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Endocrinology Vol. 148, No. 12 6134-6144
Copyright © 2007 by The Endocrine Society

Cellular Defense Mechanisms against Benzo[a]pyrene in Testicular Leydig Cells: Implications of p53, Aryl-Hydrocarbon Receptor, and Cytochrome P450 1A1 Status

Jin-Yong Chung, Ji Young Kim, Yoon-Jae Kim, Sun Ju Jung, Ji-Eun Park, Seung Gee Lee, Jin Tac Kim, Seunghoon Oh, Chang Joo Lee, Yong-Dal Yoon, Young Hyun Yoo and Jong-Min Kim

Department of Anatomy and Cell Biology (J.-Y.C., J.Y.K., Y.-J.K., S.J.J., J.-E.P., S.G.L., J.T.K., Y.H.Y., J.-M.K.), College of Medicine, Dong-A University, Busan 602-714, Korea; Department of Physiology (S.O.), College of Medicine, Dankook University, Cheonan 330-714, Korea; and Department of Life Science (C.J.L., Y.-D.Y.), College of Natural Sciences, Hanyang University, Seoul 133-791, Korea

Address all correspondence and requests for reprints to: Jong-Min Kim, Department of Anatomy and Cell Biology, College of Medicine, Dong-A University, Dongdaeshin-dong 3-1, Seo-gu, Busan 602-714, Korea. E-mail: jmkim7{at}dau.ac.kr.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Leydig cells of the mammalian testis produce testosterone and support spermatogenesis, and thereby their role in male function is fundamental. Although benzo[a]pyrene (B[a]P) has been known to exhibit carcinogenic, apoptogenic, and endocrine-disrupting activities, its potential signaling system in Leydig cells remains to be discovered. In the present study, using the TM3 Leydig cell line and primary Leydig cells, we showed that Leydig cells do not die by exposure to B[a]P and found that an increased level of X chromosome-linked inhibitor of apoptosis protein may be associated with the antiapoptotic process. The Leydig cells were shown to express p53, but its translational level was extremely low. Although a high level of p53 protein was not necessary for apoptosis induced by B[a]P-7,8-diol-9,10-epoxide (a final B[a]P metabolite) in Leydig cells, the apoptosis of primary Leydig cells appears to be p53 independent. This indicates the lack of p53 function in primary Leydig cells. Furthermore, Leydig cells were found to retain insignificant levels of endogenous aryl-hydrocarbon receptor and AhR nuclear transporter proteins in nature. Exposure to B[a]P did not result in a significant increase in aryl-hydrocarbon receptor proteins that are required for CYP1A1 transcription. CYP1A1 expression was present in Leydig cells but at levels insufficient to exhibit its activity. Finally, we have demonstrated that overexpression of CYP1A1 in Leydig cells sensitizes the cells to exhibit its activity in the presence of B[a]P and, thus, induction of apoptosis. Together, these results indicate that the deficiency of CYP1A1 activity might be a decisive condition rendering Leydig cells secure from exogenous polycyclic aromatic hydrocarbons such as B[a]P.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
LEYDIG CELLS, THE testosterone-producing cells of the mammalian testis, support spermatogenesis in seminiferous tubules (1). Although previous in vivo studies have shown that the testis is an important target organ for drugs and chemicals (2, 3), alternatively, in vitro models of Leydig cell culture systems have also been implicated in testicular toxicology (4, 5). Although direct cytotoxic effects of 2,3,7,8-tetrachlorodibenzo-p-dioxin and 7,12-dimethylbenz[a]anthracene on Leydig cells have been tested (6), the precise cellular and molecular mechanism(s) associated with benzo[a]pyrene (B[a]P) cytotoxicity and endocrine-disrupting capability in Leydig cells remain to be discovered.

B[a]P is a member of the polycyclic aromatic hydrocarbons (PAHs), compounds that are usually generated through the combustion of fossil fuels, wood, and other organic materials and are found in significant amounts in diesel exhaust, cigarette smoke, charcoal-broiled foods, and industrial waste by-products (7). B[a]P binds to and activates a ligand-dependent transcription factor termed aryl-hydrocarbon receptor (AhR) (8). Ligand-bound AhR translocates to the nucleus and associates with an AhR-related protein termed AhR nuclear transporter (ARNT). Ligand-activated AhR-ARNT complexes can interact with specific promoter elements (termed dioxin-response elements or xenobiotic-response elements), which induce the transcription of a number of genes involved in the metabolism of B[a]P, including cytochromes P450 (CYP) CYP1A1 and CYP1A2 (8, 9).

Several P450 enzymes are involved in key steps in the oxidation of B[a]P, and CYP1A1 has been demonstrated to be the most active in this oxidation in mammals (10). CYP1A1 activates B[a]P to B[a]P-7,8-oxide, which through hydration by epoxide hydrolase is metabolized to (+/–)-B[a]P-trans-7,8-dihydrodiol (DHD). B[a]P-7,8-DHD may then serve as a substrate for a second CYP-dependent oxidation reaction, generating the ultimate carcinogenic bay region metabolite r-7,t-8-dihydrodiol-t-9,10-oxy-7,8,9,10-tetrahydrobenzo[a]pyrene (BPDE-I). In the nucleus, the diol-epoxides may covalently bind to DNA, mainly forming deoxyguanoside-DNA adducts (11), which may result in misreplication and mutagenesis (12).

In addition to carcinogenic and mutagenic effects, B[a]P and other PAH compounds have been shown to induce apoptosis in vitro in Hepa1c1c7 hepatoma cells (13), Daudi human B cells (14), and A20.1 murine B cells (15). There are two major pathways of apoptosis: the death receptor pathway (16) and the mitochondrial pathway (17). In both of these pathways, caspase-3 especially plays a central role in the execution process of the cells (16, 17). Caspase-3 activation can be suppressed by up-regulation of X chromosome-linked inhibitor of apoptosis protein (XIAP) in many cell types (18).

Although the cell signaling mechanism(s) by which B[a]P induces apoptosis has been well demonstrated particularly in Hepa1c1c7 cells (13, 19), the precise biochemical and cellular mechanisms involved in cytoprotection and antiapoptosis after B[a]P exposure are relatively unknown. We hypothesized herein that the cytoprotective and antiapoptotic characteristics of Leydig cells against B[a]P might be primarily due to the insufficiency of CYP1A1 expression and activity. To test this, we examined the expression and activation of CYP1A1 in relationship to the cellular fate of Leydig cells.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Reagents and antibodies
B[a]P, {alpha}-naphthoflavone, paraformaldehyde, pififthrin-{alpha}, propidium iodide; (PI) 3-[4,5-dimethylthiazol-2-yl]-2.5-diphenyltetrazolium bromide; thiazolyl blue (MTT), anti-actin, and CYP1A1 isozyme were from Sigma Chemical Co. (St. Louis, MO). DHD and B[a]P-7,8-diol-9,10-epoxide (BPDE) were from Midwest Research Institute in National Cancer Institute Repository Dihexaoxacarbocyanine (Kansas City, MO). RNase was from QIAGEN (Valencia, CA). Dihexaoxacarbocyanine (6) was from Molecular Probes (Eugene, OR). N-Acetyl-L-aspartyl-L-glutamyl-L-valyl-L-aspartic acid amide (Ac-DEVD-AMC) and N-acetyl-L-aspartyl-L-glutamyl-L-valyl-L-aldehyde (Ac-DEVD-CHO) were from Calbiochem (La Jolla, CA). Anti-phospho-p53 (serine 15) and VDAC were from Oncogene (La Jolla, CA). Anti-AhR, ARNT, caspase-3, cytochrome c, CYP1A1, and p53 were from Santa Cruz Biotechnology (Santa Cruz, CA). Anti-cleaved caspase-3 was from Cell Signaling Technology (Beverly, MA). Anti-XIAP was from Trevigen (Gaithersburg, MD). Antirabbit and mouse Ig conjugated with horseradish peroxidase were obtained from Amersham Pharmacia Biotech (Piscataway, NJ).

Cell culture
TM3 (normal mouse Leydig cell line; American Type Culture Collection, Rockville, MD) cells were cultured in DMEM (Sigma) supplemented with 5% horse serum and 2.5% fetal bovine serum. Hepa1c1c7 (mouse hepatoma cell line; American Type Culture Collection) cells were cultured in MEM supplemented with 10% fetal bovine serum. The cells were incubated in a humidified incubator at 37 C with 5% CO2 and were exposed to B[a]P or other reagents when confluency reached 50%. For overexpression experiments, the cells were transiently transfected with CYP1A1 expression vector or empty vector (pEGFP-C2) for 12 h and subsequently exposed to B[a]P.

Leydig cell isolation and culture
Adult male Sprague Dawley rats (12 wk old) were purchased from Samtako Bio-Korea (Osan, Korea). The rats were housed in a climate-controlled (21 ± 2 C) animal room in a constant 12-h light, 12-h dark cycle, with free access to rat chow (Samtako) and water. Rats were killed by carbon dioxide asphyxiation, and testes were removed. All procedures were performed in accordance with protocols approved by the Dong-A University Animal Care and Use Committee. Decapsulated testes were incubated with dissociation buffer (M199 buffered with 8.5 mM sodium bicarbonate and 9.3 mM HEPES containing 0.1% BSA and 25 mg/liter soybean trypsin inhibitor, pH 7.4) containing 0.25 mg/ml collagenase at 34 C in a shaking water bath for 15 min. After dissociation, the seminiferous tubules were removed by filtration through 100-µm nylon mesh. The filtrate was centrifuged (250 x g for 10 min), the pellet was resuspended in buffered Hanks’ balanced salt solution, and the suspension was mixed with isoosmotic Percoll in Hanks’ balanced salt solution. After centrifugation (20,000 x g for 60 min at 4 C), fractions of 1.068 g/ml and heavier were collected and washed with dissociation buffer. Leydig cell purity was assessed by cytochemical staining for 3ß-hydroxysteroid dehydrogenase. The cell purity consistently was about 90%. For Leydig cell culture, isolated Leydig cells were resuspended in DMEM/F12 culture medium containing 15 mM HEPES, 10% fetal calf serum, 5 µg/ml gentamycin, 50 U/ml penicillin, and 50 µg/ml streptomycin. Cells were seeded on six-well culture plates (1.2 x 106 cells per well). To assess the effects of B[a]P, cells were plated for 12 h at 34 C in a humidified 5% CO2/95% air atmosphere.

Construction of CYP1A1 expression vector
Mouse cDNA was prepared from Hepa1c1c7 cells. The total RNA was prepared from Hepa1c1c7 cells using TRIzol reagent (Invitrogen, Carlsbad, CA). The cDNA was synthesized from 5 µg total RNA using oligo dT random primer (Promega, Madison, WI) and Moloney murine leukemia virus (MMLV) RNase H-reverse transcriptase (Promega). To clone full-length cDNA of mouse CYP1A1 gene, the cDNA products were subjected to PCR using specific primers for mouse CYP1A1 (M10021, GenBank). The following primers were used: forward, 5'-ATG CCT TCC ATG TAT GGA CC-3', and reverse, 5'-CTA AGC CTG AAG ATG CTG AGG A-3'. PCR amplification cycles were as follows: 94 C for 1 min, 30 sec at 94 C, 1 min at 59 C, and 2 min at 72 C for 30 cycles, and 72 C for 5 min. The PCR products were then subcloned into pGEM-T easy vector (Promega). The pGEM-T easy/mCYP1A1 and pEGFP-C2 vector DNA were digested with EcoR1, and mCYP1A1 gene encoding insert was cloned into pEGFP-C2 vector (BD Bioscience, San Jose, CA) at the EcoR1 site. The construct was confirmed by sequencing.

MTT cell viability assay
Cells were seeded in 12-well plates at a density of 5 x 105 cells per well. After treatment at an appropriate time, the culture medium was removed and replaced with a medium containing 0.5 mg MTT dissolved in PBS (pH, 7.2). After 4 h, the formed crystals were dissolved with 200 µl dimethylsulfoxide. The intensity of the color in each well was measured at a wavelength of 490 nm using a microplate reader (Bio-Tek EL-312e; Bio-Tek Instruments, Winooski, VT).

Cell cycle analysis
Cells were harvested, fixed with 95% ethanol for 24 h, incubated with 0.05 mg/ml PI and 1 µg/ml RNase A at 37 C for 30 min, and analyzed by flow cytometry using Epics XL with an analysis software (EXPO32; Beckman Coulter, Southfield, MI). The cells belonging to the sub-G1 population were considered apoptotic cells, and the percentage of each phase of the cell cycle was determined.

Annexin-V staining
The cells were stained using the annexin-V-FITC apoptosis detection kit (BD Biosciences) according to the manufacturer’s protocol. Stained cells were analyzed by flow cytometry.

Western blot analysis
Whole-cell lysates were prepared by incubating cell pellets in lysis buffer [30 mM NaCl, 0.5% Triton X-100, 50 mM Tris-HCl (pH 7.4), 1 mM Na3VO4, 25 mM NaF, 10 mM Na4P2O7] for 30 min on ice. After the insoluble fractions were removed by centrifugation at 14,000 rpm at 4 C for 30 min, the supernatants were collected and protein concentration was determined with a BCA protein assay kit (Pierce Biotechnology, Woburn, MA). The same amounts of proteins (~30 µg) were subjected to SDS-PAGE and transferred onto a nitrocellulose membrane. The membranes were incubated for 1 h at room temperature with a primary antibody in Tris-buffered saline containing 0.05% Tween 20 (pH 7.4) in the presence of 5% nonfat dry milk. After the membranes were washed in Tris-buffered saline containing 0.05% Tween 20, secondary antibody reactions were performed with an appropriate source of antibody conjugated with horseradish peroxidase. The signals were detected with an enhanced chemiluminescence detection kit (Amersham Pharmacia Biotech) in the LAS-3000 detector (Fujifilm, Tokyo, Japan). Immunoblotting for ß-actin was performed in every experiment as an internal control.

RT-PCR
Total cellular RNA was isolated from cultured cells using a Trizol reagent (Invitrogen). The cDNA was synthesized from 5 µg total RNA using oligo dT random primer (Promega) and MMLV RNase H-reverse transcriptase (Promega). cDNA (3 µl) was subjected to PCR in a 30-µl reaction mixture [10x PCR buffer, 2.5 mM dNTP, 5 U Taq polymerase (Promega), and upstream and downstream primers]. The following primers were used: AhR (20) forward, 5'-CGC TGA AAC ATG AGC AAA TTG G-3', and reverse, 5'-ACA GCT TAG GTG CTG AGT CAC AGG-3'; ARNT (20) forward, 5'-GAT GCG ATG ACC AGA TGT G-3', and reverse 5'-CGA TGA GGA AAG ATG GCT TGT AGG-3'; CYP1A1 (20) forward, 5'-CCT CTT TGG AGC TGG GTT TG-3', and reverse, 5'-TGC TGT GGG GGA TGG TGA AG-3'; and p53 (21) forward, 5'-ATA TGA GCA TCG AGC TCC CTC T-3', and reverse, 5'-CAC AAC TGC ACA GGG CAT GT-3'. Thermal cycling conditions were 94 C for 1 min, 15 sec at 94 C, 1 min at 56 C, and 30 sec at 72 C for 35 cycles, and 72 C for 5 min. The PCR products was analyzed by 2% agarose gel electrophoresis and visualized by ethidium bromide staining under UV illumination.

Immunocytochemistry and immunohistochemistry
Cells were harvested and then attached on the slide glass by cytospin centrifugation. Cells were fixed with 4% paraformaldehyde, washed with PBS, and incubated with 0.2% Triton X-100. Then, cells were incubated with the appropriate primary antibody in 1% BSA at room temperature. For secondary antibody reaction, cells were incubated with an appropriate fluorescence-conjugated secondary antibody at room temperature. For counterstaining of the nucleus, if required, cells were incubated with PI (50 µg/ml) at room temperature. Finally, cells were mounted and observed under a confocal microscope (LSM510; Carl Zeiss, Oberkochen, Germany). For p53 immunohistochemistry, deparaffinized and hydrated testis sections were treated in 3% H2O2 for 5 min and rinsed with PBS for 15 min, and subsequently the method in the instruction manual of the Vectastain ABC kit (Vector Laboratories, Burlingame, CA) was followed.

Isolation of microsomes
The cells (6 x 106) were resuspended in isotonic homogenization buffer [10 mM HEPES, 1 mM EDTA, 2 mM dithiothreitol (DTT)], spun down at 17,900 x g for 5 min, and swelled in isotonic homogenization buffer for 5 min at 4 C. After 30 strokes in a Dounce homogenizer, the lysates were centrifuged for 30 min at 7800 x g, and the supernatants were again centrifuged for 60 min at 17,900 x g. The microsomal fraction was obtained from the pellet.

Preparation of mitochondrial fractions
The cells (5 x 107) were washed in Tris-based, Mg2+/Ca2+-deficient (TD) buffer (135 mM NaCl, 5 mM KCl, 25 mM Tris-Cl, pH 7.6) and allowed to swell for 15 min in ice-cold hypotonic Ca2+ reticulocyte standard buffer (CaRSB) buffer [10 mM NaCl, 1.5 mM CaCl2, 10 mM Tris-Cl (pH 7.5), and protease inhibitors]. Cells were Dounce homogenized with 30 strokes, and mitochondria stabilization buffer (210 mM mannitol, 70 mM sucrose, 5 mM EDTA, 5 mM Tris, pH 7.6) was added. After removing nuclear contaminants (690 x g for 15 min), the supernatant was centrifuged at 20,800 x g for 15 min. Finally, the pellet (mitochondria) was directly diluted with lysis buffer [5 M NaCl, 1 M Tris-Cl (pH 7.6), 5% Triton X-100, and protease inhibitors] for protein analysis.

CYP1A1 enzyme activity assays
Cellular proteins were isolated with lysis buffer consisting of 30 mM NaCl, 50 mM Tris-HCl (pH 7.6), 5% Triton X-100, and 100 mM phenylmethylsulfonyl fluoride. The enzymatic activities of CYP1A1 was measured by P450-Glo assay kits (Promega) as per manufacturer’s instruction manual. Briefly, isolated proteins (30 µg) were mixed with the 4x cytochrome P450/KPO4/substrate reaction mixture (CYP1A1; 0.5 pM CYP1A1 isozyme, 400 mM KPO4,120 µM leuciferin-chloroethyl ether), and 2x NADPH regeneration mixture (2.6 mM NADP+, 6.6 mM glucose-6-phosphate, 0.4 U/ml glucose-6-phosphate dehydrogenase, 6.6 mM MgCl2). The sample and the 4x cytochrome P450/KPO4/substrate reaction mixture were added to a 96-well plate. After preincubating the plate at 37 C for 10 min, the 2x NADPH regeneration mixture was added to each reaction. The plate was incubated at 37 C for 30 min, and the reconstituted leuciferin detection reagent was added. Again, the plate at room temperature was incubated for 20 min, and the luminescence was recorded using a luminometer (Type392; Amersham Bioscience, Piscataway, NJ).

Caspase-3 activity assay
The collected cells were dissolved in lysis buffer (30 mM HEPES, 1 mM DTT, 1 mM EDTA, 0.1% CHAPS, pH 7.4). Caspase-3 activity was measured by the caspase-3 cellular assay kit (Calbiochem) as per manufacturer’s instruction manual. Briefly, 30 µg protein was incubated with a fluorogenic caspase-3 substrate (Ac-DEVD-AMC) in assay buffer (100 mM NaCl, 50 mM HEPES, 10 mM DTT, 1 mM EDTA, 0.1% CHAPS, 10% glycerol, pH 7.4) at 37 C. If required, 0.5 µM caspase-3-specific inhibitor (Ac-DEVD-CHO) was added to the reaction. The fluorescence was monitored by a fluorometer (Packard Bioscience, Boston, MA) with excitation at 360 nm and emission at 530 nm every 30 min.

Statistics
Data were expressed as the mean ± SD of three or four separate experiments. Where appropriate, data were subjected to ANOVA followed by Duncan’s post hoc test. Means were considered significantly different at P < 0.05.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
B[a]P exposure does not provoke cell death in TM3 Leydig cells
To evaluate the effect of B[a]P on cell viability in Leydig cells, TM3 Leydig cells were exposed to various concentrations of B[a]P (0.1–100 µM). With any concentration, cells were maintained above 50% viability, although the cell viability was gradually reduced with increasing concentrations of B[a]P (Fig. 1AGo). Nevertheless, cell death determined by sub-G1 analysis of the cell cycle showed that B[a]P treatment did not induce significant cell death with any concentration (Fig. 1BGoa), but extended duration of treatment up to 72 h showed significant cell death numerically (Fig. 1BGob). Although we recognized that TM3 cells are not susceptible to cell death significantly by exposure to B[a]P, pharmacological concentrations of B[a]P appear to cause the reduction of Leydig cell viability (measured by MTT assay) by decreasing several cellular metabolisms cytotoxically. Therefore, we chose 10 µM B[a]P as an adequate concentration for the rest of the study.


Figure 1
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  		FIG. 1. Effects of B[a]P on cell viability and death and changes of caspase-3 and XIAP in TM3 Leydig cells. A, Cell viability shown in cells treated with various concentrations of B[a]P (~0.1–100 µM) for 48 h, determined by MTT assay. At least three independent experiments were performed, and data shown are the mean ± SD. *, P < 0.05 compared with 0 µM control. B, Cell death after treatment with increasing concentrations (~0.1–100 µM) of B[a]P for 48 h (a) or at different time points with 10 µM B[a]P treatment (b). Cell death was assessed by flow cytometry. The percentage of cells with a sub-G1 DNA content was taken as a measure of cell death. At least three independent experiments were performed, and data shown are the mean ± SD. * and #, P < 0.01 and P < 0.001, respectively. C, Western blot analyses of procaspase-3, cleaved (activated) caspase-3, and XIAP after B[a]P (10 µM) treatment for the indicated periods of time up to 72 h. Equal amounts of protein (30 µg) were separated by SDS-PAGE and immunoblotted using the indicated antibodies. Actin expression was examined as a loading control. D, Representative confocal images of procaspase-3 and XIAP. Cells were treated for 48 h with 10 µM B[a]P, cytospun, fixed, and immunostained with antibodies as indicated. a, Control, procaspase-3; b, B[a]P-treated, procaspase-3; c, control, XIAP; d, B[a]P-treated, XIAP. Original magnification, x800.

 
The activation of caspase-3 and up-regulation of XIAP have been known to be involved with the induction and suppression of apoptosis, respectively (18). After B[a]P treatment, procaspase-3 and XIAP expression increased, but cleaved (active) forms of caspase-3 were not detected (Fig. 1CGo). Consistent with Western blot results, increased levels of procaspase-3 and XIAP were prominently seen in the cytoplasm of cells treated with B[a]P (Fig. 1DGo). These results confirm that apoptosis does not occur after B[a]P exposure and imply that the up-regulation of XIAP might be related to the suppressed activation (cleavage) of increased procaspase-3 that is induced by the cytotoxic effect of B[a]P.

B[a]P alters AhR, ARNT, and CYP1A1 expression in TM3 cells
It has been well demonstrated that AhR and ARNT are required for promoting CYP1A1 expression in the nucleus (8, 9). To determine whether they are regulated in TM3 cells by B[a]P, the expression of AhR and ARNT has been investigated by immunocytochemistry, Western blot and RT-PCR analyses. Figure 2AGo shows immunocytochemical detection of AhR and ARNT in TM3 cells after B[a]P treatment. Compared with the control cells (Fig. 2AGoa), more intense staining for AhR was seen in the peripheral regions of nuclei in B[a]P-treated cells (Fig. 2AGob). However, ARNT immunostaining was not noticeably changed after B[a]P exposure (Fig. 2AGo, c and d). Western blot (Fig. 2BGoa) and RT-PCR (Fig. 2BGob) analyses showed that AhR expression is up-regulated by B[a]P treatment, but the expression of ARNT appears to be up-regulated only at the transcriptional level.


Figure 2
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  		FIG. 2. Changes in AhR, ARNT, and CYP1A1 expression and CYP1A1 activity in TM3 cells after B[a]P exposure. A, Immunolocalization of AhR and ARNT. Cells were treated for 48 h with 10 µM B[a]P, cytospun, fixed, and immunostained with antibodies as indicated. The images were taken by a confocal microscope. a, Control, AhR; b, B[a]P-treated, AhR; c, control, ARNT; d, B[a]P-treated, ARNT. Original magnification, x800. B, Alterations in AhR and ARNT protein (a) and mRNA (b) expression detected by Western blotting and RT-PCR, respectively. For Western blot analysis, equal amounts of nuclear protein (30 µg) were separated by SDS-PAGE and immunoblotted using the indicated antibodies. Actin is indicated as a loading control. For RT-PCR analysis, cDNA synthesized from total RNA (5 µg) by MMLV RNase H-reverse transcriptase were amplified with the indicated primers for 30 cycles, and the PCR products were resolved by agarose gel electrophoresis. Actin expression was examined as a loading control. C, Immunocytochemical localization of CYP1A1. Cells were treated for 48 h with 10 µM B[a]P, cytospun, fixed, and immunostained with the CYP1A1 antibody. The images were taken by a confocal microscope. a, Control; b, B[a]P-treated. Original magnification, x800. D, Changes in CYP1A1 protein (a) and mRNA (b) expression detected by Western blotting and RT-PCR, respectively. For Western blot analysis, equal amounts (30 µg) of proteins from the total cell lysates (T) or microsomal lysates (M) were separated by SDS-PAGE and immunoblotted using the CYP1A1 antibody. Actin is indicated as a loading control for the total cell lysates. For RT-PCR analysis, cDNA synthesized from total RNAs (5 µg) by MMLV RNase H- reverse transcriptase were amplified with the CYP1A1 primer for 30 cycles, and the PCR products were resolved by agarose gel electrophoresis. Actin expression was examined as a loading control. E, Comparison of CYP1A1 activity between TM3 cells and Hepa1c1c7 cells after B[a]P exposure for 48 h. CYP1A1 activity was measured spectrofluorometrically using a P450-Glo assay kit (Promega), as described in Materials and Methods. At least three independent experiments were performed, and data shown are the mean ± SD. *, P < 0.05 compared with control.

 
Next, CYP1A1 expression and activity were investigated to examine whether TM3 cells have the potential to convert B[a]P to its metabolites (i.e. DHD or BPDE). Immunoreactivity for CYP1A1 was slightly increased in the cytoplasm after B[a]P treatment (Fig. 2CGo). Western blot analysis revealed that the CYP1A1 proteins were actually increased after B[a]P treatment, but they are detectable only when the isolated microsomal proteins were applied (Fig. 2DGoa). Consistent with the proteins, the exposure to B[a]P also temporally increased the gene transcripts for CYP1A1 (Fig. 2DGob). Nevertheless, CYP1A1 activity in TM3 cells after B[a]P exposure remained basal and was unchanged compared with that in Hepa1c1c7 cells, a cell line used as a positive control for B[a]P action (Fig. 2EGo) (13). These results concluded that the increased expression of CYP1A1 in response to B[a]P exposure might be insufficient to exert its activity in TM3 cells.

BPDE induces apoptosis in TM3 cells, but DHD did not
To examine whether B[a]P metabolites can induce cell death, TM3 cells have been exposed to DHD and BPDE. Like B[a]P, DHD treatment did not cause cell death at any concentration (0–10 µM); however, BPDE significantly induced cell death at concentrations above 1 µM (Fig. 3AGo). BPDE-treated cell death was evoked in apoptotic fashion as seen in caspase-3 activation (Fig. 3BGo). This feature was also confirmed by the caspase-3 activity assay (Fig. 3CGo) and immunostaining of caspase-3 using an active-form-specific antibody for caspase-3 (Fig. 3DGo).


Figure 3
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  		FIG. 3. Effects of B[a]P metabolites on cell death in TM3 Leydig cells. A, Cell death after treatment with various concentrations of DHD and BPDE (~0.1–10 µM) for 48 h. Cell death was assessed by flow cytometry. The percentage of cells with a sub-G1 DNA content was taken as a measure of cell death. At least three independent experiments were performed, and data shown are the mean ± SD. *, P < 0.001 compared with control. B, Western blot analysis of cleaved (activated) caspase-3 after B[a]P (10 µM), DHD (1 µM), and BPDE (1 µM) treatments for 48 h. Equal amounts of protein (30 µg) were separated by SDS-PAGE and immunoblotted using a cleaved-form-specific antibody for caspase-3. Actin expression was examined as a loading control (Con). C, Caspase-3 activity shown in the cells treated with B[a]P (10 µM), DHD (1 µM), and BPDE (1 µM) for 48 h. The caspase-3 activity was measured spectrofluorometrically using a caspase-3-specific substrate (Ac-DEVD-AMC), as described in Materials and Methods. DEVD-CHO, caspase-3 inhibitor. D, Representative confocal images of the activated caspase-3. Cells were treated with B[a]P (10 µM), DHD (1 µM), and BPDE (1 µM) for 48 h. The cells were then cytospun, fixed, and immunostained with an active-form-specific antibody for caspase-3. a, Control; b, B[a]P-treated; c, DHD-treated; d, BPDE-treated. Original magnification, x800.

 
The changes in mitochondrial events are important for induction of apoptosis by the intrinsic pathway (17). In this pathway, cytochrome c release occurs accompanied by a decrease in mitochondrial membrane potential (22). In this study, B[a]P did not cause cytochrome c release from the mitochondria in TM3 cells during the treatment, but it did in Hepa1c1c7 cells (supplemental Fig. S1A, published as supplemental data on The Endocrine Society’s Journals Online web site at http://endo.endojournals.org). However, BPDE exposure showed a cytochrome c release in TM3 cells (supplemental Fig. S1Bd), which appeared similar to patterns observed in B[a]P-treated Hepa1c1c7 cells (supplemental Fig. S1Bf). These results indicate that TM3 cells retain an apoptotic mechanism, but the insufficiency of CYP1A1 activity may protect TM3 Leydig cells from apoptosis by B[a]P exposure.

Sufficient levels of AhR and CYP1A1 are a prerequisite for B[a]P-induced apoptosis
Hepa1c1c7 cells, a mouse hepatoma cell line, are well known to undergo apoptosis in response to B[a]P exposure (13, 19). For this reason, we chose these cells as a positive control for studying the action of B[a]P on TM3 cells. As expected, B[a]P treatment for 48 h evoked significant cell death (~20%) in Hepa1c1c7 cells in an apoptotic manner via caspase-3 activation (supplemental Fig. S2A). Supplemental Fig. S2A shows the relative changes in AhR and CYP1A1 protein levels between TM3 and Hepa1c1c7 cells after B[a]P treatment. For this comparison, the same amount of proteins from TM3 and Hepa1c1c7 cells were loaded and analyzed for AhR and CYP1A1 protein content by Western blotting on the same blot. In Hepa1c1c7 cells, AhR proteins were endogenously abundant and tended to decrease after B[a]P treatment (supplemental Fig. S2A). The CYP1A1, however, was remarkably up-regulated at 12 h and decreased gradually afterward (supplemental Fig. S2A). In contrast, both AhR and CYP1A1 levels in TM3 cells were extremely low compared with those in Hepa1c1c7 cells (supplemental Fig. S2A). Immunocytochemical results also verified the endogenous levels of AhR and CYP1A1 proteins in TM3 (supplemental Fig. S2B, a and b) and Hepa1c1c7 cells (supplemental Fig. S2B, b and d), which were seen by Western blotting. These results indicate that TM3 cells might be incapable of expressing an adequate level of CYP1A1 due to the insufficiency of AhR protein expression even in the absence or presence of B[a]P stimulation.

B[a]P exposure results in increased levels of p53 and its phosphorylation
The occurrence of apoptosis is generally accompanied by a change in the specific phase (stage) of the cell cycle (23). Although TM3 cells did not undergo apoptosis by B[a]P exposure, the possible changes in the cell cycle during the treatment were monitored to understand the action of B[a]P. Figure 4AGo shows the cell cycle changes in TM3 and Hepa1c1c7 cells after B[a]P exposure. In both cells, common features after B[a]P exposure were maintenance of the decreased G1 phase and the increased S phase compared with each control. However, when apoptosis occurred in Hepa1c1c7 cells (at 48 and 72 h), prominent S phase arrest and the accompanying decrease in G1 phase were notable. On the contrary, these results also indicate that the increased S phase in TM3 cells after exposure to B[a]P may correlate with decreased proliferation rates without causing cell death. In addition, the comparative Western blot analyses of p53 and phosphorylated p53 at serine 15 showed that B[a]P exposure evoked remarkable increases of these proteins in Hepa1c1c7 cells but they remained much lower in TM3 cells compared with Hepa1c1c7 cells (Fig. 4BGo). Importantly, however, BPDE exposure to TM3 cells resulted in up-regulation of these proteins to almost equivalent levels in B[a]P- and BPDE-treated Hepa1c1c7 cells (Fig. 4BGo). Although both p53 and phosphorylated p53 (at serine 15) levels were relatively lower in TM3 cells than in Hepa1c1c7 cells, we were able to find in the independent Western blotting experiments that these proteins were also up-regulated in TM3 cells by B[a]P exposure (see supplemental Fig. S3). Furthermore, Fig. 4CGo clearly demonstrates that, as in Hepa1c1c7 cells, BPDE-induced cell death in TM3 was significantly reduced by pretreatment with pifithrin-{alpha}, an inhibitor of p53 activation. These results make the assumption that the TM3 cells are capable of undergoing apoptosis by B[a]P if it can be metabolized to BPDE in the cells by CYP1A1 enzyme.


Figure 4
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  		FIG. 4. Cell cycle and p53 protein in TM3 after B[a]P exposure. A, Comparison of the changes in each phase of the cell cycle between TM3 and Hepa1c1c7 cells. The cells were treated with B[a]P (10 µM) for the indicated periods of time up to 72 h, fixed, and stained with PI. The cell cycle was determined by flow cytometry. The data are expressed as mean ± SD of three independent experiments. * and **, P < 0.01 and P < 0.001, respectively (compared with each time control); # and ##, P < 0.05 and P < 0.001, respectively. B, Evaluation of the changes in p53 and phosphorylated p53 (at serine 15) [P-p53(S15)] protein levels between TM3 and Hepa1c1c7 cells. The cells were treated with B[a]P (10 µM) or BPDE (1 µM) for the indicated periods of time up to 72 h. Equal amounts of protein (30 µg) were separated by SDS-PAGE and immunoblotted using the indicated antibodies. Actin expression was examined as a loading control. C, Involvement of p53 function in the cell death of TM3 cells. The TM3 and Hepa1c1c7 cells were treated with BPDE (1 µM) in the absence or presence of pifithrin-{alpha} (PFT-{alpha}, p53 inhibitor; 5 µM) for 48 h. Cell death was assessed by flow cytometry. The percentage of cells with a sub-G1 DNA content was taken as a measure of cell death. At least three independent experiments were performed, and data shown are the mean ± SD. *, P < 0.05 compared with 0-h control.

 
Ectopic expression of CYP1A1 enables TM3 Leydig cells to undergo apoptosis by B[a]P exposure
From the above experiments, we have recognized that abundant expression of CYP1A1 is required for the induction of apoptosis in TM3 cells by B[a]P. Based on this finding, we decided to force the TM3 cells to express CYP1A1 by introducing the CYP1A1 expression vector constructed in our laboratory for this study. The ectopic overexpression of CYP1A1 gene in TM3 cells notably increased CYP1A1 protein expression (Fig. 5Go, A and B). However, CYP1A1 was activated only when B[a]P was added to the cells (Fig. 5CGo). These results suggest that CYP1A1 expression itself does not always represent its activity. Finally, we have seen that B[a]P-exposed TM3 cells undergo apoptosis in a caspase-3-dependent manner after ectopic CYP1A1 expression (Fig. 5Go, D–H). Regardless of this major finding, the transfection of either an empty (control) vector (pEGFP-C2) or a CYP1A1 expression vector alone provoked cell death (Fig. 5DGo). Indeed, there is a report that clearly shows that the cytotoxic effect of the pEGFP-C2 vector is due to residues encoded by a multiple cloning site (24). However, CYP1A1 transfection alone appeared to induce more cell death than the empty vector alone (Fig. 5Go, D–F). Even considering this effect, it is evident that the TM3 Leydig cells become susceptible to apoptosis by B[a]P exposure via ectopic CYP1A1 expression. Therefore, these results indicate that the lack of CYP1A1 activity might be a central reason why TM3 Leydig cells do not die by apoptosis after B[a]P exposure.


Figure 5
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  		FIG. 5. Effect of overexpression of CYP1A1 on its activity and apoptosis in TM3 Leydig cells. The cells were transiently transfected with CYP1A1 expression vector or empty vector (EV; pEGFP-C2) for 12 h and subsequently exposed to B[a]P (10 µM) for 48 h. A, Overexpression of CYP1A1 observed by immunocytochemistry using a confocal microscope. Original magnification, x800. B, The expression of CYP1A1 proteins detected by Western blotting. Equal amounts of protein (30 µg) were separated by SDS-PAGE and immunoblotted using a CYP1A1 antibody. Actin expression was examined as a loading control. C, Changes in CYP1A1 activity after CYP1A1 overexpression. The activity was measured spectrofluorometrically using a P450-Glo assay kit (Promega), as described in Materials and Methods. At least three independent experiments were performed, and data shown are the mean ± SD. *, P < 0.001 compared with control. D, Cell death by B[a]P exposure after CYP1A1 overexpression. Cell death was assessed by flow cytometry. The percentage of cells with a sub-G1 DNA content was taken as a measure of cell death. *, #, and ##, P < 0.05, P < 0.01, and P < 0.001, respectively. E, Flow cytograms showing cell death assessed by annexin-V-FITC and PI staining. The stained cells were analyzed by flow cytometry. In the flow cytogram, T1 represents necrotic region, T2 late apoptotic region, T3 live region, and T4 early apoptotic region. F, Graph summarizing the results shown in E. G, Western blot analysis of caspase-3 activation by B[a]P exposure after CYP1A1 overexpression. Equal amounts of protein (30 µg) were separated by SDS-PAGE and immunoblotted using a cleaved-form-specific antibody for caspase-3. Actin expression was examined as a loading control. H, Caspase-3 activity after CYP1A1 overexpression in the absence or presence of B[a]P. The activity was measured spectrofluorometrically using a caspase-3-specific substrate (Ac-DEVD-AMC), as described in Materials and Methods. At least three independent experiments were performed, and data shown are the mean ± SD. *, P < 0.001 compared with control.

 
B[a]P does not exhibit an apoptotic effect in primary Leydig cells
Although the TM3 cells originated from mouse normal Leydig cells, it is presumed that the characteristics of immortalized TM3 cells could not be representative of Leydig cells isolated from mammalian testis. Therefore, we have tested whether the primary Leydig cells of rat testis respond to B[a]P similarly to TM3 cells. Identical to TM3 cells, B[a]P exposure did not provoke cell death in the primary Leydig cells, but BPDE did (Fig. 6AGo). During the treatment, as expected, caspase-3 activation was absent, but XIAP content appeared to be slightly up-regulated (Fig. 6Go, B and C). In this study, we found that the primary Leydig cells express a very low level of p53 protein that is not detectable by Western blot analysis (Fig. 6BGo), immunocytochemistry (Fig. 6CGo), and immunohistochemistry on testis section (supplemental Fig. S4). Even BPDE treatment was not able to evoke the up-regulation of p53 in the cells, as determined by Western blotting (Fig. 6DGo). However, RT-PCR analysis revealed the p53 gene expression in Leydig cells (Fig. 6EGo). As for CYP1A1, B[a]P tended to slightly increase the protein content at 12 h, but it was not significant compared with control (Fig. 6Go, B and C). The AhR proteins were simultaneously up-regulated in culture (control), and the expression was suppressed in the presence of B[a]P. Furthermore, CYP1A1 and AhR gene expression in Leydig cells was negligible when compared with their expression in liver and spleen, respectively (Fig. 6EGo). Together, we were able to confirm that primary Leydig cells exhibit results similar to what has been observed in TM3 cells.


Figure 6
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  		FIG. 6. Evaluation of B[a]P cytotoxicity on rat primary Leydig cells. A, Cell death shown in cultured rat Leydig cells at 48 h after exposure to B[a]P (10 µM) or BPDE (1 µM). Cell death was assessed by flow cytometry. The percentage of cells with a sub-G1 DNA content was taken as a measure of cell death. B, Western blot analyses of major cell death-related proteins (XIAP, active caspase-3, and p53) and B[a]P signaling-associated proteins (AhR and CYP1A1) after B[a]P (10 µM) treatment for the indicated periods of time up to 72 h. Equal amounts of protein (30 µg) were separated by SDS-PAGE and immunoblotted using the indicated antibodies. Actin expression was examined as a loading control (Con). C, Representative confocal images of XIAP, p53, and CYP1A1. Cells were treated for 48 h with 10 µM B[a]P, cytospun, fixed, and immunostained with antibodies as indicated. a, Control, XIAP; b, control, p53; c, control, CYP1A1; d, B[a]P-treated, XIAP; e, B[a]P-treated, p53; f, B[a]P-treated, CYP1A1. Original magnification, x800. D, Effect of BPDE on p53 and caspase-3 activation, as analyzed by Western blotting. Equal amounts of protein (30 µg) were separated by SDS-PAGE and immunoblotted using the indicated antibodies. E, Expression of p53, AhR, and CYP1A1 genes in the various rat tissues including the Leydig cells. The mRNA expression was examined by RT-PCR. The cDNA synthesized from total RNA (5 µg) was amplified with the indicated primers for 30 cycles, and the PCR products were resolved by agarose gel electrophoresis. GAPDH expression was examined as a loading control. All the experiments were performed independently at least three times.

 

   Discussion
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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In this study, we have shown that Leydig cells, both TM3 and primary cells isolated from testis, are not susceptible to apoptosis by exposure to B[a]P. Indeed, Leydig cells are fully differentiated, rarely proliferate, and rarely undergo apoptosis (or attrition) in the mammalian adult testis (25). For these reasons, Leydig cells have also been considered as an important model for studying aging (26). Although a few agents such as ethane dimethanesulfonate and glucocorticoid have been known to kill Leydig cells by apoptosis in a caspase-3-dependent manner (27, 28), Leydig cells are presumed to be somewhat resistant to various types of chemotoxicants (29, 30). In response to pituitary LH, Leydig cells produce testosterone that then supports spermatogenesis by acting on the Sertoli cells in the seminiferous tubules (1, 25). Therefore, if Leydig cells are damaged or eradicated by exposure to some genotoxic agent, the male endocrine and reproductive functions can be severely disrupted.

In the present study, we showed that Leydig cells do not die by exposure to B[a]P but do so by exposure to BPDE. This indicates that Leydig cells may not have an enzymatic machinery for converting B[a]P to BPDE. As for the antiapoptotic property of Leydig cells against a cytotoxic B[a]P exposure, the involvement of increased XIAP and procaspase-3 protein contents was noticeable. The inhibitor of apoptosis proteins (IAPs) are a family of intracellular antiapoptotic proteins, first identified in baculovirus, which play a key role in cell survival by modulating death-signaling pathways at a postmitochondrial level. They include XIAP, human IAP-1 (Hiap-1), human IAP-2 (Hiap-2), neuronal apoptosis inhibitory protein (Naip), Survivin, and Livin. Among IAPs, XIAP is a direct inhibitor of caspase-3 and caspase-9 and modulates the Bax/cytochrome c (mitochondrial) pathway by inhibiting caspase-9 (18). Our results demonstrated that XIAP levels were up-regulated in Leydig cells in response to B[a]P. We believe that this would be one of the possible reasons why Leydig cells can survive even if under increased levels of procaspase-3 by exposure to B[a]P. Although the role of XIAP in Leydig cells has not been demonstrated before this study, it is tempting to speculate that XIAP might play an important role in the cellular protection mechanism of Leydig cells against apoptotic insults including genotoxic compounds.

Another important feature found in the current study was that Leydig cells appear to express the p53 gene, but its translational level seems to be extremely low. p53 is a key regulator of apoptosis, in addition to its clear effects on cell cycle arrest and its involvement in senescence, DNA repair, and differentiation (22). The expression of wild-type p53 resulted in a decrease in cell viability with characteristics of apoptosis (31). Indisputable evidence for the role of p53 in cell death was provided by studies of p53 knockout mice (32). Thymocytes from these animals are significantly more resistant to radiation-induced apoptosis than corresponding normal cells. Cellular stresses that induce p53-dependent apoptosis include DNA damage, hypoxia, and oxidative stress. Previously, several studies have demonstrated the direct association of p53 and its phosphorylation in B[a]P-induced apoptosis (19), which was also reproduced with Hepa1c1c7 cells in our study. p53 protein was detectable and up-regulated after B[a]P exposure in TM3 Leydig cells probably due to the proliferative characteristics of the immortalized cell line. However, in the case of primary rat Leydig cells isolated from testis, p53 was rarely detectable by Western blot analysis and immunocytochemical methods even using several different sources of antibodies for p53. Interestingly, our data showed that exposure to BPDE, a final metabolite of B[a]P, provoked apoptosis of both TM3 and primary Leydig cells in a caspase-3-dependent fashion but that primary Leydig cell apoptosis was likely to occur independent of p53. Currently, an alternative apoptotic mechanism for BPDE action in primary Leydig cells remains to be unveiled. Indeed, adult Leydig cells are not proliferative in the interstitial compartment of the testis (25) and generally remained in the G1 phase of the cell cycle (unpublished data). This implies that p53 protein function is not essential for the cell cycle checkpoint in the Leydig cells of the testis, and thereby the Leydig cells could be less sensitive to DNA-damaging agents that are at least p53 dependent.

In addition to the cytoprotection by the presence of XIAP as well as the lack of p53 function, Leydig cells were presumed to be safe from B[a]P-provoked cytotoxicity because they have insufficient levels of endogenous AhR and ARNT proteins. In particular, the lack or insufficiency of cellular AhR proteins may prevent the B[a]P action at the initial stage of the signaling. Although AhR expression was slightly up-regulated in TM3 cells by B[a]P exposure, it was minimal compared with the endogenous levels of AhR in Hepa1c1c7 cells that underwent apoptosis responding to B[a]P. Moreover, the isolated testicular Leydig cells were shown to express a very low level of AhR. These points indicate that Leydig cells could be less endangered from exposure to exogenous cytotoxic ligands for AhR than other cell types. However, during the primary Leydig cell culture, AhR protein expression was aberrantly up-regulated without any treatment, although it was clearly suppressed in the presence of B[a]P. Recently, a report demonstrated a similar finding and suggested that serum factor may contribute to that increase in the cells (33). Nevertheless, the nature of this phenomenon is completely unknown at present. In general, AhR is well known to bind exogenous ligands, such as 2,3,7,8-tetrachlorodibenzo-p-dioxin, B[a]P, and 3-methylcholanthrene, and mediates their biological effects (8). However, the natural ligands for AhR are unknown, and we know little about AhR function in the absence of exogenous ligands. Several studies previously demonstrated that AhR is associated with growth inhibition and cell cycle arrest (34, 35). Otherwise, the overexpression of AhR accelerated the cell proliferation of A549 lung carcinoma cells (36). Among these, Ma and Whitlock (37) have shown that stable transfection of AhR cDNA into AhR-defective (AhR-D) Hepa1c1c7 cells changes these characteristics such that the cells resemble wild-type cells. Conversely, introduction of antisense AhR cDNA into wild-type cells alters their phenotype such that they resemble AhR-D cells. And flow cytometric analysis implies that the slowed growth rate of AhR-D cells reflects prolongation of the G1 phase. Considering the facts that the primary Leydig cells are arrested in G1 and exhibit a low level of AhR, it is assumed that the AhR status of Leydig cells might be similar to AhR-D cells.

Among the cytochrome P450-dependent monooxygenases, CYP1A1 is thought to be mostly active in B[a]P oxidation steps of mammalian cells (10). Our results revealed the presence of CYP1A1 expression in Leydig cells but at levels insufficient to exhibit its activity. This also indicates that Leydig cells can be protected from the cytotoxicity-mediated cell damage/death provoked by B[a]P as well as presumably by other PAHs. In the present study, we have demonstrated that transient transfection of a CYP1A1 overexpression vector into TM3 cells sensitizes the cells to exhibit activity in the presence of B[a]P. Notably, we confirmed again from this experiment that elevated CYP1A1 levels by ectopic CYP1A1 expression do not lead to an increase in its activity. So far, there have been only two studies that have suggested the possible role of P450-dependent monooxygenases (38) and the metabolism of B[a]P in rat testis (39). Although these experiments were not performed with Leydig cells, the authors speculated that most of the metabolic actions involved may have been occurring primarily in the interstitial compartment of the testis where the Leydig cells are located. Nevertheless, based upon our data, we believe that the role of CYP1A1 in Leydig cells is effectively insignificant or not present.

Taken together, our data indicate that B[a]P-provoked apoptosis cannot occur to any significant extent in Leydig cells due to the up-regulation of XIAP, the lack of p53 function, the poor level of endogenous AhR, and the insufficient activity of CYP1A1. In particular, the deficiency in CYP1A1 activity might be a decisive condition rendering Leydig cells secure from exogenous PAHs like B[a]P. Therefore, testicular Leydig cells are believed to be strong enough to resist and survive the cytotoxic action of B[a]P. This study obviously provides the additive information on Leydig cell function that especially relates to cellular defense or tolerance mechanisms against the cytotoxic action of B[a]P. Considering the critical role of CYP1A1 activity in B[a]P metabolism in Leydig cells, it will be also important to examine the potential activities of other CYPs (i.e. CYP1A2, CYP1B1, and CYP3A4) in Leydig cell functions. However, it is now uncertain whether B[a]P exposure is disrupting the endocrine function of the Leydig cells. Indeed, B[a]P is known to have a xenoestrogenic action (40, 41) and is currently considered to be an endocrine disruptor (42). Consequently, the effect of B[a]P on Leydig cell steroidogenesis is under investigation in our laboratory.


   Footnotes
 
This study was supported by grants from the Korea Health 21 R&D Project, Ministry of Health and Welfare, Republic of Korea (A030077) and Korea Research Foundation Grant 2006-005-J03502.

Disclosure Summary: The authors have nothing to disclose

First Published Online September 20, 2007

Abbreviations: Ac-DEVD-AMC, N-Acetyl-L-aspartyl-L-glutamyl-L-valyl-L-aspartic acid amide; Ac-DEVD-CHO, N-acetyl-L-aspartyl-L-glutamyl-L-valyl-L-aldehyde; AhR, aryl-hydrocarbon receptor; AhR-D, AhR-defective; ARNT, aryl hydrocarbon receptor nuclear translocator; B[a]P, benzo[a]pyrene; BPDE, B[a]P-7,8-dihydroxy-9,10-epoxide; CYP1, cytochrome P450; DHD, (+/–)-B[a]P-trans-7,8-dihydrodiol; DTT, dithiothreitol; IAP, inhibitor of apoptosis protein; MMLV, Moloney murine leukemia virus; MTT, 3-[4,5-dimethylthiazol-2-yl]-2.5-diphenyltetrazolium bromide thiazolyl blue; PAH, polycyclic aromatic hydrocarbons; PI, propidium iodide; XIAP, X chromosome-linked inhibitor of apoptosis protein.

Received January 3, 2007.

Accepted for publication September 12, 2007.


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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
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