0090-9556/04/3208-876–883$20.00
DRUG METABOLISM AND DISPOSITION
Copyright © 2004 by The American Society for Pharmacology and Experimental Therapeutics
DMD 32:876–883, 2004
Vol. 32, No. 8
1353/1163922
Printed in U.S.A.
NADPH-DEPENDENT FORMATION OF POLAR AND NONPOLAR ESTROGEN
METABOLITES FOLLOWING INCUBATIONS OF 17␤-ESTRADIOL WITH HUMAN LIVER
MICROSOMES
Anthony J. Lee and Bao Ting Zhu
Department of Basic Pharmaceutical Sciences, College of Pharmacy, University of South Carolina, Columbia, South Carolina
Received December 10, 2003; accepted April 29, 2004
This article is available online at http://dmd.aspetjournals.org
ABSTRACT:
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 17␤-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 17␤-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.
The endogenous estrogens undergo extensive metabolism in humans (Martucci and Fishman, 1993; Zhu and Conney, 1998a), such as
oxidation (largely mediated by P450 enzymes), interconversion between 17␤-estradiol and estrone, and various conjugation-deconjugation reactions. In addition, the catechol-O-methyltransferase-mediated
O-methylation of endogenous catechol estrogens to monomethyl
ethers (Zhu and Conney, 1998b) and the acyltransferase-mediated
esterification of estrogens with fatty acyl-CoAs (Mellon-Nussbaum et
al., 1982; Hochberg, 1998) result in the formation of lipophilic estrogen derivatives. These multiple metabolic pathways not only determine the pharmacokinetic features of the endogenous estrogens in the
body and in various target tissues, but they also diversify the biological actions of endogenous estrogens in certain target sites through
metabolic formation of biologically active estrogen metabolites, such
as 4-hydroxyestradiol, 15␣-hydroxyestradiol, 16␣-hydroxyestrone,
and 2-methoxyestradiol (a predominant O-methylation product of
2-hydroxyestradiol).
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
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, 17␤-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.
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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]17␤-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,
FORMATION OF POLAR AND NONPOLAR ESTROGEN METABOLITES
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
tubes on ice followed by addition of 10 ␮l 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 ␮l of
methanol, and an aliquot (50 ␮l) 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
3
H-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 (␤-RAM; IN/US Systems, Inc.,
Tampa, FL), a Waters UV detector (model 484), and an Ultracarb 5 ODS
column (150 ⫻ 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
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 ␮M E2 (containing 2 ␮Ci [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).
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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 ␮Ci of [3H]E2) in 5 ␮l 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
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LC/MS system consisted of an Agilent 1100 HPLC apparatus (Palo Alto, CA),
an AquaSep column (50 ⫻ 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
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
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 ␮M 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.
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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 ␮l 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 ␮l, 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 ␮l) was injected into the LC/MS for determination of the mass. The
FORMATION OF POLAR AND NONPOLAR ESTROGEN METABOLITES
879
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.
Because this HPLC method was based on the detection of 3H-labled
estrogen metabolites, it required the use of ⱖ3 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.
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 estro-
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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).
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gen metabolites were substantially smaller than those shown in Fig. 1,
because this particular microsomal preparation had a much lower
activity for testosterone 6␤-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).
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
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 ␮M) 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).
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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.
FORMATION OF POLAR AND NONPOLAR ESTROGEN METABOLITES
881
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 17␤-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 17␤-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.
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 concen-
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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.
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tration (200 ␮M) 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.
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
Conclusions
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 ⱖ20 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.
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FIG. 7. Structures of M15 and M16, two novel diaryl ether dimers of 17␤-estradiol,
that were formed with human liver P450 enzymes and NADPH.
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 highresolution mass spectrometric analysis was consistent with the suggestion that M15 and M16 are E2 dimers: 2 ⫻ 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 (␦ 6⬃8). 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).
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.
FORMATION OF POLAR AND NONPOLAR ESTROGEN METABOLITES
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