Influence of ß-Naphthoflavone on 7,12-Dimethylbenz(a)anthracene Metabolism, DNA Adduction, and Tumorigenicity in Rainbow Trout

Tracy L. Weimer*, Ashok P. Reddy{dagger}, Ulrich Harttig{dagger}, David Alexander{ddagger}, S. Craig Stamm§, Michael R. Miller§,1, William Baird{dagger},||, Jerry Hendricks{dagger} and George Bailey{dagger},||

* Department of Pharmacology and Toxicology, West Virginia University, Morgantown, West Virginia 26506; {dagger} Department of Environmental and Molecular Toxicology, Oregon State University, Corvallis, Oregon 97331; {ddagger} Environmental Toxicology Center, Department of Pharmacology, University of Wisconsin, Madison, Wisconsin 53705; § Department of Biochemistry, West Virginia University, Morgantown, West Virginia 26506; CDC/NIOSH, Health Effects Laboratory Division, Physiology Pathology Research Branch, Morgantown, West Virginia 26505; and || Environmental Health Sciences Center, Oregon State University, Corvallis, Oregon 97331

Received May 11, 2000; accepted June 29, 2000


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Metabolism, DNA adduction, and tumor induction by 7,12-dimethylbenz(a)anthracene (DMBA) were examined in cultured trout liver cells and in vivo in trout. Modulating CYP1A1 activity indicated this enzyme plays a significant role in metabolizing DMBA to water-soluble compounds in isolated trout liver cells. The major DMBA metabolites identified in trout liver cells were 10-, 11-, 8,9-, and 5,6-DMBA dihydrodiols, and DMBA, 2- or 3- or 4-phenol; 7-OH-methyl-12-methyl-benz(a)anthracene and 12-OH-methyl-7-methyl-benz(a)anthracene were minor metabolites. A very small amount of DMBA-3,4-dihydrodiol was detected, and polar metabolites, which did not migrate with any DMBA metabolite standards, were observed. Incubating trout hepatocytes with DMBA-3,4-dihydrodiol produced three prominent, nonpolar adducts indistinguishable from those in mouse embryo cells. However, DMBA-DNA adducts, formed in trout in vivo or in trout liver cells exposed to DMBA, were predominantly more polar than those formed in mouse embryo fibroblasts, and levels of DMBA-DNA adducts formed in trout liver cells were not significantly altered by modulating CYP1A1 activity. No significant repair of DMBA-DNA adducts was detected in cultured trout liver cells over a 48-h period, supporting previous studies indicating that fish are less efficient than mammals in repairing polyaromatic hydrocarbon DNA adducts. Compared to animals receiving DMBA alone, ß-naphthoflavone pretreatment in vivo did not affect hepatic CYP1A1, DMBA-DNA adducts, nor hepatic tumor response; but did significantly reduce tumor response in two other target organs. These results collectively indicate that DMBA bioactivation to DNA-binding metabolites in trout liver cells and mouse embryo cells predominantly involve different metabolic pathways to form the DNA-binding intermediates.

Key Words: 7,12-dimethylbenz(a)anthracene; trout; liver cells; CYP1A1; DNA adducts.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
7,12-Dimethylbenz(a)anthracene (DMBA) is one of the most carcinogenic of the polycyclic aromatic hydrocarbons (PAHs). It is extremely efficient at inducing tumors in mammalian species and its mechanism of bioactivation and interaction with DNA in mammalian systems has been studied extensively. In mammalian cells, DMBA can be bioactivated via formation of a reactive "bay region" diol-epoxide metabolite, which subsequently adducts to adenine and guanine residues in DNA (Baird and Dipple, 1977Go; Dipple et al., 1983Go, 1984Go). In addition, DMBA can be activated by one-electron oxidation, as demonstrated by the depurinating DNA adducts observed in mouse skin (Devanesan et al., 1993Go). DMBA is also highly tumorigenic in rainbow trout (Onchorhynchus mykiss) (Fong et al., 1993Go; Reddy et al., 1995Go). While two studies have reported DMBA phase I and II metabolites in trout (Schnitz et al., 1987Go; Schnitz and O'Connor, 1992Go), the metabolic pathway(s) resulting in DMBA bioactivation and detoxification in teleosts are not well defined. DMBA is known to bind covalently to DNA in trout and there is evidence that the overall level of adduct formation is comparable to that in mammals (Schnitz and O'Connor, 1992Go), but it is unclear if these adducts are identical to those formed in mammals. Smolarek et al. (1987) used HPLC to characterize DMBA-DNA adducts formed in a trout gonadal cell line, and reported a profile of adducts very different from those produced in mammalian cells. Specifically, adducts arising from the bay region diol epoxide appeared to be absent. However, trout gonadal cells in vivo are not known targets for DMBA damage, and the relationship of these findings to DMBA-DNA binding and carcinogenesis in vivo is unclear.

Formation of the genotoxic bay region diol epoxide of DMBA in mammals proceeds through the action of the cytochrome P450 (CYP) family of enzymes. The PAH-inducible isozymes of the 1A and 1B families, as well as the phenobarbital-inducible isozymes of the 2B family have been shown to participate in mammalian metabolism of DMBA (Diamond et al., 1972Go; Dipple et al., 1984Go; McCord et al., 1988Go; Morrison et al., 1991Go). Recently, Jefcoate's laboratory (Christou et al., 1994Go; Pottenger and Jefcoate, 1990Go; Savas et al., 1993Go) has identified CYP1B1 as a prominent enzyme-metabolizing DMBA to the 3,4-diol in mammalian cells. However, {alpha}-naphthoflavone, a competitive inhibitor of mammalian CYP1A-mediated PAH metabolism, reduces DMBA-DNA adduct formation in hamster embryo cells in culture (Diamond et al., 1972Go) and inhibits binding of DMBA to DNA in the skin of NIH Swiss and C57 BL mice (Dipple et al., 1984Go). Studies have not established which CYP isozyme(s) is responsible for DMBA-DNA binding in trout. A CYP1B1 orthologue has recently been identified in teleosts (Godard et al., 1999Go); however, the capacity of this enzyme to metabolize DMBA has not yet been reported. Phenobarbital does not induce the CYP2B-related genes in teleosts (Kleinow et al., 1990Go; Miranda et al., 1990Go; Stegeman et al., 1990Go); however, phenobarbital does induce CYP1A1 expression, accompanied by aromatic hydrocarbon receptor transformation (Sadar et al., 1996Go). Mammalian CYP1A2 evolved from a relatively recent gene duplication, and a true CYP1A2 orthologue is not present in trout (Goksoyr et al., 1991Go). Trout have been shown to possess at least 2 closely related PAH-inducible CYP1A isozymes, both of which have high protein sequence homology to mammalian CYP1A1 (Berndston and Chen, 1994; Heilmann et al., 1988Go; Williams and Buhler, 1984Go). Neither the gene regulation nor the catalytic properties of these duplicated proteins have been fully characterized and, for the purposes of this communication, these isozymes will be referred to collectively as trout CYP1A1.

Recent studies indicate that liver microsomes from ß-naphthoflavone-treated trout produce several DMBA metabolites, predominantly an unidentified polar metabolite(s), 2-OH-DMBA, and 4-OH-DMBA, with lesser amounts of the DMBA 8,9-, 5,6-, and 3,4-dihydrodiols (Miranda et al., 1997Go). A purified CYP1A1, in the presence of added epoxide hydrolase produced a similar spectrum, with DMBA-8,9-dihydrodiol being the major metabolite (40% of total) and the 3,4-dihydrodiol being less than 3% of total. The objective of the present study was to test the hypothesis that CYP1A1 plays a significant role in DMBA metabolism in trout liver cells and to establish if modulation of CYP1A1 isozymes may be important in DMBA-DNA adduct formation and carcinogenicity in trout.

Removal of trout liver cell DMBA-DNA adducts was also investigated. Repair of DMBA-DNA adducts has been shown to occur at significant rates, both in mammalian target and non-target tissue, in vitro and in vivo systems (Daniel and Joyce, 1984Go; Janss et al., 1972Go; Tay and Russo, 1981Go). The susceptibility of different strains of rats to DMBA-induced tumor formation has been found to correlate less with the levels of adducts actually formed than with the length of time adducts persist (Daniel and Joyce, 1984Go). Relative to mammalian cells, fish cells have been shown to have a reduced ability to repair some types of genomic DNA damage (Bailey et al., 1988Go, 1995Go; Walton et al., 1983Go), which may increase their susceptibility to compounds that adduct to DNA. However, DMBA-DNA adduct removal in trout has not been measured.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Chemicals
Beta-naphthoflavone (BNF), {alpha}-naphthoflavone (ANF), ethoxyresorufin, aryl-sulfatase, ß-glucuronidase, dimethylsulfoxide, bovine pancreatic DNase I, snake venom phosphodiesterase, bacterial alkaline phosphomonoesterase, nuclease P1, spleen phosphodiesterase, and micrococcal endonuclease were purchased from Sigma (St. Louis, MO). DMBA was purchased from Aldrich Chemical Co. (Milwaukee, WI). [3H]-DMBA (~ 20 Ci/mmol), [3H]-benzo[a]pyrene (BaP) (~ 60 Ci/mmol), and Hyperfilm-MP were purchased from Amersham, (Arlington Heights, NJ). Reference markers for DMBA metabolites 7-hydroxy,12-methylbenzanthracene (7-OH,12-MeBa) and 3-hydroxy-dimethylbenz(a)anthracene (3-OH-DMBA) were purchased from NCI Chemical Carcinogen Radio Repository, Lenexa, KS; and (+/-)-DMBA-3,4-dihydrodiol was obtained from the National Cancer Repository (Bethesda, MD). Other metabolites were synthesized as described (Christou et al., 1986Go). Collagenase was from Worthington Biochemical. Polynucleotide kinase was from U.S. Biochemicals (Cleveland, OH). Poly(ethylenimine) (PEI)-cellulose TLC plates were from Machery and Nagel, FRG, purchased through Brinkmann Instruments (Westbury, NY). [{gamma}-32P]-ATP (6000 Ci/mmol) was purchased from NEN/Dupont (Boston, MA).

Isolation and Culture of Trout Liver Cells and Mouse Embryo Fibroblasts
Rainbow trout (Oncorhynchus mykiss) were obtained from the Bowden National Fish Hatchery, Bowden, WV, or from the Food Toxicology and Nutrition Laboratory (FTNL), Oregon State University, Corvallis, OR. Liver cell cultures were prepared and maintained at 15°C as described previously (Miller et al., 1989Go). Tumor studies were conducted at the FTNL, using duplicate groups of 100 animals by procedures described elsewhere (Bailey et al., 1995Go). Medium in cell cultures was replaced daily with the indicated concentrations of specific compounds. Mouse embryo fibroblasts were the generous gift of Dr. A. Dipple; these cells were obtained as frozen cultures and were cultured at 37°C as described (Milner et al., 1985Go).

Water-Soluble DMBA Metabolites
To investigate conversion of DMBA to water-soluble metabolites, cells were incubated with medium containing 0.1 µg/ml [3H]-DMBA for indicated times, then extracted with water, methanol, and chloroform (1:1:1), as described (Baird et al., 1978Go). Radioactivity in the aqueous and organic phases was analyzed by liquid scintillation counting, to determine the percentage of DMBA converted to water soluble metabolites.

To determine the proportion of DMBA metabolites conjugated with sulfates or glucuronides, aliquots of medium were removed at various times and submitted to either no treatment (control), ß-glucuronidase treatment, or sulfatase treatment. Additionally, samples were removed from cells that were pretreated for 24 h with 0.6 mM diethyl maleate and 1 mM buthionine sulfoximine, to deplete GSH levels prior to incubation with fresh medium containing 0.1 µg [3H]-DMBA/ml. One-ml aliquots of medium were mixed with either 2000 units of type B-10 bovine ß-glucuronidase (Sigma, G 0501) or 20 units of type V limpet sulfatase (Sigma, S 8629) and 40 mM D-saccharic acid 1,4 lactone dissolved in 1 ml of sodium acetate (0.5M, pH 5.2). The limpet sulfatase was utilized due to its low levels of contaminating ß-glucuronidase activity (< 2 Sigma units/mg solid), and the saccharic acid, 1,4 lactone, was added to the sulfatase enzymatic treatment to inhibit this small but significant amount of contaminating ß-glucuronidase activity. Control treatments utilized 1 ml of medium mixed with 1 ml of sodium acetate (0.5 M, pH 5.2). Additionally, 1 mg/ml of ascorbic acid was added to each of the control or enzymatic solutions in order to prevent auto-oxidation of [3H]-DMBA phenols to quinones. Each sample was incubated on a rotary shaker at 37° C for 4 h prior to extraction. To extract medium, 1 ml was added to an equal volume of ethyl acetate/acetone (2:1) containing 10 mM dithiothreitol. This mixture was extracted for 5 min and then centrifuged in a Beckman microfuge to generate aqueous and organic phases. The phase-separated materials were then submitted to scintillation-counting, and the percentage of aqueous and organic [3H]-DMBA was calculated and corrected against the zero time values for each treatment. Water-soluble DMBA conjugates not hydrolyzed by sulfatase or ß-glucuronidase were designated "unidentified" DMBA metabolites.

Organic-Soluble DMBA Metabolites
Phase-I metabolism of DMBA was investigated in 2 ways: initial studies partially characterized [3H]-DMBA-free primary metabolites (organic-soluble, unconjugated) produced by trout liver cells and mouse embryo fibroblasts by HPLC separation. Limited metabolites were detected this way (see Results section), and in subsequent studies, metabolites produced in trout liver cells were analyzed by hydrolysis of glucuronide conjugates, followed by HPLC separation and fluorometric analysis of the resultant DMBA hydroxylation derivatives (Christou et al., 1994Go). Free [3H]-DMBA metabolites and parent DMBA were separated by HPLC, using the conditions described by Dipple, et al. (1983). Trout liver and mouse embryo cells were incubated with medium containing 0.1 µg [3H]-DMBA/ml medium for 6 h. Media was then removed and extracted (Flowers and Miles, 1991Go) with an equal volume of acetone/ethyl acetate (1:1.33). Aqueous and organic phases (containing water-soluble, conjugated, phase II metabolites and organic-soluble, phase I metabolites, respectively), were separated; MgSO4 was added to the organic phase, which was then centrifuged (2000 x g, 5 min); the supernatant was dried with nitrogen and resuspended in methanol. This sample was then passed through a 0.45-µm filter, dried under nitrogen, resuspended in methanol, and subjected to HPLC analysis. Chromatographic analysis was performed on a Waters HPLC fitted with a Perkin-Elmer 5 µm C-18 reverse phase column, as described (Dipple et al., 1983Go). Reference DMBA metabolites were also chromatographed; UV absorbance of these compounds was monitored at 254 nm. Percent of non-aqueous radioactivity either co-eluting with DMBA or in separate peaks was detected and integrated by a Radiomatic Flo-One Beta Radioactive Detector. Peaks were defined as >= 1000 3H counts/min above background.

To identify phase-I metabolites which had been conjugated with glucuronide groups, cultured trout-liver cells were incubated with DMBA (10 µM) for 30–45 min and medium was collected, digested with ß-glucuronidase, extracted, dried under nitrogen, dissolved in DMSO, then analyzed by HPLC with fluorometric detection of parent compound and metabolites, as described in detail (Christou et al., 1986Go). These studies used short (30–45 min) exposures of cells with DMBA to minimize the production of many multiply hydroxylated metabolites, which did not co-migrate with standards, and likely represented the bulk of the polar metabolites. Additionally, these short incubation times minimize DMBA activation of the arylhydrocarbon receptor and the subsequent increase in CYP metabolic activity.

EROD Assays
CYP1A1 activity was monitored using ethoxyresorufin-O-deethylase (EROD) assays, to determine if there is a correlation between CYP1A1 activity and DMBA metabolism, DNA adduct formation, or tumorigenesis. Isolated trout liver cells were pretreated with 0.05 µg BNF/ml medium to induce CYP1A1 (Miller et al., 1993Go), and CYP1A1 activity was inhibited by dosing cells with 100 nM ANF; control cells received dimethylsulfoxide solvent. EROD activity was used to measure CYP1A1 activity, using a Shimadzu RF 5000 U Spectrofluorophotometer, as described by Burke and Mayer (1974); assays were conducted at 20°C. Due to the difficulty of obtaining microsomes from a small number of cultured cells, EROD assays were performed with whole-cell homogenates, rather than with microsomal proteins. Studies were conducted to ensure that EROD activity measured in whole cell homogenate was comparable to that measured in microsomes. To this end, isolated trout liver cells were pretreated with 0.05 µg/ml BNF or DMSO solvent for 48 h; cells were collected by centrifugation (1000 x g, 5 min), the cell pellet was resuspended in 0.1 M KCL; 0.1 M Tris-acetate, pH 7.4; 1 mM ethylenediaminetetraacetic acid (EDTA), and 0.1 mM phenylmethylsulfonyl fluoride, and was disrupted with a Branson sonicator, on ice. This sonicate was the cell homogenate. Microsomes were prepared by centrifuging an aliquot of the homogenate (10,000 x g) for 20 min, followed by centrifugation of the supernatant (100,000 x g) for 1 h, and the resulting pellet (microsomes) was suspended in resuspension buffer (50 mM Tris, 1 mM dithiothreitol, 1 mM EDTA, 20% glycerol). With control cells, EROD-specific activity was 14.2 pmol resorufin/min/mg homogenate protein in cell homogenates, and was 51.5 pmol resorufin/min/mg microsomal protein in microsomes. Total EROD activity measured in cell homogenates was comparable to that in microsome preparations. Total EROD activity in control cell homogenates was 102.0 pmol resorufin/min/total mg homogenate protein, and was 108.0 pmol resorufin/min/total mg microsomal protein in microsomes from control cells. BNF pretreatment similarly induced EROD-specific activity and total activity ~ 8.2-fold in both cell homogenates and microsomes. These data demonstrate that EROD measurements using whole cell homogenates are valid, and this preparation was used in all subsequent EROD measurements.

DNA Isolation
For most studies, DNA was isolated from cells as described by Shen et al. (1991). Aliquots of DNA isolated from trout liver cells, which had been incubated with [3H]-DMBA (0.1 µg/ml medium), were counted in a liquid scintillation counter, to quantify DMBA covalently bound to DNA (pmol DMBA/mg DNA). For experiments in which DNA was hydrolyzed for HPLC analysis, DNA was isolated by cesium chloride gradient centrifugation (Lieberman and Dipple, 1972Go).

Characterization of DMBA-DNA Adducts
Trout liver cells or mouse embryo cells were treated with 0.1 µg/ml [3H]-DMBA for 24 h, and DNA was isolated from cells and enzymatically hydrolyzed to deoxyribonucleosides (Milner et al., 1985Go). DNA hydrolysates were loaded onto Sep-Pak C18 cartridges and washed with water, then with 40% methanol to remove buffer salts and unmodified nucleosides; typical DMBA-DNA adducts were eluted with methanol. Adducts eluted from Sep Pak columns with methanol were subsequently analyzed by HPLC on a Beckman Ultrasphere octadecyl silane column with a 45–65% methanol gradient. In some studies, a more thorough DNA digestion procedure, particularly effective in digesting liver cell DNA (Eberhart et al., 1992Go), was performed prior to Sep Pak chromatography.

To determine if DMBA-DNA adducts from trout liver cells might be conjugated to sulfate or glucuronide groups, hydrolyzed DNA was incubated for 3 h at 37°C with sulfatase or ß-glucuronidase, as previously described, prior to Sep Pak column chromatography.

[32P]-Post-labeling of DMBA-DNA Adducts
A DMBA-DNA adduct standard for post-labeling was generated by exposing fetal NIH mouse cell cultures to 50 ng DMBA/ml medium for 24 h and isolating the DNA as described (Vericat et al., 1989Go). Control or adducted DNA was hydrolyzed to deoxynucleoside 3`-monophosphates by incubating 15–25 µg of DNA in with 10 µl of a mixture of 2 µg/µl spleen phosphodiesterase and 0.5 µg/µl of micrococcal endonuclease at 37°C for 4 h.

To enrich the adducts prior to chromatography, the DNA digest (20 µl, 10–20 µg DNA) was mixed with 20 µl of 10 mM tetrabutylammonium chloride, 20 µl of 100 mM ammonium formate (pH 3.5), and 140 µl of distilled water in a 1.5-ml microcentrifuge tube. The mixture was extracted twice with one volume of water-saturated 1-butanol by vortexing for 30 s and centrifuging for 3 min in a table-top microcentrifuge. The butanol layer containing DNA adducts was transferred to a clean 1.5-ml microcentrifuge tube. The combined organic phases were back-extracted thrice with 180 µl of 1-butanol-saturated, distilled, deionized water to remove trace normal nucleotides. The butanol phase after the last extraction was placed in a methanol-rinsed 1.5-ml microcentrifuge tube and evaporated in a Savant vacuum concentrator. The adduct concentrate was dissolved in 100 µl of distilled water by vortexing, then evaporated again. The adduct residue from 10–20 µg of DNA was dissolved in 15 µl of double-distilled water. To this solution was added a 10 µl aliquot (200 µCi) of a [{gamma}-32P]-ATP labeling mix containing 182 µl of labeling buffer (100 mM bicine, 100 mM MgCl2, 100 mM dithiothreitol, 10 mM spermidine, pH 9.0), 32 µl of carrier-free [{gamma}-32P]-ATP (4.4 mCi), and 6 µl of polynucleotide kinase (30 U/µl). The solution was mixed using a pipetter equipped with guarded pipette tips and incubated in a water bath at 37°C for 45 min.

To remove residual labeled, normal nucleotides, [{gamma}-32P]-ATP, [32Pi], and other radioactive contaminants and to resolve [32P]-adducts, 9.8 to 19.6 µg of DNA (~ 196 µCi) were chromatographed on a 4-directional, PEI-cellulose TLC system (Schmeiser et al., 1988Go; Vericat et al., 1989Go). Radioactive spots on developed chromatograms were located by autoradiography at –80°C for various periods of time, using Hyperfilm-MP in cassettes with Du Pont Lightning-plus intensifying screens. To aid in the matching of chromatograms to their images on the film, chromatograms were marked with alignment dots using trace amounts of [32P] in black ink.

[33P]-Post-labeling HPLC of Adducts
DNA (10 µg) from trout fed 1000 ppm DMBA in the diet (123-day sample) was digested with nuclease P1 and prostatic acid phosphatase, evaporated, and 5`-labeled with [{gamma}-33P]-ATP and polynucleotide kinase. The labeled sample was digested to 5`-deoxyribonucleotide adducts with snake venom phosphodiesterase and analyzed by HPLC (Ralston et al., 1995Go).

DNA Repair
Trout liver cells were treated with medium containing 0.1 µg/ml [3H]-DMBA or 0.1 µg/ml [3H-benzo(a)pyrene for 24 h. Medium was then removed and replaced with DMBA- and benzo(a)pyrene-free medium, respectively. Cells were collected at this time and 12, 24, and 48 h later. For all time points, DNA was isolated and adduct formation was quantitated by liquid scintillation counting, as described.

Tumor Induction
Shasta strain rainbow trout were hatched and reared in 12°C well water at the Food Toxicology and Nutrition Laboratory, Oregon State University, Corvallis, OR as previously described (Schoenhard et al., 1976Go). Duplicate groups of 100 trout were fed 1 of 3 diets: (1) Oregon Test Diet (OTD) alone; (2) OTD containing 800 ppm DMBA for weeks 2–9; or (3) OTD containing 500 ppm BNF for weeks 1–10, and also containing 800 ppm DMBA for weeks 2–9. Trout in groups 1, 2, and 3 were then returned to OTD until sacrifice at 11 months. After sacrifice, the livers, stomachs, and swim bladders of each fish were examined for gross tumors and fixed in Bouin's solution for histology. Tumors were classified according to established criteria (Hendricks et al., 1984Go). Another group of 100 trout were bath-exposed to 5 ppm DMBA for 16 h and livers removed for DNA adduct determination after the exposure.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Effects of Modulating CYP1A1 Activity on Conversion of DMBA to Water-Soluble Metabolites
Experiments were designed to characterize DMBA metabolites formed in trout liver cells and to determine if CYP1A1 plays a prominent role in metabolizing DMBA in trout. CYP1A1 activity was modulated by pre-treating trout liver cells with BNF to induce CYP1A1 and by inhibiting CYP1A1 with ANF. EROD (CYP1A1) activity (pmol resorufin/min/mg protein) in control cells was 24.7, BNF pretreatment increased EROD activity ~ 6.3-fold, and ANF inhibited EROD activity to 6.6% of control values (1.65). Figure 1Go shows that BNF induction of CYP1A1 significantly increased the initial rate of DMBA metabolism to water-soluble compounds, whereas inhibiting CYP1A1 with ANF decreased both the rate and overall extent of DMBA metabolism to water-soluble metabolites. These results indicate that CYP1A1 participates significantly in DMBA metabolism to water-soluble metabolites in trout liver cells.



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FIG. 1. Influence of BNF or ANF on DMBA metabolism by isolated trout liver cells. Trout liver cells were pretreated with BNF (0.05 µg/ml medium) or DMSO for 48 h, then dosed with fresh medium containing 0.1 µg/ml [3H]-DMBA or [3H]-DMBA + ANF (0.03 µg/ml medium). This medium was removed at indicated times and extracted. Percent of DMBA in the aqueous phase was calculated as described in Materials and Methods for DMSO control cultures (squares), cultures pretreated with BNF (circles) or cultures treated with ANF (triangles). Results shown are the average of 3 experiments, and error bars represent standard error.

 
Water-soluble metabolites formed by trout liver cells were partially characterized by their sensitivity to digestion with ß-glucuronidase or sulfatase. In these studies, glucuronide conjugates usually comprised 10–35% of total water-soluble DMBA metabolites, sulfate conjugates were <=5% total water-soluble metabolites, and in all experiments, the majority (~ 50–85%) of water-soluble DMBA metabolites were not sensitive to enzymatic digestion and were designated as "unidentified" metabolites. The unidentified phase-II metabolites were not affected by digestion with a mixture of ß-glucuronidase and sulfatase. Reports (Kirby et al., 1990Go; Nimmo and Clapp, 1979Go) suggest that a portion of the unidentified phase-II DMBA metabolites may be conjugated with glutathione. As one means of assessing the role of glutathione in DMBA phase-II metabolism, cells were pretreated with diethyl maleate and buthionine sulfoximine for 24 h as described, to deplete intracellular glutathione, resulting in ~ 90% depletion. In different experiments, glutathione levels in control cells ranged from 1.3 to 3.2 nmol/106 cells, while diethyl maleate- and buthionine sulfoximine-treated cells contained 0.05–0.38 nmol glutathione/106 cells. Depleting glutathione did not alter the percentage of DMBA metabolized to water-soluble or sulfate conjugates. Although depleting cellular glutathione marginally increased the mean percentage of glucuronide metabolites (~ 15%) and correspondingly decreased the percentage of unidentified water-soluble metabolites (~ 15%), these changes were not statistically significant (not shown).

Phase-I Metabolism of DMBA and Effects of Modulating CYP1A1
The extent to which trout liver cells metabolize DMBA to organic-soluble ("free" Phase-I) metabolites was investigated and compared to mouse embryo fibroblasts. Cells were incubated with [3H]-DMBA, and organic-soluble metabolites released into medium were extracted and analyzed by HPLC. Figure 2Go shows the HPLC profiles of major DMBA metabolites formed in trout liver and mouse embryo cells after incubation with DMBA for six h. HPLC profiles of [3H]-DMBA metabolites differed both quantitatively and qualitatively between the teleost and mammalian cells. Mouse embryo fibroblasts produced a profile consisting of the parent DMBA, and ~ 8 metabolites. These 8 metabolites, formed in mouse fibroblasts, accounted for ~ 40% of the radioactive material in the organic-soluble fraction, and DMBA-3,4-dihydrodiol comprised ~ 10% of these metabolites. In contrast, trout liver cells produced only 2 to 3 detectable DMBA metabolites, which comprised ~ 2% of the radioactive material in the organic-soluble fraction; DMBA-3,4-dihydrodiol was not detected.



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FIG. 2. Production of organic-soluble DMBA metabolites in mouse embryo cells and in trout liver cells. Trout liver cells and mouse embryo cells were dosed with medium containing [3H]-DMBA (0.1 µg/ml medium). Media was collected 6.0 h later and extracted with acetone/ethyl acetate to form organic and aqueous phases. The organic phases were blown dry with nitrogen, resuspended in methanol, injected onto a Beckman Ultrasphere ODS HPLC column, and eluted as described in Materials and Methods. Arrows 1, 2, and 3 indicate elution positions of (1) DMBA-3,4-dihydrodiol; (2) 7-OH-methylbenz-(a)-anthracene; and (3) 3-OH-7,12-DMBA standards. Total 3H-radioactivity eluting in mouse embryo cell chromatogram = 312,265 CPM; total radioactivity eluting in trout liver cell chromatogram = 348,741 CPM.

 
The lower amount of free (unconjugated) DMBA primary metabolites formed in trout liver cells, relative to mouse fibroblasts, could be due to slower overall metabolism of DMBA by trout liver cells or to more efficient conjugation of phase-I metabolites by phase-II enzymes. Direct comparisons of the rate of DMBA conversion to water-soluble metabolites in trout liver and mouse embryo cells during a 6-h period demonstrated essentially identical rates of metabolism during the first 3 h. In the subsequent 3 h, DMBA phase-II metabolism was approximately 10–20% slower in trout liver cells (not shown). These small differences in DMBA metabolism to water-soluble compounds between trout liver and mouse embryo cells can only partially account for the reduced presence of organic-soluble, phase-I metabolites in medium from trout liver cells. This suggests that phase-I metabolites are converted to phase-II metabolites more efficiently in trout liver cells than in mouse embryo cells.

To further characterize Phase-I DMBA metabolites formed in trout liver cells, cells were incubated with DMBA for 45 min, then media samples were incubated with ß-glucuronidase, prior to extraction and separation by HPLC with fluorometric detection. Table 1Go shows that the majority of these DMBA metabolites co-elute with 5,6-, 8,9- and 10,11-DMBA dihydrodiols, and with DMBA, 2- or 3- or 4-phenol. Minor metabolites co-migrated with 7-OH-methyl-12-methyl-benz(a)anthracene and 12-OH- methyl-7-methyl-benz(a)anthracene. Only a very small amount of DMBA 3,4-dihydrodiol (~ 2% of total metabolites) was detected. In addition, polar metabolites were observed that did not migrate with any DMBA standards (not shown). Because the identity of these polar metabolites is unknown, they could not be quantitated by fluorometric detection and they were not included in the analysis in Table 1Go. Pre-treating trout liver cells with BNF to induce CYP1A1 increased the rate of production of most of the identified metabolites ~ 2-fold without substantially changing the proportion of any particular metabolite formed (Table 1Go).


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TABLE 1 DMBA Metabolites Formed in Trout Liver Cells
 
Formation of [3H]-DMBA-DNA Adducts in Vitro and the Role of Cytochrome P4501A1
[3H]-DMBA formed DNA adducts when incubated with trout liver cells (Fig. 3Go). The rate of DMBA-DNA adduct formation in different experiments appeared to be correlated with the rate of DMBA metabolism. In the experiment depicted in Figure 3Go, ~ 35% of the DMBA was converted to water-soluble metabolites at the 30-min time point. In experiments where the rate of DMBA metabolism to water-soluble compounds was slower, the rate of DNA adduct formation was also reduced. However, in all experiments, the rate of DMBA-DNA adduct formation declined rapidly after 30–40% of parent DMBA was metabolized to water-soluble compounds.



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FIG. 3. Influence of BNF or ANF on formation of DMBA-DNA adducts in trout liver cells. Trout liver cells were pretreated with 0.05 µg/ml BNF or with DMSO solvent alone (control) for 48 h. This medium was then removed and replaced with medium containing 0.1 µg [3H]-DMBA/ml; one set of control cultures also received 0.1 µM ANF. At the indicated times, DNA was isolated from control cultures (squares), cultures pretreated with BNF (circles), or cultures treated with ANF (triangles), and [3H]-DMBA incorporated into DNA was quantitated by liquid scintillation counting. Results of a representative experiment are expressed as pmol DMBA/mg DNA. The 30-min DNA sample from BNF-treated cells was inadvertently lost.

 
Because CYP1A1 appears to contribute significantly to the initial rate of DMBA metabolism in trout liver cells (Table 1Go), the effect of modulating this activity with BNF or ANF on DMBA-DNA adduct formation was investigated. EROD assays assessed CYP1A1 activity following the different treatments; in BNF-treated cells, EROD was induced > 10-fold over control values, and in ANF-treated cells, EROD was reduced to 40% of control values (not shown). Altering CYP1A1 activity did not significantly change the formation of DMBA-DNA adducts in trout liver cells, depicted in a representative experiment in Figure 3Go.

Influence of BNF on Hepatic CYP1A1, DMBA-DNA Adducts and Tumor Response in DMBA-Treated Trout
Based on these findings, the effect of dietary BNF on hepatic CYP1A1, DMBA-DNA adduction, and tumor induction was investigated in vivo. Duplicate groups of 100 fingerling rainbow trout were fed 1 of 3 diets (control diet, 800 ppm DMBA, or 800 ppm DMBA plus 500 ppm BNF). A fourth group was fed 500 ppm BNF alone, to quantify CYP1A1 response, but not carried onto full tumor determination, since we have observed this treatment not to be carcinogenic or toxic (Bailey et al., unpublished results). This dose of BNF has been shown repeatedly (Takahashi et al., 1995Go) to strongly induce trout hepatic-CYP1A1 protein and EROD activity; however, the ability of DMBA alone to induce trout CYP1A1 by a dietary tumorigenic protocol has not been previously tested. Samples were taken at the end of the DMBA-exposure period for adduct and CYP1A1 assessment, and remaining trout were fed control diet until termination 7 months later. As seen in Table 2Go, DMBA, by this protocol, was weakly carcinogenic in liver, and strongly carcinogenic in stomach and swim bladder. DMBA treatment alone significantly elevated hepatic CYP1A1 levels and produced levels of DMBA-DNA adducts readily detectable by [33P]-HPLC post-labeling. While BNF alone also strongly elevated hepatic CYP1A1, combined treatment with DMBA somewhat reduced hepatic CYP1A1 from that induced by DMBA or BNF alone, but did not alter DMBA-DNA adducts or tumor response compared with animals receiving DMBA alone. BNF co-treatment strongly inhibited tumor response in stomach and swim bladder; however, DNA adducts and CYP1A1 levels were not assessed in these target tissues.


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TABLE 2 Influence of Dietary BNF on DMBA Tumorigenicity in Rainbow Trout
 
Initial Characterization of DMBA-DNA Adducts
DMBA-DNA adducts formed in mouse embryo cells have been well characterized (Bigger et al., 1983Go; Dipple and Hayes, 1979Go; Moschel et al., 1983Go; Sawicki et al., 1983Go; Vericat et al., 1989Go). In order to determine if adducts formed in trout liver cells are similar or identical to those formed in mouse embryo cells, adducts from both types of cultured cells were compared by HPLC analysis. As a first approach, trout liver cells and mouse embryo cells were incubated for 24 h in medium containing [3H]-DMBA. DNA was then isolated and enzymatically hydrolyzed to nucleosides. When hydrolyzed DNA from mouse embryo cells was chromatographed on Sep Pak cartridges, very little (~ 2%) radioactivity was eluted with water washes, ~ 53% of radioactivity eluted with 40% methanol, and ~ 45% eluted with 100% methanol. These results are similar to those published by others (Bigger et al., 1983Go; Sawicki et al., 1983Go). The [3H]-adducts eluting from Sep Pak columns with methanol were further fractionated by HPLC chromatography (Sawicki et al., 1983Go). Three peaks of radioactive material were obtained (not shown) with retention times corresponding to those of the bay region anti-dihydrodiol-epoxide adducts for deoxyguanosine and deoxyadenosine, and the bay-region syn-dihydrodiol-epoxide deoxyadenosine adducts (Dipple et al, 1983Go).

Very different results were seen when [3H]-DMBA-adducted DNA digests obtained from trout liver cells were chromatographed on Sep Pak columns. In contrast to adducts obtained from mouse embryo cells, 60–90% of the [3H]-adducts eluted from the Sep Pak column with water, 1–27% of the [3H]-adducts eluted with 40% methanol, and <= 10% of the of the [3H]-adducts eluted with 100% methanol. Further HPLC analysis of these methanol fractions was not possible due to the low amount of radioactivity contained within them.

The apparent polarity of the DMBA-DNA adducts obtained from trout liver cells could have resulted from incomplete digestion of trout liver-cell DNA, yielding adducts with oligonucleotides rather than individual deoxyribonucleosides. To determine if this were the case, a more vigorous digestion of trout liver cell DNA was undertaken, as described (Eberhart et al., 1992Go). This procedure is effective in completely digesting rat liver DNA; however, it did not alter the Sep Pak elution profile of trout liver cell DNA (not shown). These results suggest that DMBA-DNA adducts obtained from trout liver cells are more polar in nature than those formed in mammalian cells.

Further confirmation that DMBA-DNA adducts formed in trout liver cells are more polar than those formed in mouse embryo cells was obtained by extracting DNA digests in a chloroform:methanol:water mixture. Ninety five percent of [3H]-DMBA-DNA adducts from trout liver cells partitioned into the aqueous phase, whereas only 10% of [3H]-DMBA-DNA adducts from mouse embryo cells partitioned into the aqueous phase. DNA digests obtained from trout liver cells were also chromatographed on PEI cellulose TLC plates with 0.8 M (NH4)2SO4. DMBA-nucleoside adducts formed in mouse embryo cells do not migrate from the origin under these conditions. However, when DNA digests from trout liver cells were chromatographed, more than 95% of the radioactivity migrated from the origin (not shown), further demonstrating that DMBA-DNA adducts formed in trout liver cells are more polar than those formed in mouse embryo cells. To test the possibility that DMBA-DNA adducts formed in trout liver cells contain sulfate or glucuronide groups, making them hydrophilic, DNA from trout liver cells, incubated with [3H]-DMBA, was isolated and incubated with sulfatase or ß-glucuronidase prior to Sep Pak column chromatography. These enzymatic treatments did not change the elution profile of DMBA-DNA adducts (not shown).

DMBA-DNA adduction was further examined both by conventional [32P]-post-labeling and by [33P]-post-labeling-HPLC. Three major adducts were identified by [32P]-post-labeling and TLC when mouse embryo cells were incubated with DMBA (Fig. 4AGo). These adducts were barely detectable in trout liver cells incubated with DMBA (Fig. 4CGo). In contrast, when trout liver cells were exposed to DMBA-3,4-dihydrodiol, 3 prominent adducts were observed (Fig. 4BGo), which were indistinguishable from those formed in mouse embryo cells incubated with the parent DMBA (Fig. 4AGo). In vivo, exposure of trout to DMBA by the highly efficient and carcinogenic gill uptake procedure (Fig. 4DGo) did result in detectable production in the liver of one adduct that had the same Rf as one from mouse embryo cells. Based on other studies (Vericat et al., 1989Go), this adduct is tentatively assigned as the deoxyadenosine adduct from DMBA-(4S,3R)-dihydrodiol(2S,1R-epoxide). Three other unknown adducts were also observed. However, dietary exposure for 18 weeks to carcinogenic doses of DMBA was determined to elicit only trace levels of these adducts (data not shown). The DMBA-DNA adducts formed in livers by this exposure protocol were further characterized by the [33P]-HPLC post-labeling procedure and found to consist almost entirely of polar adducts (Fig. 5Go) not detected by the conventional [32P]-post-labeling, thin-layer-chromatography procedures used for other samples. This confirms that most DMBA-DNA adducts formed in trout liver are more polar than those formed in mouse embryo cells.



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FIG. 4. DMBA-DNA adducts in isolated cells detected by [32P]-post-labeling. (A) Adducts from NIH-Swiss mouse embryo cells incubated with 50 ng/ml DMBA for 24 h (film exposure = 15 min). (B) Adducts detected from trout hepatocytes incubated with 4 µg/ml (+/–)-DMBA-3,4-dihydrodiol for 24 h (film exposure = 2 h). (C) Adducts detected in trout hepatocytes exposed to 4 µg/ml of DMBA for 24 h (film exposure = 40 min). (D) Adducts detected from liver after exposure of juvenile trout to a bath of 5 ppm DMBA for 16 h by gill uptake.

 


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FIG. 5. DMBA-DNA adducts detected by [33P]-HPLC post-labeling procedure. 10 µg of trout liver DNA from trout fed 1000 ppm DMBA for 123 days was labeled and chromatographed as described in Materials and Methods. Results depict dpm [33P] as a function of retention time (min).

 
Persistence of DMBA-DNA Adducts
Because most of the DMBA-DNA adducts formed in trout liver cells appeared to be different (more polar) than those formed in mouse embryo cells, it was of interest to investigate repair of these adducts. Trout liver cells were incubated with [3H]-DMBA for 12–24 h, then placed in DMBA-free medium, and the loss of [3H]-DMBA-adducts from DNA was followed over 48 h (Fig. 6Go). In this and two other studies, no significant loss of total genomic DMBA-DNA adducts was detected over the 48-h period. In order to determine if DMBA-DNA adducts were more stable than other PAH-DNA adducts in trout liver cells, the loss of benzo(a)pyrene-DNA [BaP-DNA] adducts was also measured. Figure 6Go shows that BaP formed appreciable DNA adducts in trout liver cells, and that these DNA adducts were also quite stable. Repair of BaP-DNA adducts was not detected during a 48-h period. This is compatible with earlier indications that trout liver has low capacity for global removal of bulky DNA adducts (Bailey et al., 1988Go).



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FIG. 6. Repair of DMBA-DNA adducts or BaP-DNA adducts in trout liver cells. Trout liver cells were incubated for 24 h with medium containing 0.1 µg/ml [3H]-DMBA (squares) or 0.1 µg/ml [3H]-BaP (circles), then cultured in medium without DMBA or BaP. At the indicated times after removing [3H]-DMBA or -BaP, DNA was isolated from cells and DNA adduct formation (pmol DMBA/mg DNA or pmol BaP/mg DNA) was determined. Results depict the mean and standard deviation from data in 2 different experiments; determinations were duplicated in each experiment.

 

    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Characterization of DMBA-DNA Adduction and Persistence in Trout Liver Cells
The data presented herein demonstrate that the majority of DNA adducts formed when trout or trout hepatocytes are exposed to DMBA, are more polar than those formed in mouse embryo cells. This polarity does not appear to derive from incomplete enzymatic digestion or from conjugation with sulfate or glucuronides but could be due to conjugation with other water-soluble groups such as glutathione or its mercapturic acid breakdown products. Alternatively, DMBA-DNA adduct polarity in trout liver may derive from the formation of highly hydroxylated metabolites such as DMBA-7,12-dihydroxymethyl-5,6-dihydrodiol, a bis-diol epoxide, or other polar primary metabolites produced by trout (Fong et al., 1993Go; Miranda et al., 1997Go). We have shown that trout liver cells incubated with DMBA-3,4-dihydrodiol readily form the classic mammalian DNA adducts derived from epoxide intermediates (Fig. 4Go). Trout liver cells may convert other multiply hydroxylated DMBA metabolites into the bay region 3,4-diol-1,2-epoxides just as readily. Precedence for this type of metabolic activation has been reported for dibenz(a,h)anthracene (Carmichael et al., 1993Go; Fuchs et al., 1993Go), where highly polar bis-dihydrodiol epoxide-DNA adducts were formed. In addition, polar DMBA-DNA adducts observed in rat liver and mouse skin following in vivo treatment with DMBA have been shown to be 7-hydroxymethyl-12-methylbenz[a]anthracene-deoxyribonucleoside adducts (DiGiovanni et al., 1983Go; Joyce and Daniel, 1982Go). This indicates that additional metabolism of major hydroxylated metabolite(s) (i.e., hydroxymethyl-DMBA) occurs in mammalian liver and skin. Similar intermediates may play a role in trout tumorigenesis.

Examination of the rate of repair of DMBA-DNA adducts in trout liver cells indicated repair of these adducts from the total genome is undetectable in 48 hours (Fig. 6Go), whereas repair of DMBA-damaged DNA in mammalian cells appears to be more efficient. Tay and Russo (1981) demonstrated that rat mammary epithelial cells removed 12 to 36% of DMBA-DNA adducts in 48 h. It is unclear whether the lack of repair of DMBA-DNA adducts by trout liver cells is due to low global excision repair capacity in the teleost system or to formation of adducts that are intrinsically less repairable than those formed in mammalian cells. However, these findings are consistent with other studies indicating teleosts are less efficient in repairing bulky adducts than their mammalian counterparts (Bailey et al., 1988Go).

Enzymatic Pathways to DMBA-DNA Adduction in Trout
The enzymes involved in DMBA metabolism and bioactivation in trout are not fully characterized, and may vary depending on diet and other treatment history. Recent studies indicate that purified trout CYP1A1 metabolizes DMBA to the 3,4-diol proximate carcinogen, but only as a minor metabolite (Miranda et al., 1997Go). BNF induction of CYP1A1 in isolated trout liver cells was shown here to stimulate metabolism of DMBA to water-soluble compounds (Fig. 1Go). However, the 3,4-dihydrodiol metabolite was only a very minor metabolite formed in trout liver cells, and CYP1A1 induction did not increase the relative amount of this metabolite (Table 1Go). In trout liver cells, DMBA metabolites formed by CYP1A1, such as the 3,4-diol, may be efficiently metabolized further to detoxification products or polar DNA-binding metabolites. When cells were incubated with 4 µg/ml DMBA-3,4-diol (Fig. 4Go), the high concentration may have resulted in more metabolic activation to the diol epoxide. Neither CYP1A1 induction with BNF nor inhibition by ANF substantially affected DMBA-DNA adduct formation (Fig. 3Go). Therefore, the present results provide no evidence that CYP1A1 isoform(s) selectively catalyze DMBA metabolism to DNA-reactive intermediates in trout liver cells. These results are in apparent contrast to studies with mammalian cells, in which ANF reduced DMBA-DNA adducts (Dipple et al., 1984Go). However, ANF inhibits both CYP1A1 and 1A2 in mammalian cells, and trout appear not to possess CYP1A2 orthologues (Goksoyr et al., 1991Go). Miranda et al. (1997) demonstrated that microsomes from BNF-treated trout catalyzed DMBA metabolism to DNA-binding species in vitro; however the adducts formed were not characterized, and their quantitative importance could not be evaluated. Additional studies are needed to establish which trout CYP isozyme(s) participate in formation of stable DMBA-DNA adducts.

BNF Influence on DMBA DNA Adduction and Tumorigenicity
These are among the first studies to report DMBA carcinogenicity in any fish by dietary exposure. The results show that DMBA is highly carcinogenic by this route, with stomach and swim bladder as the major target organs, and a lesser response in liver (Table 2Go). The results also show that co-feeding of 500 ppm BNF did not significantly elevate hepatic tumor response. There are 2 opposing mechanisms through which BNF co-exposure might have potentially altered DMBA adduction and tumorigenicity: (1) by providing CYP1A1 induction additional to that seen with DMBA alone, and (2) through catalytic inhibition of CYP1A1 or other trout CYP enzymes that may bioactivate DMBA. BNF is a potent inhibitor of trout CYP1A1 and other CYP enzymes in vitro, and 500 ppm dietary BNF strongly inhibits AFB1 adduction and tumorigenicity in trout, primarily through inhibition of CYP bioactivation (Takahashi et al., 1996Go). The significant inhibition of tumor response in stomach and swim bladder may well reflect such a mechanism in these target organs, but this was not investigated directly. In liver, BNF co-treatment was found to somewhat reduce hepatic CYP1A1 activity compared with animals receiving DMBA alone, but did not alter either DMBA-DNA adduction or tumorigenicity under conditions where it does both against AFB1. In contrast, we have shown that co-treatment with Aroclor 1254, also a potent CYP1A1 inducer, strongly elevated DMBA hepatic tumor response in this model (Bailey et al., unpublished results). These results suggest that members of the trout CYP1A1 isozyme subfamily may be differentially regulated, that BNF and Aroclor 1254 induce different ratios of these isozymes, and that the isozymes have distinctly different catalytic properties for DMBA metabolism and bioactivation. In support of this, recent studies (Curtis et al., 1996Go) indicate differential regulation of 2 trout CYP1A1 genes by 2,4,5,2`,4`,5`-hexachlorobiphenyl. However, 2,4,5,2`,4`,5`-hexachlorobiphenyl is a non-coplanar polychlorinated biphenyl, which is not normally considered an inducer of CYP1A1 genes. Because some commercially available preparations of this polychlorinated biphenyl may contain a furan with potent CYP1A1-inducing ability, these conclusions may be compromised. Metabolism and induction studies with the cloned trout CYP1A1 isoforms will be necessary, in order to test this hypothesis. Finally, DMBA tumorigenesis in some systems may be driven primarily through the formation of unstable DNA adducts resulting from one-electron oxidation processes (Cavalieri and Rogan, 1992Go). We do not believe this mechanism of activation plays a major role in DMBA tumor induction in trout. While high levels of dietary antioxidant butylated hydroxyanisole did not reduce DMBA tumorigenicity in trout (Bailey et al., unpublished results), the profile of DMBA-mediated Ki-ras oncogenic mutations is not fully compatible with apurinic site mutagenesis (Reddy et al., 1995Go), and there appear to be ample stable, polar DMBA adducts in this model to account for its mutagenicity and carcinogenicity.

In summary, the cellular interactions of DMBA in trout liver cells have been initially characterized and found to differ substantially from those occurring in mouse embryo cells, particularly in the processes of bioactivation and DNA repair. Studies to further characterize the interactions of DMBA with trout liver cells may provide novel insight into mechanisms of chemical carcinogenesis.


    ACKNOWLEDGMENTS
 
This research was supported in part by the following NIH grants: CA 45131(M.R.M.), ES 03850 (G.B.), ES 04766 (G.B.) and CA 28825 (W.B.); and by ACS grant RPG-57–066–01-VM (M.R.M.).


    NOTES
 
1 To whom correspondence should be addressed at the Department of Biochemistry, WVU Health Sciences Center, PO Box 9142, Morgantown, WV 26506–9142. Fax: (304) 293-6846. E-mail: mmiller{at}hsc.wvu.edu. Back


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