ToxSci Advance Access originally published online on April 18, 2007
Toxicological Sciences 2007 98(1):137-144; doi:10.1093/toxsci/kfm089
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Microsomal Expoxide Hydrolase Is Required for 7,12-Dimethylbenz[a]anthracene (DMBA)–Induced Immunotoxicity in Mice
The University of New Mexico College of Pharmacy Toxicology Program, Albuquerque, New Mexico 87131-0001
1 To whom correspondence should be addressed at 1 University of New Mexico - MSC09 5360, Albuquerque, NM 87131-0001. Fax: (505) 272-6749. E-mail: sburchiel{at}salud.unm.edu.
Received April 3, 2007; accepted April 6, 2007
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Microsomal epoxide hydrolase (mEH, EPHX1) is involved in the metabolism of chemicals to generate dihydrodiol intermediates in the presence of the cytochrome P450. We have previously shown that 7,12-dimethylbenz[a]anthracene (DMBA) can suppress both cell-mediated and humoral immune responses in wild-type (WT) C57BL/6N mice but not in CYP1B1 null mice. In the present studies, we hypothesized the critical metabolite responsible for DMBA-induced immunotoxicity is likely to be the 3,4-dihydrodiol-1,2-epoxide metabolite of DMBA, which requires mEH for formation. Mice were gavaged orally with DMBA (0, 17, 50, and 150 mg/kg) once a day for 5 days. Immune function and other assays were performed on day 7. Our data showed that unlike WT mice, DMBA treatment of mEH null mice produced no alterations in the body weight, spleen weight, or spleen cellularity. Similarly, DMBA treatments did not affect the PFC response in mEH null mice. Natural killer activity was not altered by DMBA treatment in mEH null mice. T-cell mitogenesis was partially suppressed by 50 and 150 mg/kg DMBA treatments of mEH null mice, but B-cell mitogenesis was not affected. Finally, we assessed the biodistribution of DMBA in both C57BL/6N WT and mEH null mice in spleen, thymus, and liver after 24 h and 7 days oral gavage. The concentrations of DMBA in each organ were not significantly different in WT and in mEH null mice. Collectively, these results demonstrate that mEH (EPHX1 gene) is a crucial enzyme for metabolic activation of DMBA in vivo leading to immunosuppression of spleen cells.
Key Words: epoxide hydrolase; PAHs; immunotoxicity; DMBA.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Polycyclic aromatic hydrocarbons (PAHs) produced by incomplete combustion of organic fuels are present universally as environmental pollutants. High levels of PAHs are present in cigarette smoke, vehicle exhaust, and charcoal-grilled foods. Many of the members of the PAH family are toxicants, and can induce carcinogenic and immunotoxic effects (Conney, 1982
; Pelkonen and Nebert, 1982). As a model PAH agent, 7,12-dimethylbenz[a]anthracene (DMBA) is widely used to investigate carcinogenesis and immunotoxicity in various species, tissues, and cell types (Burchiel et al., 1988, 1990; Buters et al., 1999, 2003; Shimada and Fujii-Kuriyama, 2004; Sugiyama et al., 2002; Thurmond et al., 1988).
Previous studies have shown that PAHs such as DMBA require metabolic activation to produce hematotoxicity (Buters et al., 2003; Heidel et al., 2000). Many phase I and phase II enzymes may be involved in this activation process. The precise mechanistic pathway for this bioactivation and immunosuppression is not clearly understood. Recent data from our laboratory indicates that DMBA-induced immunotoxicity occurs in a CYP1B1-dependent manner in vivo (Gao et al., 2005). We observed that CYP1B1 null mice were protected from the cytotoxic effects of DMBA, and the cell-mediated and humoral immunosuppression induced by DMBA in WT mice was totally eliminated in CYP1B1 null mice. Therefore, it has become clear that metabolic activation by CYP1B1 is a key step in DMBA-induced immunosuppression.
In addition to CYP1B1, another key enzyme responsible for DMBA bioactivation is microsomal epoxide hydrolase (mEH, EPHX1) (Gonzalez, 2001). The epoxide hydrolases (EHs) are a family of enzymes that function to hydrate simple epoxides to vicinal diols and arene oxides to trans-dihydrodiols (Arand et al., 2005; Fretland and Omiecinski, 2000; Omiecinski et al., 2000). In mammalian cells, there are several EHs, including two widely investigated distinct xenobiotic EHs (Morisseau and Hammock, 2005): membrane-bound mEH and soluble EH (sEH). In general, sEH principally participates in the metabolism of endogenous substances, for example, epoxides of steroids, arachidonic acid derivatives (Schmelzer et al., 2005; Zeldin et al., 1995), and leukotrienes (Haeggstrom et al., 1994). In contrast, mEH is primarily associated with the metabolism of xenobiotic chemicals catalyzing hydrolysis of epoxide intermediates, and exhibiting broad substrate specificity (Friedberg et al., 1994; Newman et al., 2005). As a conserved phase I biotransformation enzyme, mEH is expressed in many tissues and cell types (Omiecinski et al., 2000). In addition to xenobiotic metabolism, mEH catalyzes hydrolysis of some endogenous lipid substrates as well (Newman et al., 2005; Vogel-Bindel et al., 1982). Thus, it plays some important physiological roles. Usually, mEH plays a critical role in detoxification of environmental contaminants because metabolism of reactive epoxides by mEH will produce less reactive and less toxic dihydrodiol intermediates. However, mEH is thought to be one of the most important enzymes for metabolic activation of DMBA resulting in carcinogenesis (Miyata et al., 1999) and immunosuppression (Miyata et al., 2001).
mEH is attached to the endoplasmic reticulum membrane at the N-terminal and is anchored toward the cell cytosol (Zhu et al., 1999). This structure supports the association of mEH with other CYPs, such as CYP1B1, to form a multienzyme complex (Seidegard and DePierre, 1983). Based on its structure, the first epoxide substrate (DMBA-3,4-epoxide) produced by CYP1B1 will be biotransformed to DMBA-3,4-dihydrodiol (Fig. 1). Finally, the ultimate metabolite, DMBA-3,4-dihydrodiol-1,2-epoxide (DMBA-DE) can be produced, leading to covalent binding to DNA and resultant DNA damage (Dipple and Nebzydoski, 1978; Tierney et al., 1978; Wislocki et al., 1980). We believe that this genotoxic pathway is the primary mechanism associated with DMBA-induced immunotoxicity. Murine skin tumorigenesis studies have shown that mEH-deficient mice are less responsive to the carcinogenic effect of DMBA (Miyata et al., 1999).
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Based on studies of the role of mEH in DMBA carcinogenicity, we hypothesized that mEH is an essential enzyme for DMBA-induced splenic immunotoxicity. In this study, we utilized mEH null mice to evaluate the role of mEH in DMBA-induced immunosuppression in vivo. Our data show that mEH null mice were highly resistant to DMBA-induced spleen cell immunosuppression, and thus a diol-epoxide mechanism is likely associated with DMBA immunotoxicity.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Chemicals, reagents and standards.
All solvents and reagents used were of analytical or high-performance liquid chromatography (HPLC) grade. DMBA (Cat. No. D3254-5G) was purchased from Sigma-Aldrich (St Louis, MO) at greater than 95% purity and was periodically checked for purity. Concanavalin A (Con A, Type IV, Cat. No. C-2010) and corn oil were also purchased from Sigma-Aldrich. Lipopolysaccharide (LPS) from Escherichia coli was purchased from Alexis Biochemicals (San Diego, CA). Cell culture materials were from Sigma-Aldrich and Invitrogen (Grand Island, NY). Methanol (HPLC grade) was purchased from Burdick & Jackson (Muskegon, MI). Hexanes (95% n-hexane) was purchased from J. T. Baker (Phillipsburg, NJ). Dibenzo[a,l]pyrene (DBP) (purity > 99%) was obtained from the National Cancer Institute chemical carcinogen reference standard repository (Kansas City, MO).
Animals.
Female wild-type (WT) C57BL/6N mice (6-8 weeks) were purchased from Harlan Laboratories (Indianapolis, IN). mEH (EPHX1 gene) null (–/–) mouse breeders (Miyata et al., 1999) were obtained from the National Institutes of Health (NIH) as a C57BL/6N F4 generation. Heterozygous mice were backcrossed with WT C57BL/6N mice to generation F11 in our AAALAC-accredited animal facility and our studies were conducted under an Institutional Animal Care and Use Committee (IACUC)–approved protocol. All of the mEH null mice were confirmed using a PCR genotyping method with DNA isolated from tail snips (Miyata et al., 1999). Female mEH null mice used for the immune function and pharmacokinetic studies were 12–16 weeks old. In the immune function study, each group (five mice) was oral gavaged with corn oil (vehicle control) or DMBA once a day for 5 days. The cumulative dose groups of DMBA were 17, 50, and 150 mg/kg. Spleens were aseptically isolated from CO2 euthanized mice 48 h following the last DMBA treatment (day 7). The preparation of spleen cell suspensions has been described previously (Gao et al., 2005). For pharmacokinetic studies, WT, and knockout mice were orally gavaged with corn oil or 30 mg/kg/day DMBA once a day for 5 days. After 24 h or 7 days oral gavage, three mice from each group were euthanized using CO2 euthanasia. Spleen, liver, and thymus were rapidly removed, frozen in liquid nitrogen, and then stored at –80°C for the HPLC biodistribution assay.
T-dependent sheep erythrocyte in vitro immunization and plaque-forming cell assay.
Single spleen cell suspensions prepared from individual mice were used to examine the induction of a primary humoral immune response to sheep red blood cells (SRBC) using a 4-day modified Mishell–Dutton culture system, as described by Bondada and Robertson (2003). Briefly, mouse spleen cells (2 x 106 cells/ml, 0.5 ml) were cocultured with equal volume of 1% SRBC (Colorado Serum, Denver, CO) or complete RPMI 1640 medium (containing 10% heat inactivated fetal bovine serum [Hyclone, Logan, UT], 50µM 2-mercaptoethanol [GIBCO, Grand Island, NY], 1mM sodium pyruvate [GIBCO, Grand Island, NY], 50 µg/ml gentamycin [GIBCO, Grand Island, NY], 2mM L-glutamine, 100 µg/ml streptomycin, and 100 Units/ml penicillin) as nonspecific plaque-forming control in 48-well, flat-bottomed plates (Corning Glass, Corning, NY). Duplicate cultures were set-up for each mouse with and without SRBC. After 4 days incubation, the plaque-forming cell (PFC) assay was performed using a glass slide modification of Jerne and Nordin, as previously described by our lab (Gao et al., 2005). Briefly, the immunized spleen cells or control cells were washed twice with complete RPMI 1640. Then the immunized spleen cells or control cells were added into 43°C prewarmed appropriate glass tubes with 400 µl of 0.8% Seaplaque® agarose (Intermountain Scientific, Kaysville, UT) and 50 µl of 50% SRBC mixture. The mixtures were poured on the appropriate 0.15% Seaplaque® agarose precoated microscope slide to determine the PFC response. The slides were incubated for 1 h at 37°C without CO2 in a humidified incubator to allow for antibody binding to the SRBC. Diluted (1:20) guinea pig complement (Colorado serum, Denver, CO) in Dulbecco's phosphate-buffered saline (DPBS with calcium and magnesium) was used to flood the slides on each slide tray. After an additional 2 h incubation at 37°C without CO2, the number of anti-SRBC PFC per culture was identified using a Bausch & Lomb dissecting microscope. The data are presented as the number of PFC/culture.
3H-thymidine incorporation mitogenesis assay.
Two mitogens, LPS and concanavillin (Con A), were used to evaluate B and T-cell proliferation, respectively. Murine spleen cells were plated at a concentration of 1 x 106 cells/ml (200 µl) with 50 µl mitogen (50 µg/ml LPS or 5 µg/ml Con A) or RPMI complete medium (no mitogen control to monitor spontaneous proliferation) (supplemented with 10% fetal bovine serum, 2mM L-glutamine, 100 µg/ml streptomycin and 100 Units/ml penicillin) for 3 days in 96 well culture plates. The replicates of six cultures were used for each mitogen and each mouse spleen sample. After 48 h incubation at 37°C in a humidified, 5% CO2 incubator, cells were pulsed with 20 µl of 50 µCi/ml [3H]-thymidine (ICN, Aurora, OH) to label the DNA of the actively dividing cells. The incubation was continued for an additional 18 h with the above conditions. Cells were harvested onto glass filters using a Brandel Model 24V cell harvester. Filter samples were dried at room temperature for approximately 30 min, and then transferred to liquid scintillation vials containing 3 ml of ScintiVerse BD cocktail (Fisher Scientific, Houston, TX). A liquid scintillation counter was used to detect the incorporated [3H]-thymidine. Proliferation in response to LPS and Con A was expressed as counts per minute (CPM).
Flow cytometric analysis.
The subtypes of cells in the spleen were characterized by specific cell surface markers. Three combinations of custom rat anti-mouse monoclonal antibody cocktails for six types of cell surface markers were obtained from BD Biosciences (BD Pharmingen, San Diego, CA): CD3 (pan T cells), CD4 (TH cells), CD8 (Tc cells), CD19 (B cells), CD16, gated (natural killer [NK] cells), Mac-1 (macrophages). Freshly isolated mouse spleen cells (1 x 106 cells/mouse) were aliquoted into three 12 x 75-mm flow tubes, and 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. A total of 20 µl of antibody cocktail containing IgG1 + IgG2a-fluorescent isothiocyanate (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 then added to the appropriate sample tube, and incubated for 30 min in the dark. To lyse red blood cells, fresh 1x ammonium chloride (2 ml per sample) was added to each tube, and incubated at room temperature for 10 min in the dark. Following incubation, samples were centrifuged at 275 x g for 10 min, supernatants were aspirated. Cell pellets were washed with 2 ml of PBS wash buffer (PBS, Sigma Chemical Co, St Louis, MO, with added 0.09% sodium azide and 1% heat inactivated fetal bovine serum) and then centrifuged as above. Supernatants were removed and cells were resuspended in 400 µl of PBS wash buffer. Tubes were capped and covered with aluminum foil for transport to the Flow Cytometry Facility for analysis. Samples were analyzed using a FACScalibur Flow Cytometer (Becton Dickinson Immunocytometry Systems, San Jose, CA). Data were acquired by gating on CD45-positive cells and acquiring 10,000 gated events for each staining analysis. The data were collected using CellQuest® software.
Natural killer cell 51Cr assay.
The NK cell activity was assessed by measuring chromium (51Cr) release from the NK sensitive Yac-1 target cells (ATCC, Manassas, VA). In brief, the Yac-1 target cells (T, target) (2 x 106 cells) were incubated with 1.0 mCi sodium chromate (51Cr) (Perkin–Elmer, Wellesley, MA) in a 50-ml centrifuge tube for 1 h in a humidified, 37°C, 5% CO2 incubator. The cells were washed twice with 10 ml of DPBS, and then resuspended at 5 x 104 cells/ml and held in a humidified, 37°C, 5% CO2 incubator. Spleen cells were plated as effector (E, effector) cells in a 96 well round-bottom culture plate (Corning Incorporated, Corning, NY) in triplicate. The first three wells of the appropriate rows were plated with 200 µl of 2 x 106 cells/ml spleen cells, and then serially diluted three wells at a time, 100 µl per well across the plate. A total of 100 µl of 51Cr labeled target cells was mixed with effector cells in each well to produce the indicated effector to target (E:T) ratios (200:1, 100:1, 50:1, and 25:1). The first 12 wells on each culture plate, contained target cells alone in complete RPMI 1640 medium; 5% Triton X-100 served for spontaneous and maximal 51Cr release. After 4 h coculturing in a humidified, 37°C, 5% CO2 incubator, plates were centrifuged at 275 x g for 5 min. The supernatant (100 µl) was collected from each well and the radioactivity was measured with a Wallac 1480 WIZARD gamma counter. The percentage of lysis was calculated by the following formula: % lysis = (mean sample 51Cr release CPM – mean spontaneous 51Cr release CPM)/(mean maximum 51Cr release CPM – mean spontaneous 51Cr release CPM) x 100.
Preparation of tissue extraction for HPLC analysis.
Tissue snips weighing 
100 mg, from the spleen, liver, and thymus (< 100 mg), were used for this study. Tissues were homogenized in 1 ml of homogenization buffer containing 10mM Tris, 0.1M K2HPO4, 0.1M KCl, and 10mM ethylenediaminetetraacetic acid (pH = 7.4). After thoroughly homogenizing the tissue using a motorized tissue grinder, the tissue suspension was centrifuged at 1000 x g for 10 min. The supernatant was collected in a 15-ml centrifuge tube. The tissue fragments were homogenized again in 1 ml of homogenization buffer and centrifuged. The supernatant was collected in the same tube as the first collection. Two milliliters of formic acid was added and mixed with the collected supernatant. The mixture was spiked with 100 µl, 5µM DBP which was used as an internal standard for HPLC analysis. Finally, samples were mixed with 2 ml of hexane and loaded on a LabQuake shaker for 15 min. Samples were centrifuged again and the upper organic phase was collected into a fresh 15-ml centrifuge tube. The samples were extracted one more time to get high extraction efficiency. Before HPLC analysis, the collected hexane was evaporated under a compressed nitrogen gas stream and then 200 µl of methanol was added to each sample.
HPLC analysis of DMBA in tissues.
HPLC analysis was performed with a P200 Spectraphysics HPLC system (Spectra Physics, Santa Clara, CA), consisting of P1500 pump, a 20-µl AS1000 autosampler and a fluorescence detector (FL 2000). The system was controlled through Spectra PC 1000 software with a data processing system. A C18 reversed-phase column (250 x 4.6 mm, 5µm) (Beckman Instruments, Fullerton, CA) with a guard protecting column was used to separate the components in each sample. DMBA was evaluated in methanol samples using a flow rate of 1 ml/min. The column was eluted with a mobile phase consisting of methanol: H2O (92:8) for 26 min. The effluent was detected by a fluorescence detector using an excitation wavelength of 276 nm and emission wavelength of 376 nm. The retention times of DMBA and DBP were 12.7 min and 20.4 min, respectively.
Statistical analysis.
All of the data were analyzed using Sigma Stat software (Jandel Science, San Rafael, CA). The statistical differences between groups were determined by one-way analysis of variance and/or Student's t-test. The data are presented as mean ± SEM. A p-value < 0.05 was used to determine statistical significance.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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DMBA Does not Produce General Toxicity in mEH Null Mice
Previous studies in our laboratory have shown that treatment of C57BL/6N WT mice for 5 days with oral DMBA at cumulative doses of 50 and 150 mg/kg results in a decrease in spleen weight and a decrease in the total lymphocytes recovered from the spleen (Gao et al., 2005
). In the current study, there were no significant changes in the spleen weights or cell recoveries from the spleen for any of the dose groups in mEH null mice (Table 1). DMBA treatment produced no changes in body weight in comparison with the corn oil control group of mice.
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DMBA-Induced Immunuosuppression of Spleen cells is Not Seen in mEH Null Mice as Measured by the SRBC PFC Response
The T-dependent antibody response to SRBC and the PFC assay has been extremely useful for studying the effects of xenobiotics on the immune system of mice (Luster et al., 1988
). We have repeatedly shown that DMBA suppresses the SRBC antibody response of male and female mice following subcutaneous dosing or oral gavage in several mouse strains (Burchiel et al., 1988, 1990), as well as mitogenic responses of T and B cells (Davis et al., 1991). Most recently we have demonstrated extreme sensitivity of the PFC response and mitogen responses in C57BL/6N WT mice treated with DMBA (Gao et al., 2005). In these previous studies, the PFC numbers were decreased starting at 17 mg/kg DMBA, and virtually completely suppressed (> 12 fold decrease) at 150 mg/kg DMBA. Because we have performed numerous studies in WT mice including the C57BL/6N strain used in the present studies, we did not perform additional immune function experiments in WT mice for the present studies, as recommended by our IACUC. In the present studies, we found that unlike WT C57BL/6N mice that are suppressed at total cumulative doses of 17–150 mg/kg, mEH null mice demonstrated no significant response to DMBA for the PFC assay as compared to the corn oil control group (Fig. 2). This result demonstrates that mEH is critical for DMBA-induced immunosuppression, and in concert with our previous work demonstrating a requirement for CYP1B1, supports the hypothesis that a critical metabolite required for DMBA immunotoxicity is the DE of DMBA (DMBA-DE, Fig. 1)
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DMBA Suppresses T Cell Mitogenesis, but not B-Cell Mitogenesis in mEH Null Mice
The proliferative responses of splenic lymphocytes were investigated using T (Con A) and B (LPS) cell mitogens. In previous studies in WT mice, we observed that DMBA induced a significant suppression of mitogenic responses of both T and B cells (Gao et al., 2005
). In the present studies with mEH null mice, DMBA treatments did not alter splenic mitogenic response to the B-cell mitogen compared with corn oil control (Fig. 3). However, the T-cell proliferative response was significantly suppressed by 50 and 150 mg/kg DMBA treatments. Previously, it has been shown that in WT mice the splenic mitogenic response to T mitogens was reduced about 57% and 87% by 17 and 50 mg/kg DMBA, respectively. In contrast, we did not observe significant T cell proliferative suppression at 17 mg/kg DMBA treatment in mEH null mice. The splenic mitogenic response to Con A for mEH null mice was reduced to 38% and 43% by 50 and 150 mg/kg DMBA, respectively. This percentage of reduction of T-cell proliferation in mEH null mice was less than WT mice. Thus, mEH null mice were protected to some extent compared to WT mice. These results indicate that mEH is necessary for DMBA to suppress mitogen-induced B-cell proliferation, and partially protects T cells from DMBA-induced splenic toxicity.
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DMBA Does Not Alter NK Activity in mEH Null Mice
To further understand the role of mEH in DMBA-induced nonspecific immunotoxicity, NK cell activity was investigated using a chromium (51Cr) release assay. Normally, we observe that the percentage of lysis of NK cells is significantly reduced by DMBA treatments in WT mice (Gao et al., 2005
). In contrast in mEH null mice, the DMBA reduction of NK activity was not observed (Fig. 4). As expected, the NK cells from mEH null mice produced 15–20% lysis at the E:T ratio of 200:1. Again, these results suggest that mEH plays a role in susceptibility of NK cells to DMBA-induced immunosuppression.
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Subpopulations of Spleen Cells in mEH Null Mice after DMBA Treatment
To examine the impact of DMBA treatments on the spleen cell subsets in mEH null mice, the following cell surface markers were used: CD3 (pan T cells), CD4 (TH cells), CD8 (cytotoxic T cells), CD19 (B cells), CD16 (NK cells), and Mac-1 (macrophages). The common leukocyte population was selected using the CD45 surface marker. As depicted in Figure 5, most of the subpopulations of splenic lymphocytes did not change between DMBA treatments and corn oil control in mEH null mice. The only significant decrease was observed in macrophages at the 150 mg/kg DMBA treatment. However, this decrease was not seen in replicate experiments. In our previous study (Gao et al., 2005
), the DMBA 5-day oral treatments also did not alter these six subsets of splenic lymphocytes in WT mice.
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DMBA Levels in Spleen, Thymus and Liver Following Oral Administration in WT and mEH Null Mice
In order to determine the difference in the biodistribution of DMBA between WT and mEH null mice, we analyzed the parent DMBA levels in three organs (spleen, thymus, and liver) from each DMBA treated mouse using HPLC. As shown in Figure 6, DMBA was detected in extracted tissue samples with 12.7-min retention time. The reasons we chose these three organs are (1) spleens were used to evaluate systemic immunosuppression in the current mEH immune function study; (2) the thymus is an essential organ for the development of T cells; and (3) the liver is an important organ in the metabolism and clearance of DMBA, and DMBA can undergo "first pass" metabolism following oral gavage.
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To determine whether the parent DMBA organ/tissue levels changed as a result of knocking out mEH expression in mice, it was necessary to make a comparison between WT and mEH null mice. Our studies utilized a 5-day dosing period for immunotoxicity studies and therefore we wanted to perform tissue analysis studies during a comparable time period. In addition, our doses of exposure ranged from 17 to 150 mg/kg. We therefore chose a 5-day oral dose of 30 mg/kg/day DMBA administered over a 5-day period, as well as a single dose study performed after 24 h on day 1.
Free DMBA levels were measured by HPLC after 24 h following a single oral dose and day 7 following 5 days oral dosing. As shown in Figure 7, we did not observe any significant differences in DMBA levels in the spleens, thymuses, or livers from WT and mEH null mice after day 1 and day 7. The recovered concentration of DMBA was slightly decreased at day 7 for mEH null mice in spleen and liver. However, there was not a significant difference with the WT mice. Overall, these results suggest that knockout of the mEH (EPHX1 gene) does not significantly alter the biodistribution of DMBA in the spleen, thymus, and liver of C57BL/6N mice.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The current study clearly demonstrates that mEH is crucial for DMBA-induced splenic toxicity in vivo. mEH null mice were protected from both DMBA-induced humoral and cell-mediated immunosuppression. Previously, we reported that CYP1B1 null mice were protected from DMBA-induced splenic immunotoxicity as well. CYP1B1 oxidizes DMBA to an unstable intermediate, DMBA-3,4-epoxide, which is then hydrolyzed by mEH to produce DMBA-3,4-dihydrodiol (Fig. 1). Thus, there are the two key steps in DMBA metabolic activation involving CYP1B1 and mEH enzymes.
Recent investigations by Miyata (1999) and his coworkers reported the detection of DMBA-3,4-dihydrodiol in WT spleen microsomes, but not in mEH null mice. Moreover, in vitro studies using mouse embryo fibroblasts (MEFs) revealed that DMBA-3,4-dihydrodiol can produce the same cytotoxic effects in both WT MEFs and mEH null MEFs (Miyata et al., 2002). It was noted that, mEH null MEFs required a 10-fold higher concentration of DMBA to produce toxicity. Although the major metabolic activation pathway for DMBA is considered to be the formation of diol-epoxides by CYP1B1 and mEH, other metabolic pathways may also be involved in bioactivation of DMBA at high concentrations (Shimada, 2006).
In the present study, mEH null mice were used to understand the role of mEH in DMBA-induced immunosuppression. After observing that mEH null mice were resistant to DMBA-induced immunotoxicity in the spleen, we questioned as to whether there was altered distribution and exposure to DMBA. Previous studies in CYP1A1 null mice have shown an altered distribution of 2,3,7,8-tetrachloro-dibenzo[p]dioxin and benzo[a]pyrene (BaP) (Diliberto et al., 1999; Uno et al. 2006). We therefore performed a direct side-by-side comparison of oral DMBA biodistribution between C57BL/6N WT and mEH null mice. We examined multiple organs and tissues and did not find significant differences in the levels of DMBA at two time points (24 h or 7 days), and report here the results for the spleen, thymus, and liver. We previously examined the in vivo biodistribution and covalent binding of DMBA in mice following a single oral ingestion after 24 h in mice given [3H]-DMBA (Archuleta et al., 1992). We found that peak absorption in most of organs occurred at 6–12 h after ingestion of DMBA. After 24 h, the recovery rate of DMBA was 1.2% and 0.22% in liver and spleen, respectively. Most of the DMBA was excreted from body via the feces and urine (70% and 20%, respectively). In the present study, recovery of free DMBA was 2% in the liver and 0.3% in the spleen after 24 h of oral gavage. Based on the analysis of multiple organs and tissues, we conclude that the distribution of DMBA is identical in WT and mEH null mice.
In previous studies, we observed that BaP produced suppression of the PFC response in C57BL/6N WT mice. In CYP1B1 null mice, we found that these mice were more protected against DMBA immunosuppression than BaP immunosuppression, indicating that BaP may be activated by other mechanisms, such as CYP1A1 (Galvan et al., 2005; Uno et al., 2006). In parallel studies performed in mEH null mice, we found that mEH null mice were protected against both DMBA- and BaP-induced splenic immunotoxicity (in preparation). Many studies have established that BaP can be metabolized by multi-CYP1 enzymes, such as CYP1A1, 1A2, and 1B1 (Nebert and Dalton, 2006). In the absence of CYP1B1 expression, BaP can still induce 1A1 and 1A2 expression in the liver and other organs by AhR-mediated signaling pathways (Nebert et al., 2004). The induced 1A1 and 1A2 can facilitate BaP metabolism (Roos et al., 2004). Both CYP1A1 and CYP1B1 require mEH for formation of the toxic diol-epoxide metabolite. Therefore, it appears that mEH is a pivotal enzyme for both DMBA and BaP bioactivation and it is not surprising that mEH null mice are protected against both DMBA and BaP immunotoxicity.
In the present studies, we observed that T-cell proliferation examined ex vivo was suppressed by 50 mg/kg and 150 mg/kg DMBA treatments of mEH null mice. Given this effect on T-cell proliferation, it is unclear why other immune function assays that utilize T cells (i.e., PFC assay) were not affected by DMBA. It is possible that T cells are more sensitive to DMBA metabolites, or that there are alternate pathways associated with immunotoxicity in these cells. In the thymus, our pharmacokinetic data showed that the parental DMBA compound is detectable, at similar levels in the WT and mEH null mouse thymus after 24 h and 7 days following oral gavage. Therefore, it is unlikely that the explanation for T-cell effects in the spleen relate to the thymic dose and exposure level.
In conclusion, this study clearly demonstrated that mEH is required for DMBA-induced immunosuppression examined ex vivo in spleen cells. It appears that the DMBA-DE metabolite of DMBA is responsible for splenic immunotoxicity. Therefore, future studies will focus on examining potential mechanisms whereby DMBA-DE produces immunotoxicity.
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ACKNOWLEDGMENTS |
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We thank Dr Jerry L. Born for his assistance with the development of the HPLC assay and for other consultations. This work was supported by NIH RO1-05495 and the New Mexico Center for Environmental Health Sciences P30-012072.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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