Change in the gene expression of hepatic tamoxifen-metabolizing enzymes during the process of tamoxifen-induced hepatocarcinogenesis in female rats
Toshihiko Kasahara,
Masamichi Hashiba1,
Tsuyoshi Harada2 and
Masakuni Degawa,3
Department of Molecular Toxicology, School of Pharmaceutical Sciences, University of Shizuoka, 52-1 Yada, Shizuoka 422-8526, Japan,
1 Institute of Applied Biochemistry, University of Tsukuba, 1-1-1, Tennodai, Tsukuba City, Ibaraki 305-8572, Japan and
2 First Department of Biochemistry, Saitama Medical School, 38 Morohongo, Moroyama-machi, Iruma-gun, Saitama 350-0495, Japan
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Abstract
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Altered gene expression of the enzymes responsible for tamoxifen metabolism during the process of tamoxifen-induced hepatocarcinogenesis in female SpragueDawley rats was examined by the RTPCR method. Treatment of rats with tamoxifen (20 mg/kg body/day) for 52 weeks, but not the 1 day, 2 or 12 week treatments, resulted in the formation of the liver hyperplastic nodules. The gene expression of CYP3A subfamily enzymes, especially CYP3A1, responsible for not only detoxification (N-demethylation) but also activation (
-hydroxylation) of tamoxifen, was increased by the tamoxifen treatments for 2 and 12 weeks, whereas after the 52 week treatment, the expression in the induced nodules returned to the control level. The gene expression of SULT2A subfamily sulfotransferases, especially HSTa, responsible for metabolic activation of
-hydroxytamoxifen was decreased to a level <20% of the control in the nodules, although no significant change in the expression was observed in the liver of rats treated with tamoxifen for 1 day, 2 or 12 weeks. On the other hand, the gene expression of CYP3A2 and flavin-containing monooxygenase 1 (FMO1), responsible for the N-demethylation and N-oxidation, respectively, of tamoxifen was increased in a time-dependent fashion up to the 52 week treatment. Although the gene expression of UDP-glucuronosyltransferase(s), which might be responsible for detoxification of tamoxifen, was also increased by the tamoxifen treatment for 2 or 12 weeks, it decreased to the control level in the nodules after the 52 week treatment. The present findings demonstrate that in the early stage of the formation of the liver hyperplastic nodules by tamoxifen, the genes of the enzymes responsible for not only detoxification but also activation of tamoxifen were activated, whereas in the later stage (in the nodules), the genes of the detoxification enzymes, CYP3A2 and FMO1, remained active, but those of the activation enzymes such as CYP3A1 and HSTa were suppressed.
Abbreviations: FMO, flavin-containing monooxygenase;; HST, hydroxysteroid sulfotransferase;; P450, cytochrome P-450;; UDPGT, UDP-glucuronosyltransferase;; UGT1A1, 1A6 and 2B1, isoforms of UDPGT.
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Introduction
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The antiestrogen tamoxifen is widely used as not only anticancer drug (1,2) but also as chemopreventive agent (3) for breast cancer. However, the potent hepatocarcinogenicity of tamoxifen in rats has also been reported (47). This hepatocarcinogenesis is thought to occur through the chemical modification of DNA by tamoxifen metabolites (813). For the formation of DNA adducts, metabolic activation of tamoxifen is indispensable; the metabolites
-hydroxytamoxifen (1419) and its O-sulfate (20,21) are characterized as proximate and ultimate carcinogens, respectively (Figure 1
). On the other hand, major metabolites such as N-desmethyltamoxifen, tamoxifen N-oxide and 4-hydroxytamoxifen are generally characterized as detoxification forms, although the further metabolites,
-hydroxyl forms of the N-desmethyltamoxifen and tamoxifen N-oxide, are able to produce the DNA adducts (22).
In humans,
-hydroxylation of tamoxifen is catalyzed mainly by CYP3A subfamily enzyme(s) (23), and N-demethylation (2426), 4-hydroxylation (27,28) and N-oxidation (29) occur mainly by the action of CYP3A4, CYP2D6 and flavin-containing monooxygenase (FMO), respectively. Likewise, rat CYP3A subfamily enzymes are reported to catalyze not only the
-hydroxylation but also the N-demethylation of tamoxifen (30). On the other hand, the rat enzymes responsible for the 4-hydroxylation and N-oxidation of tamoxifen have not yet been identified; although CYP2D subfamily enzyme(s), especially CYP2D2, and FMO1 are suggested to catalyze the 4-hydroxylation and the N-oxidation, respectively, because these rat enzymes are known to have characteristics similar to those of the corresponding human enzymes, CYP2D6 and FMO, in terms of substrate-specificity (31,32).
Concerning the phase II enzymes responsible for the metabolism of tamoxifen, rat hydroxysteroid sulfotransferases (HST), especially hydroxysteroid sulfotransferase a (HSTa), are reported to catalyze the O-sulfation of
-hydroxytamoxifen to form the ultimate carcinogen (21). UDP-glucuronosyltransferases (UDPGT) are known to mediate the O-glucuronidation of
-hydroxytamoxifens (23,33).
Alternative expression of the enzymes responsible for hepatocarcinogen metabolism during the hepatocarcinogenic process is reported to closely correlate with the carcinogenic susceptibility of animals and their organs (3438). However, few studies have been performed regarding changes in the tamoxifen-metabolizing enzymes during the hepatocarcinogenic process (39).
In the present study, we examined the tamoxifen-induced changes in the gene expression of the main enzymes, cytochrome P450 (P450) isoforms, FMO1, HST and UDPGT, responsible for tamoxifen metabolism. Herein, we present our findings and discuss the relationship between changes in the gene expression of tamoxifen-metabolizing enzymes and hepatocarcinogenesis.
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Materials and methods
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Treatment of animals
Female SpragueDawley rats were purchased from Charles River (Atsugi, Japan) and used at 6 weeks of age. They were kept individually in aluminium cages in an air-conditioned room and given CRF-1 diet (Oriental Yeast Co., Tokyo, Japan) and water ad libitum. Tamoxifen citrate (Sigma Chemical Co., St Louis, MO, USA) suspended in 0.5% carboxymethyl cellulose was used as tamoxifen. The drug (20 mg/kg body wt) was given to rats by gavage once a day for 1 day, 2, 12 or 52 weeks. To the corresponding control group rats, vehicle (0.5% carboxymethyl cellulose solution) alone was given. Twenty-four hours after the final treatment, the animals were killed by exsanguination under anesthesia, and their livers were removed quickly and stored in liquid nitrogen until processed for analysis. In the present experiments, the liver-bearing hyperplastic nodules were divided into non-nodular (nodule-surrounding tissue) and nodular parts, and they were used as separate samples.
Measurement of microsomal total cytochrome P450 (P450)
The liver was perfused with 1.15% (w/v) KCl and homogenized in 10 vol (v/w) of 50 mM TrisHCl buffer (pH. 7.4) containing 1.15% (w/v) KCl and 1 mM EDTA. The homogenate was centrifuged at 10 000 g for 20 min, and the resultant supernatant was re-centrifuged at 105 000 g for 1 h. The precipitate obtained as the microsomal fraction was resuspended in 0.1 M TrisHCl buffer (pH.7.4) containing 1 mM EDTA and stored at 80°C until used for analysis. The amount of microsomal protein was determined with a Bio-Rad DC Protein Assay kit (Bio-Rad Laboratories, Hercules, CA, USA), and that of microsomal total P450, by the method of Omura and Sato (40).
Preparation of total RNA
Total hepatic RNA was extracted with Isogen (Nippon Gene Co., Tokyo, Japan). Briefly, the liver (
0.1 g) was homogenized in 1 ml of isogen with a Stirrer MAZELA Z-1000 (Tokyo Rikakikai Co., Tokyo, Japan). To the homogenate, chloroform (0.2 ml) was added; the mixture was then shaken vigorously for 30 s, stood at room temperature for 5 min and thereafter centrifuged at 12 000 g for 15 min. The resultant aqueous layer was transferred to a fresh tube. To the aqueous solution, an equivalent volume of isopropanol was added, and the mixture was shaken gently, stood at room temperature for 10 min and then centrifuged at 12 000 g for 10 min. To the obtained precipitates, 75% ethanol was added, and the mixture was then centrifuged at 12 000 g for 5 min. The resultant precipitates were obtained as a crude RNA fraction. For further purification, the crude RNA fraction was dissolved in 0.1 ml of diethylpyrocarbonate (DEPC)-treated water and subjected again to the series of steps for RNA preparation described above. Finally, the purified RNA fraction was dissolved in 0.1 ml of DEPC-treated water, and its concentration was determined spectrophotometorically by absorbance at 260 nm: 1 OD = 40 µg/ml.
RTPCR
Complementary DNA (cDNA) synthesis from total RNA (2 µg) was performed in a reaction mixture (33 µl) containing a Not I-(dT)18 primer (Amersham Pharmacia Biotech, Buckinghamshire, UK) and a First-Strand cDNA Synthesis Kit RTG You-Prime First-Strand Beads (Amersham Pharmacia Biotech). Amplification of each cDNA (1 µl of the RT reaction mixture) with the exception of FMO1 cDNA was performed in a reaction mixture (50 µl) containing Gene Taq universal buffer (5 µl), dNTPs (0.2 mM), forward and reverse primers (each 20 pmol), Gene Taq polymerase (1.25 U; Nippon Gene). Amplification of FMO1 cDNA (1 µl of the RT reaction mixture) was performed in a 50 µl of reaction mixture containing PCR buffer for KOD-Plus (5 µl), 1 mM MgSO4, dNTPs (0.2 mM), forward and reverse primers (each 20 pmol) and KOD-Taq polymerase (1 U; TOYOBO Co., Osaka, Japan).
Amplifications of all cDNA examined were carried out with a PCR thermal cycler (Perkin-Elmer Cetus, Norwalk, CT, USA). The PCR program used was as follows: pre-treatment, at 94°C for 2 min; denaturation, at 94°C for 20 s; annealing at 57°C for 45 s; extension, at 72°C for 45 s. The primer sets used and the predicted sizes of PCR products are summarized in Table I
. A G3PDH primer was used for the G3PDH primer set (Clontech Laboratories, Palo Alto, CA, USA). The amount of product generated by each PCR increased linearly in a PCR cycle-dependent manner over the following ranges: 2125 cycles for CYP3A1, CYP3A9, CYP2D2, HST20, UGT2B1 and G3PDH; 2529 cycles for CYP3A23, HSTa, FMO1 and UGT1A1; 2834 cycles for CYP3A18 and UGT1A6 and 3236 cycles for CYP3A2. The primer set for UGT2B1 was designed by use of Genetyx-Mac software (Software Development Co., Tokyo, Japan), and the sequence of the PCR product obtained was in complete agreement with the predicted sequence (data not shown).
In addition, to assess possible contamination by genomic DNA of the RNA preparation used, we performed direct PCR (without the RT reaction) of each RNA preparation. The PCR gave no product, demonstrating no contamination of genomic DNA in the RNA preparations used.
Quantification of mRNA
At first, to confirm the molecular weight of each PCR product, we subjected the reaction solution (10 µl) after amplification of each cDNA to 2% agarose gel-electrophoresis (FMC BioProducts, Rockland, ME, USA), and visualized separated band with ethidium bromide under UV light. The molecular weights of the PCR products obtained were identical with those of the corresponding predicted products.
For quantification, a portion of each PCR product was applied on a capillary electrophoresis system (Hewlett-Packard Co., Wilmington, DE, USA), and the amount of PCR product was measured spectrophotometorically by absorbance at 254 nm and normalized to that of G3PDH, an internal standard.
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Results
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Female rats were treated with tamoxifen for 1 day, 2, 12 or 52 weeks. The 52 week treatment resulted in the formation of hyperplastic nodules in their liver (Figure 2
), whereas no nodule formation was observed in any of the other experimental groups (data not shown). In addition, in a part of one of the induced nodules, adenocarcinoma was observed by histopathological analysis (data not shown).
We first examined the effect of tamoxifen treatment on hepatic microsomal P450. As shown in Figure 3
, the amount of total P450 was not significantly changed by the treatment for 1 day, 2 or 12 weeks, whereas it was significantly decreased by the 52 week treatment, especially in the hyperplastic nodules. Therefore, we further examined the change in the gene expression of the main P450 isoforms, i.e. CYP3A and CYP2D subfamily enzymes, responsible for tamoxifen metabolism. Expression of all CYP3A genes examined was increased by the tamoxifen treatment for 2 or 12 weeks but not by that for 1 day (Figure 4
). After 52 weeks of treatment, the increased expression of CYP3A2 was observed in both hyperplastic nodules and their surrounding tissues. On the other hand, as for the expression of other CYP3A genes, no increase was observed in the hyperplastic nodules, whereas an increase in the expression of CY3A18 and CYP3A23 was observed in the nodule-surrounding tissues. Namely, the expression levels of CYP3A genes with the exception of CYP3A2 reached a maximum after 12 weeks of treatment and were
10-fold higher than the corresponding control levels (Figure 5
). On the other hand, the expression of CYP3A2 reached its maximum after 52 weeks of treatment, and its level was
18-fold higher than the control. In addition, no significant change in the expression of CYP2D2 by the tamoxifen treatment was observed at any period examined (Figures 4 and 5
), although its expression level was observed to decrease at 52 weeks later in both tamoxifen-untreated (control) and tamoxifen-treated groups (Figure 4
). Thus, a decrease in expression levels of CYP2D2 might be dependent on aging. The gene expression of FMO1, the main enzyme responsible for N-oxidation of tamoxifen, was also examined for possible changes induced by tamoxifen treatment. The expression level of FMO1 was increased by the tamoxifen treatments for 1 day, 2, 12 and 52 weeks (Figure 4
) and reached its maximum after the 52 week treatment, at which time it was >6-fold higher in both nodules and their surrounding tissues (Figure 6
).

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Fig. 3. Effect of tamoxifen-treatment on the amount of total hepatic microsomal P450. Data shown are presented as the mean ± SD (n = 35). Columns: open, control; hatched, tamoxifen treatment; closed, hyperplastic nodules. *, **Significant differences from the corresponding controls assessed by Student's t-test: *P < 0.05; **P < 0.01.
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Fig. 4. Representative profile of the agarose gel-electrophoresis of the RTPCR product for hepatic phase I enzymes, CYP3A1, CYP3A2, CYP3A9, CYP3A18, CYP3A23, CYP2D2 and FMO1, in female rats treated with tamoxifen. PCR products from five individual rats in each experimental group were combined, and the mixture was subjected to agarose gel-electrophoresis. Numbers of PCR cycles used for CYP3A1, CYP3A2, CYP3A9, CYP3A18, CYP3A23, CYP2D2, FMO1 and G3PDH were 22, 35, 25, 30, 26, 25, 28 and 23, respectively. C, control; T, tamoxifen treatment; TN, hyperplastic nodules. G3PDH was used as an internal standard.
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Fig. 5. Changes in the gene expression levels of hepatic P450 enzymes, CYP3A1, CYP3A2, CYP3A9, CYP3A18, CYP3A23 and CYP2D2, during the process of tamoxifen-induced hepatocarcinogenesis. The number of PCR cycles used for each P450 was shown in the legend for Figure 4 . Expression levels of the genes tested were calculated on a basis of that of G3PDH, an internal control, and are shown as a percentage of the corresponding controls. Amount of each RTPCR product was assayed in five individual rats in each experimental group, and the data shown are presented as the mean ± SD (n = 5). *, **Significant differences from the corresponding controls assessed by Student's t-test; *P < 0.05: **P < 0.01.
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Fig. 6. Change in the expression level of the FMO1 gene during tamoxifen-induced hyperplastic nodule formation in the liver. PCR was performed for 28 cycles. Expression levels of the gene were calculated on a basis of that level of G3PDH, an internal control, and are shown as a percentage of the corresponding controls. The amount of each RTPCR product was assayed in five individual rats in each experimental group, and the data shown are presented as the mean ± SD (n = 5). *, **Significant differences from the corresponding controls assayed by Student's t-test: *P < 0.05; **P < 0.01.
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Next we examined the tamoxifen-induced change in the gene expression of HST (HSTa and HST20) and glucuronosyltransferases (UGT1A1, UGT1A6 and UGT2B1). No significant change in the expression of the HST genes in response to the tamoxifen treatment for 1 day, 2 or 12 weeks was observed, whereas the expression was significantly decreased by the 52 week treatment, especially in the hyperplastic nodules (Figure 7
). The expression levels of HST genes in the hyperplastic nodules were <20% of those of the corresponding controls (Figure 8
). On the other hand, expression levels of the UDPGT genes examined were increased 24-fold over those of the corresponding controls by the tamoxifen treatments with the exception of the 1 day treatment (Figures 7 and 9
), although no significant increase was observed in the hyperplastic nodules.

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Fig. 7. Representative profile of the agarose electrophoresis of the PCR product of hepatic phase II enzymes (HSTa, HST20, UGT1A1, UGT1A6 and UGT2B1) in female rats treated with tamoxifen. C, control; T, tamoxifen treatment; TN, hyperplastic nodules. G3PDH was used as an internal standard. Numbers of PCR cycles used for HSTa, HST20, UGT1A1, UGT1A6, UGT2B1 and G3PDH were 28, 23, 29, 34, 23 and 23, respectively.
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Fig. 8. Changes in the gene expression levels of hepatic HSTa and HST20 during tamoxifen-induced hepatocarcinogenesis. Numbers of PCR cycles used for HSTa and HST20 were 28 and 23, respectively. Expression levels of the genes tested were calculated on a basis of that of G3PDH, an internal control, and are shown as a percentage of the corresponding controls. Amount of each RTPCR product was assayed in five individual rats in each experimental group, and the data shown are presented as the mean ± SD (n = 5). *, **Significant differences from the corresponding controls assessed by Student's t-test: *P < 0.05; **P < 0.01.
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Fig. 9. Changes in the gene expression levels of hepatic UGT1A1, UGT1A6 and UGT2B1 during the process of tamoxifen-induced hepatocarcinogenesis. Numbers of PCR cycles used for UGT1A1, UGT1A6 and UGT2B1 were 30, 34 and 23, respectively. The amount of each RTPCR product was assayed in five individual rats in each experimental group. Expression levels of the genes tested were calculated on a basis of that of G3PDH, an internal control, and are shown as a percentage of the corresponding controls. Data shown are presented as the mean ± SD (n = 5). *, **Significant differences from the corresponding controls were assessed by Student's t-test: *P < 0.05; **P < 0.01.
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Discussion
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We reported herein for the first time the overall change in the gene expression of the tamoxifen-metabolizing enzymes including phase I (P450 and FMO1) and phase II (HST and UDPGT) during the process of tamoxifen-induced hepatocarcinogenesis in female rats. Although the induction of hepatic tamoxifen-metabolizing enzymes, CYP3A1, HSTa and UGT2B1, by the short-term treatment (within 2 weeks) of rats with tamoxifen had been reported (4850), such results were further confirmed in the present study.
Moreover, in the present study we found that the amount of hepatic P450 was decreased significantly in the liver-bearing hyperplastic nodules, especially the hyperplastic nodules, after the treatment with tamoxifen for 52 weeks and further demonstrated that gene expression of CYP3A1 (8) and HSTa (21), which are thought to be the activation enzymes responsible for
-hydroxylation of tamoxifen and its O-sulfation, respectively, were remarkably decreased in the induced hyperplastic nodules. The change in the gene expression of CYP3A9, CYP3A18 and CYP3A23 enzymes was similar to that of CYP3A1, although the roles of these enzymes in the metabolism of tamoxifen have not yet been clarified. Furthermore, it is noteworthy that the gene expression of CYP3A2, which is a male-selective enzyme (42,51,52) and is thought to catalyze the detoxification (N-demethylation) of tamoxifen (26,53), increased in a treatment time-dependent manner up to the 52 week treatment, and the expression level in the hyperplastic nodules was increased
18-fold over the control. Likewise, we demonstrated for the first time that gene expression of FMO1, an enzyme responsible for detoxification (N-oxidation) of tamoxifen, was increased in a treatment time-dependent manner up to the 52 week treatment, at which time the level in the hyperplastic nodules was
10-fold higher than the control level. In addition, no significant change in the expression of CYP2D2, which is thought to catalyze the detoxification (4-hydroxylation) of tamoxifen (27,28), was observed during the process of the tamoxifen-induced hyperplastic nodule formation in the liver. The gene expression of P450 isozymes such as CYP1A2 and CYP2C6 was remarkably decreased in the tamoxifen-induced hyperplastic nodules (data not shown), although these enzymes are involved a little in the metabolism of tamoxifen (25). In addition, our preliminary experiments suggested that the microsomes from hyperplastic nodules showed higher activity for N-oxidation of tamoxifen than did those from control livers, but showed almost the same activity as the control for N-demethylation and 4-hydroxylation (data not shown).
As HSTa, a member of the SULT2A subfamily, is reported to act as an activation enzyme catalyzing O-sulfation of
-hydroxytamoxifen (21), we investigated herein a possible tamoxifen-induced change in the gene expression of HSTa. No significant change in the HSTa expression by the tamoxifen treatment for 1 day, 2 or 12 weeks was observed, whereas after 52 weeks of treatment, the expression levels in the nodules and their surrounding tissues decreased to
20 and 60%, respectively, of the control. The gene expression of HST20, a member of the SULT2A subfamily, was also decreased in the liver-bearing hyperplastic nodules, especially in the nodules, although this enzyme has no capacity for catalyzing O-sulfation of
-hydroxytamoxifen (45).
Although formation of O-glucuronide metabolite of
-hydroxytamoxifen had been reported (33), a subtype of UGT(s) responsible for this O-glucuronidation has not yet been determined. Therefore, in the present study, we focused on representative subtypes of UGTs, UGT1A1, UGT1A6 and UGT2B1, and examined the change in their expression levels after tamoxifen treatment. The expression levels of these genes were increased significantly by the treatment with tamoxifen for 2, 12 and 52 weeks; although in the hyperplastic nodules, these levels were returned to the corresponding control ones.
The present findings indicate that in the stage before the formation of hyperplastic nodules in the liver, the genes of several hepatic enzymes responsible for not only detoxification but also activation of tamoxifen were activated and that in the later stage (in the nodules), the gene activation of detoxification enzymes was selectively maintained, while that of activation enzymes was suppressed. Thus, the overall change in the gene expression of the tamoxifen-metabolizing enzymes by tamoxifen treatment appears to be reasonable for the formation and growth of the hepatic hyperplastic nodules, because the increase in detoxification enzymes in the later stage would be expected to confer tamoxifen resistance to the induced nodules. However, to further confirm this hypothesis, it will be necessary to examine the change in the activity of each enzyme responsible for the metabolism of tamoxifen, because the levels of activity and protein of a hepatic drug-metabolizing enzyme are known to be not necessarily dependent on the level of its corresponding mRNA.
Although tamoxifen is widely used for chemotherapy and prophylaxis of breast cancer, its use in long-term treatment was reported to result in an increase in incidence of endometrial cancer (54,55) and in the formation of tamoxifenDNA adducts in the endometrial tissue in humans (56). Furthermore, CYP enzyme(s) producing genotoxic intermediates from tamoxifen was reported to exist in human endometrial epithelium (57). These reports and the present findings indicate that further study on the changes in the expression of endometrial enzymes responsible for tamoxifen metabolism in tamoxifen-administered humans would be important for understanding the correlation between the use of tamoxifen and uterine carcinogenesis. Moreover, the present findings suggest that in tamoxifen-administered humans, clinical drug interactions might occur through tamoxifen-induced altered expression of drug-metabolizing enzymes.
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Notes
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3 To whom correspondence should be addressed Email: degawa{at}smail.u-shizuoka-ken.ac.jp 
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Acknowledgments
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Authors are grateful for the helpful guidance for Professor S.Hisajima, Institute of Applied Biochemistry, University of Tsukuba. This work was supported in part by Health Sciences Research Grants for Research on Environmental Health from the Ministry of Health, Labour and Welfare.
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Received September 6, 2001;
revised November 20, 2001;
accepted November 26, 2001.