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NADPH-DEPENDENT FORMATION OF POLAR AND NONPOLAR ESTROGEN METABOLITES FOLLOWING INCUBATIONS OF 17ß-ESTRADIOL WITH HUMAN LIVER MICROSOMES

Department of Basic Pharmaceutical Sciences, College of Pharmacy, University of South Carolina, Columbia, South Carolina

(Received December 10, 2003; accepted April 29, 2004)

	Abstract

Top Abstract Materials and Methods Results and Discussion Conclusions References

 
By using a versatile high-pressure liquid chromatography method (total elution time 135 min) developed in the present study, we detected the formation of some 20 nonpolar radioactive metabolite peaks (designated as M1 through M20), in addition to a large number of polar hydroxylated or keto metabolites, following incubations of [³H]17ß-estradiol with human liver microsomes or cytochrome P450 3A4 in the presence of NADPH as a cofactor. The formation of most of the nonpolar estrogen metabolite peaks (except M9) was dependent on the presence of human liver microsomal proteins, and could be selectively inhibited by the presence of carbon monoxide. Among the four cofactors (NAD, NADH, NADP, NADPH) tested, NADPH was the optimum cofactor for the metabolic formation of polar and nonpolar estrogen metabolites in vitro, although NADH also had a weak ability to support the reactions. These observations suggest that the formation of most of the nonpolar estrogen metabolite peaks requires the presence of liver microsomal enzymes and NADPH. Chromatographic analyses showed that these nonpolar estrogen metabolites were not the monomethyl ethers of catechol estrogens or the fatty acid esters of 17ß-estradiol. Analyses using liquid chromatography/mass spectrometry and NMR showed that M15 and M16, two representative major nonpolar estrogen metabolites, are diaryl ether dimers of 17ß-estradiol. The data of our present study suggest a new metabolic pathway for the NADPH-dependent, microsomal enzyme-mediated formation of estrogen diaryl ether dimers, along with other nonpolar estrogen metabolites.

During our recent analysis of the NADPH-dependent metabolism of [³H]E₂ and [³H]estrone to various hydroxylated or keto metabolites by human liver microsomes (Lee et al., 2001, 2002), we consistently detected a cluster of coeluted radioactive peaks at the end of a 60-min HPLC run, with their chromatographic polarities less than estrone. Notably, similar nonpolar radioactive peaks were also noted earlier when radioactive E₂ or estrone was incubated with rat or mouse liver microsomes (Aoyama et al., 1990; Haaf et al., 1992; Suchar et al., 1995, 1996; Zhu et al., 1997, 1998). Further characterization of these nonpolar metabolite peaks has never been pursued, largely because there was no evidence in the literature that would provide a rationale for the suggestion that highly lipophilic metabolites would be formed from steroid hormones or xenobiotics by microsomal enzymes using NADPH as a cofactor. However, in our recent study to characterize the NADPH-dependent metabolism of [³H]E₂ and [³H]estrone by 15 selectively expressed human P450 isozymes, we noted that this cluster of radioactive nonpolar metabolite peaks appeared to be selectively formed in large quantities only with certain human P450 isozymes, most notably, CYP3A4 and CYP3A5 (Lee et al., 2003). This interesting observation has prompted us to investigate further the possibility that the formation of some of these nonpolar radioactive E₂ metabolite peaks might be dependent on the presence of specific drug-metabolizing enzymes, such as certain P450 isoforms.

In the present study, we first developed a versatile HPLC method for the detection and quantification of various polar and nonpolar metabolites formed by human liver microsomes using [³H]E₂ as a substrate. By using this new method, we have studied the formation of these nonpolar estrogen metabolite peaks by human liver microsomal enzymes and NADPH. Two of the major nonpolar metabolites (M15 and M16) formed were identified to be the diaryl ether dimers of E₂.

	Materials and Methods

Top Abstract Materials and Methods Results and Discussion Conclusions References

 
Chemicals and Human Liver Microsomes. E₂, NADPH, and ascorbic acid were purchased from Sigma-Aldrich (St. Louis, MO). [2,4,6,7,16,17-³H]E₂ (specific activity of 123.0 Ci/mmol) was obtained from PerkinElmer Life and Analytical Sciences (Boston, MA). The scintillation cocktail (ScintiVerse LC) and all the solvents (HPLC grade or better) were purchased from Fisher Scientific (Atlanta, GA). Human liver microsomes used in this study were obtained from Human Biologics International (Scottsdale, AZ) and BD Gentest (Woburn, MA).

Assay of the NADPH-Dependent Metabolism of [³H]E₂ by Human Liver Microsomes. It is of note that the conditions used for the in vitro metabolic formation of the nonpolar E₂ metabolites were exactly the same as those used in our recent studies of the NADPH-dependent oxidative metabolism of estrogens (Lee et al., 2001, 2002). Specifically, the reaction mixture consisted of human liver microsomes (1 mg of protein/ml), a desired concentration of E₂ (containing 2 µCi of [³H]E₂) in 5 µl of ethanol, 2 mM NADPH, and 5 mM ascorbic acid in a final volume of 0.5 ml of 0.1 M Tris-HCl/0.05 M HEPES buffer, pH 7.4. The enzymatic reactions were initiated by the addition of liver microsomes, and the incubations were carried out at 37°C for 20 min with periodic mild shaking. The reactions were arrested by placing the reaction tubes on ice followed by addition of 10 µl of 10 mM nonradioactive E₂ substrate to reduce nonspecific adsorption of estrogens to the test tubes. The mixture was then immediately extracted with 8 ml of ethyl acetate, and the 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 an aliquot (50 µl) was injected into the HPLC apparatus for analysis of estrogen metabolite composition. Notably, all the test tubes used in our present study were silanized with 5% (v/v) dimethyldichlorosilane in toluene as described in our recent studies (Lee et al., 2001, 2002, 2003). The ³H-labeled E₂ was repurified by HPLC a day before it was used as a substrate in the in vitro metabolism experiments.

HPLC Analytical System. Analysis of radioactive E₂ metabolites was performed with an HPLC system coupled with in-line radioactivity and UV detection. The HPLC system consisted of a Waters 2690 separation module (Waters, Milford, MA), a radioactivity detector (ß-RAM; IN/US Systems, Inc., Tampa, FL), a Waters UV detector (model 484), and an Ultracarb 5 ODS column (150 x 4.60 mm; Phenomenex, Torrance, CA). The solvent system for the separation of E₂, estrone, and their metabolites consisted of acetonitrile (solvent A), water (solvent B), and methanol (solvent C).

The solvent gradient (solvent A/solvent B/solvent C) used for the selective separation of various polar estrogen metabolites (method I) was as follows: 8 min of an isocratic elution at 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 an 8.9-min isocratic period at 92:5:3. The gradient was then returned to the initial condition (16:68:16) and held for 10 min before analysis of the next sample.

The solvent gradient (solvent A/solvent B/solvent C) used for simultaneous separation of both polar and nonpolar estrogen metabolites (method II) was as follows: 8 min of an isocratic elution at 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, 32 min of a linear gradient (curve number 6) to 30:40:30, 10 min of a concave gradient (curve number 9) to 35:35:30, 10 min of a concave gradient (curve number 9) to 40:30:30, 10 min of a concave gradient (curve number 9) to 45:25:30, 10 min of a concave gradient (curve number 9) to 50:20:30, 10 min of a concave gradient (curve number 9) to 55:15:30, 5 min of a concave gradient (curve number 9) to 60:10:30, and 5 min of a concave gradient (curve number 9) to 65:5:30. The gradient was then returned to the initial condition (16:68:16) and held for 15 min before analysis of the next sample.

The solvent gradient (solvent A/solvent B/solvent C) used to selectively separate nonpolar estrogen metabolites (method III) was as follows: 20 min of an isocratic elution at 30:40:30, 10 min of a convex gradient (curve number 3) to 35:30:35; 10 min of a convex gradient (curve number 3) to 40:25:35, 10 min of a convex gradient (curve number 3) to 50:20:30, 10 min of a convex gradient (curve number 3) to 60:10:30, followed by 10 min of convex gradient (curve number 3) to 65:5:30. The gradient was then returned to the initial condition (30:40:30) and held for 5 min before analysis of the next sample.

LC/MS Analysis of the Dansyl Derivative of M15 and M16. M15 and M16 were formed following large-scale incubations of E₂ with selectively expressed human CYP3A4 in the presence of NADPH, and then were isolated by using the HPLC (methods I, II, and III). After evaporation to dryness of the collected HPLC fraction containing M15 or M16, the residues were then redissolved in 100 µl of sodium bicarbonate buffer (100 mM, pH adjusted to 10.5 with 1 N NaOH), and followed by vortex-mixing for 1 min. Dansyl chloride solution (100 µl, at 1.0 mg/ml in acetone) was then added and followed by vortex-mixing for another 1 min. The samples were incubated at 60°C for 2 h in a heat block and then cooled to room temperature. An aliquot (10 µl) was injected into the LC/MS for determination of the mass. The LC/MS system consisted of an Agilent 1100 HPLC apparatus (Palo Alto, CA), an AquaSep column (50 x 2.0 mm; ES Industries, West Berlin, NJ), and a Micromass Quattro LC triple quadruple MS detector (Waters). The solvent system consisted of water with 0.1% formic acid (A) and 95% acetonitrile with 0.1% formic acid (B) at a 0.2 ml/min flow rate. The mobile phase gradient (A/B) was as follows: 4 min of an isocratic elution at 80:20, 26 min of a linear gradient to 0:100, and 30 min of an isocratic elution at 0:100. The mass was detected using an electrospray mode at a 3.0-kV capillary voltage and a 32-V cone voltage. The source block temperature and desolvation temperature were set at 100 and 320°C, respectively.

	Results and Discussion

Top Abstract Materials and Methods Results and Discussion Conclusions References

 
Establishment of an HPLC Method for the Separation of Nonpolar Estrogen Metabolite Peaks. In several of our recent studies (Lee et al., 2001, 2002, 2003), we had used an HPLC method that could selectively separate a large number of polar metabolites (mostly hydroxylated and keto metabolites) of E₂ and estrone in a 60-min HPLC run. A representative HPLC trace is shown in Fig. 1 (as a small upper inset). With this HPLC method (as well as some other similar HPLC methods used by others), the nonpolar estrogen metabolite peaks were always eluted together at the end of a chromatogram. To develop an HPLC method that could separate both polar and nonpolar estrogen metabolites in a single run, we focused on selectively modifying the last 20 min of the HPLC mobile phase gradient while still using the original column. We chose this strategy because we desire to retain the versatile ability of our original HPLC method for the separation of all polar estrogen metabolites but, simultaneously, to also to acquire additional ability to resolve the nonpolar estrogen metabolite peaks. Experimentally, we modified each component of the mobile phase gradient in a systematic and incremental manner and then tested each modified gradient for the separation of polar and nonpolar [³H]E₂ metabolites formed with a representative human liver microsomal preparation. We found an elution gradient that could efficiently separate both polar and nonpolar [³H]E₂ metabolites in a single 135-min HPLC run. This method resolved the nonpolar estrogen metabolites (formed by human liver microsomes) into >20 well separated peaks, while it retained almost the same ability of the original HPLC method for the separation of the polar E₂ metabolites (Fig. 1, lower panel). Purely for convenience of description, major visible nonpolar estrogen metabolite peaks that were detected after incubations of [³H]E₂ with human liver microsomes and NADPH were tentatively designated as M1 through M20 (Fig. 1). Details of the mobile phase elution gradient of this method (method II) were described under Materials and Methods.

View larger version (32K): [in this window] [in a new window]   FIG. 1. Representative HPLC traces showing the separation and detection of both polar and nonpolar [3H]E2 metabolites formed by human liver microsomes and NADPH. The incubation mixtures consisted of human liver microsomes (1 mg protein/ml), 50 µM E2 (containing 2 µCi [3H]E2), 2 mM NADPH, and 5 mM ascorbic acid in a final volume of 0.5 ml of Tris-HCl (0.1 M) and HEPES (50 mM) buffer, pH 7.4. The human liver microsomes (HBI226, upper inset; HBI217, lower panel) were obtained from Human Biologics International. The method I mobile phase gradient (described under Materials and Methods) was used to generate the HPLC trace in the upper small inset, and the method II gradient was used to generate the main HPLC trace in the lower panel. The polar estrogen metabolites were identified by comparing their HPLC and gas chromatography retention times as well as the gas chromatography/MS mass fragmentation pattern of their trimethylsilyl derivatives against those of authentic reference compounds as described in our recent study (Lee et al., 2001).

Because this HPLC method was based on the detection of ³H-labled estrogen metabolites, it required the use of 3 ml/min of the scintillation cocktail, and a single 135-min HPLC run would produce a rather large volume (400 ml) of radioactive organic waste. We thus also carefully looked into the option of markedly shortening the HPLC elution time by selectively separating the nonpolar estrogen metabolites while quickly eluting out all polar estrogen metabolites in clusters. After gradual alterations of the mobile phase gradients, we developed a shortened HPLC method (method III) for the selective separation of nonpolar radioactive estrogen metabolite peaks (Fig. 2C). With this method, all polar E₂ metabolites were eluted within the first 10 min as clusters, followed by the elution of nonpolar estrogen metabolites in the next 60 min.

View larger version (40K): [in this window] [in a new window]   FIG. 2. Detection of various polar and/or nonpolar metabolites of [3H]E2 formed by human liver microsomes using three different HPLC methods. The composition of the incubation mixtures was the same as that described in the legend to Fig. 1, except that 20 µM E2 was used as substrate. Human liver microsomes (HG42, obtained from BD Gentest) were used in this study. Details of the three HPLC mobile phase gradients (methods I, II, and II) were described under Materials and Methods.

It is also of note that when the polar and/or nonpolar estrogen metabolite peaks formed by a representative human liver microsomal preparation were analyzed using all three HPLC methods described here (methods I, II, and III), the metabolite profiles and the absolute rates of their formation appeared to be quite consistent (representative HPLC traces shown in Fig. 2). Although we did not have authentic standards to match with these metabolite peaks, attempts were still made to tentatively label most of the peaks, largely based on comparisons of the relative quantity ratios between the metabolites.

NADPH-Dependent, Human Liver Microsome-Mediated Formation of Nonpolar E₂ Metabolites. Using the new HPLC methods developed in this study, we determined whether the formation of the nonpolar estrogen metabolite peaks was dependent on the presence of human liver microsomal enzymes. We first compared the formation of these nonpolar radioactive estrogen metabolites with control and boiled human liver microsomes (Fig. 3, A and B). Our data showed that incubation of [³H]E₂ with control human liver microsomes in the presence of NADPH formed various polar and nonpolar estrogen metabolites. Notably, the peaks of various polar and nonpolar estrogen metabolites were substantially smaller than those shown in Fig. 1, because this particular microsomal preparation had a much lower activity for testosterone 6ß-hydroxylation (an indicator of human CYP3A activity) compared with the one used for generating the HPLC trace shown in Fig. 1. Nevertheless, the overall metabolite profiles produced by these two human liver microsomal preparations were quite similar. The same liver microsomes after boiling for 10 min did not show any detectable activity for the formation of either polar or nonpolar estrogen metabolites (Fig. 3B).

View larger version (33K): [in this window] [in a new window]   FIG. 3. Effect of heat treatment and carbon monoxide on the formation of polar and/or nonpolar metabolites of [3H]E2 by human liver microsomes. The composition of incubation mixtures was the same as that described in the legend to Fig. 1. In this experiment, human liver microsomes (HG42, obtained from BD Gentest) were assayed. To deactivate the microsomal enzymes, the microsomes were boiled for 10 min before assay of their metabolizing activity (panel B). To inhibit microsomal monooxygenases, CO was bubbled into the reaction mixture during the incubation (panel C). The selectively expressed human CYP3A4 was also determined for its catalytic activity for the formation of various nonpolar estrogen metabolites, and the data are shown for comparison (panel D).

To probe whether human liver microsomal monooxygenases (mainly the P450 enzymes) were involved in the formation of various nonpolar estrogen metabolites, we also compared the effects of carbon monoxide (CO) on the formation of both polar and nonpolar estrogen metabolites. When CO was bubbled into the reaction mixture (at 2 small bubbles per second) during the reaction, the formation of polar and nonpolar metabolites from [³H]E₂ was almost completely inhibited (Fig. 3C). Only some residual activity for the conversion of E₂ to estrone and for the formation of M9 was detected. Thus, it was suggested that the formation of most of the nonpolar E₂ metabolites (except M9) requires the presence of microsomal monooxygenases (likely P450 enzymes), which is analogous to the formation of polar oxidative estrogen metabolites.

Notably, in our recent study to characterize the NADPH-dependent metabolism of [³H]E₂ and [³H]estrone by 15 selectively expressed human P450 isozymes (Lee et al., 2003), we noted that a cluster of nonpolar estrogen metabolite peaks (not separated) appeared to be formed only with a few human P450 isozymes, most notably CYP3A4 and CYP3A5. To probe the role of human CYP3A enzymes in the formation of nonpolar estrogen metabolites, we also determined the formation of nonpolar estrogen metabolites following incubations of [³H]E₂ with the selectively expressed human CYP3A4 or 3A5 plus NADPH. Our data showed that the overall profiles of the nonpolar estrogen metabolites formed by human liver microsomes and CYP3A4 or 3A5 were quite similar (a representative data set with CYP3A4 is shown in Fig. 3D). Interestingly, although peak M9 was a major nonpolar metabolite peak formed with human liver microsomes, it was essentially not formed with human CYP3A4 or 3A5. Here it should also be noted that, under the same in vitro reaction conditions, human CYP1A2 (a P450 isoform which we recently confirmed to have the highest activity for the oxidative metabolism of E₂ to 2-hydroxyestradiol) did not have any appreciable catalytic activity for the formation of nonpolar estrogen metabolites (data not shown). This observation suggested that the overall catalytic activity of different human P450 isoforms for the oxidative metabolism of E₂ did not necessarily correlate with their ability for the formation of nonpolar estrogen metabolites. It appeared that only certain P450 isoforms were involved in the formation of the nonpolar estrogen metabolite peaks.

Since it is known that P450 enzymes use NADPH as their optimum physiological cofactor, we, thus, also compared the ability of various other cofactors (NAD, NADP, NADH, and NADPH) to support the metabolic formation of nonpolar estrogen metabolites (Fig. 4). In the absence of any exogenously added cofactor, the human liver microsomes (HG42) only had a weak catalytic activity for the conversion of E₂ to estrone and for the formation of the metabolite peak M9. When NAD was used as a cofactor, E₂ was rapidly converted to estrone by human liver microsomes, suggesting that the 17ß-hydroxysteroid dehydrogenase was the predominant E₂-metabolizing activity in human liver microsomes when NAD was used as a cofactor. This observation is consistent with the earlier reports that NAD is a preferable cofactor for the metabolic conversion of E₂ to estrone by human type II 17ß-hydroxysteroid dehydrogenase (Luu-The et al., 1995). Similarly, NADP did not have an appreciable role in supporting liver microsomal metabolism of E₂, except a weak activity for the conversion of E₂ to estrone and the formation of M9. However, in the presence of NADH, human liver microsomes had weak but detectable activity for the formation of several hydroxylated and nonpolar metabolites, but the overall activities were far weaker compared with the activity seen with NADPH as a cofactor. In conclusion, the formation of various nonpolar E₂ metabolites was NADPH-dependent, similar to the formation of various polar estrogen metabolites.

View larger version (35K): [in this window] [in a new window]   FIG. 4. Effect of different cofactors (NAD, NADP, NADP, and NADPH) on the formation of polar and nonpolar metabolites of [3H]E2 by human liver microsomes. The composition of the incubation mixtures was the same as described in the legend to Fig. 1, except that different cofactors (as indicated) were used. The human liver microsomes (HG42, from BD Gentest) were used.

We also determined the effect of different [³H]E₂ substrate concentrations on the formation of various nonpolar metabolites with human liver microsomes, and some representative data are shown in Fig. 5. Although the rates of conversion of [³H]E₂ to most nonpolar metabolites did not show saturation at the highest substrate concentration (200 µM) tested, it should be noted that very similar curve patterns were also observed in our recent study (Lee et al., 2001) when [³H]E₂ was used as the substrate for the NADPH-dependent metabolism to various polar E₂ metabolites by the same human liver microsomes.

View larger version (38K): [in this window] [in a new window]   FIG. 5. Effect of different substrate concentrations on the formation of major nonpolar [3H]E2 metabolites by human liver microsomes. The experimental conditions were the same as those described in the legend to Fig. 1, except that varying concentrations (1, 10, 20, 50, 100, and 200 µM) of E2 were used. In this experiment, human liver microsomes (HH18, obtained from BD Gentest) were used. Each point was the mean of duplicate determinations, and each curve was obtained by using the nonlinear regression program of Prism software (GraphPad Software Inc., San Diego, CA).

To provide information about the structural identity of some of these nonpolar estrogen metabolites, we compared the HPLC retention times of all nonpolar metabolite peaks with various catechol estrogen monomethyl ethers as well as several E₂-17-fatty acid esters. As we had predicted, these known nonpolar estrogen metabolites had very different HPLC retention times from those formed with human liver microsomes, suggesting that the nonpolar estrogen metabolites observed in the present study were a group of previously unknown metabolites. Since M15 and M16 (two representative nonpolar estrogen metabolites) are selectively formed with human CYP3A4, we thus incubated E₂ with human CYP3A4 and NADPH to produce small amounts of M15 and M16 (two representative nonpolar estrogen metabolites) for structural analysis by LC/MS. M15 and M16 were isolated to purity by sequentially using the HPLC methods (methods I, II, and III). They were then converted to dansyl derivatives according to a method described in a recent study (Anari et al., 2002) and analyzed with LC/MS. Initial LC/MS analysis of the dansyl derivatives of the metabolically formed M15 and M16 showed that each had a specific peak with a m/z of 777 (as M + H). This information suggested that the molecular weights of M15 and M16 could be 542, based on the following calculation: 777 (M + H) - 1 (H) - 234 (a dansyl group) = 542. Additional mass spectrometric analysis using a direct-probe approach confirmed that the molecular weights of both M15 and M16 were 542. Given the fact that the molecular weight of E₂ is 272, a molecular weight of 542 would thus give a possibility that M15 and M16 might be certain E₂ dimers. High-resolution mass spectrometric analysis showed that the molecular weights of M15 and M16 were 542.3400 and 542.3397, respectively. Based on the precise molecular weight information, the same chemical formula, C₃₆H₄₆O₄ (with its calculated molecular weight of 542.3396), was automatically suggested for both M15 and M16, with only 0.7 and 0.2 ppm of error, respectively. Notably, the formula (C₃₆H₄₆O₄) derived from high-resolution mass spectrometric analysis was consistent with the suggestion that M15 and M16 are E₂ dimers: 2 x C₁₈H₂₄O₂ (E₂) - 2H = C₃₆H₄₆O₄.

Definitive evidence for the confirmation of M15 and M16 as E₂ dimers was obtained from multiple NMR analyses. The ¹H NMR spectra (full scale) for M15 and M16 are shown in Fig. 6 (panels B and C, respectively). Overall, the ¹H NMR spectra of M15 and M16 were quite similar to the spectrum of E₂, except for the aromatic region ( 68). Because only very small amounts of metabolically formed M15 and M16 were available, a few additional peaks coming from impurities and solvents were also shown in the NMR spectra. However, these additional peaks could be rationally excluded by analyzing their two-dimensional NMR spectra. According to the NMR analyses, the structures of M15 and M16 were identified to be the dimers of E₂, which were connected together through a diaryl ether bond between a phenolic oxygen atom of one E₂ molecule and the 2- or 4-position aromatic carbon of the other E₂ molecule (structures shown in Fig. 7). Detailed information on the structural identification of M15 and M16 is desribed elsewhere (Lee et al., 2004a).

View larger version (37K): [in this window] [in a new window]   FIG. 6. Structural determination of M15 and M16 by NMR spectrometric analyses. The NMR spectra were recorded on a Varian Inova 500 spectrometer operating at a proton frequency of 500 MHz and a carbon frequency of 125 MHz. Chemical shifts were given as values with reference to CD3OD as an internal standard. d, doublet; d, doublet with smaller coupling constant; dd, doublet of doublet; s, singlet. A, partial structures (only showing the aromatic rings of two E2 molecules) of three possible E2 dimers with a diaryl ether bond. Their theoretical 1H NMR peak patterns for various aromatic protons are labeled. B, 1H NMR spectrum of M15. C, 1H NMR spectrum of M16. D, 1H NMR spectrum (aromatic region) of M15 and peak assignments with the suggested structure. E, 1H NMR spectrum (aromatic region) of M16 and peak assignments with the suggested structure.

View larger version (14K): [in this window] [in a new window]   FIG. 7. Structures of M15 and M16, two novel diaryl ether dimers of 17ß-estradiol, that were formed with human liver P450 enzymes and NADPH.

Lastly, it is also of note that we have recently successfully synthesized small amounts of M15 and M16 using E₂ as the starting material (Lee et al., 2004). The chemically synthesized M15 or M16, when mixed together with the metabolically formed M15 or M16 and then injected into our HPLC system, had the same retention time as a single HPLC peak. In addition, the ¹H and ¹³C NMR spectra of the metabolically formed and chemically synthesized M15 and M16 matched perfectly with each other (data not shown), further confirming the structural identity of M15 and M16 as novel E₂ dimers.

	Conclusions

Top Abstract Materials and Methods Results and Discussion Conclusions References

 
A versatile HPLC method was developed in the present study for the selective separation and quantification of various nonpolar estrogen metabolite peaks formed by human liver microsomes. We detected 20 nonpolar radioactive metabolite peaks (M1–M20), in addition to a large number of polar hydroxylated or keto metabolites following incubations of [³H]E₂ with human liver microsomes in the presence of NADPH as a cofactor. The formation of most of the nonpolar metabolite peaks (except M9) was dependent on the presence of microsomal proteins and could be selectively inhibited by the presence of carbon monoxide. NADPH was the optimum cofactor for their metabolic formation, although NADH also had a weak ability to support the reactions. Data from these experiments suggested that the formation of most nonpolar estrogen metabolite peaks was dependent on the presence of certain microsomal monooxygenases (such as the P450 enzymes). Efforts are presently underway to determine the structures of these nonpolar estrogen metabolites, and our initial studies showed that M15 and M16, two representative nonpolar metabolites, are the diaryl ether dimers of E₂. It will be of interest to determine the biological activities that may be associated with these novel nonpolar E₂ metabolites.

	Footnotes

 
This study was supported, in part, by a Department of Defense Breast Cancer Research Program Predoctoral Training Award (DAMD17-02-1-0566, to A.J.L.), a National Institutes of Health grant (RO1 CA 92391, to B.T.Z.), and an American Cancer Society research grant (RSG-02-143-01-CNE).

ABBREVIATIONS: P450, cytochrome P450; E₂, 17ß-estradiol; NADP and NADPH, nicotinamide dinucleotide phosphate and its reduced form, respectively; NAD and NADH, nicotinamide dinucleotide and its reduced form, respectively; HPLC, high-pressure liquid chromatography; LC/MS, liquid chromatography/mass spectrometry; CO, carbon monoxide; NMR, nuclear magnetic resonance.

Address correspondence to: Bao Ting Zhu, Department of Basic Pharmaceutical Sciences, College of Pharmacy, University of South Carolina, 700 Sumter Street, Columbia, SC 29208. E-mail: BTZhu{at}cop.sc.edu

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Top Abstract Materials and Methods Results and Discussion Conclusions References

 


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