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Department of Basic Pharmaceutical Sciences (A.J.L., B.T.Z.), College of Pharmacy, University of South Carolina, Columbia, South Carolina 29208; and Susan Lehman Cullman Laboratory for Cancer Research (M.X.C., P.E.T., A.H.C.), Department of Chemical Biology, College of Pharmacy, Rutgers, The State University of New Jersey, Piscataway, New Jersey 08854
Address all correspondence and requests for reprints to: Bao Ting Zhu, Department of Basic Pharmaceutical Sciences, College of Pharmacy, University of South Carolina, Columbia, South Carolina 29208. E-mail: BTZhu{at}cop.sc.edu.
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-, 6-, 4-, and 7-hydroxylation. However, when estrone was the substrate, CYP1A1 formed more 4-hydroxyestrone than 15- or 6-hydroxyestrone, with 2-hydroxyestrone as the major metabolite. CYP1A2 had the highest activity for the 2-hydroxylation of both 17ß-estradiol and estrone, although it also had considerable activity for their 4-hydroxylation (9–13% of 2-hydroxylation). CYP1B1 mainly catalyzed the formation of catechol estrogens, with 4-hydroxyestrogens predominant. CYP2A6, 2B6, 2C8, 2C9, 2C19, and 2D6 each showed a varying degree of low catalytic activity for estrogen 2-hydroxylation, whereas CYP2C18 and CYP2E1 did not show any detectable estrogen-hydroxylating activity. CYP3A4 had strong activity for the formation of 2-hydroxyestradiol, followed by 4-hydroxyestradiol and an unknown polar metabolite, and small amounts of 16- and 16ß-hydroxyestrogens were also formed. The ratio of 4- to 2-hydroxylation of 17ß-estradiol or estrone with CYP3A4 was 0.22 or 0.51, respectively. CYP3A5 had similar catalytic activity for the formation of 2- and 4- hydroxyestrogens. Notably, CYP3A5 had an unusually high ratio of 4- to 2-hydroxylation of 17ß-estradiol or estrone (0.53 or 1.26, respectively). CYP3A4 and 3A5 also catalyzed the formation of nonpolar estrogen metabolite peaks (chromatographically less polar than estrone). CYP3A7 had a distinct catalytic activity for the 16-hydroxylation of estrone, but not 17ß-estradiol. CYP4A11 had little catalytic activity for the metabolism of 17ß-estradiol and estrone. In conclusion, many human CYP isoforms are involved in the oxidative metabolism of 17ß-estradiol and estrone, with a varying degree of catalytic activity and distinct regioselectivity. |
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Several earlier studies have examined the catalytic activity of a number of CYP isoforms from animals and humans for the NADPH-dependent oxidative metabolism of E2 and E1 (21, 22, 23, 24, 25, 26, 27, 28, 29, 30). In most of these studies, usually only a few estrogen metabolites (e.g. products of estrogen 2-, 4-, and 16- hydroxylation) were determined. However, it is now known that a large number of hydroxylated or keto metabolites of E2 and/or E1 are present in biological samples (e.g. urine, tissues, and blood) or formed during in vitro incubations of estrogen substrates with microsomal enzymes from either animals or humans (1). Some of the estrogen metabolites (such as 4-OH-E2, 15-OH-E2, 16-OH-E1, and 2-methoxyestradiol) may have unique biological functions that are not associated with their parent hormones E2 and E1 (1, 31, 32, 33). It is, therefore, important to characterize the complete profiles of the hydroxylated or keto metabolites of endogenous E2 and E1 that are formed by each of the human CYP isoforms present in liver and particularly those selectively present in estrogen target tissues or cells, and also to identify the potential biological functions associated with these estrogen metabolites. Detailed knowledge on this would be useful for our better understanding of their biological functions in different target cells in which some of these CYP isoforms are selectively expressed under certain physiological or pathological conditions.
Recently, we have characterized a large number of oxidative metabolites of E2 and E1 formed by human liver (34, 35) and human term placenta (36). We describe here our data on the systematic characterization of the NADPH-dependent metabolites of E2 and E1 formed by 15 human CYP isoforms. These human CYP isoforms were selectively expressed in insect cells that were transfected with a baculovirus expression system containing the cDNA for each of the desired human CYP isoforms. Our data demonstrate that a variety of human CYP isoforms are catalytically active for the NADPH-dependent oxidative metabolism of E2 and E1 to multiple metabolites, and many of them have distinct regioselectivity for the catalysis of metabolic reactions.
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-OH-E2, 12ß-OH-E2, 12-keto-E2, 14-OH-E1, 14-OH-E2, 15-OH-E2, and 15ß-OH-E2 were described in an earlier paper (37). Several hydroxy-E1 metabolites, including 6-OH-E1, 7-OH-E1, 7ß-OH-E1, 12ß-OH-E1, 15-OH-E1, 15ß-OH-E1, and 16ß-OH-E1, were biosynthetically prepared in our laboratory from their respective hydroxy-E2 metabolites through incubations with human liver microsomes in the presence of NAD+ as cofactor. Each of the products formed was extracted with ethyl acetate and then isolated by HPLC (described in HPLC analysis of [3H]E2 or [3H]E1metabolites). The reference compounds for all other estrogen metabolites used in this study were obtained from Steraloids, Inc. (Newport, RI). N,O-bis(Trimethylsilyl)trifluoroacetamide containing 1% trimethylchlorosilane was obtained from Pierce Chemical Co. (Rockford, IL). Most of the organic solvents used in this study were of HPLC grade or better, and they were obtained from Fisher Scientific (Atlanta, GA). [2,4,6,7,16,17-3H]E2 and [2,4,6,7-3H]E1 (numerically labeled, specific radioactivity of 123 and 100 Ci/mmol, respectively) were purchased from Perkin-Elmer Life Sciences (Boston, MA). There was no published information available that would enable us to ascertain whether each of the designated positions was evenly labeled. However, a comparison of several tritium-labeled E2 and E1 products such as [6,7-3H]E2, [2,4,6,7,-3H]E2, and [2,4,6,7,16,17-3H]E2 prepared by the same company showed that their highest specific activities (curies per millimole) increased almost proportionally with increasing positions labeled with tritium, which suggested that each position likely was quite evenly labeled. In addition, it should be noted that when we compared the estrogen metabolites formed by various human CYP isoforms with 20 µM of [2,4,6,7,16,17-3H]E2 or [4-14C]E2 as substrate, the overall profiles of the major E2 metabolites formed were found to be highly similar, but most of the quantitatively minor metabolites of E2 could not be detected with [4-14C]E2 because of its low specific radioactivity.
Selectively expressed human CYP isoforms
The selectively expressed human CYP isoforms were obtained from BD Gentest Co. (Woburn, MA). These human CYP isoforms were expressed in insect cells that were selectively transfected with a baculovirus (Autographa californica) expression system containing the cDNA for each of the desired human CYP isoforms. The total microsomal protein concentration, CYP content, CYP reductase activity, and cytochrome b5 content, and the specific catalytic activity for the marker substrate for each expressed CYP isoform in the microsomes are summarized in Table
1.
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3 µCi of [3H]E2 or [3H]E1), 2 mM NADPH, and 5 mM ascorbic acid in a final volume of 0.5 ml of 0.1 M Tris-HCl buffer (pH 7.4) containing 0.05 M HEPES. The presence of 5 mM ascorbic acid in the incubation mixture has previously been shown to protect catechol estrogen metabolites from oxidative degradation without significantly altering the enzyme activity (4). The enzymatic reaction was initiated by addition of microsomes, and the incubations were carried out at 37 C for 20 min with mild shaking. The microsomal reaction was arrested by placing test tubes on ice and was then immediately extracted with 8 ml of ethyl acetate. The organic supernatants were transferred to another set of test tubes and dried under a stream of nitrogen. The resulting residues were redissolved in 100 µl of methanol, and a 40-µl aliquot was injected into the HPLC for analysis of estrogen metabolite composition. It should also be noted that all of the glass test tubes used in our study were silanized with 5% (vol/vol) dimethyldichlorosilane in toluene for 10 min, followed by rinses in pure toluene once and pure methanol twice. The test tubes were allowed to dry at room temperature and then were thoroughly rinsed with distilled water. Our earlier analyses of the NADPH-dependent [3H]E2 metabolism by human and rat liver microsomes using seven different types of unsilanized glass test tubes obtained from three different manufacturers showed that even under exactly the same incubation, extraction, and HPLC analytical conditions, the results were very different for each of the hydroxylated [3H]E2 metabolites detected. Based on measuring the radioactivity associated with [3H]2-OH-E2 and [3H]4-OH-E2 peaks, their overall recoveries with unsilanized test tubes were only 30–67% of the recoveries with the silanized test tubes. The reason for the increased recoveries of hydroxyestrogen metabolites with silanized glass tubes likely was because pretreatment of the glassware surface with dimethyldichlorosilane deactivated the active chemical groups, thereby reducing physical adsorption of the hydroxylated estrogen metabolites to the test tubes (38).
HPLC analysis of [3H]E2 or [3H]E1 metabolites
Analysis of [3H]E2 and [3H]E1 metabolites was performed with an HPLC system coupled with in-line UV and radioactivity detections as described earlier (37). The HPLC system consisted of a Waters 2690 separation module (Milford, MA), a Waters UV detector (model 484), an IN/US ß-RAM radioactivity detector (Tampa, FL), and an Ultracarb 5 ODS column (150 x 4.60 mm, Phenomenex, Torrance, CA). The solvent system for separation of E2, E1, and their metabolites consisted of acetonitrile (solvent A), 0.1% acetic acid in water (solvent B), and 0.1% acetic acid in methanol (solvent C). The solvent gradient (solvent A/solvent B/solvent C) used for eluting estrogen metabolites was as follows: 8 min of isocratic at an initial composition of 16:68:16, 7 min of a concave gradient (curve number 9) to 18:64:18, 13 min of a concave gradient (curve number 8) to 20:59:21, 10 min of a convex gradient (curve number 2) to 22:57:21, 13 min of a concave gradient (curve number 8) to 58:21:21, followed by a 0.1-min step to 92:5:3 and a 8.9-min isocratic period at 92:5:3. The gradient was then returned to the initial composition (16:68:16) and held for 10 min before analysis of the next sample.
The HPLC retention times for all authentic estrogen metabolites were determined by using in-line UV detection, whereas the [3H]E2 or [3H]E1 metabolite peaks formed with selectively expressed human CYP isoforms were determined by using in-line radioactivity detection. The calculation of the amount of each estrogen metabolite formed was based on the amount of radioactivity detected for each corresponding metabolite peak. Here, it should also be noted that CYP isoform-mediated formation of hydroxylated or keto metabolites of [3H]E2 or [3H]E1 at any of their 3H-labeled positions (namely, 2, 4, 6, 7, 16, and 17 for [3H]E2 and 2, 4, 6, and 7 for [3H]E1) was known to remove tritium from the substrate, resulting in the formation of [3H]H2O. In the present study, therefore, the calculated final rates for the formation of hydroxylated or keto metabolites at the 3H-labeled positions were adjusted according to the estimated loss of radioactivity in each of these products. The rate for the formation of an estrogen metabolite by an expressed CYP isoform was expressed as "picomoles of the estrogen metabolite formed per nanomole of CYP isoform per minute," abbreviated as "pmol/nmol·min."
Structural identification of E2 or E1 metabolites formed by selectively expressed human CYP isoforms
The identity of most of E2 or E1 metabolites formed by selectively expressed CYP isoforms was confirmed through comparisons of their HPLC retention times, gas chromatography/mass spectrometry (GC/MS) retention times, and mass fragmentation spectra with all authentic reference compounds. We had a total of 49 authentic metabolites of E2 and E1, which included: 2-OH-E1, 2-OH-E2, 2-OH-E3, 2-methoxy-E1, 2-methoxy-E2, 4-OH-E1, 4-OH-E2, 4-methoxy-E1, 4-methoxy-E2, 6-OH-E1, 6-OH-E2, 6ß-OH-E1, 6ß-OH-E2, 6-keto-E1, 6-keto-E2, 6-keto-E3, 6- dehydro-E1, 6-dehydro-E2, 7-OH-E1, 7-OH-E2, 7ß-OH-E1, 7ß-OH-E2, 7-dehydro-E1, 7-dehydro-E2, 9(11)-dehydro-E2, 11-OH-E1, 11-OH-E2, 11ß-OH-E1, 11ß-OH-E2, 11-keto-E1, 12ß-OH-E1, 12ß-OH-E2, 12-keto-E2, 14-OH-E1, 14-OH-E2, 15-OH-E1, 15-OH-E2, 15-OH-E3, 15ß-OH-E1, 15ß-OH-E2, 16-OH-E1, 16-OH-E2, 16-OH-17-E2, 16ß-OH-E1, 16ß-OH-E2, 16ß-OH-17-E2, 16-keto-E1, 6-keto-E2, and 17-E2. For the purpose of comparison, the mass spectrum for each trimethylsilylated reference compound was obtained with our GC/MS system under the same analytical conditions for metabolically formed estrogen metabolites.
Experimentally, the collected HPLC fractions containing suspected estrogen metabolite(s) were first evaporated to dryness under a stream of nitrogen gas and then incubated at 60–65 C for 30 min in the presence of 50 µl of N,O-bis(trimethylsilyl)trifluoroacetamide containing 1% trimethylchlorosilane. A Hewlett Packard model-5890 gas chromatograph and a model-5970 mass spectrometer (Hewlett Packard, Palo Alto, CA) were used with an RTX-5MS capillary column (0.25 mm x 30 m; 0.25-µm film thickness; Restek Corp., Bellafonte, PA), with helium as the carrier gas. The mass spectrometer was operated in the electron impact mode (70 eV), and the mass abundance was determined by scanning masses from 50–600 m/z at 1.4 times/sec. The injector and detector temperatures were 260 and 280 C, respectively. During analysis, the column temperature was increased from 180–260 C at a rate of 4 C/min and then maintained isothermal at 260 C for the remainder of the run. For spectrum match-up between the metabolically formed E2 or E1 metabolites and the authentic standards, we used both the built-in library search function of our GC/MS system and the manual comparison of their mass spectra. Notably, selected ion monitoring method was also used in some cases for the identification of certain quantitatively minor estrogen metabolites.
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Blank microsomes without the expressed human CYP isoforms
When 20 µM of [3H]E2 or [3H]E1 was incubated with microsomes prepared from the insect cells infected with the wild-type baculovirus without a human CYP gene, several very small baseline radioactive peaks were detected by our HPLC system (data not shown). Notably, when some of the baseline metabolite peaks were collected from the HPLC column and further analyzed by GC/MS after trimethylsilyl (TMS) derivatization, we found that a few of them did not consistently match any of the 49 authentic E2 or E1 metabolites on the basis of their HPLC retention times, GC/MS retention times, and mass fragmentation spectra. Notably, the blank control microsomes also had weak but detectable activity for catalyzing the interconversions between E2 and E1, likely due to the presence of weak 17ß-hydroxysteroid dehydrogenase activity in these microsomes. When we quantified each of the estrogen metabolites formed with the expressed human CYP isoforms, its metabolite peak was compared against the metabolite profile of blank microsomes, and usually only the metabolite peak with its radioactivity substantially above the control baseline peak was considered and quantified.
CYP1 family isoforms
CYP1A1.
When 20 µM [3H]E2 was used as substrate, CYP1A1 had high catalytic activity for 2-OH-E2 formation (2523 ± 208 pmol/nmol·min), followed by 15-OH-E2 formation (927 ± 45 pmol/nmol·min) (Fig. 1). 4-OH-E2, 6-OH-E2, and 7-OH-E2 were also formed in significant quantities. Notably, because 7-OH-E2 was coeluted with or very near 6-OH-E2 and 15-OH-E2 on our HPLC system, its identification was based on further GC/MS analysis. We analyzed the collected HPLC fraction from 8–13 min (corresponding to 6-OH-E2, 7-OH-E2, and 15-OH-E2) and identified 7-OH-E2 as a metabolite (Fig. 2). The estimated ratio between 6-OH-E2, 7-OH-E2, and 15-OH-E2 was approximately 4.7:1:7.1. Small amounts of 6ß-OH-E2 were detected on the HPLC (Fig. 1), which was further confirmed by GC/MS analysis. Two radioactive HPLC peaks, with their retention times matched for 16-OH-E2 (E3) and y-OH-E2, 1were also detected. In addition, substantial amounts of E1 as well as small amounts of a few hydroxy-E1 metabolites (2-OH-E1, 4-OH-E1, and 15-OH-E1) were also detected. It is likely that these hydroxy-E1 metabolites were largely formed through 17ß- oxidation of their corresponding hydroxy-E2 metabolites. The ratio of E2 4-hydroxylation to 2-hydroxylation was approximately 7% with CYP1A1 (Table 2). Notably, when [3H]E2 was the substrate, large peaks D (eluted right before E2) were detected on the HPLC for all three CYP1 family enzymes. Further GC/MS analysis of these radioactive peaks (from 43–47 min) formed by CYP1A1 showed the presence of 6- dehydro-E2 and a few unknown compounds (Fig. 3, top). The quantification of the dehydroestrogen metabolites formed was not attempted in the present study, largely because smaller amounts of them were also formed with blank microsomes, and more importantly, because we did not have various dehydroestrogens as standards.
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-OH-E1 and 7-OH-E2 (combined rate of 172 ± 11 pmol/nmol·min), 6-OH-E1 (168 ± 5 pmol/nmol·min), and 16-OH-E1 (101 ± 6 pmol/nmol·min) (Fig. 1
). Again, small amounts of 7-OH-E1 were found to be coeluted with 15-OH-E1 on the HPLC, and the structures for both metabolites were further confirmed by GC/MS analysis (data not shown). The ratio of E1 4-hydroxylation to 2-hydroxylation was 19% (Table 2), which is more than twice as high as the ratio with E2 as substrate. Similarly, small amounts of several hydroxy-E2 metabolites (2-OH-E2, 6-OH-E2, and 15-OH-E2) were also detected during the incubation with E1 as substrate. Notably, although the dehydro-E2 metabolites (peak D) were readily detected with E2 as substrate, the corresponding dehydro-E1 peaks were not identified with E1. Because we found that 6-dehydro-E1 and 7-dehydro-E1 (two standards we have) were not clearly separated from E1 on the HPLC, it is thus very likely that some of those dehydro-E1 metabolites formed were not identified because they were not separated from E1 and thus were not visible on the HPLC traces.
CYP1A2.
At 20 µM [3H]E2, CYP1A2 had the highest catalytic activity for 2-hydroxylation (4065 ± 156 pmol/nmol·min), and it also had high absolute activity for 4-hydroxylation (343 ± 24 pmol/nmol·min) (Fig. 1 and Table 2). The ratio of 4-OH-E2 formation to 2-OH-E2 formation by this isoform was 9% (Table 2). Two other radioactive HPLC peaks with their retention times matched for y-OH-E2 and 6ß-OH-E2 were detected at very small quantities (<70 pmol/nmol·min). In addition, the formation of considerable amounts of 2-OH-E1 and 4-OH-E1 (357 ± 14 and 138 ± 3 pmol/nmol·min, respectively) was also detected, but the rate for the conversion of E2 to E1 with CYP1A2 was not significantly different from that with control microsomes.
When 20 µM [3H]E1 was the substrate, CYP1A2 had the highest catalytic activity for E1 2-hydroxylation (8503 ± 317 pmol/nmol·min). Notably, the rate of E1 2-hydroxylation by CYP1A2 was more than twice the rate of E2 2-hydroxylation under the same reaction conditions. Large amounts of 4-OH-E1 (1109 ± 34 pmol/nmol·min) were also formed. The ratio of E1 4- to 2-hydroxylation was approximately 13%, somewhat higher than the ratio with E2 as substrate (Table 2). In addition, small amounts of E2, 2-OH-E2, and 4-OH-E2 were also detected, likely due to metabolic conversion from E1 or their corresponding E1 metabolites.
CYP1B1.
At 20 µM [3H]E2, CYP1B1 had a distinct selectivity for the formation of 4-OH-E2 (Fig. 1). Although the rate of 4-OH-E2 formation was 371 ± 16 pmol/nmol·min, the rate of 2-OH-E2 formation was only 108 ± 3 pmol/nmol·min, giving a ratio of E2 4- to 2-hydroxylation of 3.4 (Table 2). Very small metabolite peaks with retention times matched for 15-OH-E2, 6ß-OH-E2, and 16ß-OH-E2 were also detected with E2 as substrate. In addition, substantial amounts of E1 as well as small amounts of 4-OH-E1 (99 ± 11 pmol/nmol·min) were also detected, likely due to the 17ß-oxidation of E2 and 4-OH-E2.
At 20 µM E1, CYP1B1 had similar but slightly lower catalytic activity for the formation of 4-OH-E1 and 2-OH-E1 compared with that with E2 as substrate. The rates of the 4- and 2-hydroxylation of E1 were 366 ± 25 and 149 ± 28 pmol/nmol·min, respectively, giving a ratio of E1 4- to 2- hydroxylation of 2.5 (Table 2). In addition, substantial amounts of E2 as well as small amounts of 4-OH-E2 and 2-OH-E2 were also detected.
CYP2 family isoforms
CYP2A6.
At 20 µM [3H]E2 as the substrate, CYP2A6 had no detectable activity for the formation of catechol estrogen metabolites, but a small peak with retention time matched for 16ß-OH-E2 (58 ± 4 pmol/nmol·min) was detected (Fig. 4). CYP2A6 had substantial activity for the conversion of E2 to E1 and to metabolites less polar than E1. In addition, a few radioactive peaks (collectively labeled as D) eluted shortly before E2 were detected with CYP2A6. Notably, it appeared that all other CYP2 family isoforms tested also had a varying degree of catalytic activity for the formation of these metabolite peaks. Further GC/MS analysis of the isolated HPLC fractions (from 43–51 min) that were generated with CYP2A6 showed the presence of 6-dehydro-E2, 9(11)-dehydro-E2, and a few unknown compounds (Fig. 3, middle).
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CYP2B6.
At 20 µM [3H]E2 or [3H]E1, CYP2B6 only had weak detectable activity for the formation of 2-OH-E2 and 2-OH-E1 (data not shown), at rates of 98 ± 3 and 133 ± 10 pmol/nmol·min, respectively. CYP2B6 had little or no catalytic activity for the formation of 4-hydroxylated E2 or E1. In addition, CYP2B6 had some catalytic activity for the interconversion between E1 and E2 and for the formation of metabolites less polar than E1.
CYP2C8.
At 20 µM [3H]E2, CYP2C8 showed a weak activity for the formation of 2-OH-E2 (51 ± 3 pmol/nmol·min) (Fig. 4). Interestingly, small radioactive HPLC peaks with retention times matched for 16-OH-E2 and 16ß-OH-E2 were also detected (26 ± 3 and 38 ± 3 pmol/nmol·min, respectively). This isoform had no detectable activity for the 4-hydroxylation of E2 or E1. Formation of E1 and metabolites less polar than E1 was also observed.
At 20 µM [3H]E1, CYP2C8 had a weak activity for the formation of 2-OH-E1 (29 ± 3 pmol/nmol·min) and 16-OH-E1/16ß-OH-E1 (two coeluted metabolites, with a combined rate of 31 ± 2 pmol/nmol·min) (Fig. 4). However, the structures of 16-OH-E1 and 16ß-OH-E1 were not further confirmed by GC/MS analysis in this case because of their relatively small quantities. Formation of E2 and metabolites less polar than E1 were also observed.
CYP2C9.
At 20 µM of [3H]E2 or [3H]E1, CYP2C9 showed mainly 2-hydroxylase activity at the rate of 114 ± 21 or 137 ± 11 pmol/nmol·min, respectively (Fig. 4). Notably, this estrogen 2-hydroxylation activity was considered to be very high among all the CYP2 family isoforms tested. CYP2C9 also had a weak but detectable activity for the 4-hydroxylation of E2 and E1 (16 ± 2 and 59 ± 8 pmol/nmol·min, respectively). In addition, CYP2C9 had some catalytic activity for the interconversion between E1 and E2.
CYP2C18.
At 20 µM [3H]E2 or [3H]E1, CYP2C18 had little or no catalytic activity for the oxidative metabolism of estrogens, but had a weak activity for the interconversion between E2 and E1 (data not shown). Metabolites less polar than E1 were also observed.
CYP2C19.
At 20 µM [3H]E2 or [3H]E1, CYP2C19 had a weak activity for their 2-hydroxylation, 123 ± 33 or 64 ± 25 pmol/nmol·min, respectively (data not shown). CYP2C19 also had a weak activity for the conversion of E2 to E1, but the activity for the conversion of E1 to E2 was negligible. Metabolites less polar than E1 were also observed.
CYP2D6.
At 20 µM [3H]E2 or [3H]E1, CYP2D6 showed weak 2-hydroxylase activity, at the rate of 113 ± 17 or 131 ± 14 pmol/nmol·min, respectively (data not shown). Some interconversion between E2 and E1 and the formation of metabolites less polar than E1 were also observed.
CYP2E1 + b5.
At 20 µM [3H]E2 or [3H]E1, CYP2E1 had little or no detectable catalytic activity for the hydroxylation of estrogens, but it had a weak activity for the conversion of E2 to E1 and the formation of metabolites less polar than E1 (data not shown).
CYP3A family isoforms
CYP3A4.
At 20 µM [3H]E2, CYP3A4 had high catalytic activity for the formation of multiple hydroxylated estrogen metabolites (Fig. 5). 2-OH-E2 (355 ± 41 pmol/nmol·min) was the major hydroxy-E2 metabolite formed, followed by y-OH-E2 and 4-OH-E2 (98 ± 10 and 78 ± 11 pmol/nmol·min, respectively). Several other hydroxylated metabolites (6-OH-E2, 6ß-OH-E2, 12ß-OH-E2, 15-OH-E2, 16-OH-E2, and 16ß-OH-E2) were also formed from E2 with CYP3A4. The structures of these metabolites were confirmed by GC/MS analyses of the isolated fractions from the HPLC (data not shown). In addition, GC/MS analysis of the peak(s) labeled as D in Fig. 5 showed that 9(11)-dehydro-E2 was a major component (Fig. 3, bottom). In addition, large amounts of nonpolar metabolites (collectively labeled as X in Fig. 5) were formed during incubations of E2 with CYP3A4.
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). 2-OH-E1 was the major hydroxy-E1 metabolite formed, followed by 4-OH-E1 and y-OH-E1. Quantitatively, the rates for the formation of 2-OH-E1 and 4-OH-E1 were 312 ± 6 and 159 ± 14 pmol/nmol·min, respectively, which gave the ratio of E1 4- to 2-hydroxylation approximately 51% (Table 2). The KM and VMAX values determined for the formation of 2-OH-E1 by CYP3A4 were 7.7 µM and 168 pmol/nmol·min, respectively, and those for the formation of 4-OH-E1 were 7.2 µM and 80 pmol/nmol·min, respectively (Fig. 6).
Several other hydroxy-E1 metabolites (6-OH-E1, 6ß-OH-E1, 16-OH-E1, and 16ß-OH-E1) and 6-keto-E1 were also formed in substantial quantities by CYP3A4, and their structures were confirmed by GC/MS analyses. In addition, large amounts of nonpolar metabolites (collectively labeled as X in Fig. 5) were formed during incubations of E1 with CYP3A4.
Notably, we also analyzed estrogen metabolism by microsomes with either CYP3A4 alone or CYP3A4 combined with cytochrome b5 expression. We found that the overall profiles of estrogen metabolites formed with CYP3A4 plus cytochrome b5-expressed microsomes were almost the same as those formed with CYP3A4-expressed microsomes, but the rate for the formation of each metabolite was much higher with the CYP3A4/cytochrome b5 microsomes (Fig. 5). This observation showed that the presence of higher levels of cytochrome b5 accelerated the rate of estrogen metabolism under the same reaction conditions, although it did not significantly alter the overall profile of estrogen metabolites formed.
CYP3A5.
At 20 µM [3H]E2, CYP3A5 catalyzed the formation of several hydroxylated estrogen metabolites. The overall profile formed with this CYP isoform was very similar to that formed with CYP3A4 (Fig. 5). 2-OH-E2 was the major hydroxy-E2 metabolite formed (at the rate of 125 ± 22 pmol/nmol·min), followed by y-OH-E2 (82 ± 7 pmol/nmol·min) and 4-OH-E2 (67 ± 5 pmol/nmol·min). The ratio of E2 4- to 2-hydroxylation was 0.53 (Table 2). The KM and VMAX values determined for the formation of 2-OH-E2 by CYP3A5 (a separate batch) were 52.5 µM and 627 pmol/nmol·min, respectively, and those for the formation of 4-OH-E2 were 46.0 µM and 298 pmol/nmol·min, respectively (Fig. 6).
In addition, several other hydroxy-E2 metabolites (6ß-OH-E2, 16-OH-E2, and 16ß-OH-E2) were also formed by CYP3A5. The formation of all these metabolites was confirmed by GC/MS analyses of the isolated fractions from the HPLC (data not shown). Considerable amounts of a few dehydro-E2 metabolites (labeled as D) and nonpolar metabolites (collectively labeled as X in Fig. 5) were also detected during incubations of E2 with CYP3A5.
At 20 µM [3H]E1, CYP3A5 had a slightly different profile for E1 metabolism than did CYP3A4 (Fig. 5). CYP3A5 at the same molar concentration formed much less catechol estrogen metabolites than CYP3A4. However, 4-OH-E1 became the major hydroxyestrogen metabolite with CYP3A5, followed by 2-OH-E1 and y-OH-E1. Quantitatively, the rates for the formation of 2-OH-E1 and 4-OH-E1 were 67 ± 11 and 84 ± 8 pmol/nmol·min, respectively, giving a ratio of 4- to 2- hydroxylation of 1.26 (Table 2). Further enzyme kinetic analysis showed that the KM and VMAX values for the formation of 2-OH-E1 were 15.0 µM and 103 pmol/nmol·min, respectively, and those for the formation of 4-OH-E1 were 27.8 µM and 156 pmol/nmol·min, respectively (Fig. 6).
In addition, several other hydroxy-E1 metabolites (6ß-OH-E1, 16-OH-E1, and 16ß-OH-E1) were also formed in substantial quantities by CYP3A5 (Fig. 5). The formation of these metabolites was confirmed by GC/MS analyses of the isolated fractions from the HPLC (data not shown). Large amounts of nonpolar metabolites (collectively labeled as X in Fig. 5) were formed during incubations of E1 with CYP3A5.
CYP3A7 + b5.
At 20 µM [3H]E2, CYP3A7 (coexpressed with cytochrome b5) had a weak catalytic activity for E2 2- hydroxylation (146 ± 17 pmol/nmol·min). In addition, small metabolite peaks with retention times matched for 6ß-OH-E2, 16-OH-E2, and y-OH-E2 as well as the dehydro-E2 metabolites (peak D) were detected (Fig. 5). Appreciable amounts of E1 and metabolites less polar than E1 were also formed.
At 20 µM [3H]E1, CYP3A7 had a modest catalytic activity for the formation of 2-OH-E1 and 4-OH-E1, with a combined rate of 368 ± 18 pmol/nmol·min (Fig. 5). Further GC/MS analysis showed that the ratio of E1 4- to 2-hydroxylation was 10% (Table 2). Notably, CYP3A7 plus cytochrome b5 had the highest catalytic activity for the formation of 16-OH-E1 and 6ß-OH-E1 (two coeluted metabolites with a combined rate of 346 ± 3 pmol/nmol·min) (Fig. 6). GC/MS analysis showed that 16-OH-E1 accounted for more than 90%.
CYP4 family isoform
CYP4A11.
At 20 µM [3H]E2 or [3H]E1, CYP4A11 did not have detectable catalytic activity for the oxidative metabolism of the estrogens (data not shown). When E2 was the substrate, substantial amounts of E1 and metabolites less polar than E1 were formed. Similarly, E2 and metabolites less polar than E1 were formed from E1 as substrate.
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CYP1 family isoforms
CYP1A1.
Human CYP1A1 is essentially expressed in extrahepatic tissues, and the level of its expression is readily inducible by exposure to environmental chemicals or toxins, such as 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) and polycyclic aromatic hydrocarbons (40). The results of our present study showed that CYP1A1 catalyzed the formation of several hydroxylated estrogen metabolites from E2 or E1. When E2 was the substrate, CYP1A1 had high catalytic activity for its 2-hydroxylation, followed by 15-, 6-, and 4-hydroxylation. The formation of small amounts of 6ß-OH-E2, 7-OH-E2, and 16-OH-E2 was also detected. In addition, CYP1A1 also catalyzed the formation of 6-dehydro-E2.
When E1 was the substrate, CYP1A1 had high catalytic activity for the formation of 2-OH-E1, 4-OH-E1, 15-OH-E1, 6-OH-E1, and 16-OH-E1. Small amounts of 7-OH-E1 were also formed. Notably, although the ratio of 4- to 2-hydroxylation with CYP1A1 was approximately 7% when E2 was the substrate, this ratio was increased almost three times (19%) with E1 as substrate.
Although there is little information in the literature on CYP1A1-mediated metabolism of E1, the results of our present study on its metabolism of E2 are in agreement with several earlier studies (14, 24, 30). Spink et al. (14) reported earlier that exposure of MCF-7 human breast cancer cells to TCDD markedly increased the microsomal formation of 2-OH-E2, 4-OH-E2, 15-OH-E2, and 6-OH-E2 from E2 as a substrate. Additional studies by these authors using transfected human CYP1A1 gene confirmed that CYP1A1 catalyzed the 2-, 4-, 6-, and 15-hydroxylation of E2 (24). In a recent study, it was also reported that CYP1A1 had high activity for E2 2-hydroxylation but lower activity for E2 4- and 16-hydroxylation (30).
It is of note that among the 15 human CYP isoforms analyzed in the present study, CYP1A1 was the only CYP isoform that had a distinctly high catalytic activity for the 15-hydroxylation of estrogens. When E2 was the substrate, 15-OH-E2 was the second major hydroxy-E2 metabolite formed (next to 2-OH-E2). In comparison, the rate for the 15-hydroxylation of E1 by CYP1A1 was markedly lower than the rate for the 15-hydroxylation of E2. This difference in the rate of 15-hydroxylation of E2 vs. E1 likely is due to the differential influence of the 17ß-hydroxyl group (in the case of E2) and the 17-keto group (in the case of E1) on the CYP1A1-mediated 15-hydroxylation.
It is of interest to note that the 15-hydroxylated estrogen metabolites are known to be formed in large amounts during human pregnancy (41), and the amount of urinary 15- hydroxyestriol excretion by the expectant mother was found to be a reliable indicator for fetal well-being (42, 43). However, it is yet to be determined whether CYP1A1 is the major or sole CYP isoform that is responsible for the 15-hydroxylation of endogenous estrogens during human pregnancy.
It has been suggested for many years that 16-OH-E1 plays an important role in mammary carcinogenesis (31, 44, 45). This estrogen metabolite is not only hormonally active but also chemically reactive and may bind covalently to the estrogen receptor, possibly resulting in sustained hormonal stimulation of the ER-positive target cells (31, 46). Our recent studies showed that human liver microsomes form only very minute amounts of 16-OH-E2 and 16-OH-E1 when [3H]E2 or [3H]E1 was used as the substrate (34, 35). However, the results of our present study showed, for the first time, that human CYP1A1 (an extrahepatic isoform) has relatively high catalytic activity for the 16-hydroxylation of E1, but not so with E2. Although CYP1A1 has a preference for the 15-hydroxylation of E2 over E1, its opposite preference for the 16-hydroxylation of E1 over E2 may also be due to a different influence of the 17-keto or 17ß-hydroxyl groups to the neighboring C16-positions.
CYP1A2.
CYP1A2 constitutes approximately 13% of the total CYP enzymes contained in human liver (47), and it metabolizes E2 and/or E1 to 2-hydroxylated metabolites (22, 27, 29, 30). The results of our present study showed that among all human CYP isoforms tested, CYP1A2 had the highest 2- hydroxylase activity for E2 and E1 (4065 or 8503 pmol/nmol·min, respectively), with E1 2-hydroxylation more than twice as fast as E2 2-hydroxylation. The rate of 4-hydroxylation of E2 and E1 by CYP1A2 was 9 and 13%, respectively, of the rate of their 2-hydroxylation (Table 2). It should be noted that similar high rates for the 2-hydroxylation of E2 and E1 by a cDNA-expressed human CYP1A2 were also reported earlier (22, 27, 29, 30).
Notably, an earlier study suggested that human CYP1A2 had weak activity for the 16-hydroxylation of E2 but not E1 (29). In addition, it was suggested that the 16-hydroxylation of E2 by human liver microsomes (which contained CYP1A2) was partially inhibited by anti-CYP1A2 antibody, whereas the 16-hydroxylation of E1 was not affected by anti-CYP1A2 antibody (29). However, the results of our present study clearly showed that human CYP1A2 had little or no catalytic activity for the formation of 16-hydroxylated metabolites of E2 or E1.
CYP1B1.
CYP1B1 is another extrahepatic CYP1 family isoform that is chemically inducible and often is overexpressed in tumor tissues (48). This CYP isoform is known to play an important role in the 4-hydroxylation of estrogens. Hayes et al. (26) reported earlier that the elevated E2 4-hydroxylase activity in TCDD-treated MCF-7 breast cancer cells was, in part, due to an elevated expression of the CYP1B1 gene. In addition, selective expression of human CYP1B1 gene in Saccharomyces cerevisiae produced an enzyme that predominantly catalyzed E2 4- and 2-hydroxylation (26). The ratios of E2 4- to 2-hydroxylation for the expressed proteins of four human CYP1B1 gene variants were reported earlier to range from 1.8–3.6 (49). Our results also showed a predominant hydroxylase activity for CYP1B1 in the C4-positions of E2 and E1, giving the highest ratios (3.4 and 2.5) for estrogen 4- to 2-hydroxylation among the 15 CYP isoforms tested (Table 2). However, despite a distinct activity for catechol estrogen formation and a high ratio for estrogen 4- to 2-hydroxylation, it should be pointed out that the absolute amounts of 4-OH-E2 and 4-OH-E1 formed with CYP1B1 were not high when compared with the amounts formed with some other CYP isoforms, such as CYP1A1 and 1A2.
In summary, whereas CYP1A1 had high catalytic activity for the 2-hydroxylation of E2 and E1 (with 4-hydroxylation a minor metabolic pathway), this isoform had a highly distinct activity for the D-ring hydroxylation of estrogens at the C15-positions (for E2 and E1) and at the C16-position (for E1). CYP1A2 was catalytically most active for the 2-hydroxylation of E2 and E1. CYP1B1 had a distinct, selective activity for the 4-hydroxylation of E2 and E1 when compared with the 2-hydroxylation of these steroids.
CYP2 family isoforms
CYP2A6.
Our results showed that CYP2A6 had no detectable activity for catechol estrogen formation from E2 or E1, which is in agreement with an earlier suggestion that CYP2A6 did not have estrogen-metabolizing activity (28). Interestingly, our results showed that a few dehydro-E2 metabolites, such as 6-dehydro-E2 and 9(11)-dehydro-E2, were formed with this CYP isoform. All other CYP2 family isoforms also formed varying amounts of the peak D (which presumably also contained dehydro-E2 metabolites). Notably, the peak D was earlier suggested to be impurities from the radioactive E2 (37). We indeed noted that a small amount of the peak D was contained in the radioactive [3H]E2 substrate, and incubation of [3H]E2 with control blank microsomes also formed small amounts of the peak D. When microsomes with most of the selectively expressed CYP 2 family isoforms were tested, the formation of peak D appeared to be increased to varying degrees.
CYP2B6.
CYP2B6 was reported to be present in 12 of 17 human liver microsomes tested, ranging from 3–74 pmol/mg microsomal protein (50). An earlier study suggested that CYP2B6 did not form any significant amounts of estrogen metabolites (27). Our data showed that CYP2B6 had a moderate activity for the 2-hydroxylation of E2 and E1 (98 and 133 pmol/nmol·min, respectively), but this CYP isoform had no detectable activity for estrogen 4-hydroxylation. The discrepancy between our data and an earlier report may be attributable to the greatly improved sensitivity of our in vitro assay system.
CYP2C8.
Several earlier studies suggested that CYP2C8 had no detectable activity for the oxidative metabolism of estrogens (22, 27). Our data showed that CYP2C8 had a weak but detectable activity for the 2-hydroxylation of E2 and E1. In addition, small amounts of estrogen 16/16ß-hydroxylation products were also detected with CYP2C8.
CYP2C9.
Earlier studies suggested that human CYP2C9 had a moderate catalytic activity for catechol estrogen formation from E2 (22, 29) or E1 (27). Our results showed that CYP2C9 had a relatively high activity for the 2-hydroxylation of E2 and E1 among the CYP2 family isoforms tested, and it also had a weak but detectable activity for estrogen 4-hydroxylation. Notably, an earlier study showed that the presence of anti-CYP2C9 antibody or sulfaphenazole did not significantly affect liver microsomal hydroxylation of E2 or E1 (29), likely suggesting the relatively minor contribution of CYP2C9 to the overall estrogen metabolism in human liver.
Although it was also suggested earlier that CYP2C9 had a weak activity for the 16-hydroxylation of E2 (29), little or no estrogen 16-hydroxylase activity was detected in the present study.
CYP2C18.
Our results showed that CYP2C18 had little or no detectable activity for the oxidative metabolism of E2 and E1.
CYP2C19.
CYP2C19 has been suggested to have a weak catalytic activity for the 16-hydroxylation of E2 (29). Our results showed that CYP2C19 had a relatively high 2- hydroxylase activity for E2 or E1, but the formation of 16-hydroxylated estrogen metabolites was not detected.
CYP2D6.
An earlier study suggested that CYP2D6 had no detectable activity for the formation of estrogen metabolites (22). We found that CYP2D6 had a modest activity for the formation of 2-OH-E2 and 2-OH-E1 (at rates of 113 and 131 pmol/nmol·min).
CYP2E1 + b5.
Although an earlier study suggested that CYP2E1 was among the important hepatic CYP isoforms for estrogen 2-hydroxylation (23), most other studies (22, 25, 27) have reported that CYP2E1 had no detectable activity for the formation of estrogen metabolites. Our results also showed that CYP2E1 had no detectable activity for the hydroxylation of E2 or E1.
In summary, the results of our present study showed that CYP2 family isoforms overall did not have distinctly high activity for the oxidative metabolism of E2 and E1, but a varying degree of estrogen 2-hydroxylase activity was detected with several of the isoforms. Because the CYP2 family isoforms account for approximately 30% of human hepatic CYP enzymes (with 2C subfamily isoforms predominant; Ref. 47), their collective contribution to the catechol estrogen formation (particularly 2-hydroxyestrogen) may still be of considerable importance. In addition, CYP2 family isoforms showed a varying degree of catalytic activity for the formation of certain dehydro-E2 metabolites from E2.
CYP3 family isoforms
CYP3A4.
CYP3A family enzymes are the most abundant CYP isoforms present in human liver (51, 52), and it was estimated that CYP3A4 accounts for approximately 30% of total CYP content (47). Several earlier studies showed that CYP3A4 is involved in the 2- and 4-hydroxylation of E2 by human liver microsomes (22, 25, 29, 30, 34, 53).
Our results showed that CYP3A4 had high catalytic activity for the formation of multiple hydroxylated metabolites of E2 and E1. The catechol estrogen metabolites were the major metabolites formed from 20 µM E2 and E1, with apparent ratios of 4- to 2-hydroxylation of 0.22 and 0.51, respectively. Notably, the ratio of E1 4- to 2-hydroxylation was almost twice as high as the ratio for E2. Further enzyme kinetic analysis showed that although the capacity (VMAX) of the CYP3A4-mediated 2- and 4-hydroxylation of E2 was approximately eight times higher than its catalytic capacity for E1, its affinity (1/KM) for the 2- and 4-hydroxylation of E2 was only approximately 20% of its affinity for E1. Hence, although CYP3A4 has a lower affinity but a higher capacity for the 2- and 4-hydroxylation of E2, it has a higher affinity but a lower capacity for the 2- and 4-hydroxylation of E1. The causes for the opposite kinetic features of CYP3A4-mediated 2- and 4-hydroxylation of E2 vs. E1 are not known. Interestingly, despite the dramatic differences in the VMAX and KM values for the 2- and 4-hydroxylation of E2 and E1, the VMAX/KM ratios of CYP3A4-mediated 2- and 4-hydroxylation of these two estrogens were very similar. This likely would make the CYP3A4-mediated metabolism of physiological concentrations of E2 or E1 in vivo proceed at comparable rates.
Notably, our results with radioactive E2 and E1 as substrates showed that CYP3A4 only had a very weak catalytic activity for the 16-hydroxylation of E1 and E2. This observation is in contrast to a few earlier studies that showed that CYP3A4 had high catalytic activity for the 16-hydroxylation of E2 and E1, based on the detection of nonradioactive estrogen metabolites (27, 28, 29, 30).
Besides 2-, 4-, and 16-hydroxy-E2 metabolites, we also showed that several other hydroxy-E2 metabolites (y-OH-E2, 6-OH-E2, 6ß-OH-E2, 12ß-OH-E2, 15-OH-E2, and 16ß-OH-E2) and a dehydroestrogen metabolite, 9(11)-dehydro-E2, were also formed from E2 by CYP3A4. Similarly, several other hydroxy-E1 metabolites (y-OH-E1, 6-OH-E1, 6ß-OH-E1, and 16ß-OH-E1) were formed from E1 in substantial quantities. In addition, our results showed that large amounts of nonpolar metabolites (collectively labeled as X in Fig. 5) were formed during incubations of CYP3A4 with either E2 or E1 as substrate.
Lastly, it is of interest to note that when a higher level of cytochrome b5 was present, CYP3A4 showed a markedly enhanced activity for the formation of multiple hydroxylated estrogen metabolites, but it did not alter the overall estrogen metabolite profiles. Mechanistically, because cytochrome b5 (a mediator of second electron from NADPH) has also been shown to stimulate CYP-mediated metabolism of certain other substrates (54, 55), this stimulation likely is due to a nonspecific mechanism, such as the increased availability of the second electron required for the reactions. In light of our observations, it should be noted that when the metabolic rates determined in this study are used for comparing the rate of each CYP isoform-mediated metabolism of E2 or E1, it is under the assumption that the cytochrome b5 content as well as other supporting factors (such as CYP reductase) in the expressed microsomes were the same. Because their variability was almost unavoidable, it is thus advised to take into careful consideration their possible influence when making definitive comparisons of the absolute metabolic rates for each of the CYP isoforms.
CYP3A5.
Our results showed that when E2 was the substrate, CYP3A5 formed estrogen metabolites with a similar profile as that of CYP3A4, with 2-OH-E2 as the major hydroxy-E2 metabolite, followed by y-OH-E2 and 4-OH-E2. In comparison, CYP3A5 formed much less catechol estrogen metabolites from E1 than did CYP3A4, but 4-OH-E1 became the major hydroxyestrogen metabolite formed by CYP3A5, followed by 2-OH-E1 and y-OH-E1. Accordingly, although the ratio of E2 4- to 2-hydroxylation by CYP3A5 was 0.53, its ratio for E1 4- to 2-hydroxylation was 1.26, which was the second highest ratio for E1 (next to CYP1B1) among the 15 isoforms tested.
Our enzyme kinetic analysis showed that the capacity (VMAX) of CYP3A5-mediated 2- and 4-hydroxylation of E2 was approximately 6 and 2 times higher, respectively, than its catalytic capacity for E1, but its affinity (1/KM) for the 2- and 4-hydroxylation of E2 was significantly lower than its affinity for E1. Therefore, similar to CYP3A4, CYP3A5 had a relatively lower affinity but a higher capacity for the 2- and 4-hydroxylation of E2, but it had a relatively higher affinity but a lower capacity for the 2- and 4-hydroxylation of E1. The VMAX/KM ratio of CYP3A5-mediated 2-hydroxylation of E2 was higher than that for the 2-hydroxylation of E1, but the VMAX/KM ratios for the 4-hydroxylation of E2 and E1 were similar.
Our results showed that CYP3A5 also formed a few other hydroxy-E2 metabolites (6ß-OH-E2, 16-OH-E2, and 16ß-OH-E2) from E2. Similarly, several other hydroxy-E1 metabolites (6ß-OH-E1, 16-OH-E1, and 16ß-OH-E1) were formed from E1 in substantial quantities. In addition, we found that large amounts of nonpolar metabolites (collectively labeled as X in Fig. 5) were also formed during incubations of CYP3A5 with E2 or E1 as substrate. It is noteworthy that CYP3A4 and CYP3A5 had much higher activity for the formation of these nonpolar metabolite peaks than the other CYP isoforms.
Notably, a recent study suggested that hepatic CYP3A5 is expressed in approximately one third of Caucasians and approximately 60% of African-Americans tested (56). Quantitatively, CYP3A5 has been found in 10–30% of hepatic samples and estimated to be 10–30% of the CYP3A4 content (57), and in certain individuals it might account for as high as approximately 50% of the total CYP3A content (56). Because it is known that CYP3A5 in human liver has a polymorphic distribution (57, 58), this CYP isoform likely is one of the genetic factors that may affect the interindividual and interracial differences in the hepatic formation of catechol estrogens, particularly the 4-hydroxylated estrogens. In light of the mounting evidence for an important etiological role of 4-hydroxyestrogen metabolites in estrogen-induced cancer (32), we believe that it would be of considerable interest to determine whether this polymorphism correlates with the amounts of 4-hydroxylated estrogens produced in humans as well as the risk for estrogen-associated human cancers.
CYP3A7 + b5.
CYP3A7 was originally found in human fetal liver where it accounted for 30–50% of the total CYP content (59, 60). Additional studies also suggested the presence of constitutive or induced expression of CYP3A7 in adult human liver (61, 62, 63), although quantitative data on its level is still not available, partly due to the lack of specific antibodies.
Our results showed that CYP3A7 (coexpressed with cytochrome b5) had a very different metabolite profile from those of CYP3A4 and 3A5. When E2 was the substrate, CYP3A7 had moderate activity for its 2-hydroxylation (146 pmol/nmol·min), and small amounts of 6ß-OH-E2, 16-OH-E2, and y-OH-E2 were also formed. However, with E1 as substrate, CYP3A7 had modest catalytic activity for the formation of 2-OH-E1 (335 pmol/nmol·min), but it had the highest catalytic activity for the formation of two coeluted metabolites 16-OH-E1 and 6ß-OH-E1 (with the former accounting for >90%) at a combined rate of 346 pmol/nmol·min. This is the first demonstration that human CYP3A7 has a distinct high activity for the 16-hydroxylation of E1, but this was not observed with E2.
Our data showed that although E1 was a very good substrate for CYP3A7-mediated 16-hydroxylation, E2 was essentially not 16-hydroxylated by this CYP isoform. We hypothesize that the presence of a 17-keto group in the steroid is essential for it to be a suitable substrate for CYP3A7. In support of this hypothesis, an earlier study also showed that although CYP3A7 catalyzed the 16-hydroxylation of dehydroepiandrosterone and its 3-sulfate (both have a 17-keto group), it did not catalyze the 16-hydroxylation of testosterone or cortisol (both lack a 17-keto group) (54, 64).
It is of interest to point out that the long-held view that 16-hydroxylation only occurred with E1 as the substrate (65) appears to be largely true in the case of CYP3A7 as catalyst. However, it is also worth noting that when the 16-hydroxylation of E2 or E1 was recently analyzed with 33 adult human liver microsomes (34, 35), we found that the average rates for the 16-hydroxylation of these two estrogens were very low and similar, thus suggesting that the contribution of CYP3A7 to the overall hepatic estrogen metabolism in humans is very minor. Nonetheless, because CYP3A7 is known to be also expressed in human uterine endometrium and placenta (66), the extent of its contribution to the in situ as well as systematic formation of 16-OH-E1 remains to be determined. In this context, it is also of note that because CYP3A7 expression in human liver and intestine appears to have a polymorphic distribution, with an estimated approximately 11% of Caucasians belonging to a distinct subgroup of high expression phenotype (67), it may be of interest to determine whether individuals with the high expression phenotype produce more 16-OH-E1 and whether they have an increased risk for breast cancer.
In summary, CYP3A4 and 3A5 catalyzed the metabolism of E2 to various hydroxylated estrogens, generally with 2- hydroxyestrogens as the predominant metabolites. However, when E1 was substrate, CYP3A5 formed more 4- hydroxyestrone than 2-hydroxyestrone, giving an unusually high ratio of E1 4- to 2-hydroxylation (1.26). CYP3A4 and 3A5 also catalyzed the formation of large amounts of nonpolar metabolites from E2 and E1. CYP3A7 had moderate activity for the 2-hydroxylation of E2 and E1, but it had distinct catalytic activity for the 16- and 6ß-hydroxylation of E1.
CYP4A11
CYP4A11 did not have detectable activity for the oxidative metabolism of E2 or E1.
Conclusions
The results of our present study showed that a variety of human CYP isoforms were catalytically active for the NADPH-dependent oxidative metabolism of E2 and E1, with a varying degree of catalytic activity and distinct regioselectivity (summarized in Fig. 7). For the 2-hydroxylation of E2 and E1, CYP1A2 had the highest catalytic activity, but CYP1A1, 3A4, and 3A5 also had relatively high catalytic activity. Several CYP2 family enzymes also had modest activity for the 2-hydroxylation of estrogens. For the 4- hydroxylation of E2 and E1, CYP1B1 had selective activity, but CYP3A5 also had distinct activity for the 4-hydroxylation of estrogens (E1 in particular). CYP1A1, 1A2, and 3A4 had a modest activity for estrogen 4-hydroxylation. Except for CYP1B1 and CYP3A7, most of CYP enzymes had a higher ratio of 4- to 2-hydroxylation with E1 as substrate than with E2. For the 15-hydroxylation of E2 and E1, CYP1A1 was the only isoform that had high catalytic activity, although very small amounts of 15-OH-E2 were also formed with CYP1B1 and 3A4. For the 16-hydroxylation of E1, CYP3A7 had distinct high activity, but it only had very weak activity for the 16-hydroxylation of E2. CYP1A1, 3A4, and 3A5 showed a weak activity for the 16-hydroxylation of both E2 and E1. In addition to the quantitatively major metabolic pathways at the C-2, C-4, C-15, and C-16 positions with certain CYP isoforms studied here, E2 and E1 were also oxidatively metabolized at several other positions (such as C-6, C-6ß, C-7, C-16ß) by some of the CYP isoforms (summarized in Fig. 7). Given the fact that some of the estrogen metabolites (such as 4-OH-E2, 15-OH-E2, 16-OH-E1, and 2-methoxyestradiol) may have unique biological functions that are not associated with their parent hormones E2 and E1 (1, 31, 32, 33), we believe that the detailed knowledge provided in this study regarding the complete profiles of estrogen metabolites formed by various human CYP isoforms would be very helpful to the understanding of their biological functions in different target cells that are known to preferentially express certain CYP isoforms.
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Acknowledgments |
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Footnotes |
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A.H.C. is the William M. and Myrle W. Garbe Professor of Cancer and Leukemia Research.
Abbreviations: CYP, Cytochrome P450; GC/MS, gas chromatography/mass spectrometry; E1, estrone; E2, 17ß-estradiol; NADPH, nicotinamide adenine dinucleotide phosphate, reduced form; TCDD, 2,3,7,8-tetrachlorodibenzo-p-dioxin; TMS, trimethylsilyl.
1 The structures for y-hydroxylated E2 and E1 metabolites (y-OH-E2 and y-OH-E1, respectively) described in this study are still unidentified. Notably, in our recent studies (34 35 ), we suggested that the y-hydroxylated estrogens might be 1-, 8-, 9-, 12-, or 18-hydroxyestrogens on the basis of the HPLC and GC/MS analyses of these two unknown metabolite peaks with all available estrogen standards. 
Received February 10, 2003.
Accepted for publication May 5, 2003.
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J. F. Brown Jr, B. A. Mayes, J. B. Silkworth, and S. B. Hamilton Polychlorinated Biphenyls Modulated Tumorigenesis in Sprague Dawley Rats: Correlation with Mixed Function Oxidase Activities and Superoxide (O2* ) Formation Potentials and Implied Mode of Action Toxicol. Sci., August 1, 2007; 98(2): 375 - 394. [Abstract] [Full Text] [PDF]
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Y.-L. Low, A. M. Dunning, M. Dowsett, E. Folkerd, D. Doody, J. Taylor, A. Bhaniani, R. Luben, K.-T. Khaw, N. J. Wareham, et al. Phytoestrogen Exposure Is Associated with Circulating Sex Hormone Levels in Postmenopausal Women and Interact with ESR1 and NR1I2 Gene Variants Cancer Epidemiol. Biomarkers Prev., May 1, 2007; 16(5): 1009 - 1016. [Abstract] [Full Text] [PDF]
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E. Higashi, M. Nakajima, M. Katoh, S. Tokudome, and T. Yokoi Inhibitory Effects of Neurotransmitters and Steroids on Human CYP2A6 Drug Metab. Dispos., April 1, 2007; 35(4): 508 - 514. [Abstract] [Full Text] [PDF]
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H. Sakai, H. Iwata, E.-Y. Kim, O. Tsydenova, N. Miyazaki, E. A. Petrov, V. B. Batoev, and S. Tanabe Constitutive Androstane Receptor (CAR) as a Potential Sensing Biomarker of Persistent Organic Pollutants (POPs) in Aquatic Mammal: Molecular Characterization, Expression Level, and Ligand Profiling in Baikal Seal (Pusa sibirica) Toxicol. Sci., November 1, 2006; 94(1): 57 - 70. [Abstract] [Full Text] [PDF]
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 Y. Tsuchiya, M. Nakajima, S. Takagi, T. Taniya, and T. Yokoi MicroRNA Regulates the Expression of Human Cytochrome P450 1B1. Cancer Res., September 15, 2006; 66(18): 9090 - 9098. [Abstract] [Full Text] [PDF]
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K. A. Usmani, T. M. Cho, R. L. Rose, and E. Hodgson Inhibition of the Human Liver Microsomal and Human Cytochrome P450 1A2 and 3A4 Metabolism of Estradiol by Deployment-Related and Other Chemicals Drug Metab. Dispos., September 1, 2006; 34(9): 1606 - 1614. [Abstract] [Full Text] [PDF]
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 B. T. Zhu, G.-Z. Han, J.-Y. Shim, Y. Wen, and X.-R. Jiang Quantitative Structure-Activity Relationship of Various Endogenous Estrogen Metabolites for Human Estrogen Receptor {alpha} and {beta} Subtypes: Insights into the Structural Determinants Favoring a Differential Subtype Binding Endocrinology, September 1, 2006; 147(9): 4132 - 4150. [Abstract] [Full Text] [PDF]
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A. E. Cribb, M. J. Knight, D. Dryer, J. Guernsey, K. Hender, M. Tesch, and T. M. Saleh Role of polymorphic human cytochrome p450 enzymes in estrone oxidation. Cancer Epidemiol. Biomarkers Prev., March 1, 2006; 15(3): 551 - 558. [Abstract] [Full Text] [PDF]
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 T. M. Sissung, D. K. Price, A. Sparreboom, and W. D. Figg Pharmacogenetics and Regulation of Human Cytochrome P450 1B1: Implications in Hormone-Mediated Tumor Metabolism and a Novel Target for Therapeutic Intervention Mol. Cancer Res., March 1, 2006; 4(3): 135 - 150. [Abstract] [Full Text] [PDF]
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D. A. Symonds, K. P. Miller, D. Tomic, and J. A. Flaws Effect of Methoxychlor and Estradiol on Cytochrome P450 Enzymes in the Mouse Ovarian Surface Epithelium Toxicol. Sci., February 1, 2006; 89(2): 510 - 514. [Abstract] [Full Text] [PDF]
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J. Thibaudeau, J. Lepine, J. Tojcic, Y. Duguay, G. Pelletier, M. Plante, J. Brisson, B. Tetu, S. Jacob, L. Perusse, et al. Characterization of Common UGT1A8, UGT1A9, and UGT2B7 Variants with Different Capacities to Inactivate Mutagenic 4-Hydroxylated Metabolites of Estradiol and Estrone Cancer Res., January 1, 2006; 66(1): 125 - 133. [Abstract] [Full Text] [PDF]
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B. W. Ogilvie, D. Zhang, W. Li, A. D. Rodrigues, A. E. Gipson, J. Holsapple, P. Toren, and A. Parkinson GLUCURONIDATION CONVERTS GEMFIBROZIL TO A POTENT, METABOLISM-DEPENDENT INHIBITOR OF CYP2C8: IMPLICATIONS FOR DRUG-DRUG INTERACTIONS Drug Metab. Dispos., January 1, 2006; 34(1): 191 - 197. [Abstract] [Full Text] [PDF]
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A. Wellejus, A. Olsen, A. Tjonneland, B. L. Thomsen, K. Overvad, and S. Loft Urinary Hydroxyestrogens and Breast Cancer Risk among Postmenopausal Women: A Prospective Study Cancer Epidemiol. Biomarkers Prev., September 1, 2005; 14(9): 2137 - 2142. [Abstract] [Full Text] [PDF]
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J. R. Starr, C. Chen, D. R. Doody, L. Hsu, S. Ricks, N. S. Weiss, and S. M. Schwartz Risk of Testicular Germ Cell Cancer in Relation to Variation in Maternal and Offspring Cytochrome P450 Genes Involved in Catechol Estrogen Metabolism Cancer Epidemiol. Biomarkers Prev., September 1, 2005; 14(9): 2183 - 2190. [Abstract] [Full Text] [PDF]
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L. Le Marchand, T. Donlon, L. N. Kolonel, B. E. Henderson, and L. R. Wilkens Estrogen Metabolism-Related Genes and Breast Cancer Risk: The Multiethnic Cohort Study Cancer Epidemiol. Biomarkers Prev., August 1, 2005; 14(8): 1998 - 2003. [Abstract] [Full Text] [PDF]
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Z.-J. Liu, W. J. Lee, and B. T. Zhu Selective Insensitivity of ZR-75-1 Human Breast Cancer Cells to 2-Methoxyestradiol: Evidence for Type II 17{beta}-Hydroxysteroid Dehydrogenase as the Underlying Cause Cancer Res., July 1, 2005; 65(13): 5802 - 5811. [Abstract] [Full Text] [PDF]
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A.-M. Yu, K. Fukamachi, K. W. Krausz, C. Cheung, and F. J. Gonzalez Potential Role for Human Cytochrome P450 3A4 in Estradiol Homeostasis Endocrinology, July 1, 2005; 146(7): 2911 - 2919. [Abstract] [Full Text] [PDF]
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Y.-K. Leung, K.-M. Lau, J. Mobley, Z. Jiang, and S.-M. Ho Overexpression of Cytochrome P450 1A1 and Its Novel Spliced Variant in Ovarian Cancer Cells: Alternative Subcellular Enzyme Compartmentation May Contribute to Carcinogenesis Cancer Res., May 1, 2005; 65(9): 3726 - 3734. [Abstract] [Full Text] [PDF]
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P. Kisselev, W.-H. Schunck, I. Roots, and D. Schwarz Association of CYP1A1 Polymorphisms with Differential Metabolic Activation of 17{beta}-Estradiol and Estrone Cancer Res., April 1, 2005; 65(7): 2972 - 2978. [Abstract] [Full Text] [PDF]
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M. Delaforge, A. Pruvost, L. Perrin, and F. Andre CYTOCHROME P450-MEDIATED OXIDATION OF GLUCURONIDE DERIVATIVES: EXAMPLE OF ESTRADIOL-17{beta}-GLUCURONIDE OXIDATION TO 2-HYDROXY-ESTRADIOL-17{beta}-GLUCURONIDE BY CYP 2C8 Drug Metab. Dispos., March 1, 2005; 33(3): 466 - 473. [Abstract] [Full Text] [PDF]
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J. A. Doherty, N. S. Weiss, R. J. Freeman, D. A. Dightman, P. J. Thornton, J. R. Houck, L. F. Voigt, M. A. Rossing, S. M. Schwartz, and C. Chen Genetic Factors in Catechol Estrogen Metabolism in Relation to the Risk of Endometrial Cancer Cancer Epidemiol. Biomarkers Prev., February 1, 2005; 14(2): 357 - 366. [Abstract] [Full Text] [PDF]
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 M. J. Ronis, Y. Chen, C.-H. Jo, P. Simpson, and T. M. Badger Diets Containing Soy Protein Isolate Increase Hepatic CYP3A Expression and Inducibility in Weanling Male Rats Exposed during Early Development J. Nutr., December 1, 2004; 134(12): 3270 - 3276. [Abstract] [Full Text] [PDF]
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 V K Turan, R I Sanchez, J J Li, S A Li, K R Reuhl, P E Thomas, A H Conney, M A Gallo, F C Kauffman, and S Mesia-Vela The effects of steroidal estrogens in ACI rat mammary carcinogenesis: 17{beta}-estradiol, 2-hydroxyestradiol, 4-hydroxyestradiol, 16{alpha}-hydroxyestradiol, and 4-hydroxyestrone J. Endocrinol., October 1, 2004; 183(1): 91 - 99. [Abstract] [Full Text] [PDF]
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 J. Lepine, O. Bernard, M. Plante, B. Tetu, G. Pelletier, F. Labrie, A. Belanger, and C. Guillemette Specificity and Regioselectivity of the Conjugation of Estradiol, Estrone, and Their Catecholestrogen and Methoxyestrogen Metabolites by Human Uridine Diphospho-glucuronosyltransferases Expressed in Endometrium J. Clin. Endocrinol. Metab., October 1, 2004; 89(10): 5222 - 5232. [Abstract] [Full Text] [PDF]
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A. J. Lee and B. T. Zhu NADPH-DEPENDENT FORMATION OF POLAR AND NONPOLAR ESTROGEN METABOLITES FOLLOWING INCUBATIONS OF 17{beta}-ESTRADIOL WITH HUMAN LIVER MICROSOMES Drug Metab. Dispos., August 1, 2004; 32(8): 876 - 883. [Abstract] [Full Text] [PDF]
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V. H. Black and L. C. Quattrochi MOLECULAR CLONING OF THE GUINEA PIG CYP1A2 GENE 5'-FLANKING REGION: IDENTIFICATION OF FUNCTIONAL AROMATIC HYDROCARBON RESPONSE ELEMENT AND CHARACTERIZATION OF CYP1A2 EXPRESSION IN GPC16 CELLS Drug Metab. Dispos., June 1, 2004; 32(6): 595 - 602. [Abstract] [Full Text] [PDF]
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Y. Tsuchiya, M. Nakajima, S. Kyo, T. Kanaya, M. Inoue, and T. Yokoi Human CYP1B1 Is Regulated by Estradiol via Estrogen Receptor Cancer Res., May 1, 2004; 64(9): 3119 - 3125. [Abstract] [Full Text] [PDF]
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 R. K. Dubey, S. P. Tofovic, and E. K. Jackson Cardiovascular Pharmacology of Estradiol Metabolites J. Pharmacol. Exp. Ther., February 1, 2004; 308(2): 403 - 409. [Abstract] [Full Text] [PDF]
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