellular oxidation because of hydrogen peroxide generation. Peroxidase reaction, in which there is hydrogen transfer from donor to hydrogen peroxide, could account for at least some of the oxidative reactions which are presumed to be necessary for the activation of chemicals in cells lacking cytochrome P450 activity. The dynamic in vitro environment is complicated when $9 is added to these systems. This is an extracellular source of metabolic function, the products of which may be further metabolized should they enter a cell. These consecutive metabolic steps (one extracellular, followed by an intracellular reaction before reaching the trans-

formation target), may explain why polycyclic aromatic hydrocarbons have a greater transforming potential in C 3 H 1 0 T 1 / 2 ceils in the absence than in the presence of $9 (McGregor, unpublished observation), whereas the other cell lines (with their different analytical end-points) are refractory to these chemicals unless $9 is present. Explanations for other cell line differences are not so obvious. The unresponsiveness of Chinese hamster V79 cells to aromatic amines in both the presence and absence of $9 (O'Donovan, 1985; Langenbach et al., 1986; Oglesby et al., 1983) is such an example and the results reported here do not appear to provide any explanation for the phenomenon, unless the greater glutathione-Stransferase activity in Chinese hamster V79 cells is sufficient to provide protection. In vitro assays can play a role in the assessment of the potential hazard of chemicals to humans but it must never be forgotten that in vitro systems are artificial in that they are designed to measure certain biological endpoints under tissue culture conditions. It is only with such understanding that we shall be able to properly evaluate in vitro data and reasonably apply them in chemical toxicology.

Acknowledgements The authors wish to thank Drs. Robert Langenbach and Thomas Eling for their comments on this manuscript.

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