Application of an epithelial liver cell line, metabolically competent, for mutation studies of promutagens

Mutation Research, 271 (1992) 79-88 © 1992 Elsevier Science Publishers B.V. All rights reserved 0165-1161/92/$05.00

79

MUTENV 08811

Application of an epithelial liver cell line, metabolically competent, for mutation studies of promutagens G. T u r c h i

1, A.

N a r d o n e 1 and F. Palitti 2

1 Istituto di Mutagenesi e Differenziamento del C.N.R., Pisa and 2 Dipartimento di Agrobiologia Agrochimica, Universit~ della Tuscia, Viterbo (Italy)

(Received 13 December 1990) (Revision received 12 September 1991) (Accepted 13 September 1991)

Keywords: Promutagens; Chinese hamster epithelial liver cells

Summary Recently numerous attempts have been made to reduce the use of vertebrate animals in laboratory experiments to evaluate general and acute toxicity, mutagenesis and teratogenesis of new drugs or chemicals. One common approach is to use established, proliferating cell lines that preserve differentiated functions such as the competence to metabolize xenobiotics, To this end a continuous Chinese hamster epithelial liver cell line (CHEL cells) was established, cultured as used for mutagenesis studies. Structurally different promutagens, such as 7,12-dimethylbenz[a]anthracene (7,12-DMBA), benzo[a]pyrene (B(a)P), aflatoxin B 1 (AB 1) and cyclophosphamide (CP), were used in order to check and validate the test system, anti-Chrysene-l,2-diol 3,4-epoxide (CDE) and mitomycin C (MMC) were taken as representatives of direct mutagens. The genetic change induced by the mutagens was quantified by measuring mutation frequencies at the HGPRT locus. Several parameters, such as mutant expression time for each chemical, cell density for selection of mutants and enzymatic characterization for HGPRT phenotype, were examined to establish the optimal assay conditions. All promutagens analyzed significantly affected either the cloning efficiency and/or the mutant frequency of CHEL cells after 24 h of exposure. In addition, various enzyme activities involved in the metabolism of the promutagens were determined in CHEL cells, under the experimental conditions of chemical exposure used in the

Correspondence: Dr. G. Turchi, Institute of Mutagenesis and Differentiation of C.N.R., Via Svezia 10, 56100 Pisa (Italy). Abbreviations: 8-AZA, 8-azaguanine; 6-TG, 6-thioguanine;

7,12-DMBA, 7,12-dimethylbenz[a]anthracene; AB1, aflatoxin B1; CP, cyclophosphamide; MMC, mitomycin C; B(a)P, benzo[a]pyrene; CDE, anti-chrysene-l,2-diol 3,4-epoxide; HGPRT, hypoxanthine-guanosine phosphoribosyl transferase;

ECOD, 7-ethoxycoumarin O-deethylase; EROD, 7-ethoxyresorufin O-deethylase; DCPIP, dichlorophenolindophenol; CDNB, 1-chloro-2,4-dinitrobenzene; HYP, hypoxanthine; PRIB-PP, a-5-phosphoribosyl-l-pyrophosphate; PAH, polycyclic aromatic hydrocarbon; EDTA, ethylenediaminetetraacetic acid; DMSO, dimethyl sulfoxide; PBS, Dulbecco's phosphate-buffered saline.

80 mutagenesis assay. The enzyme activities were compared with those found in uninduced Chinese hamster liver.

Mammalian cell cultures have become a fundamental tool in the study of the molecular mechanisms by which environmental contaminants exert their toxic, mutagenic and carcinogenic effects (Glatt et al., 1989; Autrup, 1990; Pool et al., 1990). These in vitro results must be compared with in vivo assays using rodents. Given the high degree of species and organ specificity observed in experimental carcinogenesis it is clear that model cell lines for in vitro tests should display characteristics of the original tissue and retain their organ-specific sensitivity to genotoxic materials. The majority of cell lines commonly used in short-term tests have, however, lost most of their differentiated characteristics, including the capacity to metabolize xenobiotics (Wiebel et al., 1980; Bradley et al., 1981; Glatt et al., 1989). It is believed that factors such as embryonic origin and differentiation state are responsible for the organospecific metabolic competence to carcinogens (Kleihues et al., 1983; Guengerich et al., 1982; Domin et al., 1986; Gould et al., 1986; Hsu et al., 1987; Oesch, 1987; Autrup, 1988; Guengerich, 1988; Bengtsson et al., 1988). It is well known that the majority of human neoplasias occur in cells of epithelial, or lymphoblastic origin, and yet most tissue culture assays for mutation/transformation have used fibroblasts as the indicator cell lines (Cairns, 1975; Peto, 1977). It is possible that replacement of fibroblasts by epithelioid cells would make the in vitro assay more reflective of the in vivo situation. In consequence we have developed and characterized a Chinese hamster epithelial liver cell line. As previously reported, this cell line is able to metabolically activate numerous promutagens, and the reactive metabolites generated can modify the DNA of activating cells and that of proximal indicator cells (Turchi et al., 1987; De Salvia et al., 1988). In the present study the genetic effect of reactive metabolites generated from promutagens was monitored as induced mutation frequency at the HGPRT locus in the activating liver cell line.

Direct-acting mutagens were selected and analyzed as positive controls. Material and methods

Material The promutagens aflatoxin B 1 (AB1), 7,12-dimethylbenz[a]anthracene (7,12-DMBA), the purine analogues 6-thioguanine (6-TG) and 8azaguanine (8-AZA), the a-5-phosphoribosyl-1pyrophosphate (P-RIB-PP) and insulin were purchased from Sigma Chemical Co., while cyclophosphamide (CP) and mitomycin C (MMC) were obtained from Asta-Werke (Bielefeld, Germany), benzo[a]pyrene (B(a)P) gold label grade, 99% pure was supplied by Lancaster Synthesis Ltd (U.K.). anti-Chrysene-l,2-diol 3,4-epoxide (CDE), 89% pure was synthesized and kindly provided by Dr. A. Seidel of the Institute of Toxicology, University of Mainz (Germany). AB 1, 7,12-DMBA, B(a)P and CDE were dissolved in spectrograde dimethyl sulfoxide (DMSO) prior to addition to the medium and the final concentration of DMSO in the medium did not exceed 0.5%. CP and MMC were dissolved in serum-free Williams m e d i u m (Flow Laboratories). Dichlorophenolindophenol (DCPIP) and resorufin were obtained from Fluka (Buchs, Switzerland); dicoumarol and 7-ethoxycoumarin were purchased from EGA-Chemie (Steinheim, Germany); 1chloro-2,4-dinitrobenzene (CDNB) was obtained from Eastman Kodak (Rochester, U.S.A.) and NAPDH was purchased from Buerchingen (Munich, Germany). 7-Ethoxyresorufin was synthesized from resorufin by ethylation with ethyl iodide (Klotz et al., 1984). [8-14C]Hypoxanthine (2.74 Ci/mole) was purchased from the Radiochemical Centre (Amersham). Aquasol-2 liquid scintillation cocktail was from New England Nuclear. PEI-cellulose precoated thin-layer plastic sheets (0.1 mm thick) were obtained from Merck. All other chemicals and solvents were obtained from common commercial sources.

81

Cells Chinese hamster epithelial liver cells (CHEL cells) were obtained as previously described (Turchi et al., 1987a). Cells were routinely cultured in Williams E medium supplemented with 10% fetal bovine serum (FBS) and antibiotics (complete medium). Williams E medium containing 5% FBS was used for treatment with test compounds. To minimize possible variations in responses to mutagens, CHEL cells between the 7th and the 15th passage were used. The Chinese hamster fibroblast cell line V79 as originally obtained from Dr. C.F. Arlett (MRC Cell Mutation Unit, University of Sussex, U.K.) and usually cultured in Dulbecco's modified Eagle's medium (Gibco, Scotland) with 5% FBS and antibiotics. Cells were routinely tested (Chen, 1977) and found to be free of mycoplasma. Cytotoxicity assay Test concentrations for the mutagenesis experiments were chosen after a preliminary rangefinding cytotoxicity assay. For this assay, 1000 cells were seeded as single-cell suspensions into each of five 60-mm dishes in 5 ml of Williams E medium and treated 1 day later with test compounds dissolved in an appropriate solvent. When necessary, control cultures received DMSO alone. After 24 h the chemical-containing medium was removed and the plates were washed 3 times with Dulbecco's phosphate-buffered saline (PBS). Cultures were then continued in complete medium for 1 week, at which time the colonies were fixed, stained with Giemsa and counted. Only colonies containing 30 or more cells were scored. Cloning efficiencies ranged between 17 and 20% in untreated controls. Enzyme assays For induction experiments cells were plated at a density of 6000 cells/cm 2 on 140-mm dishes containing 20 ml of Williams E medium. After 24 h the medium was changed and the test compound dissolved in the medium or in the minimal amount of DMSO was added. The cultures were exposed to the test compounds for 24 h (slightly more than one cell generation time, 22 h). At the end of this time the culture medium was removed and monolayers (5 dishes per experiment) were

washed twice with 3 ml PBS. The cells were scraped off the plate with a rubber policeman, collected in a conical tube and centrifuged at 800 x g for 10 min at 4 ° C. Cell homogenates were obtained by sonicating the cells, resuspended in 2 ml of cold PBS and incubated in ice, by three 15-s bursts with a Sonipred 150 sonicator set at 25% of maximum intensity. Microsomes were prepared from homogenate as previously described (Longo et al., 1986). The washed microsomal pellets were resuspended in K phosphate buffer (0.1 M, pH 7.4), 1 mM ethylenediaminetetraacetic acid (EDTA) and stored at - 80 ° C. The postmicrosomal supernatant was further centrifuged for 60 min at 100,000 x g and used for cytosolic enzyme assays. 7-Ethoxycoumarin O-deethylase (ECOD) activity was determined according to the method of Aitio (1978). 7-Ethoxyresorufin O-deethylase (EROD) was assayed by measuring the formation of the corresponding hydroxy product in a Perkin-Elmer spectrofluorimeter as described by Krijgsheld and Gram (1984). Glutathione S-transferase activity was determined spectrophotometrically using CDNB as substrate (Habig et al., 1974). DT-Diaphorase was assayed following the reduction of DCPIP (as electron acceptor) inhibitable by 1 /zM dicoumarol (Wermuth et al., 1986). HGPRT activity was analyzed both in the mutants and in the wild-type cells. 6-TG r and 8-AZAr clones, chosen randomly, were isolated from untreated cells or from mutagenized ceils, individually cultured for 10 days in non-selective medium and subcultured and expanded in medium containing selective agents. HGPRT activity was assayed in a crude extract of 5 x 106 cells by monitoring conversion of [8-14C]hypoxanthine to inosine monophosphate as described (Camici et al., 1988). Cytosolic and microsomal protein concentrations were measured by the method of Lowry et al. (1951) with bovine serum albumin fraction V (Sigma) as standard.

Mutagenesis assay One day after seeding 2 x 10 6 CHEL cells into each of four 140-mm dishes, when the ceils were in exponential growth, the complete medium was removed and the cultures were refed with the Williams E medium used for treatment (5% FBS).

82 Subsequently the test compound was added. After 24 h exposure, the medium containing the chemical was removed, the cultures rinsed twice with PBS and then immediately trypsinized. The mutant selection and the cloning efficiency were determined following the experimental schedule previously described (Turchi et al., 1981). At least 2 separate experiments were performed for each compound. Briefly, 5 x 105 cells were seeded in 4 dishes for selection of 6-TGr mutants. 6-TG was added to the dishes immediately after seeding to give a final concentation of 10 /zg/ml. Fourteen days after the initial addition of selective agent, colonies were fixed with methanol, stained with Giemsa and counted. The optimal expression time of 6-TG r mutants was determined by analyzing colony formation at 5 different times after treatment. For cloning efficiency 500-1000 cells were seeded in quadruplicate 60-mm dishes. The colonies developed after 7 days were evaluated as described above. Mutation frequency values were expressed as mutants per 106 clonable cells.

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Quantitative mutagenesis studies in C H E L ceils were carried out optimizing several parameters affecting the mutation rate. (I) Optimal concentration of selectiue agent. Inhibition growth curves of wild-type C H E L cells in the presence of increasing concentrations of 6-TG indicated that 10 ~zg/ml was the optimal concentration of the selective agent in the medium. (II) Effect of cell density on mutant recouery. Contact and crossfeeding of toxic 6-TG nucleotides by wild-type cells to H G P R T mutants were observed in fibroblasts and resulted in a decreased recovery of induced 6-TG r mutants from high-density cultures (Jacobs and Demars, 1978). Reconstruction experiments were carried out to establish optimal conditions for maximum mutant recovery and minimum effect of metabolic cooperation. We measured the recovery of spontaneous or induced 6-TG r mutants as a function of increasing number of co-cultured wild-type cells. The maximum

83 recovery was obtained when < 5 × 105 cells ( < 3200 cells/cm 2) were seeded per 100-mm dish with 500 mutants. (III) Phenotypic expression time. The kinetics of mutant expression represents a critical parameter and influences the quantitative response of the test. In our system the maximal expression time ranged between 5 and 8 days, and then, in some cases, declined slightly. (IV) Biochemical characterization of mutants and stability ofphenotype. Four spontaneous and 4 chemically induced 6-TG r and 8-AZAr mutants were randomly isolated by encirclement with a stainless steel cylinder, individually cultured and characterized for HGPRTase activity. The resistant phenotypes of each clone were stable on culture in purine analogue-free medium for > 10 passages. The HGPRTase activity was measured utilizing cell-free extracts to convert hypoxanthine to inosine monophosphate. The reaction was linear with time over a range of protein concentrations of 0.8-1.2 mg/ml for at least 50 min. The activity in the wild-type cells was 0.304 m U / m g protein, whereas all mutants analyzed showed greatly reduced activity. Four clones showed less than 3% of wild-type activity, the other 4 clones between 4 and 18%.

Xenobiotic metabolizing enzyme activities Various enzyme activities of the CHEL cells were investigated and the results, which confirmed those previously reported (Turchi et al., 1987; Glatt et al., 1990), are summarized in Table 1. The cytochrome P-450-dependent mono-oxygenase activities determined as ECOD and EROD activities were much lower than those found in the liver of untreated Chinese hamsters (Table 1). However, high levels of NADPH cytochrome P-450 reductase were found in CHEL cells (Turchi et al., 1987; Glatt et al., 1990). This enzyme is functionally coupled with all forms of cytochrome P-450 involved in xenobiotic metabolism and at the same time its distribution has been suggested to be consistent with the major sites of tumor formation (Keyes et al., 1984). The low activities of mono-oxygenases in CHEL ceils, associated with high levels of reductase, and the high mutagenic responses to various promutagens discussed below, may indicate the

potential induction of the multiple forms of cytochrome P-450 present in these cells. DT-Diaphorase, a cytosolic flavoprotein NADPH: (quinone-acceptor) oxidoreductase that catalyzes the 2-electron reduction of quinones and may therefore represent an indirect detoxifying pathway of the cells, was also determined. The levels were relatively high, about 1/5 of these detected in non-induced Chinese hamster livers. The enzyme activities were also examined under conditions of the mutagenesis assay (namely after 24 h of chemical exposure) in the presence of 3 promutagens (CP, B(a)P and 7,12-DMBA), phenobarbital (PB) and insulin (Table 1). PB was chosen as a classic inducer of mono-oxygenases and insulin because it had been shown to affect numerous hepatic functions in vivo and in vitro (Grimm, 1976). The ECOD and EROD activities were about 2-fold increased by 7,12-DMBA and PB, respectively. DT-diaphorase was not affected by exposure to insulin and 7,12-DMBA, whereas it was inhibited by CP and B(a)P (44.8 and 46.1% of its control value).

Cytotoxicity and mutagenicity of promutagens and direct mutagens in CHEL cells Preliminary cytotoxicity studies were performed to determine the concentration dependence of the toxicity both of the direct-acting mutagens (CDE, MMC) and of the mutagens requiring metabolic activation. On the basis of the respective ICs0 (concentration that inhibits clonal growth by 50%), the chemicals can be divided into 3 groups. One group, which was extremely cytotoxic, included the two direct agents CDE and MMC, and 7,12DMBA (their ICs0 ranged between 0.1 and 1.8 /zM). It is worthy of note that, in the same concentration range, the cytotoxicity of 7,12DMBA was of the same order of magnitude as that of CDE, which does not require metabolic activation. A second group include A11 and B(a)P, showing ICs0 of 22 and 30 /zM respectively. Finally, CP was about 1 thousand-fold less toxic than the most active compound. The role played by metabolizing enzymes in the cytotoxic effect induced by chemicals was tested by parallel experiments with V79 cells, which notoriously do not express NADPH-cytochromes P-450 whereas

84 TABLE2

TABLE 3

MUTAGENIC EFFECT OF STRUCTURALLY DIFFERE N T P R O M U T A G E N S A N D D I R E C T M U T A G E N S ON C H E L CELLS

M U T A G E N I C I T Y O F A N A L Y Z E D P R O M U T A G E N S IN C H E L A N D V79 CELLS C O - C U L T U R E D W I T H C H E L CELLS

Compound

Compound

Concentration (p.M)

Mutation frequency (6-TG r m u t a n t s / 1 0 6 survivors) C H E L cells

V 7 9 / C H E L cells a

Control ABj B(a)P CP 7,12-DMBA

30 30 5mM 1

8.5+ 6.7 92.7+_19.6 51.7+ 2.3 101.7_+ 7.6 477.6_+97.1

7.5+ 3.8 54.4+ 5.1 38.2+_ 6.2 61.7_+ 4.3 336.8_+31.4

Control MMC CDE AB I B(a)P CP 7,12-DMBA

Concentration (/xM)

0.1 5 30 30 5 mM 0.01 0.1 0.5 1

Relative C F E (%)

100 45 4.5 18 45 45 115 110 93.5 65

Mutation frequency ~' (6-TG r mutants/106 survivors) 8.5 + 6.7 48.5 + 3.5 2220 -+ 95.3 92.7-+ 19.6 51.7-+ 2.3 101.7_+ 7.6 15.1_+ 5.8 395.8 _+26.5 350.4 + 96.3 477.6 _+97.1

The values constitute the m e a n s and SD of > 2 separate experiments. Mutation frequencies were calculated from the plateau of the expression curves.

they do express other enzymes such as NADPHcytochrome P-450 reductase, glutathione transferase, DT-diaphorase, etc. Fig. la reports the concentration dependence of the clonal growth inhibition of C H E L and V79 cells in the presence of 7,12-DMBA. The resistance of V79 cells to large concentrations of this

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a Mutation frequency induced in V79 cells co-cultured with C H E L cells as metabolizing source (Turchi et al., 1987). Data are shown for the optimal concentrations only.

chemical suggest that the cellular metabolic competence was a necessary prerequisite for the genotoxic activity of 7,12-DMBA. The sensitivity manifested by the cells at high doses of the chemical could be attributed to non-specific cytotoxic effects. Fig. lb shows the cytotoxic activity of MMC in C H E L and V79 cells. MMC, a direct mutagen, would be expected to manifest similar dose-response curves in both kind of cells. At MMC doses exceeding 0.1/xM the epithelial liver ceils were instead more resistant that V79 cells. A detoxification mechanism could explain this

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85 finding: DT-diaphorase in association with cytochrome P-450 mono-oxygenases may influence the degree of metabolic activation of MMC in CHEL cells by cytochrome P-450 reductase. The promutagens and direct mutagens were investigated for induction of gene mutations (Table 2). Table 3 summarizes the mutation frequencies observed in CHEL cells, as determined in this study, and in V79 cells as indicator in a cell-mediated mutagenesis assay (Turchi et al., 1987). Discussion

In previous studies, CHEL cells have been extensively characterized for growth properties and biochemical activities (Turchi et al., 1987). Applied in a cell-mediated mutagenesis assay, as external metabolizing source, and in a sister-chromatid exchange assay, the CHEL cells revealed a stable metabolic competence to activate different classes of promutagens into biologically active metabolites (Turchi et al., 1987; De Salvia et al., 1988; Giatt et al., 1990). Moreover, they showed a selective capacity to discriminate in vivo recognized carcinogens from non-carcinogens (De Salvia et al., 1988). The presence of high levels of 5'-nucleotidase, associated with moderate HGPRTase activity, in cultures of rat liver fibroblasts was seen to confer cellular resistance to toxic purine analogues through the active catabolism of the mononucleotides (Berman et al., 1977). The lack of any correlation between drug sensitivity, HGPRTase and substantial 5'-nucleotidase activity in CHEL cells suggests that purine metabolism may be regulated by different mechanisms in these ceils (Simili et al., 1989). These features facilitate the optimal recovery of resistant variants with lower concentrations of purine analogues and indicate that CHEL cells are suitable for quantitative mutagenesis studies. The observed mutagenic effects indicate that the promutagens studied were biotransformed to active metabolites in CHEL cells (Table 2). 7,12-DMBA is a very potent mutagen and carcinogen which elicits these properties in different species and organs only after metabolic activation (Yang and Dower, 1975; Huberman et al., 1979).

The cytochrome P-450 mono-oxygenases play a critical role in the reactions that lead to ultimate carcinogenic forms of 7,12-DMBA. The monooxygenases are not exclusive in this multi-step process because other enzymes, such as epoxide hydrolase (Cristou et al., 1989) and liver sulfotransferase (Watabe et al., 1987), are additionally involved in the activation and might be rate-limiting. Our results demonstrate that 7,12-DMBA mutagenizes CHEL cells highly efficiently, causing a 56-fold increase in HGPRT mutant frequency, and that the ECOD activity in these cells was induced almost 2-fold by 7,12-DMBA during the 24 h of exposure (Table 1). Analogously, other mammaliam cell lines, derived from hepatic or extrahepatic tissues, were particularly responsive to the genetic action of 7,12-DMBA (Meyer and Dean, 1981; Allen-Hoffmann and Rheinwald, 1984; Tong et al., 1984; Gould et al., 1986; Steele et al., 1989; Mane et al., 1990). All these cell lines, characterized by a common embryonic endodermal origin, express very low or undetectable levels of constitutive mono-oxygenase activity toward common substrates, but nevertheless show a high mutagenic response with 7,12-DMBA (Cristou et al., 1987). The existence in the CHEL cells of specific cytochrome P-450 monooxygenases with high affinity for the promutagen remains the most likely explanation for these findings, even though it is not possible to exclude the inducibility of specific form(s) during chemical exposure. CHEL cells actively metabolize CP, as indicated by the cytotoxic and mutagenic response (12-fold the control). Since CP is activated in vivo by phenobarbital-inducible mono-oxygenase (Hales, 1983), it appears that the cells retain significant constitutive levels of this P-450 enzymatic form(s). The mutagenic response obtained with the mycotoxin AB 1 was significant, the mutation frequency increasing ll-fold over the control. The high concentration of AB 1 necessary to increase the mutation frequency was comparable to those generally required when intact cells are used, as external metabolizing source or indicator ceils, in mammalian cell mutagenesis assays (Madie et al., 1986; Ray-Chaudhuri et al., 1980). It is known that a large spectrum of cytochrome P-450 mono-oxygenases is involved in

86

the metabolic activation of AB I (K~irenlampi, 1987; Shimada et al., 1987). The poor inducibility of ECOD activity (double the background) by 7,12-DMBA and the slight effect on E R O D activity of PB would suggest that constitutive monooxygenases stably expressed by the C H E L cells or PAH-inducible are responsible for the AB 1 activation and that these forms display low affinity for ABe. Again, the limited mutagenic activity of ABe, in C H E L cells, could be a consequence of detoxification mechanisms where the main reactive form, suggested to be the 2,3-oxide, may be efficiently deactivated by conjugation to glutathione (Degen and Neumann, 1981). The high level of glutathione S-transferase activity, assayed with CDNB as the substrate (330 n m o l e / m g protein x min) in uninduced C H E L cells could support this hypothesis. B(a)P increased the mutation frequency 6-fold over the control. Since B(a)P is not considered to be a liver carcinogen, the poor mutagenic activity with hepatic cells is in agreement with the in vivo carcinogenicity data (Salt et al., 1983). The weak activity that we found was also observed in other cell lines of hepatic and extrahepatic origin, but numerous conflicting results are reported in the literature. For example, rat embryonic fibroblasts efficiently activate B(a)P to mutagenic metabolites, whereas rat liver cells do not (Langenbach et al., 1978; Borek and Williams, 1980; Murison et al., 1984). A rat lung epithelial cell-mediated system converted B(a)P to mutagenic intermediates (Tompa and Langenbach, 1979). Furthermore, Murison et al. (1984) have established a rat liver cell line, RL-12, unusually sensitive to the genotoxic effects of B(a)P and 7,12-DMBA. The results provided by the direct mutagens tested in C H E L cells did not diverge from the expectation, since CDE was highly mutagenic and cytotoxic as found by Glatt et al. (1986) in several short-term tests, whereas MMC was a weak mutagen (5.7-fold the control). Cytochrome P-450 reductase, whose activity was higher in C H E L cells (Glatt et al., 1990) than in V79 cells (Keyes et al,, 1984), can be responsible for the reductive activation of MMC to genotoxic metabolites (Keyes et al., 1984). DT-Diaphorase and cytochrome P-450 (the latter undetectable in V79 cells) do not seem to be directly

involved in the bioactivation of MMC, although they may influence the degree of activation to reactive species by modifying the activity of P-450 reductase (Keyes et al., 1984). Finally, it is interesting to note (Table 3) that the mutagenic effect induced in C H E L cells by the promutagens used in this study was higher than that revealed by V79 cells, when C H E L cells were applied as external metabolizing sources in a cell-mediated mutagenesis assay (Turchi et al., 1987), but did not differ qualitatively. This suggests that the ultimate forms that mutagenize C H E L cells are the same that, in the cell-mediated mutagenesis assay, leave the metabolizing cells and permeate the co-cultured indicator cells. The quantitative difference observed in the two assays could be due to the different amount of electrophylic metabolites that reside in the proximity of the DNA target in the two indicator cell systems.

Acknowledgements The authors wish to express their gratitude to Dr. V. Longo for his contribution to determine enzyme activities, and thank Prof. G. Bellucci for helpful discussion and critical reading of the manuscript, and Mr. M. Cini for technical assistance. This investigation was supported in part by the STEP project, Grant EV4V-0045-I of the Commission of the European Community, and in part by National Research Council (CNR) Targeted Project 'Prevention and Control of Disense Factors, Subproject 2'.

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