Drug Metabolism and Disposition Fast Forward
First published on June 6, 2007; DOI: 10.1124/dmd.107.015602
0090-9556/07/3509-1482-1488$20.00
DMD 35:1482-1488, 2007
Kinetic Studies of 25-Hydroxy-19-nor-vitamin D3 and 1
,25-Dihydroxy-19-nor-vitamin D3 Hydroxylation by CYP27B1 and CYP24A1
Naoko Urushino,
Sachie Nakabayashi,
Midori A. Arai,
Atsushi Kittaka,
Tai C. Chen,
Keiko Yamamoto1,
Keiko Hayashi,
Shigeaki Kato,
Miho Ohta,
Masaki Kamakura,
Shinichi Ikushiro, and
Toshiyuki Sakaki
Division of Food Science and Biotechnology, Graduate School of Agriculture, Kyoto University, Kitashirakawa, Oiwake-cho, Sakyo-ku, Kyoto, Japan (N.U.); Department of Biotechnology, Faculty of Engineering, Toyama Prefectural University, Kurokawa, Imizu, Toyama, Japan (S.N., K.H., S.I., M.K., T.S.); Faculty of Pharmaceutical Sciences, Teikyo University, Sagamiko, Kanagawa, Japan (M.A.A., A.K.); Boston University School of Medicine, Boston, Massachusetts (T.C.C.); Institute of Biomaterials and Bioengineering, Tokyo Medical and Dental University, Kanda-Surugadai, Chiyoda-ku, Tokyo, Japan (K.Y.); Institute of Molecular and Cellular Biosciences, Tokyo University, Yayoi, Bunkyo, Tokyo, Japan (S.K.); and Development Nourishment Department, Soai University, Nankonaka, Suminoe, Osaka, Japan (M.O.)
(Received March 5, 2007;
Accepted June 5, 2007)
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Abstract
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Our previous study demonstrated that 25-hydroxy-19-nor-vitamin D3 [25(OH)-19-nor-D3] inhibited the proliferation of immortalized noncancerous PZ-HPV-7 prostate cells similar to 1
,25-dihydroxyvitamin D3 [1,25(OH)2D3], suggesting that 25(OH)-19-nor-D3 might be converted to 1,25-dihydroxy-19-nor-vitamin D3 [1,25(OH)2-19-nor-D3] by CYP27B1 before exerting its antiproliferative activity. Using an in vitro cell-free model to study the kinetics of CYP27B1-dependent 1-hydroxylation of 25(OH)-19-nor-D3 and 25-hydroxyvitamin D3 [25(OH)D3] and CYP24A1-dependent hydroxylation of 1,25(OH)-19-nor-D3 and 1,25(OH)2D3, we found that kcat/Km for 1-hydroxylation of 25(OH)-19-nor-D3 was less than 0.1% of that for 25(OH)D3, and the kcat/Km value for 24-hydroxylation was not significantly different between 1,25(OH)2-19-nor-D3 and 1,25(OH)2D3. The data suggest a much slower formation and a similar rate of degradation of 1,25(OH)2-19-nor-D3 compared with 1,25(OH)2D3. We then analyzed the metabolites of 25(OH)D3 and 25(OH)-19-nor-D3 in PZ-HPV-7 cells by high-performance liquid chromatography. We found that a peak that comigrated with 1,25(OH)2D3 was detected in cells incubated with 25(OH)D3, whereas no 1,25(OH)2-19-nor-D3 was detected in cells incubated with 25(OH)-19-nor-D3. Thus, the present results do not support our previous hypothesis that 25(OH)-19-nor-D3 is converted to 1,25(OH)2-19-nor-D3 by CYP27B1 in prostate cells to inhibit cell proliferation. We hypothesize that 25(OH)-19-nor-D3 by itself may have a novel mechanism to activate vitamin D receptor or it is metabolized in prostate cells to an unknown metabolite with antiproliferative activity without 1-hydroxylation. Thus, the results suggest that 25(OH)-19-nor-D3 has potential as an attractive agent for prostate cancer therapy.
1,25(OH)2D3, the active form of vitamin D3, regulates at least 200 genes, including those involved in cellular proliferation and differentiation, bone and calcium metabolism, immune responses, etc. (Zhang et al., 2005
). Because it can inhibit cancer cell proliferation, 1,25(OH)2D3 has been used to treat cancer patients in clinical trials. However, because systemic administration of 1,25(OH)2D3 can cause hypercalcemia and hypercalciuria, it is not suitable as a therapeutic agent for cancer treatment. Consequently, several laboratories have attempted to synthesize analogs of 1,25(OH)2D3 with less or noncalcemic property (Abe-Hashimoto et al., 1993; Llach et al., 1998; Plum et al., 2004).
Since the discovery of 1-hydroxylase in prostate cells in 1998, several groups have demonstrated that 25(OH)D3 has the ability to inhibit cell proliferation in cultured prostate cells and concluded that 25(OH)D3 was converted to 1,25(OH)2D3 by prostatic CYP27B1 in an autocrine fashion before exerting antiproliferative activity (Barreto et al., 2000; Chen et al., 2000; Hsu et al., 2001). Whitlatch et al. (2002) further established the role of CYP27B1 by demonstrating that LNCaP prostate cancer cells, which were not responsive to 25(OH)D3 inhibition, became responsive after the cells were transfected with the CYP27B1 cDNA plasmid. We recently demonstrated that the growth of PZ-HPV-7 cells, immortalized normal prostate cells that express a high level of CYP27B1 (Wang et al., 2003), could be inhibited in the presence of 25(OH)-19-nor-D3. The data therefore led us to hypothesize that 25(OH)-19-nor-D3 might be converted to 1,25(OH)2-19-nor-D3 within the prostate cells by CYP27B1 and that 1,25(OH)2-19-nor-D3 was responsible for the observed antiproliferative activity (Arai et al., 2005).
We have previously revealed the enzymatic properties of CYP27A1 (Sawada et al., 2000), CYP27B1 (Sakaki et al., 1999b; Sawada et al., 1999; Uchida et al., 2004), CYP24A1 (Sakaki et al., 1999a, 2000; Sawada et al., 2004), and the roles of certain amino acid residues responsible for their enzyme reaction by combination of site-directed mutagenesis and computer modeling (Yamamoto et al., 2005; Hamamoto et al., 2006; Urushino et al., 2006). A remarkable species-based difference was observed in the CYP24A1-dependent metabolism of 1,25(OH)2D3 (Sakaki et al., 2000) and its analogs (Sakaki et al., 2000; Kusudo et al., 2004) between humans and rats.
In this study, we investigated the kinetic characteristics of 1-hydroxylation of 25(OH)D3 and 25(OH)-19-nor-D3 by CYP27B1 and 24-hydroxylation of 1,25(OH)2D3 and 1,25(OH)2-19-nor-D3 by CYP24A1. We found that the kcat/Km for 25(OH)-19-nor-D3 was too low [less than 0.1% of that for 25(OH)D3] to be physiologically significant, whereas the kcat/Km values for 1,25(OH)2-19-nor-D3 and 1,25(OH)2D3 were not significantly different. A docking model for CYP27B1 with 25(OH)-19-nor-D3 was proposed to explain its extremely low activity toward 25(OH)-19-nor-D3. Furthermore, we describe a species-based difference in the CYP24A1-dependent metabolism of 1,25(OH)2-19-nor-D3 between human and rats. To confirm our observed CYP27B1 kinetic data, we then studied the metabolism of 25(OH)D3 and 25(OH)-19-nor-D3 in PZ-HPV-7 prostate cells. Our results indicate that while 25(OH)D3 was hydroxylated to 1,25(OH)2D3,no1,25(OH)2-19-nor-D3 was detected in the prostate cells incubated with 25(OH)-19-nor-D3 under the same culture conditions. Thus, the present data do not support our previous hypothesis that 25(OH)-19-nor-D3 must be converted to 1,25(OH)2-19-nor-D3 by CYP27B1 in prostate cells to exhibit its antiproliferative activity.
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Materials and Methods
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Materials. DNA-modifying enzymes, restriction enzymes, a DNA sequencing kit, and Escherichia coli JM109 (used as a host strain) were obtained from Takara Shuzo Co., Ltd. (Kyoto, Japan). 25(OH)D3 and 1,25(OH)2D3 were purchased from Wako Pure Chemical Industries, Ltd. (Osaka, Japan). [26,27-Methyl-3H]1,25(OH)2D3 (specific activity, 180 Ci/mmol) was purchased from GE Healthcare (Little Chalfont, Buckinghamshire, UK). NADPH was obtained from Oriental Yeast Co. (Tokyo, Japan). Terrific broth was purchased from Invitrogen (Carlsbad, CA). 25(OH)-19-nor-D3 and 1,25(OH)2-19-nor-D3 were synthesized as described previously (Arai et al., 2005). Other chemicals used were of the highest quality commercially available. PZ-HPV-7, a noncancerous prostate cell line, was purchased from the American Type Culture Collection (Manassas, VA).
Substrate Docking. Graphical manipulations were performed using SYBYL 7.1 (Tripos, St. Louis, MO). As reported previously (Urushino et al., 2006), we constructed a three-dimensional model of mouse CYP27B1 by the replacement method using human CYP27B1 as a template (Yamamoto et al., 2005). Based on the experimental results described below, the substrate, 25(OH)D3 with a ß-form of the A-ring, was manually docked into the substrate-binding pocket by superposition with the 25(OH)D3 in human CYP27B1 (Yamamoto et al., 2005), whereas 25(OH)-19-nor-D3 with an -form of the A-ring was docked in the same pocket by superposition with 25(OH)D3, except for the A-ring part with a different conformation. Each substrate-mouse CYP27B1 complex was minimized on Tripos force field 100 iterations.
Constructs of Expression Plasmids. The expression plasmids pKH27A1 (Sawada et al., 2000) for human CYP27A1, pKCHis-m1 (Uchida et al., 2004) for mouse CYP27B1, pKSN24R2 (Sakaki et al., 1999a) for rat CYP24A1, and pKH24 (Sakaki et al., 2000) for human CYP24A1 were constructed as described previously.
Cultivation of the Recombinant E. coli Cells. Recombinant E. coli cells were grown in terrific broth containing 50 µg/mL ampicillin at 26°C under good aeration produced by bubbling. The induction of transcription of cDNAs for CYP27A1, CYP27B1, and CYP24A1 under the control of the tac promoter was initiated by the simultaneous addition of isopropyl-thio-ß-D-galactopyranoside at a final concentration of 1 mM and 
-aminolevulinic acid at a final concentration of 0.5 mM when the cell density (OD660) reached 0.5. The recombinant cells were gently shaken at 26°C under sufficient aeration by bubbling.
Expression of P450s in E. coli. The expression levels of human CYP27A1, mouse CYP27B1, and human and rat CYP24A1 determined by the CO difference spectrum were approximately 1000, 300, 100, and 80 nM culture, respectively. Membrane fractions were prepared from the recombinant E. coli JM109/pKH27A1 cells for human CYP27A1 (Sawada et al., 2000), JM109/pKH24 cells for human CYP24A1, and JM109/pKSN24R2 cells for rat CYP24A1. The human CYP27A1, human CYP24A1, and rat CYP24A1 contents in the membrane fractions were calculated to be 1.10, 0.102, and 0.074 nmol/mg protein, respectively.
Measurement of Reduced CO Difference Spectra. The reduced CO difference spectra were measured with a Shimadzu UV-2200 (Shimadzu, Kyoto, Japan). The concentration of human CYP27A1 and mouse CYP27B1was determined from the reduced CO difference spectrum using a difference of the extinction coefficients at 446 nm and 490 nm of 91 mM-1cm-1 (Omura and Sato, 1964). The absorption coefficient difference between 445 and 490 nm (105 mM-1cm-1) was used for the calculation of the CYP24A1 hemoprotein concentration as described previously (Akiyoshi-Shibata et al., 1994).
Measurement of Enzyme Activity of CYP27A1, CYP27B1, and CYP24A1. CYP27A1-dependent hydroxylation toward 25(OH)-19-nor-D3 and CYP24A1-dependent hydroxylation toward 1,25(OH)2-19-nor-D3 were measured in the reconstituted system containing the membrane fraction, whereas CYP27B1-dependent hydroxylation toward 25(OH)-19-nor-D3 was measured in the reconstituted system containing 0.1% CHAPS as described previously (Uchida et al., 2004). The activities were measured under the conditions as follows: 1) 0.32 µM human CYP27A1, 2.0 µM ADX, 0.02 µM ADR, and 2.0 µM 25(OH)-19-nor-D3; 2) 0.05 µM mouse CYP27B1, 2.0 µM ADX, 0.02 µM ADR, and 0 to 2.0 µM 25(OH)-19-nor-D3 or 25(OH)D3;3) 0.02 µM rat or human CYP24A1, 5.0 µM ADX, 0.5 µM ADR, and 5.0 µM 1,25(OH)2-19-nor-D3; and 4) 0.002 µM human CYP24A1, 0.1 µM ADX, 0.01 µM ADR, 0 to 1.6 µM1,25(OH)2-19-nor-D3 in 100 mM Tris-HCl (pH 7.4), and 1 mM EDTA at 37°C.
For analyzing the products of CYP24A1-dependent 1,25(OH)2-19-nor-D3 hydroxylation, the third condition as defined above was used. On the other hand, for the determination of kinetic parameters, the reaction was performed under the fourth condition containing extremely small amounts of ADX and ADR to avoid successive reaction by CYP24A1 as described previously (Hamamoto et al., 2006). The reaction was initiated by the addition of NADPH at a final concentration of 0.5 mM. Aliquots of the reaction mixture were collected at various time intervals and extracted with 4 volumes of chloroform/methanol (3:1). The organic phase was recovered and dried down under reduced pressure. The resultant residue was dissolved in acetonitrile and applied to HPLC under the following conditions: column, YMC-Pack ODS-AM (5 µm; 4.6 mm x 300 mm) (YMC Co., Kyoto, Japan); UV detection, 265 nm; flow rate, 1.0 ml min-1; column temperature, 40°C; mobile phase, linear gradient of 70 to 100% acetonitrile aqueous solution per 15 min followed by 100% acetonitrile for 10 min for the metabolism of 25(OH)D3, and 20 to 100% acetonitrile aqueous solution per 25 min followed by 100% acetonitrile for 10 min for the metabolism of 1,25(OH)2D3.
LC/MS Analysis of the Metabolites. Isolated metabolites from HPLC effluents were subjected to mass spectrometric analysis using a Finnegan Mat TSQ-70 with atmospheric pressure chemical ionization, positive mode. The conditions of liquid chromatography were as follows: column, reverse phase ODS column (µBondapak C18, 5 µm; Waters, Milford, MA) (6 mm x 150 mm); mobile phase, 80% methanol aqueous solution per 25 min; flow rate, 1.0 ml min-1; UV detection, 265 nm.
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FIG. 1. HPLC profiles of 25(OH)-19-nor-D3 and its metabolites generated by mouse CYP27B1 (A) and human CYP27A1 (B). The metabolites were designated as M1, M2, and M3.
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Binding Assay for Calf Thymus VDR. Displacement of [3H]1,25(OH)2D3 from calf thymus cytosol receptor (Yamasa Shoyu, Choshi, Japan) by the metabolites of 25(OH)D3-19-nor-D3 following its incubation with CYP27B1 was determined as described previously (Sawada et al., 2004). Various amounts of 1,25(OH)2D3,1,25(OH)2-19-nor-D3, and the metabolites of 25(OH)-19-nor-D3 in 20 µl of ethanol were added to 500 µl of the calf thymus cytosol diluted with 50 mM potassium phosphate buffer (pH 7.4) containing 0.3 M KCl and incubated for 1 h at 20°C. Next, 34 fmol of [3H]1,25(OH)2D3 in 25 µl of ethanol was added and incubated for 1 h at 20°C. The 200 µl of dextran/charcoal (0.05% dextran T-150, 0.5% Charcoal Decolorizing Neutral; Yamasa Shoyu) in 50 mM sodium phosphate buffer (pH 7.5) was added to the mixture, followed by centrifugation at 1600g for 10 min at 4°C to separate bound from free [3H]1,25(OH)2D3. The radioactivity in the supernatant was measured with a liquid scintillation counter.
Metabolism of 25(OH)D3 and 25(OH)-19-nor-D3 in Prostate Cells. PZ-HPV-7 cells were grown on 100-mm culture dishes and maintained on a serum free-defined medium as described previously (Young et al., 2004). When cells reached confluence, they were treated with either 1 µM 25(OH)D3 or 25(OH)-19-nor-D3 for 2, 4, 8, and 12 h. At the end of treatment, media were removed, and cells were extracted twice with methanol. A dish of untreated cells was also extracted with methanol and served as a background control. The two methanol fractions were combined and dried down under a stream of nitrogen gas. The resultant residue was redissolved in acetonitrile and applied to HPLC under the following conditions: column, YMC-Pack ODS-AM (5 µm; 4.6 mm x 300 mm) (YMC Co.); UV detection, 265 nm; flow rate, 1.0 ml min-1; column temperature, 40°C; mobile phase, linear gradient of 70 to 100% acetonitrile aqueous solution per 15 min and 100% acetonitrile for 10 min.
Other Methods. The concentration of vitamin D3 and 19-nor-D3 derivatives was determined by their molar extinction coefficient of 1.8 x 104 M-1cm-1 at 264 nm (Sakaki et al., 1999b) and 1.7 x 104 M-1cm-1 at 254 nm (Harmata and Barnes, 1990), respectively.
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Results
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Metabolism of 25(OH)-19-nor-D3 by CYP27B1 and CYP27A1. The reconstituted system containing CYP27B1 or CYP27A1 together with ADX and ADR was examined for the metabolism of 25(OH)-19-nor-D3. Figure 1 shows the HPLC profiles of the substrates and their metabolites produced by CYP27B1 and CYP27A1 catalyses. Unexpectedly, three metabolites, M1, M2, and M3, were observed in CYP27B1-dependent metabolism. Retention time of M1 coincided with that of 1,25(OH)2-19-nor-D3, suggesting that CYP27B1 can catalyze 1-hydroxylation toward not only 25(OH)D3 but also 25(OH)-19-nor-D3. However, the putative 1-hydroxylated metabolite is a minor product, and the major metabolite is M3, which is a single metabolite produced by CYP27A1.
Affinity of the Metabolite M1 for VDR. The calf thymus VDR-binding assay demonstrated that 1,25(OH)2D3-19-nor-D3 had significantly lower affinity than 1,25(OH)2D3. The concentrations of 1,25(OH)2D3 and 1,25(OH)2D3-19-nor-D3 for the 50% B/B0 binding were 19 and 480 pM, respectively. Thus, the affinity of 1,25(OH)2-19-nor-D3 for VDR was estimated to be 4% compared with the affinity of 1,25(OH)2D3. As shown in Fig. 2, the metabolite M1 produced by CYP27B1 showed an affinity for VDR similar to 1,25(OH)2-19-nor-D3. The concentration of M1 for 50% B/B0 binding was 590 pM. HPLC analysis and VDR assay strongly suggest that M1 may be 1,25(OH)2-19-nor-D3. Unexpectedly, the metabolite M2 showed a VDR affinity similar to M1, whereas M3 showed no affinity for VDR (data not shown).
Kinetics of CYP27B1-Dependent Metabolism of 25(OH)-19-nor-D3. When the activity of CYP27B1 toward 25(OH)2D3 was measured at the substrate concentrations of 0 to 2.0 µM, the reaction followed Michaelis-Menten-type kinetics. The kinetic parameters were calculated by the nonlinear regression analysis using Kaleida-Graph (Synergy Software, Reading, PA) (Table 1). However, the activity toward 25(OH)-19-nor-D3 did not show Michaelis-Menten-type kinetics but linearly increased with increasing substrate concentration. This result suggests that the Km value is much higher than the highest substrate concentration used in this study (2 µM), and the Michaelis-Menten equation v = kcat[E0]S/(Km + S) is simplified to v = kcat[E0]S/Km. Although kcat and Km are not estimated separately, kcat/Km can be calculated as shown in Table 1. Note that kcat/Km for M1 production from 25(OH)-19-nor-D3 is approximately 0.035% of kcat/Km for 1,25(OH)2D3 formation from 25(OH)D3. Based on these results, we assume that conformation of 25(OH)-19-nor-D3 in the substrate-binding pocket of CYP27B1 is significantly different from that of 25(OH)D3. Figure 3 shows our proposed model, which demonstrates that 25(OH)D3 prefers the ß-form, whereas 25(OH)-19-nor-D3 prefers the -form. Distances between hydrogen at the C-1 position and heme iron for 25(OH)D3 and 25(OH)-19-nor-D3 were calculated to be 3.9 and 5.3 Å, respectively, resulting in the extremely low activity of CYP27B1 for 25(OH)-19-nor-D3. This conclusion was supported by the finding that no 1,25(OH)2-19-nor-D3 was detected in cell extracts obtained from PZ-HPV-7 cells incubated with 1 µM 25(OH)-19-nor-D3 for 2 or 12 h at 37°C (data not shown).
Metabolism of 1,25(OH)2-19-nor-D3 by CYP24A1. The reconstituted system containing human CYP24A1 or rat CYP24A1, ADR, and ADX was examined for the metabolism of 1,25(OH)2-19-nor-D3. Figure 4 shows HPLC profiles of 1,25(OH)2-19-nor-D3 as the substrate and its metabolites obtained by incubation with human CYP24A1 and rat CYP24A1. Note that the HPLC profiles derived from incubating 1,25(OH)2-19-nor-D3 with human CYP24A1 resembles those of 3-epi-1,25(OH)2D3 metabolites incubated with human CYP24A1 (Kusudo et al., 2004). LC/MS analysis suggests that at least eight metabolites were produced by human CYP24A1. On the other hand, five metabolites were observed in the metabolism by rat CYP24A1. Thus, significant species difference was observed in CYP24A1-dependent metabolism of 1,25(OH)2-19-nor-D3 between humans and rats. The metabolites numbered 1 through 8 were considered to be 23,26(OH)2, 24-oxo-23(OH), tetranor-23(OH)-, 23(OH)-, 24(OH)-, 23-oxo, 24-oxo-, and 25,26,27-trinor-24-ene derivatives of 1,25(OH)2-19-nor-D3, respectively, based on HPLC pattern compared with the corresponding metabolites of 3-epi-1,25(OH)2D3 (Kusudo et al., 2004).
To confirm the chemical structures of the metabolites, we collected the metabolites in the effluents from HPLC and subjected them to mass spectrometric analysis. Relative intensities (%) of major ion fragments of the metabolites were as follows: M1: m/z 365 (M+H-4H2O), 29%; m/z 383 (M+H-3H2O), 27%; m/z 401 (M+H-2H2O), 100%; m/z 419 (M+H-H2O), 48%; m/z 437 (M+H) 29%, 466 (M+H +29), 11% of which are consistent with our assumption that M1 is 1,23,25,26(OH)4-19-nor-D3. M2: m/z 399 (M+H-2H2O), 60%; m/z 417 (M+H-H2O), 100%; m/z 435 (M+H), 56%; m/z 464 (M+H + 29), 28% of which are consistent with our assumption that M2 is 24-oxo-1,23,25(OH)3-19-nor-D3. M3: m/z 331 (M+H-H2O), 100%; m/z 349 (M+H), 41%; m/z 385 (M+H + 29), 20% of which is consistent with our assumption that M3 is 24,25,26,27-tetranor-1,23,25(OH)3-19-nor-D3. The mixed metabolites M4 and M5: m/z 293, 7%; m/z 311, 16%; m/z 329, 22%; m/z 349 (M+H-4H2O), 7%; m/z 367 (M+H-3H2O), 42%; m/z 385 (M+H-2H2O), 100%; 403 (M+H-H2O), 65%; m/z 421 (M+H), 30%; m/z 450 (M+H + 29), 11% of which are consistent with our assumption that M4 and M5 are 1,23,25(OH)3-19-nor-D3 and 1,24,25(OH)3-19-nor-D3, respectively. The fragment ions at 329, 311 (329-H2O), and 293 (329-2H2O), which result from the cleavage between C-23 and C-24, are characteristic of the 23-hydroxylated compound as described by Horst et al. (1983). The corresponding metabolites produced by rat CYP24A1 shown in Figure 4 contained a faint amount of these ions, suggesting that the ratio of 23-hydroxylated compound to 24-hydroxylated compound is significantly low. M6: m/z 325 (343-H2O), 30%; m/z 343, 49%; 383 (M+H-2H2O), 39%; 401 (M+H-H2O), 58%; m/z 419 (M+H) 100%, which is consistent with our assumption that M6 is 23-oxo-1,25(OH)2-19-nor-D3. The fragment ions at 343 and 325 (343-H2O), which result from the cleavage between C-24 and C-25, are characteristic of the 23-oxo compounds as described by Horst et al. (1983). M7: m/z 365 (M+H-3H2O), 42%; m/z 383 (M+H-2H2O), 100%; m/z 401 (M+H-H2O), 99%; m/z 419 (M+H), 57%; m/z 448 (M+H + 29), 32%, which is consistent with our assumption that M7 is 24-oxo-1,25(OH)2-19-nor-D3. M8: m/z 309 (M+H-2H2O), 59%; m/z 327 (M+H-H2O), 100%; m/z 345 (M+H), 92%; m/z 374 (M+H + 29), 50%, which is consistent with our assumption that M8 is 25,26,27-trinor-23-ene-1 (OH)-19-nor-D3. Based on these data, metabolic pathways of 1,25(OH)2-19-nor-D3 by human CYP24A1 are proposed in Fig. 5, although we have not detected the final products of C-23 and C-24 pathways.
Kinetic Analysis of the Metabolism of 1,25(OH)2D3 and 1,25(OH)2-19-nor-D3 by Human CYP24A1. When the CYP24A1 activity toward 1,25(OH)2D3 and 1,25(OH)2-19-nor-D3 was measured at the substrate concentrations of 0 to 2.0 µM, the reaction followed Michaelis-Menten-type kinetics. The kinetic parameters were calculated by the nonlinear regression analysis using Kaleida-Graph (Synergy Software) (Table 2). The Km and kcat values for the human CYP24A1 were calculated to be 1.60 µM and 2.25 min-1 for 1,25(OH)2-19-nor-D3 and 0.26 µM and 0.39 min-1 for 1,25(OH)2D3, respectively. Although both values for 1,25(OH)2-19-nor-D3 were significantly higher than those for 1,25(OH)2D3, the kcat/Km value for 1,25(OH)2D3-19-nor-D3 was not significantly different from that for 1,25(OH)2D3. Thus, it is possible that 1,25(OH)2-19-nor-D3 is as good a substrate as 1,25(OH)2D3 for human CYP24A1.
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Discussion
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A large number of vitamin D analogs have been synthesized and studied for potential clinical application. (Binderup et al., 1991; Bishop et al., 1994; Bouillon et al., 1995; Yamada et al., 2003). Among the analogs, A-ring-modified 19-nor vitamin D compounds have received most of the attention in recent years (DeLuca et al., 2005). Kittaka's group recently synthesized A-ring-modified vitamin D3 analogs and demonstrated that this class of analogs had unique biological activity (Kittaka et al., 2000; Konno et al., 2000; Suhara et al., 2001; Ono et al., 2003; Arai et al., 2005) and can alter the VDR-coactivator interaction, resulting in selective potentiation of the transcription function (Fujishima et al., 2003; Arai and Kittaka, 2006). One of the 19-nor vitamin D analogs, 1,25-dihydroxyvitamin 19-nor-D2 [1,25(OH)2-19-nor-D2], a synthetic analog of 1,25(OH)2D2, is approved by the Food and Drug Administration for the treatment of secondary hyperparathyroidism. Several randomized controlled clinical trials have shown that 1,25(OH)2-19-nor-D2 is noncalcemic (Schwartz et al., 2005). Because of its noncalcemic property and its structural similarity to 1,25(OH)2D3, a series of studies was initiated to compare the antiproliferative activity of 1,25(OH)2-19-nor-D2 against 1,25(OH)2D3 in primary cultures and cell lines of human prostate cancer, as well as its VDR transactivation potential in a prostate cancer cell line, PC-3, that was stably transfected with VDR (Chen et al., 2000). In addition, because prostate cells express CYP27B1 and can convert 25(OH)D3, a prohormone, to 1,25(OH)2D3 within the cells, avoiding the problem of systemic hypercalcemia, the antiproliferative and VDR transactivation activities between 25(OH)D3 and 1,25(OH)2D3 were also compared. The studies confirmed that the behavior of 1,25(OH)2-19-nor-D2 and 25(OH)D3 in prostatic cells was similar to that of 1,25(OH)2D3. Thus, 25(OH)D3 and 1,25(OH)2-19-nor-D2 would be attractive candidates for human clinical trials in prostate cancer treatment, especially since both drugs have been approved by the Food and Drug Administration for human use (e.g., for treating vitamin D deficiency due to liver disease, or secondary hyperparathyroidism). Along this line, our laboratory has synthesized and studied a prodrug of the 19-nor vitamin D analog, 25(OH)-19-nor-D3, and showed that it had antiproliferative activity similar to 1,25(OH)2D3 in immortalized prostate cells. Thus, 25(OH)-19-nor-D3 would be attractive for prostate cancer treatment. The data also lead us to conclude that 25(OH)-19-nor-D3 must be hydroxylated to 1,25(OH)2-19-nor-D3, catalyzed by CYP27B1, which is present in high concentration in PZ-HPV-7 cells, and that 1,25(OH)2-19-nor-D3, not 25(OH)-19-nor-D3 itself, is responsible for the inhibitory effect of 25(OH)-19-nor-D3 on prostate cell growth. This conclusion was supported by the findings that LNCaP cells were not growth-inhibited in the presence of 25(OH)D3 (Skowronski et al., 1995), but the cells became responsive to 25(OH)D3 after the transfection of an expression plasmid for CYP27B1 to induce the synthesis of 1,25(OH)2D3 (Whitlatch et al., 2002). To prove whether this is the case for 25(OH)-19-nor-D3, in this report we first performed kinetic analysis of CYP27B1 on the metabolism of 25(OH)-19-nor-D3 using a cell-free model system established in our laboratory (Uchida et al., 2004). Because we have not successfully overproduced human CYP27B1 in E. coli cells (Sawada et al., 1999), we used mouse CYP27B1 in this study. Three metabolites of 25(OH)-19-nor-D3, designated as M1, M2, and M3, were detected in the HPLC chromatograms (Fig. 1). Among them, M1, a minor peak, had retention time identical to standard 1,25(OH)2-19-nor-D3. Analyses by mass spectroscopy and VDR-binding affinity assay (Fig. 2) further confirmed that M1 is 1,25(OH)2-19-nor-D3. Interestingly, no M1 metabolite was detected in the HPLC chromatogram obtained from the methanol extracts of PZ-HPV-7 cells incubated with 10-6 M 25(OH)-19-nor-D3 for up to 12 h (data not shown), indicating no 1-hydroxylation by CYP27B1 or rapid metabolism of the produced 1,25(OH)2-19-nor-D3 in the prostate cells in cultures.
To explain the lack of enzymatic conversion of 25(OH)-19-nor-D3 to 1,25(OH)2-19-nor-D3 by CYP27B1, we performed AutoDock (Goodsell et al., 1996) to predict the bound conformations of 25(OH)D3 and 25(OH)-19-nor-D3 to CYP27B1. Our docking model of CYP27B1 with 25(OH)D3 suggests that 25(OH)D3 binds to the substrate-binding pocket of CYP27B1 in a ß-form, which is the most favorable configuration. If 25(OH)-19-nor-D3 were in the same ß-form, the distance between the heme iron and hydrogen at the C-1 position of 25(OH)-19-nor-D3 would be nearly the same as that for 25(OH)D3, and similar kcat/Km values would have been obtained for the enzyme regardless of whether 25(OH)D3 or 25(OH)-19-nor-D3 was used as its substrate. To explain such an extremely low activity of CYP27B1 toward 25(OH)-19-nor-D3, we hypothesize that 25(OH)-19-nor-D3 may bind to the substrate-binding pocket of CYP27B1 not in ß-form but in -form. As shown in Fig. 3, the distance between heme iron and hydrogen at the C-1 position of 25(OH)-19-nor-D3 is 5.3 Å, which is significantly longer than the distance between heme iron and hydrogen at the C-1 position of 25(OH)D3 (3.9 Å). Note that the distances between heme iron and hydrogens at the C-1ß and C-2 positions of 25(OH)-19-nor-D3 are 4.7 and 4.4 Å, respectively. Based on the docking model shown in Fig. 3B, we could speculate that the metabolite M2 is 1ß,25(OH)2-19-nor-D3 or 2,25(OH)2-19-nor-D3. Even if M2 is taken into consideration, the kcat/Km value for 25(OH)-19-nor-D3 is less than 0.1% of 25(OH)D3 (Table 2). In addition, the affinity of M1 and M2 for VDR is only 4% of 1,25(OH)2D3, and the major metabolite M3 showed no affinity for VDR. It is likely that more than 60% of the total metabolites of 25(OH)-19-nor-D3 is M3, which showed no affinity for VDR.
Next we considered the possibility that extremely low catabolism of the resultant 1,25(OH)2-19-nor-D3 through the CYP24A1-dependent pathway might explain the high antiproliferative activity of 25(OH)-19-nor-D3. As initially reported by Miller et al. (1995), normal prostate cells and prostate cancer cells express high levels of CYP24A1. Thus, we examined CYP24A1-dependent metabolism of 1,25(OH)2-19-nor-D3 in our cell-free model. Although both Km and kcat values for 1,25(OH)2-19-nor-D3 are considerably higher than those for 1,25(OH)2D3, the ratio kcat/Km is not significantly different between 1,25(OH)2-19-nor-D3 and 1,25(OH)2D3, suggesting that 1,25(OH)2-19-nor-D3 is metabolized to the same extent as 1,25(OH)2D3 by CYP24A1. Thus, it is possible that a small amount of 1,25(OH)2-19-nor-D3 produced by CYP27B1 in PZ-HPV-7 cells is rapidly metabolized by CYP24A1 (Miller et al., 1995), as shown in Fig. 6. Based on the analysis of metabolites, the metabolic pathways of 1,25(OH)2-19-nor-D3 seemed quite similar to those of 3-epi-1,25(OH)2D3 (Kusudo et al., 2004). Although the ratio between the C-23 and C-24 oxidation pathways in the metabolism of the native form of 1,25(OH)2D3 with the 3ß-hydroxyl group is approximately 1:4, the ratio for 1,25(OH)2-19-nor-D3 is 1:0.8, similar to 3-epi-1 ,25(OH)2D3, which has a 3-hydroxyl group (Kusudo et al., 2004). Note that the major circulating, prohormonal form of vitamin D, 25(OH)D3,hasa3ß-hydroxyl group. The distance from the activated oxygen atom to C-23 and C-24 seems to determine the ratio between the C-23 and C-24 oxidation pathways. Thus, it is possible that 1,25(OH)2-19-nor-D3 has a conformation somewhat different from 1,25(OH)2D3 in the substrate-binding pocket of human CYP24A1. The rapid metabolism of 1,25(OH)-19-nor-D3 by CYP24A1 eliminates the possibility that extremely low catabolism of the resultant 1,25(OH)2-19-nor-D3 through the CYP27B1-dependent pathway is responsible for the high antiproliferative activity of 25(OH)-19-nor-D3.
In summary, the present results demonstrate the absence of CYP27B1-dependent 1-hydroxylation toward 25(OH)-19-nor-D3 by kinetic and metabolic studies. Thus, the findings that 25(OH)-19-nor-D3 exerts its antiproliferative activity without further conversion to 1,25(OH)2-19-nor-D3, in conjunction with the report by Whitlatch et al. (2002) that prostate cancer cells have no or reduced CYP27B1 activity, suggest that 25(OH)-19-nor-D3 could be an attractive therapeutic and chemopreventive agent for prostate cancer.
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Acknowledgments
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We express gratitude to Hideki Hara and Ryuji Tsutsumi of Teikyo University for the synthesis of 25(OH)-19-nor-D3 and 1,25(OH)2-19-nor-D3.
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Footnotes
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This work was partly supported by Health Science Research Grants from the Ministry of Health Labour and Welfare of Japan and a Grant-in-Aid for Scientific Research from the Ministry of Education, Science, Sports and Culture of Japan.
doi:10.1124/dmd.107.015602.
ABBREVIATIONS: 1,25(OH)2D3,1,25-dihydroxyvitamin D3; 25(OH)D3, 25-hydroxyvitamin D3; 25(OH)-19-nor-D3, 25-hydroxy-19-nor-vitamin D3;1,25(OH)2-19-nor-D3,1,25-dihydroxy-19-nor-vitamin D3;1,25(OH)2-19-nor-D2,1,25-dihydroxyvitamin 19-nor-D2; CHAPS, 3-[(3-cholamidopropyl)dimethylammonio]propanesulfonate; ADX, adrenodoxin; ADR, NADPH-adrenodoxin reductase; HPLC, high-performance liquid chromatography; ODS, octadecylsilane; AM, acetoxymethyl ester; LC/MS, liquid chromatography/mass spectometry; VDR, vitamin D receptor.
1 Current affiliation: Laboratory of Drug Design and Medicinal Chemistry, Showa Pharmaceutical University, Higashi-tamagawagakuen, Machida, Tokyo, Japan. 
Address correspondence to: Dr. Toshiyuki Sakaki, Department of Biotechnology, Faculty of Engineering, Toyama Prefectural University, 5180 Kurokawa, Imizu, Toyama 939-0398, Japan. E-mail: tsakaki{at}pu-toyama.ac.jp
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Abstract
Materials and Methods
),1
), and the metabolite M1 (
) shown in 
