Cytochrome P450 1B1 Is Required for 7,12-Dimethylbenz(a)-anthracene (DMBA) Induced Spleen Cell Immunotoxicity

Abstract

7,12-Dimethylbenz(a)anthracene (DMBA) is a potent carcinogen that induces immunosuppression of both humoral and cell-mediated immunity in mice and other species. Previous studies have shown that CYP1B1 is required for bone marrow toxicity produced by DMBA in mice. Therefore, the purpose of these studies was to determine whether CYP1B1 was required for spleen cell immunotoxicity. Female C57BL/6N wild-type (WT) and CYP1B1 knockout (-/-) mice were treated with 0, 17, 50, or 150 mg/kg (cumulative dose) DMBA in corn oil by oral gavage once a day for five days. Several immunotoxicological assays were used to assess the effects of DMBA on systemic immunity. These included the in vitro T-dependent antibody response to sheep red blood cells (SRBC) measured using a direct plaque forming cell (PFC) assay, T- and B-cell mitogenesis induced by Con A and LPS, and nonspecific cell-mediated immunity was evaluated using an NK cytotoxicity assay. In addition, lymphocyte subpopulations were measured by flow cytometry using specific cell surface markers. Following five days of DMBA treatment, the body weights and spleen cell surface markers of the WT and CYP1B1 (-/-) mice showed no significant changes. A decrease in NK activity was found at the 50 mg/kg DMBA dose in WT mice, but not in the CYP1B1 (-/-) mice. Interestingly, at the 150 mg/kg dose of DMBA, CYP1B1 null mice had decreased NK activity, whereas WT mice did not. The SRBC PFC response demonstrated that the IgM antibody response was suppressed by DMBA in WT mice in a dose-dependent manner (significant at 50 and 150 mg/kg). However, there were no changes in the SRBC PFC responses in any DMBA test group in the CYP1B1 (-/-) mice. Similarly, while DMBA suppressed B- and T-cell mitogenesis at the 50 and 150 mg/kg dose levels in C57BL/6N WT mice, no effect was seen in CYP1B1 (-/-) mice. Thus, CYP1B1 appears to be critical for the immunosuppression of DMBA in mice, suggesting a role for bioreactive metabolites in the spleen cell immunotoxicity produced by DMBA.

Polycyclic aromatic hydrocarbons (PAHs) are environmental pollutants generated by incomplete combustion of fuels, wood, and other organic matter. PAHs are present in cigarette smoke, grilled foods, automobile exhaust, and woodsmoke (Burchiel et al., 2005). Most of the members of the PAH family are potent carcinogens for humans and other species (Pelkonen and Nebert, 1982). Some of these compounds are immunotoxicants. DMBA is a prototypic PAH with carcinogenic and immunosuppressive effects in various species (Burchiel et al., 1988, 1990; Buters et al., 1999, 2003; Shimada and Fujii-Kuriyama, 2004; Thurmond et al., 1987). Because of photolysis and photooxidation properties, DMBA is not found in the natural environment. However, previous studies aimed at understanding mechanisms of DMBA immunotoxicity have provided important clues into the actions of environmentally prevalant PAHs.

DMBA has been widely evaluated for its carcinogenicity, and is often used as a model compound for breast, skin, and other cancers in rodents (Ethier and Ullrich, 1982). Rodent cancer studies have demonstrated the critical importance of cytochrome P450 metabolism (Nebert and Russell, 2002) leading to the formation of reactive metabolites that bind to DNA causing mutations and cancer initiation. Two key enzymes responsible for DMBA bioactivation are cytochrome P450 1B1 (CYP1B1) and microsomal epoxide hydrolase (EPHX1) (Gonzalez, 2001). Together, these enzymes form the ultimate carcinogen of DMBA, DMBA-3,4-dihydrodiol-1,2-epoxide (DMBA-DE) (Cavalieri and Rogan, 1992; Kleiner et al., 2002; Slaga et al., 1979). CYP1B1 is thought to be the key enzyme responsible for DMBA metabolism in humans and rodents (Buters et al., 2003). Historically, CYP1B1 was first found in mouse embryonic fibroblast cells and rat adrenal glands (Otto et al., 1992; Savas et al., 1997). Human CYP1B1 was first isolated and cloned in 1994 by Sutter et al. (1994). It is expressed in many extrahepatic tissues such as lung, mammary gland, spleen, kidney, prostate, uterus, and heart (Choudhary et al., 2003). The expression of CYP1B1 is also associated with many cancers, such as breast, lung, and ovarian cancers in rodents treated with PAHs (Buters et al., 2003). CYP1B1 knockout mice were found to be resistant to DMBA-induced lymphomas (Buters et al., 1999) and ovarian cancers (Buters et al., 2003).

DMBA-induced bone marrow toxicity has been found to be dependent on CYP1B1 expression in mice (Heidel et al., 1999). The molecular mechanism of DMBA bone marrow toxicity has been extensively studied during the last few years (Heidel et al., 1998, 1999, 2000; Page et al., 2003, 2004). Bone marrow stromal cell CYP1B1 is required for pre-B cell apoptosis induced by DMBA in vitro. An in vivo study has shown that DMBA treatment (ip injection, 50 mg/kg DMBA) can induce the release of cytokines by bone marrow stromal cells, such as, TNF-α; leading to down-regulation of IL-2 (Pallardy et al., 1989). TNF-α is able to activate TNFR death receptors, and initiates the caspase-8 signaling pathway (Page et al., 2003). Deletion of TNFR totally blocks DMBA toxicity in bone marrow in vivo. In addition, TNFR also activates dsRNA-dependent protein kinase (PKR) after TNF-α ligand binding following DMBA treatment. Upregulated PKR induces the phosphorylation of p53 (Page et al., 2003). The p53 protein induces cell cycle arrest and apoptosis.

In previous studies in our laboratory, we found that DMBA can immunosuppress both humoral and cell-mediated immunity (Burchiel et al., 1988). However, the mechanism of DMBA induced splenic immunotoxicity is not clear. Based on previous CYP1B1 knockout mice studies in bone marrow, we hypothesized that CYP1B1 is an essential enzyme for DMBA-induced spleen cell immunotoxicity. In this report, we utilized female C57BL/6N WT and CYP1B1 (-/-) mice to analyze the role of CYP1B1 in spleen cell immunotoxicity. Our data showed that CYP1B1 knockout mice were protected from DMBA-induced spleen cell toxicity and immunosuppression compared to WT mice.

MATERIALS AND METHODS

Chemicals and reagents.

7,12-Dimethylbenz(a)anthracene (Sigma Cat. No. D3254-5G) at greater than 95% purity and Concanavalin A (Con A, Type IV, Cat. No. C-2010) were obtained from Sigma-Aldrich (St. Louis, MO). Cell culture materials were from Sigma-Aldrich and Invitrogen (Grand Island, NY). Lipopolysaccharide (LPS) from E. Coli was purchased from Alexis Biochemicals (San Diego, CA).

Animals.

Female WT C57BL/6N mice (8–10 weeks old) were purchased from Harlan Laboratories (Indianapolis, IN). CYP1B1 knockout mice were received as gift from National Institutes of Health (NIH) and were backcrossed to generation F10 or greater in our AAALAC-accredited animal facility under an IACUC-approved protocol. Age-matched 10–14 week old female knockout mice were used for these immunotoxicity studies. The results reported in these studies were verified in two or more replicate experiments, with similar results except as noted. Briefly, WT or knockout mice were gavaged using corn oil. DMBA was given once a day for five days using equal daily doses (i.e., total cumulative dose divided by 5). The total cumulative doses of DMBA were 17, 50, and 150 mg/kg. Control groups of mice were gavaged with corn oil only. Spleens were aseptically obtained from CO₂ euthanized mice on day 7, 48 h after the last DMBA dose. Mouse body weights and spleen weights were recorded at the time of euthanasia.

Spleen cell preparation.

Single cell suspensions were prepared from five individual mice per treatment group. Spleen cells were harvested as described previously (Burchiel et al., 2004). In brief, spleens were isolated in RPMI 1640 complete medium supplemented with 10% fetal bovine serum, 2 mM L-glutamine, 100 μg/ml streptomycin and 100 Units/ml penicillin, and centrifuged at 280 × g for 10 min. Cell pellets were resuspended and maintained in 2 ml RPMI 1640 complete medium on ice. Viable spleen cell counts were obtained by the trypan blue (Sigma Chemical Co., St. Louis, MO) exclusion method using a hemacytometer.

Lymphocyte mitogenesis assay.

Lipopolyssaccharide (LPS) and concanavillin (Con A) were used to evaluate B and T cell proliferation, respectively. Spleen cells from individual mice were exposed to mitogens for three days in 96 well culture plates (200 μl @ 1 × 10⁶ cells/ml) in replicates of six containing 50 μl of 50 μg/ml of LPS or 5 μg/ml Con A. Complete RPMI 1640 medium lacking LPS or Con A was used as the no mitogen control. Plates were incubated at 37°C in a humidified, 5% CO₂ incubator for 48 h, pulsed with 20 μl of 50 μCi/ml [³H]-thymidine (ICN, Aurora, OH), and then incubated at the above conditions for an additional 18 h. The cells were harvested on glass filters using a Brandel Model 24V cell harvester. Filter samples were allowed to dry at room temperature for 30 min, and were then transferred to liquid scintillation vials with 3 ml ScintiVerse BD cocktail (Fisher Scientific, Houston, TX). The incorporated [³H]-thymidine was measured using a Wallac 1410 liquid β scintillation counter.

In vitro plaque-forming cell assay.

Mouse spleen cells collected sterilely (2 × 10⁶ cell/ml, 0.5 ml) were cultured for four days with 0.5 ml of washed 1% sheep red blood cells (SRBC) (Colorado Serum, Denver, CO) in 48-well, flat-bottomed plates (Corning Glass, Corning, NY) with RPMI 1640 medium [containing 10% heat inactive fetal bovine serum (Hyclone, Logan, UT), 50 μM 2-mercaptoethenol (GIBCO, Grand Island, NY), 1 mM sodium pyruvate (GIBCO, Grand Island, NY) and 50 μg/ml gentamycin (GIBCO, Grand Island, NY)]. The plates were placed in a humidified, 37°C, 5% CO₂ incubator. RPMI 1640 medium without SRBC were added into the spleen cells as control using a modified Mishell and Dutton (1967) approach (Bondada and Robertson, 2003). Triplicate cultures were run for each mouse with and without SRBC. Four days later, a glass slide modification of Jerne and Nordin (1963) PFC assay was performed. Briefly, the immunized spleen cells were collected from individual cultures, and washed twice with RPMI 1640. The immunized spleen cells with 50 μl 50% SRBC were then added into the appropriate glass tubes. These tubes were placed in a 43°C constant temperature water bath with 400 μl 0.8% Seaplaque agarose (Intermountain Scientific, Kaysville, UT). SRBC were added to the tubes and one slide was used for each culture (triplicate) to determine the PFC response. The mixture of spleen cells and SRBC was poured onto 3 × 1 × 1 mm, 0.15% Seaplaque agarose precoated microscope slide and allowed to cool. The slides were incubated for 1.5 h at 37°C in a humidified without CO₂ incubator. Guinea pig complement (Colorado Serum, Denver, CO) in Dulbecco's phosphate buffered saline (DPBS) with calcium (1:20) was used to flood the slides on each tray. Following an additional 1.5 h incubation at 37°C, the number of anti-SRBC plaque-forming cells (PFC) per culture were identified. The data are presented as the number of PFC/culture (10⁶ cells per culture on day 0).

Flow cytometric analysis.

After spleen cells were harvested, 1 × 10⁶ cells were aliquoted into three 12 × 75 mm tubes, and their surface marker expression was analyzed using a FACS Calibur Flow Cytometry system (Becton Dickinson Immunocytometry Systems, San Jose, CA). Three combinations of custom rat anti-mouse monoclonal antibody cocktails were custom ordered from BD Biosciences (BD Pharmingen, San Diego, CA). Splenic cells were first incubated with purified rat anti-mouse CD16/CD32 monoclonal antibody (Fc block antibody) (BD Pharmingen, San Diego, CA) for 10 min at room temperature in the dark. To detect the lymphatic subpopulations, 20 μl of antibody cocktail containing IgG1+IgG2a-FITC/IgM-PE/CD45- PerCP/IgG2a-APC, or CD3-FITC/CD8a-PE/CD45 -PerCP/CD4-APC or CD3+CD19-FITC/Pan NK-PE/CD45-PerCP/Mac-1APC was added to the appropriate sample tube. After 30 min incubation in the dark, 2 ml of 1X fresh ammonium chloride was added to each sample to lyse red blood cells, samples were then incubated at room temperature for 10 min in the dark. Samples were centrifuged at 275 × g for 10 min, supernatants were aspirated and the pellets were washed with 2 ml of the DPBS wash buffer (Sigma Chemical Co, St. Louis, MO) [contains sodium azide and fetal bovine serum] and then centrifuged as above. Cells were then resuspended in 400 μl of PBS wash buffer, tubes were capped and covered by aluminum foil for transport to the Flow Cytometry Facility for analysis. Data were acquired by gating on CD45 positive cells and acquiring 10,000 gated events. CellQuest software was used to analyze the data.

Natural killer cell assay.

To measure the nonspecific immunity of natural killing cells (NK), NK cell activity was quantitated by determining the ability of NK cells to lyse the NK sensitive Yac-1 target cells (ATCC, Manassas, VA). Briefly, the Yac-1 target cells, 2 × 10⁶ cells/ml were suspended in complete RMPI 1640 medium and radiolabeled with sodium chromate (⁵¹Cr) (Perkin-Elmer, Wellesley, MA), for 1 h in a humidified, 37°C, 5% CO₂ incubator. Following loading of Yac-1 target (T) cells with ⁵¹Cr, excess ⁵¹Cr was washed away and cells were resuspended at 5 × 10⁴ cells/ml and held in a humidified, 37°C, 5% CO₂ incubator. Spleen cells were plated as effector (E) cells at 2 × 10⁶ cells/ml and then serially diluted in triplicate. Dilutions were made by plating, 200 μl of cells onto the first three wells of the appropriate rows of 96 well round-bottom culture plate (Corning Incorporated, Corning, NY), and then serially diluted three wells at a time, 100 ml per well across the plate. One-hundred Ml ⁵¹Cr labeled target cell was then added to each well. The resulting effector/target (E/T) ratios were 200:1, 100:1, 50:1, and 25:1. The cells were co-cultured for 4 h in a humidified, 37°C 5% CO₂ incubator. Following incubation, plates were centrifuged and 100 μl of supernatant was harvested from each well. The amount of intracellular ⁵¹Cr released into the supernatant was measured with the Wallac 1480 WIZARD gamma counter. The first 12 wells on each culture plate were control wells used to measure the spontaneous ⁵¹Cr release and maximum ⁵¹Cr release by target cells. To measure the maximum ⁵¹Cr release by target cells, 100 μl of 5% Triton X-100 solution was added to the appropriate wells. The percentage of lysis was calculated by (mean sample CPM − mean spontaneous CPM)/(mean total release CPM-mean spontaneous CPM) × 100.

Statistical analysis.

All of the data reported in this paper were analyzed by SigmaStat software (Jandel Scientific, San Rafael, CA). The statistical differences were determined by one-way analysis of variance (ANOVA) followed by a Bonferroni multiple comparison test. A p-value of ≤ 0.05 was considered significant. The student's t-test was used to assess the statistical significance of the NK cytotoxicity effects.

RESULTS

DMBA Produces Spleen Cell Toxicity in C57BL/6 WT, but Not CYP1B1 (-/-) Mice

As shown in Table 1, after 5 days DMBA treatment of both WT and CYP1B1 (-/-) mice, the body weight did not significantly change (18.3 ± 3.4 g to 21.4 ± 1.5 g). No differences between treatment groups of the CYP1B1 (-/-) mice were observed. However, the spleen weights were significantly smaller after the 50 mg/kg and 150 mg/kg DMBA treatments in WT mice, while there was no change in the CYP 1B1 (-/-) mice (Table 1). We have previously shown that oral exposure of mice to DMBA produces apoptosis in murine spleen cells (Burchiel et al., 1990). These results demonstrate that CYP1B1 is required for spleen cell cytotoxicity.

DMBA Suppresses T- and B-Cell Mitogenesis in WT, but Not CYP1B1 (-/-) Mice

In the mitogenesis assay, B-cell and T-cell proliferation were analyzed by stimulation with LPS or Con A mitogens, respectively. In female C57BL/6 WT mice, DMBA was found to produce a dose-dependent inhibition of B cell proliferation at all exposure levels (Fig. 1, top). DMBA at low doses produced more suppression of T-cell proliferation than the response of B cells to mitogens. However, in the present experiment there was a slight increase in the amount of T-cell proliferation at 150 mg/kg compared to 50 mg/kg in this experiment, which was not seen in replicate experiments. CYP1B1 (-/-) mice showed no significant differences between the corn oil and the DMBA treatment groups for either Con A or LPS induced cell proliferation (Fig. 1, bottom). These findings indicate that DMBA suppresses B and T cell proliferation in the presence of CYP1B1, but this effect is lost in the absence of CYP1B1.

DMBA Suppression of the in Vitro Spleen Cell Response to SRBC is CYP1B1-Dependent

T-dependent humoral immune responses are one of the most sensitive and frequently used methods to evaluate the immunotoxicity of xenobiotics. In our studies, we utilized an in vitro immunization to sheep erythrocytes (SRBC) followed by a direct plaque-forming cell (PFC) assay. In WT mice treated with DMBA, a dose-dependent decrease in the number of PFC was seen starting at the 17 mg/kg dose and the trend reached statistical significance at the 50 and 150 mg/kg DMBA doses compared to the corn oil group (Fig. 2, top). The dose-dependent suppression of the PFC response was not seen in CYP1B1 (-/-) mice (Fig. 2, bottom). These results demonstrate that CYP1B1 is necessary for DMBA-induced immunosuppression of humoral immunity.

Effect of DMBA on NK Cytotoxicity Activity

Figure 3 (top) depicts NK cell cytotoxic activity in WT mice using a ⁵¹Cr release assay. A significant decrease in NK activity was observed in the 150 mg/kg DMBA treated CYP1B1 (-/-) group compared to the control corn oil group. Because this effect was not seen in the WT group, we conclude that other metabolic pathways may be involved in this effect (peroxidase?). At the 50 mg/kg dose of DMBA, there was significant suppression of the NK cytotoxic activity compared to the corn oil group. This suppression was not seen in CYP1B1 (-/-) mice (Fig. 3, bottom). Collectively, these results suggest that CYP1B1 and non-CYP1B1 metabolic pathways may contribute to the suppression of NK activity by DMBA.

DMBA Treatment Dose Not Alter the Cell Surface Markers

We utilized custom antibody cocktails to analyze spleen cell subpopulations. A purified rat anti-mouse CD16/CD32 monoclonal antibody was used to block the nonspecific Fc binding sites. Spleen cells were immunostained with different cell surface marker antibodies as described in Materials and Methods. A common leukocyte surface marker (CD45) was used in all staining reactions. Lymphocyte subsets were detected by flow cytometry based on the following cell surface markers: total T cells (CD3), T_H cells (CD4), cytotoxic T cell (CD8), B cell (CD19), NK cell (CD16), and macrophage (Mac-1). There were no significant changes in any of cell surface markers expressed in the WT or CYP1B1 (-/-) mice after DMBA treatment (Fig. 4). These findings are somewhat different than our previous results obtained in B6C3F1 mice, where we observed a slight decrease in splenic T cells four and eight weeks after DMBA exposure (Burchiel et al., 1988). The delay in analysis in the previous work is likely responsible for the different result.

DISCUSSION

The present study was designed to investigate the mechanism of DMBA immunotoxicity in mice. CYP1B1 has previously been implicated in the hematotoxicity induced by DMBA in murine bone marrow (Heidel et al., 1999; Page et al., 2003). CYP1B1 has not previously been assessed or implicated in systemic immunotoxicity. Knockout mice have proved to be very useful in unraveling the mechanisms of action of xenobiotics (Gonzalez, 2001). We used CYP1B1 (-/-) mice bred on a C57BL/6 background as our animal model and compared immune effects to WT mice to characterize the role of CYP1B1 in DMBA-induced spleen cell toxicity. After DMBA treatment, no obvious physical changes, such as body weight, or animal morbidity or mortality were observed in either the WT or CYP1B1 (-/-) groups of mice. However, we found that spleen weight in WT mice was significantly decreased in the 50 mg/kg and 150 mg/kg DMBA treatment groups demonstrating the cytotoxicity of DMBA at these two doses. There was no effect of DMBA and spleen cell recovery or organ weights (spleen) in the CYP1B1 (-/-) mice.

Based on our own biodistribution studies of oral DMBA exposure in mice (Archuleta et al., 1992) and the recent results of Galvan et al. (2005), we do not believe that the lack of CYP1B1 expression in mice alters DMBA pharmacokinetics. Previous studies with benzo(a)pyrene (BaP) in CYP1A1 null mice showed a dramatic increase in the levels of BaP in the blood and liver of mice, and pretreatment with TCDD accelerated BaP clearance in WT mice (Uno et al., 2004). However, because CYP1B1 is not expressed in the liver, there is no first pass metabolism that occurs following gavage administration in mice. In addition, we performed a broad toxicogenomic analysis of expressed genes in the spleens of mice reported in the present studies and found that there was no CYP1A1 induction in WT or CYP1B1 null mice, nor CYP1B1 induction in WT mice spleens (manuscript in preparation). Therefore, we hypothesize that the lack of susceptibility of CYP1B1 null mice to DMBA results from a change in the local peripheral metabolism in CYP1B1-expressing organs and tissues, such as the spleen (Choudhary et al., 2003). Galvan et al. (2005) also found that low levels of CYP1B1 expressed locally in the bone marrow was likely responsible for the hematotoxicity of DMBA.

The present studies assessed immune function endpoints that have previously been shown to be sensitive to DMBA treatment in murine models (Burchiel et al., 1988; Davis et al., 1991; Thurmond et al., 1987). We observed significant suppression of B- and T-cell mitogenesis and the in vitro plaque-forming cell response to SRBC in WT mice. However, the CYP1B1 (-/-) mice were protected from this immunosuppression produced by DMBA. We found no effects on surface marker expression for B cells, T cells, NK cell, and monocytes in either WT or CYP1B1 (-/-) mice. NK activity was suppressed by DMBA at the 50 mg/kg dose level, and this was not observed in the CYP 1B1 (-/-) mice. These results clearly demonstrate that CYP1B1 is required for DMBA-induced immunotoxicity in the spleen.

Previous studies have found that the aromatic hydrocarbon receptor (AhR) is involved in the induction CYP1B1 expression after xenobiotic exposure, such as PAHs (Hankinson, 1995). However, Thurmond et al. (1987) found that DMBA was equally immunosuppressive in AhR high and low affinity strains and concluded that DMBA acts via AhR-independent mechanisms. Recent studies in our lab support this conclusion as we found that AhR (-/-) mice are not protected against the immunotoxicity of DMBA (Burchiel et al., in preparation). A possible explanation for these findings is that constitutive expression of CYP1B1 occurs in many tissues including mouse spleen (Choudhary et al., 2003) that is adequate for activation of DMBA independent of any AhR induction.

DMBA has been shown to be a preferred substrate for CYP1B1 metabolism leading to the formation of reactive metabolites that bind to DNA (Buters et al., 1999, 2003; Kleiner et al., 2002; Savas et al., 1997; Shimada and Fujii-Kuriyama, 2004). In the presence of CYP1B1 and EPHX1, DMBA forms reactive metabolites that bind to DNA and produce genotoxicity (Gonzalez, 2001; RamaKrishna et al., 1992). Jefcoate and colleagues found that DMBA was metabolized by CYP1B1 to DMBA-3,4-dihydrodiol in mouse bone marrow stem cells (Heidel et al., 1998), and this metabolite has been implicated in DNA adduction (Slaga et al., 1979). DMBA-3,4-diol-1,2-epoxide (DMBA-DE) is considered to be the ultimate carcinogen of DMBA requiring two rounds of metabolism by CYP1B1 with an intervening conversion of DMBA-3,4-epoxide to DMBA-3,4-diol by the EPHX1 enzyme. Thus, we believe that EPHX1 may also be required for the immunotoxicity of DMBA. To confirm the role of EPHX1 in DMBA immunotoxicity, studies are currently underway in EPHX1 (-/-) mice (Miyata et al., 1999) to further support the role of this metabolic pathway in the immunotoxicity of DMBA. Our tentative conclusion is that DMBA-DE will likely be an important metabolite in the immunotoxicity of DMBA.

In conclusion, we have found that DMBA produces immunosuppression of both humoral and cell-mediated in a dose dependent manner in WT mice. Splenic T and B cells are both susceptible to immunosuppression by DMBA, and at some doses, NK cells may also be adversely affected. CYP1B1 deficiency protects mice against DMBA-induced immunosuppression in the spleen. Hence, CYP1B1 is a dominant enzyme involved in the formation of DMBA metabolites that appear to target the immune system.

The authors would like to thank Dr. Frank Gonzalez and the National Institutes of Health for providing the CYP1B1 knock out mice. This study was supported by RO1 ES05495 and P30 ES012072.

References