Benzylic hydroxylation of 1-methylpyrene and 1-ethylpyrene by human and rat cytochromes P450 individually expressed in V79 Chinese hamster cells

Wolfram Engst, Robert Landsiedel, Heino Hermersdörfer, Johannes Doehmer1 and Hansruedi Glatt2

Deutsches Institut für Ernährungsforschung (DIfE), Department of Toxicology, Arthur-Scheunert-Allee 114–116, D-14558 Bergholz-Rehbrücke and
1 Institut für Toxikologie und Umwelthygiene, Technische Universität München, Lazarettstraße 62, D-80636 München, Germany


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Alkyl-substituted polycyclic aromatic hydrocarbons may be metabolized to highly reactive benzylic sulfuric acid esters via benzylic hydroxylation and subsequent sulfonation. We have studied the benzylic hydroxylation of 1-methylpyrene (MP), a hepatocarcinogen in rodents, and 1-ethylpyrene (EP), whose benzylic hydroxylation would produce a secondary alcohol ({alpha}-HEP), in contrast to the primary alcohol ({alpha}-HMP) formed from MP. The hydrocarbons were incubated with hepatic microsomal preparations from humans and rats, as well as with V79-derived cell lines engineered for the expression of individual cytochrome P450 (CYP) forms from human (1A1, 1A2, 1B1, 2A6, 2E1, 3A4) and rat (1A1, 1A2, 2B1). All microsomal systems and CYP-expressing cell lines used, but not CYP-deficient V79 cells, showed biotransformation of both hydrocarbons. Formation of the benzylic alcohol was detected in each case. {alpha}-HMP and its oxidation product, 1-pyrenylcarboxylic acid (COOH-P), accounted for a major part of the total amount of the metabolites formed from MP in the presence of human liver microsomes (38–64%) and cells expressing human 3A4, 2E1 or 1B1 (80–85%). Likewise, cells expressing human 1A1 showed a higher contribution of {alpha}-HMP and COOH-P to the total metabolites (45%) than cells expressing the orthologous enzyme of the rat (3%). EP was metabolized at a higher rate and with modified regioselectivity compared with MP, although {omega}-hydroxylation of the side chain was not detected with the cell lines and only accounted for a small percent of the biotransformation by the microsomal preparations. The highest contributions of {alpha}-HEP to the total metabolites from EP were detected with the cells expressing human 1A1, 1B1 and 3A4 (38–51%). {alpha}-HEP accounted for 16% of the metabolites formed in the presence of human hepatic microsomes. Thus, benzylic hydroxylation is a major initial step in the metabolism of MP and EP. This pathway appears to be even more important in humans than in rats. Previously, we had shown that the second step of the activation, the sulfonation of {alpha}-HMP and {alpha}-HEP, is also efficiently catalysed by various forms of human sulfotransferases.

Abbreviations: {alpha}-HEP, 1-(1-pyrenyl)ethanol; {alpha}-HMP, 1-hydroxymethylpyrene; B[a]P, benzo[a]pyrene; COOH-P, 1-pyrenylcarboxylic acid; CYP, cytochrome P450; EP, 1-ethylpyrene; MC, 3-methylcholanthrene; MP, 1-methylpyrene; PAH, polycyclic aromatic hydrocarbon; {omega}-HEP, 2-(1-pyrenyl)ethanol.


    Introduction
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Polycyclic aromatic hydrocarbons (PAHs) are a major class of environmental carcinogens, which are among likely risk factors for a number of different cancer types in humans (15). They are formed by incomplete combustion of organic materials and diagenesis of organic sediments (e.g. mineral oil, coal). At low combustion temperatures and diagenesis, alkylated PAHs are predominantly formed (68). Rogge et al. (9) found that 56% of the PAHs present in the exhaust of diesel-engined lorries were alkylated. The formation of methylated PAHs has also been observed when unsubstituted PAHs were incubated in the presence of subcellular hepatic preparation and cells in culture (1012). In rats treated with benzo[a]pyrene (B[a]P), a small amount of 6-methylbenzo[a]pyrenyl–DNA adducts were detected, in addition to larger levels of the well known dihydrodiol-epoxide-derived adducts (13).

PAHs require metabolic activation to electrophilic species to exert their mutagenic and carcinogenic effects. The best known reactive intermediates are epoxides, in particular bay-region dihydrodiol-epoxides, formed by cytochromes P450 (CYPs) and epoxide hydrolases (14,15). In particular, in the case of alkyl-substituted PAHs, the metabolic introduction of a good leaving group at the benzylic site via hydroxylation and subsequent O-sulfonation (also termed sulfation) is an alternative mechanism for the formation of reactive intermediates (1618). The spontaneous heterolytic cleavage of the leaving group leads to the formation of a benzylic carbocation, which is stabilized by the aromatic system and avidly reacts with numerous electrophiles, including DNA (19,20). The potent hepatocarcinogenicity of 6-hydroxymethylbenzo[a]pyrene and the moderate hepatocarcinogenicity of 1-methylpyrene (MP; structural formula in Figure 1Go) appear to be caused by the formation of the corresponding sulfuric acid esters (18,2126). The chemically synthesized sulfuric acid esters are carcinogenic and mutagenic and form the same patterns of DNA adducts if incubated with DNA in vitro, as have been observed in the liver of rats treated with 6-hydroxymethylbenzo[a]pyrene and MP, respectively. For the study of this activation pathway, we have used MP and its congeners, because the structure of these compounds does not allow the formation of vicinal dihydrodiol-epoxides, an alternative activation which would complicate the investigations. Moreover, the benzylic carbocation formed from 1-sulfooxymethylpyrene is stabilized by the same aromatic system as that formed from anti-benzo[a]pyrene-7,8-dihydrodiol-9,10-oxide upon opening of the oxirane ring. anti-Benzo[a]pyrene-7,8-dihydrodiol-9,10-oxide is the major ultimate carcinogen of B[a]P, the most thoroughly studied PAH (27).



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Fig. 1. Structural formulas of MP, EP and their side chain hydroxylated metabolites.

 
MP has been detected, for example, in cigarette smoke condensates at levels which are 3.3 times higher than those of B[a]P (28), in car exhausts and cellulose pyrolysates (29), and also as a pollutant in oysters, crabs and finfish (30).

Understanding of a bioactivation pathway involves, among other things, the identification of enzymes which may catalyse critical reactions. In previous studies we have demonstrated that at least three rat sulfotransferases and seven human sulfotransferases can catalyse the activation of 1-hydroxymethylpyrene ({alpha}-HMP; structural formula in Figure 1Go) to a mutagen (3134). Rat hydroxysteroid sulfotransferase a, human hydroxysteroid sulfotransferase and human estrogen sulfotransferase were the enzymes showing the highest activities. Much less is known about the first step of the activation of MP, its benzylic hydroxylation. Our observation that i.p. administration to rats of MP and {alpha}-HMP, at equimolar doses, leads to identical patterns and very similar levels of hepatic DNA adducts (Z.Papanikolaou and H.R.Glatt, unpublished data) suggests that benzylic hydroxylation is an important pathway in this species. Likewise, Rice et al. (35) have found that {alpha}-HMP is a major metabolite of MP incubated with rat hepatic homogenates. However, no data are available on the metabolism of MP by human enzymes.

Next to the methyl group, ethyl is the most common alkyl side chain in PAHs which occur in the environment, in particular in organic sediments (6). The additional methyl group at the benzylic carbon may alter the steric interaction with enzymes and provide an alternative site for side chain hydroxylation. The benzylic alcohols derived from ethyl-substituted PAHs are secondary alcohols in contrast to the primary benzylic alcohols derived from methylated PAHs. This difference may affect the further metabolism, as, for example, primary alcohols may be oxidized via the aldehyde to carboxylic acid, whereas secondary alcohols can undergo only the first oxidation step, resulting in the formation of a ketone. Esterification of the hydroxyl groups introduced in the {omega}-position of ethyl-substituents of PAHs should not lead to the formation of reactive products, since this position is not conjugated with the aromatic system. Indeed, {omega}-sulfooxy-1-ethylpyrene is stable in aqueous media and is not mutagenic in the Ames assay (unpublished data).

We have therefore incubated MP and its ethyl homologue, 1-ethylpyrene (EP; structural formula in Figure 1Go), with various sources of rat and human CYPs, and analysed the formation of metabolites oxidized at the benzylic position. Primarily, we used V79-derived cell lines which express an individual rat or human CYP. These cells do not appear to express any endogenous CYPs, sulfotransferases or UDP-glucuronosyltransferases (3639), a fact which minimizes background problems and facilitates the identification of phase-I metabolites.


    Materials and methods
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Chemicals
{alpha}-HMP, 1-pyrenylcarboxaldehyde, 1-pyrenylcarboxylic acid (P-COOH), 1-pyrenylacetic acid, 1-hydroxypyrene and pyrene were purchased from Aldrich (Steinheim, Germany). {alpha}-HEP was synthesized as described (40,41) and purified by flash chromatography using silica gel as solid phase and a mixture of dichloromethane and acetone (19:1, v/v) as eluent. The primary alcohol of EP, 2-(1-pyrenyl)ethanol ({omega}-HEP) was synthesized by esterification of 1-pyrenylacetic acid with dimethyl sulfate and reduction of the resulting methyl ester with lithium aluminium hydride. The raw product was purified by flash chromatography using silica gel as solid phase and hexane/ethyl acetate (10:1, v/v) as eluent. MP and EP were prepared by reduction from 1-pyrenylcarboxaldehyde and 1-acetylpyrene, respectively, as described elsewhere (40) and purified by preparative high-performance liquid chromatography (HPLC) on silica gel using cyclohexane as eluent. 1-Acetylpyrene was kindly provided by Dr A.Seidel (Department of Toxicology, University of Mainz, Germany). The purity of all compounds was >98%, as determined by gas chromatography with mass spectrometry (GC-MS), HPLC and 300 or 400 MHz 1H-NMR analyses. All other chemicals were of analytical grade. The solvents used for the clean-up and for HPLC (Fisher Scientific, UK) were of high-grade purity for spectroscopy or were freshly distilled.

Cell lines and cell culture
The specific clone of Chinese hamster V79 cells, now termed V79-MZ, which was used for the construction of CYP-expressing cell lines, has been characterized with regard to the expression of various endogenous xenobiotic-metabolizing enzymes and other properties. In particular, V79-MZ cells do not appear to express any endogenous CYP, UDP-glucuronosyltransferase, sulfotransferase or N-acetyltransferase, but they express CYP reductase, glutathione transferase and microsomal epoxide hydrolase (3639). Besides this parental V79-MZ cell line, V79-derived cell lines were used which were engineered for the expression of human (h) or rat (r) CYPs. For reasons of lucidity, we are using a uniform nomenclature here. The original designations of the clones used are given in brackets: V79-h3A4 [clone V79MZh3A4hOR-1 (42)], V79-h2E1 [V79MZh2E1 (43)], V79-h1A1 [V79MZh1A1 (44)], V79-h1A2 [XEMh1A2-MZ#1 (45)], V79-h1B1 [V79MZh1B1, clones 3–6 (46)], V79-h2A6 (J.Doehmer, manuscript in preparation), V79-r1A1 [XEM2 (37)], V79-r1A2 [XEMd-MZ (47)], V79-r2B1 [SD1 (36)]. Human CYP reductase was co-expressed in V79-h3A4 cell line and in V79-h1B1 clone 6, but not in any of the other cell lines used. The reasons are historical, since several of the CYPs used have not yet been co-expressed with heterologous CYP reductase. They use the endogenous CYP reductase, whose expression level is moderate and may not be optimal for each CYP. Co-expression of human CYP reductase had only minor effects on the enzyme activity using standard substrates with human CYP1A2 and 1B1 expressed in V79 cells (46,48), but substantially enhanced the activity of expressed human CYP3A4 (42). For this reason, a cell line which co-expresses human CYP reductase was used to study the metabolism of MP and EP by CYP3A4. For human CYP1B1, cell lines with and without co-expressed CYP reductase were available. They were used to study the influence of enhanced CYP reductase activity on the metabolite profiles formed by a representative CYP.

The cells were grown in Dulbecco's modified Eagle's medium supplemented with fetal bovine serum (5%), penicillin (100 U/ml) and streptomycin (100 µg/ml). The cultures were maintained at 37°C in a humidified atmosphere containing 5% CO2.

Liver microsomes
Rat liver microsomes were prepared from adult male Wistar rats (Schönwalde, Germany) as described elsewhere (49). Some animals were treated with 3-methylcholanthrene (MC), a potent inducer of CYP1 enzymes. MC was dissolved in sunflower oil (32 mg/ml) and administered i.p. at a dose of 80 mg/kg body weight 24 h before the animals were killed. Protein content was determined using a bicinchonic acid assay kit of Pierce (Rockford) with bovine serum albumin as a standard. Microsomal aliquots were stored at –80°C.

Samples of resected human livers were a generous gift of Dr M.Arand (Department of Toxicology, University of Mainz, Germany). The liver samples were stored at –80°C and microsomes were prepared as described with rat liver.

Metabolism studies with genetically engineered V79 cells
A total of 1.5x106 cells and 10 ml medium were added to 75 cm2 culture flasks and cultured for 14 h. The medium was then replaced with 3 ml new medium containing MP (10 µM) or EP (5 µM). The test compounds had been added using DMSO as the solvent (final concentration 0.0125–0.025%). Control cultures only received the solvent. After incubation for 10 h (unless specified otherwise), medium and cells were harvested and combined. At the end of each incubation, the cell number was determined from a separate, parallel incubation using a Casy-1 cell counter (Schärfke System, Reutlingen, Germany). Under the conditions used, the test compounds showed no influence on the number or morphology of the cells.

Metabolism studies with liver microsomes
MP or EP (6 µM unless specified otherwise) was incubated at 37°C for 15 min with microsomal suspensions (0.4 mg protein/ml) from humans, control and MC-treated rats. The reactions were started by adding the test compound dissolved in DMSO (final concentration 0.6%). The incubations were performed in a final volume of 1 ml of 50 µM potassium phosphate buffer (pH 7.4) containing 3 mM MgCl2, 1 mM EDTA and an NADPH-generating system consisting of 1 mM NADP, 5 mM glucose 6-phosphate and 0.7 U glucose 6-phosphate dehydrogenase (Merck, Darmstadt, Germany). Reactions were stopped by cooling on ice. The NADPH-generating system was omitted in negative control incubations.

Chemical analysis of the metabolites of MP and EP
The incubation mixtures were adjusted to pH 2.0 with phosphoric acid. When quantification of the metabolites was sought, 12 nmol pyrene was added as an internal standard. The metabolites were isolated under nitrogen by liquid–liquid extraction on Extrelut 20 columns (Merck) using ethyl acetate (3x5 ml) as the eluent. The solvent was evaporated by centrifugation at 30°C under reduced pressure. The residue was dissolved in 500 µl DMSO/acetonitrile (0.5:10, v/v) and subjected to HPLC analysis.

Separation of metabolites was achieved on a Waters 600E delivery system equipped with a NovaPakC18 column (3.9x300 mm, 4 µm) and a Waters 996 photodiode array detector coupled to a Shimadzu RF 551 fluorescence detector using the following solvent gradient of eluent A (water adjusted with phosphoric acid to pH 2.3) and eluent B (acetonitrile/methanol 1:1 v/v): A from 80 to 16% within 20 min, then a 15 min linear gradient to 10% A, and finally a 5 min linear gradient to 2% A. The flow rate was 1 ml/min at 30°C column temperature. In general, 20 µl of the sample was injected using a Waters 717 autosampler.

Metabolites were quantified from their fluorescence signal ({lambda}ex 273 nm/{lambda}em 385 nm). Response factors and recoveries were determined for MP, EP, {alpha}-HMP, 1-pyrenylcarboxaldehyde, COOH-P, {alpha}-HEP, 1-acetylpyrene, {omega}-HEP, 1-pyrenylacetic acid and 1-hydroxypyrene (used as a model for phenolic metabolites of MP and EP). The recoveries were 90–97%. The response factors were within a range of 100 ± 15% of the value determined for {alpha}-HMP, except for the carbonyl derivatives (1-pyrenylcarboxaldehyde and 1-acetylpyrene), whose response factors were markedly lower.

Co-chromatography with synthetic standards under different HPLC conditions, photodiode array HPLC recording the UV spectra in the range between 220 and 500 nm and matching with standard spectra were used for the identification of metabolites. GC-MS and HPLC-MS analyses were used for the corroboration of the identification of the metabolite {alpha}-HMP. For HPLC-MS analysis, the HPLC system was coupled with a single quadrupole mass spectrometer (VG Platform, Fisons Instruments, UK). The analytes were separated on a LiChrospher-PAH column (4x250 mm; Merck) with acetonitrile as eluent and analyzed by positive atmospheric pressure chemical ionization under the following conditions: corona 3.5 kV, cone 17 eV, HV lens 40 eV, source temperature 150°C, probe temperature 550°C. For GC-MS analysis, the analytes, dissolved in n-heptane, were separated on a capillary column with cross-linked 5% phenylmethyl silicone gum phase using helium as carrier gas. Eluted analytes were detected by mass spectrometry with electron impact ionization at 70 eV performed in a quadrupole MS-detector (MSD 5972; Hewlett Packard).


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 Materials and methods
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Metabolism of MP in cell lines
When the parental (control) V79 cell line was used, formation of metabolites from MP was not detected. Other materials with fluorescent or UV-absorbing properties similar to those of potential metabolites (comprising the phenanthrene or pyrene chromophore) were also not present. However, the formation of metabolites from MP was detected in all CYP-expressing cell lines. The pattern of metabolites formed substantially differed between cell lines expressing different CYPs, as shown in Figure 2Go for three cell lines. Peak 5 was identified as {alpha}-HMP and peak 4 as its oxidation product COOH-P, whereas the intermediate aldehyde, 1-pyrenylcarboxaldehyde, was not detected (limit of detection: 0.1 pmol/10 h/106 cells, {lambda}ex 269 nm/{lambda}em 446 nm). Peaks 6, 8 and 9 were also formed when {alpha}-HMP, rather than MP, was incubated with a source of CYP (V79-r2B1 cells or liver microsomes from MC-treated rat), suggesting that these metabolites were oxidized at both the ring and the benzylic position. Peaks 3 and 7 were formed from MP, but not from {alpha}-HMP, indicating that they are oxidized only at the ring, but not at the benzylic position. The UV spectra of these metabolites (Figure 3Go) show that the pyrene chromophore is maintained in metabolites 3–6 and 8, whereas the UV spectrum of peak 7 is virtually identical to that published for the trans-4,5-dihydrodiol of MP (35).



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Fig. 2. HPLC profiles of metabolites of MP formed in V79-derived cell lines expressing rat CYP1A1 (upper panel), human CYP1A1 (middle panel) and human CYP3A4 (lower panel). Peak 1, MP; peak 2, pyrene (internal standard); peak 3, ring-oxidized metabolite, pyrene chromophore; peak 4, COOH-P; peak 5, {alpha}-HMP; peaks 6, 8 and 9, ring- and side-chain-oxidized metabolites showing the pyrene chromophore (these peaks were also observed when {alpha}-HMP, rather than MP, was incubated with cells or microsomes); peak 7, ring-oxidized metabolite, phenanthrene chromophore; unnumbered peaks, not classified, as the amounts formed were too small to produce a useful UV spectrum. Peaks with retention times >15 min were only detected in incubations of MP with CYP-expressing cell lines or liver microsomes, but not with control V79 cells. Therefore, it is probable that the peaks represent metabolites.

 


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Fig. 3. UV spectra of metabolites of MP formed in V79-r1A1 cells. The spectra were recorded using a photodiode array detector from the peaks marked with the indicated number in the upper panel of Figure 2Go. Virtually identical UV spectra were observed for the metabolite peaks marked with the same number (middle and lower panels of Figure 2Go, and Figure 6Go) using other metabolizing systems.

 
Time course studies of the formation of metabolites were performed using V79-h1A1, V79-h3A4 and V79-r1A1 cell lines. These studies demonstrated linearity of the amount of product formed with the incubation time, with correction for the increasing cell number, in the range selected for further investigations (10 h). Some changes in the metabolite profile were detected with varying incubation times, but the same metabolites dominated the profile at all time points. For example, the contribution of peak 3 (ring-oxidized metabolite) to the total metabolites in V79-r1A1 cells always amounted to >70% (Figure 4Go). In the other cell lines in which the metabolite profile was studied using varying incubation times (V79-h1A1, V79-h3A4), {alpha}-HMP was the most prominent metabolite at all time points. Its contribution to the total metabolites was usually decreased somewhat with increasing incubation times in favour of COOH-P and/or the putative ring- plus and side-chain-oxidized metabolites (peaks 6, 8 and 9). These secondary metabolites are indicated by solid symbols in Figure 4Go.



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Fig. 4. Influence of the incubation time on the level and profile of metabolites formed from MP formed in V79-r1A1 cells. The left scale of the ordinate is used for the major metabolite, peak 3 (as indicated in the legend to Figure 2Go) (open triangles). The right scale is used for the minor metabolites: {alpha}-HMP (open circles), COOH-P (solid circles), peaks 7 (open squares), 8 (solid triangles) and 9 (solid squares). For the assignment of the peaks see legend to Figure 2Go. Values are means and SD of three cultures.

 
For the estimation of the total rates of metabolism, it was assumed that the fluorometric properties of the metabolites which were not available as standards were equal to those of {alpha}-HMP. The error resulting from this inaccuracy should be small, since all major metabolites, except metabolite 7, contain the pyrene chromophore. The hitherto calculated rates of total MP metabolism varied between 16 and 415 [pmol/10 h/106 cells] between the cell lines used (Table IGo). These rates may be influenced not only by the kind of CYP expressed, but also by differences in their expression levels. The expression levels are not exactly known on a molar level, although the enzyme activity levels using characteristic substrates suggest that they vary within a relatively small range. Therefore, the results indicate that rat 1A1, human 1B1 and human 1A1 are particularly active in metabolizing MP.


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Table I. Formation of the benzylic alcohol metabolite of MP and EP in cell lines expressing individual cytochromes P450a
 
{alpha}-HMP was formed in all CYP-expressing cell lines tested. However, the rate of formation of {alpha}-HMP and the relative importance of this pathway differed substantially between the cell lines (Table IGo). The highest absolute rates of benzylic hydroxylation were detected in the cells expressing human 1B1, human 1A1 and human 3A4, in this order. The relative rates of benzylic hydroxylation ({alpha}-HMP and COOH-P as percent of total metabolites) were highest for human 3A4 (83%), human 2E1 (81%), human 1B1 (80–85%) and human 1A1 (45%), rat 1A2 (45%) and rat 2B1 (45%). It is worth emphasizing that rat 1A1 differed substantially in its regioselectivity from its human orthologue, as {alpha}-HMP only accounted for 3% of the total amount of metabolites formed. Included in these figures are further benzylic oxidation products of MP (aldehyde and carboxylic acid), but not metabolites involving both ring and side chain oxidation (0.5–13% of the total amount of metabolites formed, depending on the cell line used).

Among the further benzylic oxidation products of {alpha}-HMP, only COOH-P, but not the intermediate aldehyde, was observed (Figure 1Go). In general, the amount of COOH-P formed was markedly lower than that of {alpha}-HMP; however, cell line V79-h1A2 formed both metabolites in similar quantities.

Four separate recombinant cell lines expressing human CYP1B1 were used. In one of them, human CYP reductase was co-expressed, in the other three cell lines only the endogenous CYP reductase was present. All four cell lines showed similar rates of metabolism and indistinguishable metabolite profiles from MP, indicating no influence of co-expressed human CYP reductase.

Metabolism of EP in cell lines
EP, as observed with MP, was not metabolized in the V79 control cell line, but the formation of metabolites was detected in all CYP-expressing cell lines. As found with MP, the pattern of metabolites substantially differed between cell lines expressing different CYPs (Figure 5Go). Some metabolites were oxidized only at the ring (peaks 2, 4, 5 and 7), others only at the side chain (peak 6), or others at both the ring and the side chain (peaks 8–10).



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Figure 5. HPLC profiles of metabolites of EP formed in V79-r1A1 (upper panel) and V79-h3A4 cells (lower panel). Peak 1, EP; peaks 2, 4 and 5, ring-oxidized metabolites, pyrene chromophore; peak 3, pyrene (internal standard); peak 6, {alpha}-HEP and {omega}-HEP (shoulder marked by an arrow in Figure 7Go). Further analyses of the peak using chromatographic conditions which lead to separation of the isomeric alcohols showed that only {alpha}-HEP was formed in the cell lines used, whereas in microsomal systems small amounts of {omega}-HEP were additionally formed; peak 7, ring-oxidized metabolite, phenanthrene chromophore; peaks 8–10, ring- and side-chain-oxidized metabolites showing the pyrene chromophore (these peaks were also observed when {alpha}-HEP or {omega}-HEP, rather than EP, was incubated with cells or microsomes); unnumbered peaks, not classified, as the amounts formed were too small to produce a useful UV spectrum. Peaks with retention times >15 min were only detected in incubations of EP with CYP-expressing cell lines or liver microsomes, but not with control V79 cells. Therefore, it is probable that the peaks represent metabolites.

 
The ring-oxidized metabolite eluting in peak 2 (Figure 5Go) was observed with several human CYPs (1A1, 1A2, 1B1 and 3A4), but not with any rat enzyme studied (1A1, 1A2 and 2B1). The UV spectrum of peak 7 is very similar to that of the peak 7 metabolite of MP (Figure 2Go, panel 7), suggesting that this metabolite is a K-region (4,5- or 9,10-)dihydrodiol of EP.

Peak 6 represents {alpha}-HEP, which, however, co-elutes with {omega}-HEP. Separation of the isomeric alcohols was achieved under isocratic HPLC conditions using a mixture of 40% eluent A and 60% eluent B and a flow rate of 0.7 ml/min (retention times of 18.8 min and 19.7 min, respectively). Using these additional analyses, it was shown that only {alpha}-HEP, but no {omega}-HEP, was formed in the cell lines used (limit of detection: 0.1 pmol/10 h/106 cells). The oxidation products of {omega}-HEP (1-pyrenylacetic acid) and {alpha}-HEP (1-acetylpyrene) were not detected either [limits of detection: 0.2 pmol/10 h/106 cells ({lambda}ex 273 nm/{lambda}em 385 nm) and 1 pmol/10 h/106 cells (UV absorption at 240 nm), respectively]. Incubation of {omega}-HEP with either V79-h1A1 cells or liver microsomes from MC-treated rats led to the formation of only small quantities of secondary oxidation products. These products showed the same elution times and UV spectra as peaks 8–10 formed in the metabolism of EP (Figure 5Go). Therefore, it is not likely that substantial amounts of {omega}-HEP were formed from EP in the CYP-expressing cell lines, but were not detected owing to further metabolism.

The total rates of metabolism of EP were estimated assuming that the intensity of the fluorescence of the unidentified metabolites is similar to that of {alpha}-HEP (Table IGo). In V79-h3A4 cells, MP and EP were metabolized at similar rates. In the other eight CYP-expressing cell lines used, EP was metabolized faster than MP by a factor of 1.3 (V79-h2E1) to 11 (V79-h1B1 clone 6). The total rates of metabolism of EP varied between 22 and 1039 pmol/10 h/106 cells in the order: human 1B1 (with co-expressed human CYP reductase) > rat 1A1 > human 1B1 > human 1A1 > human 1A2 > rat 2B1 >> human 3A4 (with co-expressed human CYP reductase) {approx} rat 1A2 {approx} human 2A6 {approx} human 2E1. This order is similar to that observed for MP, except that the relative activity of human in particular if co-expressed with human CYP reductase, was increased with EP (Table IGo).

Each CYP-expressing cell line formed {alpha}-HEP in addition to varying amounts and patterns of other metabolites of EP (Table IGo). The highest absolute rates of formation of {alpha}-HEP were detected with the cells expressing human 1A1, human 1B1 and rat 1A1. The order of the cell lines according to the rate of this reaction was similar to that for the biotransformation of MP to {alpha}-HMP, except for rat 1A1. This was due to an alteration in the regioselectivity in the metabolism of EP (15% {alpha}-HEP of the total metabolites) as compared with MP (only 3% {alpha}-HMP of the total metabolites). However, the highest relative rates of benzylic hydroxylation of EP (as percent of total metabolism) were observed with h1A1 (51%), h1B1 (41–43%) and h3A4 (38%).

Like with MP, co-expression of human CYP reductase did not affect the pattern of metabolites formed from EP in V79-h1B1 cells. However, unlike with MP, it increased the rate of metabolism of EP (Table IGo). This may be due to the fact that EP was metabolized much faster than MP, requiring a higher rate of CYP reduction.

Metabolism of MP and EP by rat and human hepatic microsomes
After incubation of MP with NADPH-supplemented human liver microsomes, three major product peaks were detected (Figure 6Go, upper panel). The pyrene chromophore was retained in all product peaks. Peaks 5 and 4 showed the retention time and UV spectra of {alpha}-HMP and COOH-P, respectively. Formation of {alpha}-HMP was further confirmed by GC-MS and LC-MS analyses. It is therefore concluded that the third peak (peak 3) must comprise a ring-oxidized, phenolic metabolite(s). This simple pattern of metabolites is dominated by the side-chain-oxidized metabolites (38–64% of total metabolites in incubations using microsomes from four subjects, Table IIGo). It is similar to metabolite patterns found with the V79-derived cells expressing h3A4, the quantitatively predominant CYP in human liver, or human 1A1 and 1B1, the forms showing the highest activity with MP (Table IGo).



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Fig. 6. HPLC profiles of metabolites of MP formed in hepatic microsomal systems from human (upper panel), control rat (middle panel) and MC-treated rat (lower panel). For the assignment of the peaks see legend to Figure 2Go

 

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Table II. Formation of the benzylic alcohol metabolite of MP and EP in hepatic microsomesa
 
A similar pattern of metabolites was observed when MP was incubated with liver microsomes from untreated rats (Figure 6Go, middle panel), rather than from humans. However, the use of microsomes from MC-treated rats led to a strong increase in the amount of ring-oxidized products formed (peaks 3 and 6–9) and a marked decrease in the side-chain-oxidized metabolites (Figure 6Go, lower panel). The resulting metabolite profile was similar to that observed with V79 cells expressing rat 1A1 (Figure 2Go). This similarity is plausible since 1A1 is strongly induced by MC, and it metabolizes MP very efficiently (Table IGo).

Incubation of EP with human hepatic microsomes resulted in the formation of several product peaks (Figure 7Go, upper panel). One of the peaks (peak 6) showed the retention time and the UV spectrum of {alpha}-HEP (which are virtually identical to those of its isomer, {omega}-HEP). Rechromatography of this peak, using conditions which lead to separation of the isomers, indicated that peak 6 is comprised of both alcohols (~80% {alpha}-HEP and 20% {omega}-HEP). Incubation of EP with NADPH-fortified liver microsomes from MC-treated rats produced a relatively complex pattern of metabolites (Figure 7Go, lower panel), which was very similar to that found with the V79-derived cell line expressing rat 1A1 (Figure 2Go). However, further analysis of peak 6 demonstrated the formation of {alpha}-HEP (90% of peak 6) as well as {omega}-HEP (10% of peak 6) in the presence of the rat liver microsomal system, whereas only the {alpha}-isomer was found in incubations with V79-r1A1 cells, as well as with all other cell lines used (see above).



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Fig. 7. HPLC profiles of metabolites of EP formed in hepatic microsomal systems from human (upper panel) and from MC-treated rat (lower panel). For the assignment of the peaks see legend to Figure 5Go

 
Increase of the substrate concentration of MP and EP from 6 to 60 µM in incubations with human liver microsomes led to enhanced rates of metabolism, but the profiles of metabolites remained virtually unaltered (Table IIGo).


    Discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Our results demonstrate that the initial step in the bioactivation of MP and EP, the hydroxylation at the benzylic position, is extensively catalysed by both rat and human CYPs. Indeed, all six human and three rat CYPs tested were capable of forming the benzylic alcohols from both hydrocarbons.

Using human hepatic microsomes, {alpha}-HMP was the predominant primary metabolite of MP. This correlates with the observation that 3A4, the most abundant CYP in human liver, mediated the formation of {alpha}-HMP (including its oxidation product, COOH-P) with high selectivity (83% of total metabolites). This preference of 3A4 for oxidation at the exocyclic position is in good agreement with results published for the metabolism of 5-methylchrysene and 6-methylchrysene (50). The pattern of metabolites formed from MP by hepatic microsomes from untreated rats was similar to that formed by human liver microsomes. However, since the major constitutive rat CYPs have not yet been expressed in V79 cells, it could not be examined whether the activity of these forms could explain the formation of the metabolites detected.

Treatment of rats with MC, which induces CYP1 enzymes, led to a profound change in the pattern of microsomal metabolites. Ring oxidation was strongly enhanced, whereas formation of {alpha}-HMP was decreased both in percent of total metabolites and absolute rate. A part of the decrease in the amount of {alpha}-HMP detected may be due to its further metabolism at the ring system, as ~9% of the total metabolites appeared to be ring- as well as side-chain-oxidized. Moreover, it is known that treatment of rats with MC leads to down-regulation of 2C9, the major constitutive form in male rat liver (51). It is also possible that induced CYPs compete with constitutive forms for CYP reductase, leading to reduced metabolic rates by the constitutive forms. The profile of metabolites observed with microsomes from MC-treated rats was very similar to that found in the presence of V79-r1A1 cells, suggesting a major role of 1A1 in the metabolism of MP by microsomes from MC-treated rats. It is unlikely that a similarly dramatic shift in the metabolism of MP would be evoked in humans by enzyme induction of the MC type (e.g. by autoinduction by PAHs), as human 1A1 substantially differed in its regioselectivity from rat 1A1. With all three human CYP1 enzymes, {alpha}-HMP was a major metabolite formed from MP.

It should be noted that very similar metabolite profiles were detected in microsomal systems and in living cells expressing a corresponding individual CYP, i.e. human liver microsomes and cells expressing human 3A4, or liver microsomes from MC-treated rats and cells expressing rat 1A1. These results suggest that no major phase-I metabolites were undetected in the cellular systems due to conjugation to phase-II metabolites, which were not analyzed in the present study.

Elongation of the side chain from methyl to ethyl led to a general increase in the metabolism rates. This effect is not the consequence of enhanced metabolism at the side chain due to the presence of an additional carbon which may be hydroxylated. The ethyl group was preferentially hydroxylated in the benzylic ({alpha}) position rather than in the terminal ({omega}) position. In fact, {omega}-hydroxylation of EP was not detected in the cell lines expressing individual CYPs, but was found to be a minor metabolic pathway in the microsomal systems. It is probable that other CYP forms than those expressed in V79 cells in the present study catalysed the {omega}-hydroxylation of EP. The observation that the {alpha}-position of EP is the preferred position for side chain hydroxylation agrees with results obtained in other studies, using other ethyl-substituted aromatic compounds, such as ethylbenzene (52) and 7-ethylbenz[a]anthracene (53).

The results of the present study demonstrate that genetically engineered cell lines are useful to compare the metabolism of xenobiotics by enzymes from humans and laboratory animals, and to assess the possible role of individual enzyme forms. Our results suggest that benzylic hydroxylation is a major initial step of metabolism of MP and EP in humans. As shown in previous studies, the second step of the bioactivation of MP and EP leading to potent mutagens, the O-sulfonation, is also efficiently carried out by human enzymes (3134,54).

We are now constructing cell lines which co-express CYP and sulfotransferase forms which mediate sequential steps in the activation of alkylated PAHs. These cells can be used not only for metabolism studies, but also for the detection of toxicological effects, such as the formation of DNA adducts and the induction of mutations or cytotoxic effects. These systems may be useful for the detection of those alkylated PAHs which are most efficiently activated via this pathway by human enzymes. The enzymology of the activation may be dissected and then used for the prediction of tissues and subjects at high risks, as well as for the production of an adequate animal model for in vivo studies.


    Acknowledgments
 
We thank Mrs Brigitte Knuth for excellent technical assistance, and Dr Charles Falany (University of Alabama at Birmingham) for critical reading of the manuscript. This work was financially supported by the Bundesministerium für Bildung, Wissenschaft, Forschung und Technologie (BMBF; grant 0311243).


    Notes
 
2 To whom correspondence should be addressed Email: glatt{at}www.dife.de Back


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

  1. IARC (1983) IARC Monographs on the Evaluation of the Carcinogenic Risk of Chemicals to Humans. Polynuclear Aromatic Compounds. Part 1. Chemicals, Environmental, and Experimental Data. Vol. 32. IARC Scientific Publications, IARC, Lyon.
  2. IARC (1984) IARC Monographs on the Evaluation of the Carcinogenic Risk of Chemicals to Humans. Polynuclear Aromatic Compounds. Part 2. Carbon Blacks: Mineral Oils and some Nitroarenes. Vol. 33. IARC Scientific Publications, IARC, Lyon.
  3. IARC (1984) IARC Monographs on the Evaluation of the Carcinogenic Risk of Chemicals to Humans. Polynuclear Aromatic Compounds. Part 3. Industrial Exposures in Aluminium Production, Coal Gasification, Coke Production, and Iron and Steel Founding. Vol. 34. IARC Scientific Publications, IARC, Lyon.
  4. IARC (1985) IARC Monographs on the Evaluation on the Carcinogenic Risk of Chemicals to Humans. Polynuclear Aromatic Compounds. Part 4. Bitumens: Coal-Tars and Derived Products, Shale-Oils and Soots. Vol. 35. IARC Scientific Publications, IARC, Lyon.
  5. IARC (1986) IARC Monographs on the Evaluation of the Carcinogenic Risk of Chemicals to Humans. Tobacco Smoking. Vol. 38. IARC Scientific Publications, IARC, Lyon.
  6. Radke,M. (1987) Organic geochemistry of aromatic hydrocarbons. In Brooks, J. and Welte, D.H. (eds) Advances in Petroleum Geochemistry. Academic Press, London, UK, vol. 2, pp. 141–207.
  7. Radke,M., Garrigues,P. and Willsch,H. (1990) Methylated dicyclic and tricyclic aromatic hydrocarbons in crude oils from the Handil field, Indonesia. Org. Geochem., 15, 17–34.[ISI]
  8. Schenck,P.A. and de Leeuw,J.W. (1982) Molecular organic geochemistry. In Hutzinger,O. (ed.) Handbook of Environmental Chemistry. Springer-Verlag, Berlin, vol. B/1B, pp. 111–129.
  9. Rogge,W.R., Hildemann,L.M., Mazurek,M.A., Cass,G.R. and Simoneit, B.R.T. (1993) Sources of fine organic aerosol: 2. Noncatalyst and catalyst-equipped automobiles and heavy-duty diesel trucks. Environ. Sci. Technol., 27, 636–651.[ISI]
  10. Flesher,J.W., Myers,S.R. and Stansbury,K.H. (1990) The site of substitution of the methyl group in the bioalkylation of benzo[a]pyrene. Carcinogenesis, 11, 493–496.[Abstract]
  11. Flesher,J.W., Myers,S.R. and Blake,J.W. (1986) Bioalkylation of polynuclear aromatic hydrocarbons: a predictor of carcinogenic activity. In Cooke,M. and Dennis,A.J. (eds) Polynuclear Aromatic Hydrocarbons: Chemistry, Characterization and Carcinogenesis. Battelle Press, Columbus, OH, pp. 271–284.
  12. Jacob,J., Grimmer,G., Mohr,U., Emura,M., Riebe-Imre,M., Raab,G. and Knebel,J. (1993) Metabolic activation of chrysene and benzo[a]pyrene in hamster, rat and human in vitro lung cell cultures. Polycyclic Aromatic Comp., 3 (suppl.), 1175–1182.
  13. Stansbury,K.H., Flesher,J.W. and Gupta,R.C. (1994) Mechanism of aralkyl-DNA adduct formation from benzo[a]pyrene in vivo. Chem. Res. Toxicol., 7, 254–259.[ISI][Medline]
  14. Glatt,H.R. and Oesch,F. (1986) Structural and metabolic parameters governing the mutagenicity of polycyclic aromatic hydrocarbons. In de Serres,F.J. (ed.) Chemical Mutagens: Principles and Methods for Their Detection. Plenum Press, New York, NY, vol. 10, pp. 73–127.
  15. Thakker,D.R., Yagi,H., Levin,W., Wood,A.W., Conney,A.H. and Jerina, D.M. (1985) Polycyclic aromatic hydrocarbons: metabolic activation to ultimate carcinogens. In Anders,M.W. (ed.) Bioactivation of Foreign Compounds. Academic Press, Inc., pp. 177–242.
  16. Watabe,T., Ishizuka,T., Isobe,M. and Ozawa,N. (1982) A 7-hydroxymethyl sulfate ester as an active metabolite of 7,12-dimethylbenz[a]anthracene. Science, 215, 403–405.[ISI][Medline]
  17. Miller,J.A. and Surh,Y.-J. (1994) Sulfonation in chemical carcinogenesis. In Kauffman,F.C. (ed.) Handbook of Pharmacology, Vol. 112, Conjugation-Deconjugation Reactions in Drug Metabolism and Toxicity. Springer-Verlag, Berlin, pp. 429–458.
  18. Cavalieri,E., Roth,R. and Rogan,E. (1979) Hydroxylation and conjugation at the benzylic carbon atom: a possible mechanism of carcinogenic activation for some methyl-substituted aromatic hydrocarbons. In Jones, P.W. and Leber, P. (eds) Polynuclear Aromatic Hydrocarbons. Ann Arbor Science Publishers, Ann Arbor, MI, pp. 517–529.
  19. Landsiedel,R., Engst,W., Seidel,A. and Glatt,H.R. (1996) Physico-chemical properties and mutagenicity of benzylic compounds. Exp. Toxicol. Pathol., 48 (Suppl. 2), 215–222.
  20. Landsiedel,R., Engst,W., Scholtyssek,M., Seidel,A. and Glatt,H.R. (1996) Benzylic sulphuric acid esters react with diverse functional groups and often form secondary reactive species. Polycyclic Aromat. Comp., 11, 341–348.[ISI]
  21. Rice,J.E., Rivenson,A., Braley,J. and LaVoie,E.J. (1987) Methylated derivatives of pyrene and fluorene: evaluation of genotoxicity in the hepatocyte/DNA repair test and tumorigenic activity in newborn mice. J. Toxicol. Environ. Health, 21, 525–532.[ISI][Medline]
  22. Surh,Y.-J., Liem,A., Miller,E.C. and Miller,J.A. (1990) The strong hepatocarcinogenicity of the electrophilic and mutagenic metabolite 6-sulfooxymethylbenzo[a]pyrene and its formation of benzylic DNA adducts in the livers of infant male B6C3F1 mice. Biochem. Biophys. Res. Commun., 172, 85–91.[ISI][Medline]
  23. Surh,Y.-J., Blomquist,J.C. and Miller,J.A. (1990) Activation of 1-hydroxymethylpyrene to an electrophilic and mutagenic metabolite by rat hepatic sulfotransferase activity. In Witmer,C.M., Snyder,R.R., Jollow,D.J., Kalf,G.F., Kocsis,J.J. and Sipes,I.G. (eds) Biological Reactive Intermediates IV: Molecular and Cellular Effects and Their Impact on Human Health. Plenum Press, New York, NY, pp. 383–391.
  24. Glatt, H.R., Werle-Schneider,G., Enders,N., Monnerjahn,S., Pudil,J., Czich,A., Seidel,A. and Schwarz,M. (1994) 1-Hydroxymethylpyrene and its sulfuric acid ester: toxicological effects in vitro and in vivo, and metabolic aspects. Chem. Biol. Interact., 92, 305–319.[ISI][Medline]
  25. Flesher,J.W., Horn,J. and Lehner,A.F. (1997) 6-Sulfooxymethylbenzo[a]pyrene is an ultimate electrophilic and carcinogenic form of the intermediary metabolite 6-hydroxymethylbenzo[a]pyrene. Biochem. Biophys. Res. Commun., 234, 554–558.[ISI][Medline]
  26. Flesher,J.W. and Sydnor,K.L. (1973) Possible role of 6-hydroxymethylbenzo[a]pyrene as a proximate carcinogen of benzo[a]pyrene and 6-methylbenzo[a]pyrene. Int. J. Cancer, 11, 433–437.[ISI][Medline]
  27. Conney,A.H., Chang,R.L., Jerina,D.M. and Wei,S.J.C. (1994) Studies on the metabolism of benzo[a]pyrene and dose-dependent differences in the mutagenic profile of its ultimate carcinogenic metabolite. Drug Metab. Rev., 26, 125–163.[ISI][Medline]
  28. Grimmer,G. (1988) Polycyclische aromatische Kohlenwasserstoffe. In Eisenbrand,G., Frank,H.K., Grimmer,G., Hapke,H.-J., Thier,H.-P. and Weigert,G. (eds) Derzeitige Belastung und Trends bei der Belastung der Lebensmittel durch Fremdstoffe. Verlag W. Kohlhammer GmbH, Karlsruhe, pp. 151–187.
  29. Okamoto, H. and Yoshida,D. (1981) The isolation identification of mutation enhancing principles from cellulose pyrolysate. Agric. Biol. Chem., 45, 1291–1294.[ISI]
  30. Pancirov,R.J. and Brown,R.A. (1977) Polynuclear aromatic hydrocarbons in marine tissues. Environ. Sci. Technol., 11, 989–992.[ISI]
  31. Glatt,H.R., Pauly,K., Czich,A., Falany,J.L. and Falany,C.N. (1995) Activation of benzylic alcohols to mutagens by rat and human sulfotransferases expressed in Escherichia coli. Eur. J. Pharmacol., 293, 173–181.[Medline]
  32. Glatt,H.R. (1997) Bioactivation of mutagens via sulfation. FASEB J., 11, 314–321.[Abstract/Free Full Text]
  33. Glatt,H.R., Bartsch,I., Christoph,S., Coughtrie,M.W.H., Falany,C.N., Hagen,M., Landsiedel,R., Pabel,U., Phillips,D.H., Seidel,A. and Yamazoe,Y. (1998) Sulfotransferase-mediated activation of mutagens studied using heterologous expression systems. Chem. Biol. Interact., 109, 195–219.[ISI][Medline]
  34. Hagen,M., Pabel,U., Landsiedel,R., Bartsch,I. and Glatt,H.R. (1998) Expression of human estrogen sulfotransferase in Salmonella typhimurium: differences between hHST and hEST in the enantioselective activation of 1-hydroxyethylpyrene to a mutagen. Chem. Biol. Interact., 109, 249–253.[ISI][Medline]
  35. Rice,J.E., Geddie,N.G., DeFloria,M.C. and LaVoie,E.J. (1988) Structural requirements favoring mutagenic activity among methylated pyrenes in S. typhimurium. In Cooke,M. and Dennis,A. (eds) Polynuclear Aromatic Hydrocarbons: A Decade of Progress. Battelle Press, Columbus, OH, pp. 773–785.
  36. Doehmer,J., Dogra,S., Friedberg,T., Monier,S., Adesnik,M., Glatt,H.R. and Oesch,F. (1988) Stable expression of rat cytochrome P450IIB1 cDNA in Chinese hamster cells (V79) and mutagenicity of aflatoxin B1. Proc. Natl Acad. Sci. USA, 85, 5769–5773.[Abstract]
  37. Dogra,S., Doehmer,J., Glatt,H.R., Mölders,R., Siegert,P., Friedberg,T., Seidel,A. and Oesch,F. (1990) Stable expression of rat cytochrome P450IA1 cDNA in V79 Chinese hamster cells and their use in mutagenicity testing. Mol. Pharmacol., 37, 608–613.[Abstract]
  38. Glatt,H.R., Gemperlein,I., Setiabudi,F., Platt,K.-L. and Oesch,F. (1990) Expression of xenobiotic-metabolising enzymes in propagatable cell cultures and induction of micronuclei by 13 compounds. Mutagenesis, 5, 241–249.[Abstract]
  39. Glatt,H.R., Hornhardt,S., Pauly,K., Piée-Staffa,A., Seidel,A. and Czich,A. (1994) Activation of promutagens by endogenous and heterologous sulfotransferases expressed in continuous cell cultures. Toxicol. Lett., 72, 13–21.[ISI][Medline]
  40. Autorenkollektiv (1988) Organikum. VEB Deutscher Verlag der Wissenschaften, Berlin.
  41. Enders,N., Seidel,A., Monnerjahn,S. and Glatt,H.R. (1993) Synthesis of 11 benzylic sulfate esters, their bacterial mutagenicity and its modulation by chloride, bromide and acetate anions. Polycyclic Aromatic Comp., 3 (suppl.), 887–894.
  42. Schneider,A., Schmalix,W.A., Siruguri,V., de Groene,E.M., Horbach,G.J., Kleingeist,B., Lang,D., Bocker,R., Belloc,C., Beaune,P., Greim,H. and Doehmer,J. (1996) Stable expression of human cytochrome P450 3A4 in conjunction with human NADPH-cytochrome P450 oxidoreductase in V79 Chinese hamster cells. Arch. Biochem. Biophys., 332, 295–304.[ISI][Medline]
  43. Schmalix,W.A., Barrenscheen,M., Landsiedel,R., Janzowski,C., Eisenbrand,G., Gonzalez,F., Eliasson,E., Ingelman-Sundberg,M., Perchermeier,M., Greim,H. and Doehmer,J. (1995) Stable expression of human cytochrome P450 2E1 in V79 Chinese hamster cells. Eur. J. Pharmacol., 293, 123–131.[Medline]
  44. Schmalix,W.A., Maser,H., Kiefer,F., Reen,R., Wiebel,F.J., Gonzalez,F., Seidel,A., Glatt,H.R., Greim,H. and Doehmer,J. (1993) Stable expression of human cytochrome P450 1A1 cDNA in V79 Chinese hamster cells and metabolic activation of benzo[a]pyrene. Eur. J. Pharmacol., 248, 251–261.[Medline]
  45. Wölfel,C., Heinrich-Hirsch,B., Schulz-Schalge,T., Seidel,A., Frank,H., Ramp,U., Wächter,F., Wiebel,F.J., Gonzalez,F., Greim,H. and Doehmer,J. (1992) Genetically engineered V79 Chinese hamster cells for stable expression of human cytochrome P450IA2. Eur. J. Pharmacol., 228, 95–102.[Medline]
  46. Luch,A., Coffing,S.L., Tang,Y.M., Schneider,A., Soballa,V., Greim,H., Jefcoate,C.R., Seidel,A., Greenlee,W.F., Baird,W.M. and Doehmer,J. (1998) Stable expression of human cytochrome P450 1B1 in V79 Chinese hamster cells and metabolically catalyzed DNA adduct formation of dibenzo[a,l]pyrene. Chem. Res. Toxicol., 11, 686–695.[ISI][Medline]
  47. Wölfel,C., Platt,K.-L., Dogra,S., Glatt,H.R., Wächter,F. and Doehmer,J. (1991) Stable expression of rat cytochrome P450IA2 cDNA and hydroxylation of 17b-estradiol and 2-aminofluorene in V79 Chinese hamster cells. Mol. Carcinog., 4, 489–498.[ISI][Medline]
  48. Schmalix,W.A., Lang,D., Schneider,A., Böcker,R., Greim,H., and Doehmer,J. (1996) Stable expression and co-expression of human cytochrome P450 oxidoreductase and CYP1A2 in V79 Chinese hamster cells: sensitivity to quinones and biotransformation of 7-alkoxyresorufin and triazines. Drug Metab. Dispos., 24, 1314–1319.[Abstract]
  49. Bücker,M., Golan,M., Schmassmann,H.U., Glatt,H.R., Stasiecki,P. and Oesch,F. (1979) The epoxide hydratase inducer trans-stilbene oxide shifts the metabolic epoxidation of benzo[a]pyrene from the bay- to the K-region and reduces its mutagenicity. Mol. Pharmacol., 16, 656–666.[Abstract]
  50. Koehl,W., Amin,S., Staretz,M.E., Ueng,Y.F., Yamazaki,H., Tateishi,T., Guengerich,F.P. and Hecht,S.S. (1996) Metabolism of 5-methylchrysene and 6-methylchrysene by human hepatic and pulmonary cytochrome P450 enzymes. Cancer Res., 56, 316–324.[Abstract]
  51. Safa,B., Lee,C. and Riddick,D.S. (1997) Role of the aromatic hydrocarbon receptor in the suppression of cytochrome P450 2C11 by polycyclic aromatic hydrocarbons. Toxicol. Lett., 90, 163–175.[ISI][Medline]
  52. McMahon,R.E. and Sullivan,H.R. (1966) Microsomal hydroxylation of ethyl benzene: stereo-specifity and the effect of phenobarbital induction. Life Sci., 5, 921–926.[Medline]
  53. McKay,S., Farmer,P.B., Cary,P.D. and Grover,P.L. (1987) The metabolism of 7-ethylbenz[a]anthracene by rat liver microsomal preparations. Drug Metab. Dispos., 15, 682–694.[Abstract]
  54. Glatt,H.R., Seidel,A., Harvey,R.G. and Coughtrie,M.W.H. (1994) Activation of benzylic alcohols to mutagens by human hepatic sulphotransferases. Mutagenesis, 9, 553–557.[Abstract]
Received August 14, 1998; revised May 17, 1999; accepted June 4, 1999.