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College of Pharmacy, Chung-Ang University, Seoul, South Korea
(Received July 12, 2005; Accepted August 23, 2005)
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Abstract |
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). CYP1B1 is mainly found in extrahepatic steroidogenic tissues (e.g., ovary, testis, and adrenal gland) and in steroid-responsive tissues, e.g., breast, uterus, and prostate. The enzyme is also found in many other extrahepatic tissues, including kidney, thymus, lung, spleen, brain, heart, colon, and intestine (Sutter et al., 1994; Shimada et al., 1996).
CYP1B1 genes have been cloned and characterized from mice, rats, and humans (Savas et al., 1994; Shen et al., 1994; Walker et al., 1995; Tang et al., 1996). The human CYP1B1 gene is located on chromosome 2 at 2p21-22, spanning approximately 12 kilobases of DNA, and contains three exons and two introns (Tang et al., 1996). Expression of CYP1B1 is induced by polycyclic aromatic hydrocarbons, which act via the Ah receptor-mediated pathway. 2,3,7,8-Tetrachlorodibenzo-p-dioxin (TCDD) is a well known agonist for activating transcription of the CYP1B1 gene (Sutter et al., 1991). Several positive and negative regulatory DNA elements have been identified in the 5' region of the CYP1B1 gene and may be involved in Ah receptor-mediated gene regulation (Wo et al., 1997).
A major interest in CYP1B1 arises from the fact that it is the most catalytically efficient 17ß-estradiol (E2) 4-hydroxylase characterized to date, and 4-hydroxy E2 has been suggested to be carcinogenic (Hayes et al., 1996; Shimada et al., 1999). Moreover, CYP1B1 is expressed at a higher level in various human cancers, including brain, skin, testis, and breast (McKay et al., 1995; Murray et al., 1997). Because of the postulated significant role of CYP1B1 on carcinogenicity of E2, CYP1B1 is regarded as a target enzyme for blocking tumor initiation, and selective inhibition of CYP1B1 may prevent E2-related tumor formation (Liehr et al., 1986; Li and Li, 1987; Liehr, 1997; Shimada et al., 1997).
Many trans-stilbene analogs have been shown to inhibit the CYP1 family of proteins, which function in the metabolic activation of procarcinogens. Resveratrol inhibits CYP1A1 and CYP1B1 (Chun et al., 1999; Chang et al., 2000; Chen et al., 2004). Rhapontigenin and 3,5,3',4',5'-pentamethoxystilbene exhibit a potent and selective inhibition of CYP1A1, and 3,4'-dimethoxy-5-hydroxystilbene showed inhibition of EROD activity of both CYP1A1 and CYP1B1 with an IC50 value of 0.1 µM (Chun et al., 2001b; Lee et al., 2004). trans-Stilbene analogs also modulate expression of CYP1A1 and CYP1B1 in cultured human cells (Chen et al., 2004; Lee et al., 2004). Previously, we reported that 2,4,3',5'-tetramethoxystilbene (TMS), a methoxy derivative of oxyresveratrol, is a potentially selective inhibitor of CYP1B1 (Chun et al., 2001a; Kim et al., 2002; Chun and Kim, 2003). TMS was a competitive inhibitor of recombinant human CYP1B1 with a Ki of 3 nM (Chun et al., 2001a). It also strongly inhibited 4- and 2-hydroxylation of E2 by CYP1B1-expressing membranes or purified CYP1B1. The activation of 2-amino-3,5-dimethylimidazo[4,5-f]quinoline (MeIQ) in an Escherichia coli lac-based mutagenicity tester system containing functional human CYP1B1 was strongly inhibited by TMS. TMS also inhibited human CYP1B1-mediated melatonin 6-hydroxylation with an IC50 value of 30 nM (Ma et al., 2005).
The aim of the present study was to elucidate the effect of TMS on CYP1B1 activity and expression in human tumor cells and to determine the possibility of TMS as an adjuvant agent for cancer protection. The results show that TMS is able to inhibit activity and expression of CYP1B1 induced by TCDD in human tumor cells. Enhancement of sensitivity to TMS by TCDD in tumor cells emphasizes the possibility that CYP1B1-dependent metabolism may be involved in TMS-mediated cytotoxicity.
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Materials and Methods |
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EROD Enzyme Assay. EROD activity was determined for the measurement of CYP1 activities (Burke et al., 1985). The reaction mixture contained 0.1 M potassium phosphate buffer (pH 7.4), 2 mg/ml bovine serum albumin, 2 µM ethoxyresorufin, and rat lung microsomes or cellular sonicates. The reaction mixtures were preincubated at 37°C for 3 min, and the reactions were initiated by addition of 120 µM NADPH. Incubations were performed in a shaking water bath at 37°C for 20 min and terminated by addition of 1 ml of methanol. The formation of resorufin was determined fluorometrically with a PerkinElmer LS 5 spectrofluorimeter (PerkinElmer Life and Analytical Sciences, Boston, MA), with excitation and emission wavelengths of 550 nm and 585 nm, respectively. Protein concentrations were estimated using the bicinchoninic acid method according to the supplier's recommendations (Pierce Chemical, Rockford, IL) using bovine serum albumin as a standard.
Western Blot Analysis. Proteins were separated by 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis and transferred to nitrocellulose membranes. Membranes were blocked in 5% (w/v) nonfat dried milk in Tris-buffered saline with 0.05% Tween 20 overnight at 4°C. Membranes were then incubated for 1 h with rabbit anti-CYP1B1 polyclonal antibodies at a 1:1000 dilution in Tris-buffered saline with 0.05% Tween 20. After subsequent incubation with a horseradish peroxidase-conjugated goat anti-rabbit IgG antibody, proteins were visualized by an enhanced chemiluminescence method.
RT-PCR. Total RNA was extracted using TRIzol reagent (Invitrogen, Carlsbad, CA). RT-PCR was performed using an Access RT-PCR system purchased from Promega (Madison, WI) according to the manufacturer's instructions. Human CYP1B1 cDNA was amplified by PCR using a sense primer (5'-AACGTCATGAGTGCCGTGTGT-3') and an antisense primer (5'-GGCCGGTACGTTCTCCAAATC-3') with 30 cycles by denaturation at 94°C for 20 s, annealing at 58°C for 20 s, and extension at 72°C for 40 s (Li et al., 1998). Human Ah receptor cDNA was amplified using a sense primer (5'-CATGCTTGGTCTTTTATGC-3') and an antisense primer (5'-TTCCCTTTCTTTTTCTGTCC-3') with 30 cycles by denaturation at 94°C for 20 s, annealing at 52°C for 20 s, and extension at 72°C for 40 s. Human ARNT cDNA was amplified using a sense primer (5'-GGAACAAGATGACAGCCTAC-3') and an antisense primer (5'-CAGAAAGCCATCTGCTGCC-3') with 30 cycles by denaturation at 94°C for 20 s, annealing at 60°C for 20 s, and extension at 72°C for 40 s. Human ß-actin cDNA was amplified using a sense primer (5'-CTACAATGAGCTGCGTGTGG-3') and an antisense primer (5'-TAGCTCTTCTCCAGGGAGGA-3') with 30 cycles by denaturation at 94°C for 20 s, annealing at 52°C for 20 s, and extension at 72°C for 40 s. The number of amplification cycles was optimized in preliminary experiments to ensure that the PCR had not reached its plateau. The amplified PCR products were analyzed by 2% agarose gel electrophoresis and ethidium bromide staining.
Determination of Cell Toxicity. Cells were plated onto 96-well plates and incubated at 37°C in a 5% CO2 atmosphere. After incubation for the designated time, 10 µg of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) was added per well. After incubating at 37°C for 4 h, cells was centrifuged at 1000g for 5 min, the medium was removed by aspiration, and then the MTT formazan crystal formed was dissolved by adding 0.15 ml of DMSO and shaking for 15 min. The absorbance at 540 nm was measured using a microplate reader. The percentages of cells surviving from each group relative to control, defined as 100% survival, were calculated.
Apoptosis Assay. Cells were harvested and washed twice with ice-cold phosphate-buffered saline (pH 7.4). Cells were resuspended in 0.19 ml of binding buffer (pH 7.4) containing 10 mM HEPES, 140 mM NaCl, and 2.5 mM CaCl2. Ten microliters of Annexin V-FITC (Zymed Laboratories, South San Francisco, CA) was added per well and cells were incubated for 10 min at room temperature. Cells stained with Annexin V-FITC were washed with the binding buffer. Apoptotic cells were determined using fluorescence microscopy with excitation and emission wavelengths of 488 nm and 518 nm, respectively.
DNA Fragmentation Assay. Cells were harvested and lysed in 0.15 ml of lysis buffer containing 50 mM Tris-HCl (pH 8.0), 10 mM EDTA, 0.5% (w/v) sodium dodecyl sulfate, and 0.5 mg/ml proteinase K. Proteolytic digestion was allowed to proceed at 50°C for 12
16 h. DNA was extracted with phenol/chloroform/isoamyl alcohol (25:24:1) and precipitated with 1 volume of 7.5 M ammonium acetate and 2 volumes of absolute ethanol. Extracted chromosomal DNA was dissolved in 50 µl of Tris-EDTA buffer (pH 8.0) containing 0.2% sodium dodecyl sulfate, and 0.2 mg/ml DNase-free RNase. After incubation at 37°C for 1 h, DNA samples were analyzed in 1% agarose gel.
Expression of Recombinant Human CYP1B1. Coexpression (bicistronic) plasmids for human CYP1B1 and NADPH-P450 reductase were transformed into E. coli DH5
(Parikh et al., 1997). A single ampicillin-resistant colony of transformed cells was selected and grown in overnight culture to saturation at 37°C in LB medium containing 100 µg/ml ampicillin. A 10-ml aliquot was used to inoculate each liter of Terrific Broth containing 0.2% bactopeptone (w/v), 100 µg/ml ampicillin, 1 mM thiamine, trace elements, 0.5 mM 
-aminolevulinic acid, and 1 mM isopropyl ß-D-thiogalactoside. The cultures were grown at 30°C with shaking at 200 rpm for 48 h. After incubation, cells were harvested by centrifugation at 6500g for 20 min. Spheroplasts were prepared using lysozyme and disrupted by sonication. The cellular sonicates were centrifuged at 10,000g for 20 min, and the membranes were collected by centrifugation at 110,000g for 90 min and were resuspended in 10 mM Tris-HCl buffer (pH 7.4) containing 1.0 mM EDTA and 20% glycerol (v/v) (Guengerich et al., 1996). P450 content of whole E. coli cells and membranes was determined by the spectral method of Omura and Sato (1964) using an extinction coefficient of 91 mM–1 cm–1 with a Shimadzu UV-160A spectrophotometer (Shimadzu, Kyoto, Japan) at ambient temperature. The isolated membrane fractions were stored at –70°C.
HPLC. TMS (50 µM) was incubated in 0.1 M potassium phosphate buffer (pH 7.4), containing bacterial membranes having 10 pmol of human CYP1B1 and NADPH-P450 reductase in a final volume of 100 µl. After 5 min of preincubation at 37°C, the reaction was initiated by the addition of 1 mM NADPH and continued for 20 min with shaking. The same experiments were performed without NADPH or with heat-inactivated membranes. The reaction was terminated by the addition of 1 ml of dichloromethane. The reaction products were separated by HPLC on a 5-µm Nucleosil C18 reverse phase column (4.6 mm x 250 mm) with the mobile phase consisting of 40% solvent A (glacial acetic acid in water, pH 2.0) and 60% solvent B (20% solvent A and 80% acetonitrile). The absorbance at 306 nm was measured using a Jasco UV-975 UV/visible detector (Jasco, Tokyo, Japan).
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Results |
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Enhancement of Sensitivity to TMS by TCDD. Treatment with 2 nM TCDD for 72 h significantly decreased the cellular proliferation of MCF-7 and HL-60 cells (Table 1). TCDD (2 nM) treatment for 72 h caused 43% of the suppression in MCF-7 cell proliferation, whereas the same concentration of TCDD suppressed 55% of HL-60 cell proliferation. The effect of TCDD on TMS-mediated cytotoxicity was determined in MCF-7 and HL-60 cells (Table 1). Interestingly, cotreatment with TCDD significantly enhanced the sensitivity to TMS in human tumor cells. Treatment with TMS (1 µM) for 72 h caused 95% of the suppression of MCF-7 cell proliferation in the presence of 2 nM TCDD, whereas the same concentration of TMS suppressed 92% of HL-60 cell proliferation. Treatment with 1 µM TMS alone for 72 h was able to block the cell growth for 66% and 73% in MCF-7 and HL-60 cells, respectively (Table 1).
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Metabolism of TMS by CYP1B1. In vitro metabolism studies were performed with bacterial membranes of human CYP1B1 coexpressed with human NADPH-P450 oxidoreductase to examine whether CYP1B1 is able to metabolize TMS to its metabolite(s). As shown in Fig. 4A, TMS was metabolized by human CYP1B1, and one product (tR, 7.2 min) was formed when the reaction was supported by NADPH (1 mM). The product peak was not formed when NADPH was not added (Fig. 4B) or membranes were heat-inactivated before added in reaction mixtures (Fig. 4C). Addition of -naphthoflavone (-NF) (10 µM), a well known CYP1 inhibitor, blocked the formation of a TMS metabolite (tR, 7.2 min) by CYP1B1 (Fig. 5). We found that -NF was also metabolized by CYP1B1 to produce a major metabolite (tR, 8.4 min).
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Discussion |
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). TMS significantly prevented CYP1B1-dependent P450 activities such as EROD and E2 hydroxylation in CYP1B1-expressing E. coli membranes and the purified CYP1B1 enzyme. TMS also blocked CYP1B1-dependent MeIQ genotoxicity. In this study, to evaluate the mechanisms of anticarcinogenesis by TMS in cells, we examined the effect of TMS on the activity and expression of CYP1B1 in human cells. The results of these experiments indicate that TMS inhibits TCDD-induced EROD activity and also the expression of CYP1B1 in MCF-7 cells. In MCF-10A, an immortalized nontumorigenic breast epithelium cell, and HL-60, an acute promyelocytic leukemic cell, TMS also down-regulates CYP1B1 mRNA and protein expression. Interestingly, suppression of TCDD-induced CYP1B1 expression by TMS was much more significant in MCF-10A and HL-60 cells than in MCF-7 cells. Western blot and RT-PCR analyses showed a similar concentration dependence (Fig. 2, A and B). The induction of CYP1B1 expression by TCDD at 1 µM concentration was almost completely blocked by TMS in HL-60 cells. As previously reported (McKay et al., 1995; Murray et al., 1997), we also found the basal expression of CYP1B1 in human cell lines such as MCF-7, MCF-10A, and HL-60. Although TMS significantly suppressed TCDD-induced CYP1B1 expression, Ah receptor and ARNT mRNA expression was not affected by TMS treatment. The recent demonstration that resveratrol acts as an Ah receptor antagonist and suppresses the Ah receptor-mediated dioxin-responsive element binding activity suggests that TMS may also suppress the Ah receptor signal pathway (Ciolino et al., 1998; Casper et al., 1999; Chen et al., 2004). To determine the detailed mechanism of CYP1B1 modulation by TMS in cells, it will be necessary to investigate whether TMS may down-regulate CYP1B1 mRNA expression by competing with TCDD at the Ah receptor binding site.
Because we have considered TMS as an adjuvant agent for cancer prevention, the cytotoxic effects of TMS were determined. TMS alone prevents cell proliferation in MCF-7 and HL-60 cells in a concentration- and time-dependent manner. We were also able to detect the indications of apoptotic cell death such as phosphatidylserine exposure measured by Annexin V-binding or chromosomal DNA fragmentation in cells treated with TMS at less than 10 µM. Our prior work (Lee et al., 2002) and data from other laboratories (Clement et al., 1998; Huang et al., 1999; Wolter et al., 2002; Estrov et al., 2003) provide evidence that many trans-stilbene compounds also induce apoptotic cell death in various human tumor cells, although cytotoxic effects were relatively lower than with TMS. Geahlen and McLaughlin (1989) previously suggested that piceatannol (3,4,3',5'-tetrahydroxystilbene) acts as an inhibitor of protein tyrosine kinase p72Syk. The ability of TMS to inhibit protein tyrosine kinases will need to be determined to understand the cytotoxic mechanism of TMS.
The enhancement of sensitivity to TMS by TCDD in tumor cells is interesting. From in vitro metabolism studies with recombinant CYP1B1, we found a significant metabolism product of TMS using HPLC. Previously, we suggested that the metabolic product might be an O-demethylated TMS (Chun et al., 2001a). We considered the possibility that the cytotoxic effect of O-demethylated metabolite(s) may be more potent in tumor cells. Because induction of CYP1B1 expression by TCDD increases the production of TMS metabolite(s), simultaneous treatment with TCDD may significantly enhance TMS-mediated protection of tumor cell proliferation. The effect of TCDD on TMS cytotoxicity is stronger in MCF-7 cells than in HL-60 cells, although TCDD induction of CYP1B1 expression is substantially higher in HL-60 than in MCF-7, likely due in part to the relatively stronger metabolic ability of MCF-7 cells than HL-60 cells.
Estrogen is a known risk factor in various human tumors, especially in breast tumor. Because CYP1B1 has been detected in over 70% of the human breast tumor samples analyzed and is known as a major enzyme responsible for carcinogenic metabolism of estrogen, this enzyme is regarded as a target enzyme for blocking tumor initiation and its inhibition remains a logical target for cancer protection. The data reported here suggest that TMS acts as a modulator of CYP1B1 gene expression as well as a potently selective inhibitor in vitro and in cultured cells. The ability of TMS to induce apoptotic cell death in tumor cells as well as CYP1B1 inhibition may be beneficial in cancer prevention. A more detailed understanding of the precise mechanism(s) by which TMS inhibits CYP1B1 and induces apoptosis may facilitate the development of a new strategy for cancer chemoprevention.
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Acknowledgments |
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Footnotes |
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Article, publication date, and citation information can be found at http://dmd.aspetjournals.org.
ABBREVIATIONS: Ah, aryl hydrocarbon; ARNT, aryl hydrocarbon receptor nuclear translocator; P450, cytochrome P450; DMBA, 7,12-dimethylbenz[a]anthracene; EROD, 7-ethoxyresorufin O-deethylation; MTT, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide; -NF, -naphthoflavone; TCDD, 2,3,7,8-tetrachlorodibenzo-p-dioxin; TMS, 2,4,3',5'-tetramethoxystilbene; E2, 17ß-estradiol; MeIQ, 2-amino-3,5-dimethylimidazo[4,5-f]quinoline; RT-PCR, reverse transcription-polymerase chain reaction; DMSO, dimethyl sulfoxide; FITC, fluorescein isothiocyanate; HPLC, high-performance liquid chromatography.
Address correspondence to: Prof. Young-Jin Chun, Ph.D., College of Pharmacy, Chung-Ang University, 221 Huksuk-Dong, Dongjak-Gu, Seoul 156-756, South Korea. E-mail: yjchun{at}cau.ac.kr
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  H. Park, S. E. Aiyar, P. Fan, J. Wang, W. Yue, T. Okouneva, C. Cox, M. A. Jordan, L. Demers, H. Cho, et al. Effects of Tetramethoxystilbene on Hormone-Resistant Breast Cancer Cells: Biological and Biochemical Mechanisms of Action Cancer Res., June 15, 2007; 67(12): 5717 - 5726. [Abstract] [Full Text] [PDF]
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