Liquid chromatography-mass spectrometry (LC-MS) of steroid hormone metabolites and its applications

Article abstract of DOI:10.1016/j.jsbmb.2010.01.005

Advances in liquid chromatography-mass spectrometry (LC-MS) can be used to measure steroid hormone metabolites in vitro and in vivo. We find that LC-electrospray ionization (ESI)-MS using a LCQ ion trap mass spectrometer in the negative ion mode can be used to monitor the product profile that results from 5α-dihydrotestosterone (DHT)-17β-glucuronide, DHT-17β-sulfate, and tibolone-17β-sulfate reduction catalyzed by human members of the aldo-keto reductase (AKR) 1C subfamily and assign kinetic constants to these reactions. We also developed a stable isotope dilution LC-electron capture atmospheric pressure chemical ionization (ECAPCI)-MS method for the quantitative analysis of estrone (E1) and its metabolites as pentafluorobenzyl (PFB) derivatives in human plasma in the attomole range. The limit of detection for E1-PFB was 740. attomole on column. Separations can be performed using normal-phase LC because ionization takes place in the gas phase rather than in solution. This permits efficient separation of the regioisomeric 2- and 4-methoxy-E1. The method was validated for the simultaneous analysis of plasma E2 and its metabolites: 2-methoxy-E2, 4-methoxy-E2, 16α-hydroxy-E2, estrone (E1), 2-methoxy-E1, 4-methoxy-EI, and 16α-hydroxy-E1 from 5. pg/mL to 2000. pg/mL. Our LC-MS methods have sufficient sensitivity to detect steroid hormone levels in prostate and breast tumors and should aid their molecular diagnosis and treatment.

Full text of DOI:10.1016/j.jsbmb.2010.01.005

Journal of Steroid Biochemistry & Molecular Biology 121 (2010) 546–555
Contents lists available at ScienceDirect
Journal of Steroid Biochemistry and Molecular Biology
journal homepage: www.elsevier.com/locate/jsbmb
Liquid chromatography–mass spectrometry (LC–MS) of steroid hormone
metabolites and its applicationsଝ
Trevor M. Penning∗, Seon-Hwa Lee1, Yi Jin, Alejandro Gutierrez2, Ian A. Blair
Centers of Excellence in Environmental Toxicology and Cancer Pharmacology, Department of Pharmacology, University of Pennsylvania School of Medicine,
Philadelphia, PA 19104-6084, USA
a r t i c l e i n f o
a b s t r a c t
Article history:
Advances in liquid chromatography–mass spectrometry (LC–MS) can be used to measure steroid hor-
mone metabolites in vitro and in vivo. We find that LC–electrospray ionization (ESI)-MS using a LCQ
ion trap mass spectrometer in the negative ion mode can be used to monitor the product profile
that results from 5␣-dihydrotestosterone (DHT)-17␤-glucuronide, DHT-17␤-sulfate, and tibolone-17␤-
sulfate reduction catalyzed by human members of the aldo–keto reductase (AKR) 1C subfamily and assign
kinetic constants to these reactions. We also developed a stable isotope dilution LC–electron capture
atmospheric pressure chemical ionization (ECAPCI)-MS method for the quantitative analysis of estrone
(E1) and its metabolites as pentafluorobenzyl (PFB) derivatives in human plasma in the attomole range.
The limit of detection for E1-PFB was 740 attomole on column. Separations can be performed using
normal-phase LC because ionization takes place in the gas phase rather than in solution. This permits
efficient separation of the regioisomeric 2- and 4-methoxy-E1. The method was validated for the simulta-
neous analysis of plasma E2 and its metabolites: 2-methoxy-E2, 4-methoxy-E2, 16␣-hydroxy-E2, estrone
(E1), 2-methoxy-E1, 4-methoxy-EI, and 16␣-hydroxy-E1 from 5 pg/mL to 2000 pg/mL. Our LC–MS meth-
ods have sufficient sensitivity to detect steroid hormone levels in prostate and breast tumors and should
aid their molecular diagnosis and treatment.
Received 27 September 2009
Received in revised form 9 January 2010
Accepted 11 January 2010
Keywords:
Steroid conjugates
Catechol estrogens
Electron capture atmospheric pressure
chemical ionization
Normal-phase HPLC
© 2010 Elsevier Ltd. All rights reserved.
1. Introduction
Radioimmunoassay or ELISA based methods were once consid-
ered state-of-the-art methods for measuring steroid metabolites
Abbreviations: 3␣-Diol-17G, 3␣-hydroxy-5␣-androstane-17␤-glucuronide; 3␤-
Diol-17G, 3␤-hydroxy-5␣-androstane-17␤-glucuronide; 3␣-Diol-17S, 3␣-hydro-
xy-5␣-androstane-17␤-sulfate; 3␤-Diol-17S, 3␤-hydroxy-5␣-androstane-17␤-
sulfate; DHT, dihydrotestosterone; DHTG, DHT-17␤-glucuro- nide; DHTS, DHT-
17␤-sulfate; E1, estrone; E2, 17␤-estradiol; Tibolone, [7␣,17␣]-17-hydroxy-7-
methyl-19-norpregn-5(10)-en-20-yn-3-one; TibS, tibolone-17␤-sulfate; 3␣-OH-
Tib, 3␣-hydroxy-tibolone; 3␤-OH-Tib, 3␤-hydroxy-tibolone; 3␣-OH-TibS, 3␣-
hydroxy-tibolone-17␤-sulfate; 3␤-OH-TibS, 3␤-hydroxy-tibolone-17␤-sulfate; 2-
methoxy-E1, 2-methoxy-estrone; 4-methoxy-E1, 4-methoxy-estrone; 2-methoxy-
E2, 2-methoxy-17␤-estradiol; 4-methoxy-E2, 4-methoxy-17␤-estradiol; 2-OH-E1,
2,3-dihydroxy-estrone; 2-OH-E2, 2,3-dihydroxy-17␤-estradiol; 4-OH-E1, 3,4-
dihydroxyestrone; 4-OH-E2, 3,4-dihydroxy-17␤-estradiol; AcN, acetonitrile; AKR,
aldo–keto reductase; PFB, pentafluorobenzyl; HQC, high quality control sample;
LLQC, lower limit of quantitation; LQC, lower quality control sample; MQC, middle
quality control sample; QC, quality control sample.
in biospecimens. These approaches now appear to be fraught with
difficultly. First, they can only measure one analyte at a time, thus
multiple assays are required to measure all the metabolites from
a single steroid hormone. Second, steroid metabolites have highly
related structures and in a biospecimen mixtures of stereoisomers,
regioisomers or compounds that differ by only the substitution
of a carbonyl group for an alcohol exist. It is thus not possible
to control for interference in the immunoassay from both known
and unknown structurally related steroids that may be present in
the biological matrix. Third, these immunological approaches do
not give any structural validation of the analyte. Fourth, in many
instances antisera do not exist for all the steroid metabolites of
interest to allow immunodetection in the first place. This is cer-
tainly true for the detection of conjugated steroids.
ଝ
Article from special issue on “Steroid profiling and analytics: going towards
Sterome”.
∗
Corresponding author at: Department of Pharmacology, University of Penn-
The reliability of radioimmunoassays for steroid hormones has
also been questioned by position papers which have documented
the large inter-laboratory variability that exists in measuring
plasma testosterone [1–3] and the difficulty in measuring 17␤-
estradiol (E2) and its metabolites in plasma and urine [4–6]. In
contrast, gas chromatography–mass spectrometry (GC–MS) cou-
sylvania, 130C John Morgan Building, 3620 Hamilton Walk, Philadelphia, PA
19104-6084, USA. Tel.: +1 215 898 9445; fax: +1 215 573 2236.
E-mail address: penning@upenn.edu (T.M. Penning).
1
Current address: Department of Bioanalytical Chemistry, Graduate School of
Pharmaceutical Science, Tohoku University, Aoba-ku, Sendai 980-8578, Japan.
2
Current address: Department of Pediatric Oncology, Dana-Farber Cancer Insti-
tute, Boston, MA 02115, USA.
0960-0760/$ – see front matter © 2010 Elsevier Ltd. All rights reserved.
doi:10.1016/j.jsbmb.2010.01.005

T.M. Penning et al. / Journal of Steroid Biochemistry & Molecular Biology 121 (2010) 546–555
547
pled with stable isotope dilution methodology is sensitive, specific,
and accurate and has been used for the quantitative analysis of
steroid hormones in biological samples such as urine and plasma
[7,8]. Unfortunately this method requires extremely tedious extrac-
tion and derivatization procedures for each sample. However, when
used in conjunction with electron capture negative chemical ion-
ization and tandem MS, very low detection limits can be obtained
for plasma E2 (0.063 pg/mL) [9]. A further drawback of the GC–MS
methods is that, steroid conjugates cannot be analyzed directly.
LC–MS can now circumvent many of these problems. We have
developed negative ion LC–ESI/MS in order to analyze multiple
steroid conjugates directly. We demonstrate the utility of the
method by showing that it can be used to conduct product pro-
filing of the enzymatic reduction of endogenous DHT-conjugates
and conjugates derived from the hormone replacement therapeutic
tibolone catalyzed by members of the aldo–keto reductase (AKR)
1C subfamily. By contrast, LC–ESI/MS of underivatized estrogens
are relatively insensitive in both positive and negative ionization
modes so it cannot be used to determine the concentrations of
estrogens and their metabolites which are present in the low pg/mL
range [10]. To circumvent this problem, we have developed sta-
ble isotope dilution LC–ECAPCI/MS methodology, which can detect
estrogen PFB derivatives in the attomole range on column [10].
This has made it possible to quantify multiple estrogen metabo-
lites with high sensitivity in the same chromatographic run. We
demonstrate the utility of this approach by analyzing E1 and E2,
together with their corresponding 2- and 4-methoxy and 16 ␣-
hydroxy metabolites in plasma. These methods can be adapted to
measure targeted steroid metabolomes within prostate and breast
tumor biopsy samples.
or better. Recombinant AKR1C enzymes were over-expressed and
purified to homogeneity as previously described [11].
2.2. Identification of steroid conjugates produced by AKRs using
LC–MS
Products formed during the reduction of conjugated steroids
catalyzed by AKR1C enzymes were prepared for LC–MS analyses
as follows. Reaction mixtures contained 100 mM potassium phos-
phate buffer (pH 7.0), steroid (36 ␮M DHTG, 45 ␮M DHTS or TibS),
0.5 mM NADPH, and 4% methanol in 450 ␮l of total volume. The
reaction was initiated by the addition of purified enzyme (buffer for
no-enzyme control, 13–33 ␮g of AKR1C1–AKR1C4) and incubated
for 0–90 min at 37 ◦C. Reaction mixtures were extracted twice with
1.5 mL water-saturated ethyl acetate. The pooled organic extracts
were vacuum dried and the residues were re-dissolved in 200 ␮l
50% methanol.
2.3. LC separation of steroid conjugates
Chromatography was performed using a Waters Alliance 2690
HPLC system (Waters Corporation, Milford, MA) coupled to the
mass spectrometer. For DHTG and DHTS, a SunFire C8 column
(4.6 mm × 150 mm, 3.5 ␮m Waters), total run time 30 min was
employed. Solvent A was 5 mM aqueous ammonium acetate in
water, and solvent B was 5 mM ammonium acetate in acetonitrile.
The linear gradient used was as follows: 30% solvent B at 0 min
and 3 min, 50% solvent B at 13 min, 80% solvent B at 14 min and
19 min, and 30% solvent B at 20 min and 30 min with a flow rate
of 0.3 mL/min. For TibS, a Jupiter C18 column (2.0 mm × 150 mm,
5 ␮m, Phenomenex, Torrance, CA) total run time 22 min was
employed. Solvent A was 5 mM aqueous ammonium acetate, and
solvent B was 5 mM ammonium acetate in acetonitrile. The linear
gradient was as follows: 20% B at 0 min and 2 min, 30% B at 8 min
and 12 min, and 20% B at 14 min with a flow rate of 0.5 mL/min. All
separations were performed at ambient temperature.
2. Methods
2.1. Materials
DHT, DHT-17␤-glucuronide (DHTG), DHT-17␤-sulfate (DHTS),
3␣-hydroxy-5␣-androstane-17␤-glucuronide
(3␣-Diol-17G),
and 3␤-hydroxy-5␣-androstane-17␤-sulfate (3␤-Diol-17S) were
obtained from Steraloids (Wilton, NH, USA). The latter compound
was custom synthesized by Steraloids. E2, E1, 2-methoxy-E2, 4-
methoxy-E2, 16␣-hydroxy-E2, 2-methoxy-E1, 4-methoxy-E1, and
16␣-hydroxy-E1 were obtained from Steraloids Inc. (Newport, RI).
16,16,17-[2H3]-E2, 2,4,17-[2H3]-16␣-hydroxy-E2, and 2,4,16,16-
[2H4]-E1 were obtained from C/D/N isotopes (Pointe-Claire,
Quebec, Canada). [2H3]-2-methoxy-E2, [2H3]-4-methoxy-E2,
[2H3]-2-methoxy-E1, [2H3]-4-methoxy-E1, and 2,4,16-[2H3]-E2
were synthesized using standard procedures from [2H3]-methyl
iodide (C/D/N isotopes) and unlabeled E2 or E1. 2,4,16-[2H3]-
16␣-hydroxy-E1 was prepared by sodium borodeuteride/PdCl2
(Sigma–Aldrich, St. Louis, MO) reduction in CH3OD (Sigma–Aldrich)
of 2,4-dibromo-17,17-ethylenedioxy-1,3,5(10)-estratriene-3,16␣-
diol (prepared by standard procedures from estrone) followed by
acid hydrolysis. Tibolone (Tib), Tib-17␤-sulfate (TibS), 3␣- and
3␤-hydroxy-Tib (3␣- and 3␤-OH-Tib), and 3␣- and 3␤-OH-Tib-17S
were provided by N.V. Organon (Oss, Netherlands). Pentafluo-
robenzyl bromide (R-bromo-2,3,4,5,6-pentafluorotoluene) was
obtained from Sigma–Aldrich Chemical Co. (Milwaukee,WI).
Pyridine nucleotides were purchased from Roche Applied Sci-
ence (Indianapolis, IN, USA). HPLC grade water, Optima grade
acetonitrile, methanol, hexane, isopropanol, water, ethyl acetate,
and potassium bromide were purchased from Fisher Scientific
Co. (Fair Lawn, NJ). Ethanol was from Pharmco (Brookfield, CT).
Ammonium acetate was obtained from J.T. Baker (Phillipsburg, NJ).
Heparinized male human plasma was from Biological Specialty
Corporation (Colmar, PA). All other reagents were purchased from
Sigma–Aldrich and were of ACS (American Chemical Society) grade
2.4. MS of steroid conjugates
Analyses were conducted using an LCQ ion trap mass spectrom-
eter (Thermo Fisher, San Jose, CA) equipped with an electrospray
ionization source. The mass spectrometer was operated in the neg-
ative ion mode with a potential of 4.5 kV applied to the electrospray
ionization needle. Operating conditions for DHTG and DHTS were
as follows: heated capillary temperature 220 ◦C, capillary voltage
−4 V, tube lens offset 10 V, nitrogen was used for the sheath gas at
80 psi, and for the auxiliary at 10 (arbitrary units). Operating condi-
tions for TibS were as follows: heated capillary temperature 230 ◦C,
capillary voltage −23 V, tube lens offset −25 V, nitrogen was used
for the sheath gas at 50 psi, and for the auxiliary gas at 30 (arbi-
trary units). Full scanning analyses were performed in the range
of m/z 100–600. Products of the reactions were identified based on
their LC retention times and mass spectra relative to those observed
with the authentic standards. The molecular ions monitored for the
steroid conjugates were as follows: DHTG [M−H−; m/z = 465]; 3␣-
and 3␤-Diol-17G [M−H−; m/z = 467]; DHTS [M−H−; m/z = 369]; 3␣-
and 3␤-Diol-17S [M−H−; m/z = 371]; TibS [M−H−; m/z = 391.5];
3␣- and 3␤-OH-TibS [M−H−; m/z = 393.5].
2.5. LC separation of estrogen PFB derivates
Chromatographic separation of the estrogen PFB derivatives
was conducted using a YMC Silica column (250 mm × 4.6 mm i.d.,
5 ␮m particle size, 120 Å pore size, Waters) heated to 40 ◦C. Sol-
vent A was hexane and solvent B was isopropanol/hexane (30:70,
v/v). The LC conditions were as follows: 2% solvent B at 0 min

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T.M. Penning et al. / Journal of Steroid Biochemistry & Molecular Biology 121 (2010) 546–555
and 2 min, 10% solvent B at 5 min, 20% solvent B at 12 min, 50%
solvent B at 22 min, and 24 min, 2% solvent B at 28 min and
35 min with a flow rate of 0.9 mL/min. A post-column flow-rate
of 0.5 mL/min was used to prevent the source from becoming con-
taminated with carbon deposits. Pre-column filters (2 ␮m; Alltech,
Deerfield, IL) were used to protect the column from particulate
matter.
under nitrogen, and reconstituted in 200 ␮l of isopropanol/hexane
(2:98, v/v), 90 ␮l of which was injected on the LC.
2.9. Data analysis
Calibration curves ranged from 5 pg/mL to 1000 pg/mL. For each
curve, eight different concentrations, distributed over the entire
concentration range, were used. Peak height ratios between the
analytes and their respective internal standard were calculated
from each sample using Finnigan LCquan version 1.2 software.
The data were fit to a linear least-squares regression curve with
a weighting index of 1/x. A water blank, plasma blank, and plasma
blank spiked with internal standard were also processed with each
calibration curve.
2.6. ECAPCI of estrogen PFB derivates
A TSQ 7000 (Thermo Fisher) mass spectrometer was operated
in the APCI negative ion mode under the following condi-
tions: vaporizer temperature 500 ◦C, heated capillary temperature
200 ◦C, corona needle discharge current 15 ␮A, sheath gas pres-
sure (nitrogen) 20 psi, auxiliary gas (nitrogen) 15 (arbitrary units).
The parent and daughter resolution settings were of 0 V for
the methoxy compounds and 10 V for all other analytes and
internal standards. The [M−PFB]− ions of the analytes were
filtered through the first quadrupole, and collision-induced dis-
sociation (CID) was performed using argon as the collision gas
at 3.0 mTorr in the second (rf-only) quadrupole. Product ions
were then detected in the third quadrupole. Collision energies
were optimized for each analyte, and ranged from 25 eV to
45 eV (Table 3). MRM transitions were performed as shown in
Table 3.
2.10. Accuracy and precision
Five replicates of each quality control (QC) sample at each con-
centration level were processed and analyzed along with the 8
standard curve samples. The lower quality control sample (LQC),
middle quality control sample (MQC), and high quality control sam-
ple (HQC) were analyzed on three separate days, while the lower
limit of quantitation (LLOQ) samples were analyzed on 1 day only.
Assay accuracy was assessed by comparing means of the measured
analyte concentrations with the theoretical concentrations in the
QC samples. Within-day precision was expressed as percent rela-
tive standard deviation (% RSD) and was obtained by calculating
the percent ratio between the relative standard deviation of five
replicates (n = 5) and their mean at each concentration within the
same validation run. Inter-day precision, defined as percent rela-
tive standard deviation (% RSD) of three different validation runs
(n = 3) was also assessed.
2.7. Sample preparation for LC–ECAPCI/MS analysis
Heparinized male human plasma was allowed to thaw. Plasma
aliquots (1 mL) were transferred to 10 mL glass centrifuge tubes.
An aliquot (10 ␮l) of analyte solution of the appropriate concen-
tration was added to each sample using a dedicated microsyringe,
followed by vortex mixing. This was followed by an aliquot (10 ␮l;
4 ng) of internal standard solution, which was added using another
dedicated microsyringe. Samples were vortex mixed for 1 min, and
4 mL of ethyl acetate was added. This mixture was mechanically
shaken for 1 h, followed by centrifugation at 4000 RPM for 5 min.
The samples were then placed at −20 ◦C and the lower phase was
allowed to freeze, after which the supernatant was transferred by
decantation into a new 5 mL glass centrifuge tube and evaporated to
dryness under nitrogen at 37 ◦C using an N-Evap Analytical Evap-
orator (Organomation, Berlin, MA). Methanol (200 ␮l) was added
to the dried samples, followed by vortex mixing for 2 min. Water
(800 ␮l) was then added, and the samples were vortex-mixed again.
The reconstituted solution was then placed on a C18 solid phase
extraction (SPE) cartridge (7 mm/3 mL Empore C18 Standard Den-
sity Extraction Disk Cartridges, 3 M, St. Paul, MN), which had been
pre-conditioned with 0.5 mL of methanol and 0.5 mL of water.
This cartridge was placed inside a 15 mL plastic centrifuge tube
(Fisher Scientific). All samples and washes were eluted through
the SPE cartridge by centrifugation at 1800 RPM for 3 min. The car-
tridge was washed sequentially with 1 mL of water and 0.3 mL of
methanol/water (20:80, v/v). The cartridges were then transferred
to a new 15 mL plastic centrifuge tube and the analytes were eluted
with 0.3 mL of methanol. The resulting eluate was transferred to a
1.8 mL plastic centrifuge tube (Fisher Scientific), and evaporated to
dryness under nitrogen at 37 ◦C.
2.11. Determination of steady state kinetic parameters
Initial rates of the NADPH dependent reduction of ketosteroids
and their conjugates catalyzed by AKR1C were measured using
a Hitachi F-2500 fluorescence spectrophotometer (Hitachi Amer-
ica Ltd., New York, NY) by monitoring the change in fluorescence
emission of NADPH. Excitation and emission wavelengths were
set at 340 nm and 450 nm, respectively. Changes in fluorescence
units were converted to nanomoles of cofactor by using standard
curves of fluorescence emission versus known NADPH concentra-
tions. Data were analyzed by nonlinear least-squares fitting to the
equation:
kcat[E][S]
ꢀ =
Km + [S]
where ꢀ is the initial velocity, [E] and [S] are the total molar
concentrations of the enzyme and steroid substrate, respectively,
kcat (s−1) is the turnover number, and Km (␮M) is the apparent
Michaelis–Menten constant for the steroid substrate.
3. Results
3.1. Metabolism of endogenous steroid conjugates by aldo–keto
reductase (AKR) 1C subfamily enzymes
The four aldo–keto reductase (AKR) 1C subfamily members
found in humans (AKR1C1–AKR1C4) have been shown to catalyze
the NADPH dependent reduction of 3-ketosteroids to yield 3␣-
or 3␤-hydroxysteroids with different stereochemical preferences
based on steroid substrate. For example, AKR1C2 is predominately
a 3␣-HSD with DHT; while AKR1C1 is predominately a 3␤-HSD with
the same substrate [12]. By contrast the same two enzymes con-
vert tibolone (a hormone replacement therapeutic pro-drug) only
2.8. PFB derivatization
After drying, samples were reconstituted in 100 ␮l of acetoni-
trile, and were treated with 100 ␮l of pentafluorobenzyl bromide in
acetonitrile (1:19, v/v), followed by 100 ␮l of ethanolic potassium
hydroxide (2:100, w/v). After vortex mixing, samples were allowed
to react at room temperature for 15 min, evaporated to dryness

T.M. Penning et al. / Journal of Steroid Biochemistry & Molecular Biology 121 (2010) 546–555
549
Fig. 1. LC–MS analysis of the reduction of DHTG catalyzed by human AKR1C iso-
forms. A, the ion chromatogram (m/z 450–500) of a mixture of authentic standards
of DHTG and 3␣-Diol-17G; B–F, corresponding ion chromatograms of reaction sam-
ples containing no enzyme (B) and AKR1C1–AKR1C4 (C–F). Samples were prepared
as described in Section 2. Reproduced with permission from the American Society
of Biochemists and Molecular Biologists.
Fig. 2. LC–MS analysis of the reduction DHTS catalyzed by human AKR1C isoforms.
A, the ion chromatogram (m/z 350–400) of a mixture of authentic standards of
DHTS and 3␤-Diol-17-S; B–F, corresponding ion chromatograms of reaction sam-
ples containing no enzyme (B) and AKR1C1–AKR1C4 (C–F). Samples were prepared
as described in Section 2. Reproduced with permission from the American Society
of Biochemists and Molecular Biologists.
to its 3␤-hydroxysteroid (estrogenic metabolite) form [13]. Sev-
eral other human AKRs, not involved in steroid metabolism, were
shown to catalyze the reduction of glutathionyl conjugates of 4-
hydroxy-2-nonenal [14]. These observations led us to ask whether
human AKR1C1–AKR1C4 isoforms were capable of metabolizing
steroid conjugates e.g., glucuronides and sulfates. This required the
development of LC–ESI/MS methods using negative ionization for
product profiling so that kinetic constants could be assigned to the
respective reactions with confidence.
The LC–ESI/MS method for detecting steroid conjugates was val-
idated as follows: linear standard curves were obtained in the range
0.5–50 ␮g/mL in the reaction buffer for steroid conjugates (data not
shown). Replicate determinations (n = 3) of quality control samples
were conducted at the low concentration (1.0 ␮g/mL), mid-range
concentration (25 ␮g/mL), and high concentration (50 ␮g/mL) on
three separate days. The intra- and inter-day accuracy of all quality
control samples was within 100 15% of theoretical with a preci-
sion (coefficient of variation) better than 15%.
Using DHTG as substrate LC–ESI/MS product profiling showed
that AKR1C1–AKR1C3 all converted this substrate to 3␤-Diol-
17G (RT = 7.4 min); while AKR1C4 converted DHTG to 3␣-Diol-17G
(RT = 12.6 min), Fig. 1. An authentic synthetic standard only existed
for 3␣-Diol-17G but the 3␤-isomer was positively identified based
on its molecular ion which is identical to that for the 3␣-isomer
and its retention time (see below). DHTS was next tested as
substrate for the AKR1C1–AKR1C4 isoforms. LC–ESI-MS product
profiling showed that the product profile for the sulfate conjugate
was identical to that observed for free DHT. Thus AKR1C1 pro-
duced 3␤-Diol-17S (RT = 13.6 min); AKR1C2 produced 3␣-Diol-17S
(RT = 18.1 min); and AKR1C3 and AKR1C4 produced mixtures of the
3␣- and 3␤-isomers, Fig. 2. An authentic synthetic standard only
existed for 3␤-Diol-17S but the 3␣-isomer was positively identi-
fied based on its molecular ion which is identical to that for the
3␤-isomer. Moreover, the elution order in the ion chromatogram
for the glucuronide and sulfate conjugate products was the same:
3␤-isomer > parent conjugate > 3␣-isomer. The unexpected find-
ings are that both glucuronide and sulfate conjugates could be
reduced by AKR1C enzymes. Moreover, with DHTG and DHTS there
was an inversion of stereochemical preference for AKR1C2 produc-
ing a 3␤-isomer with the first substrate and a 3␣-isomer with the
second substrate.
Assignment of catalytic efficiencies (kcat/Km) to these reactions
for each enzyme isoform was then made based on fluorimetric
assays which monitored the concurrent consumption of NADPH
associated with the appearance of products followed by LC–ESI-MS,
Table 1. It was also found that the kcat/Km values for the reduction of
DHT and DHTG for AKR1C4 were unaffected by the presence of the
glucuronide group but were depressed for the reduction of DHTG
by AKR1C1, and AKR1C2. It was also found that the kcat/Km val-
ues for the reduction of DHT and DHTS for AKR1C1, AKR1C2 and
AKR1C4 were unaffected by the presence of the sulfate group. This
led to the conclusion that AKR1C enzymes can work equally well
on conjugated as well as free DHT [15].
3.2. Metabolism of conjugates of synthetic steroids by aldo–keto
reductase (AKR) 1C subfamily enzymes
We extended our studies on the reduction of steroid conjugates
to include therapeutically relevant steroids. Tibolone is a hormone
replacement therapeutic and pro-drug [16,17]. It has been previ-
ously shown that AKR1C isoforms catalyze the reduction of tibolone
to its 3␣- and 3␤-hydroxy metabolites [13,18]. These reactions are
important since these are the estrogenic metabolites of the pro-
drug and their tissue specific formation may contribute to the tissue
selective estrogen properties of tibolone. By contrast TibS, 3␣-OH-
TibS and 3␤-OH-TibS are the major inactive metabolites of tibolone
[19].

550
T.M. Penning et al. / Journal of Steroid Biochemistry & Molecular Biology 121 (2010) 546–555
Table 1
Table 2
Kinetic parameters for the reduction of DHT, Tib and their conjugates catalyzed by
AKR1C isoforms.
Limits of detection using APCI and ECAPCI for selected estrogen metabolites.
Analyte
Negative APCIa
LOD (fmol)
ECAPCIb LOD (amol)
Increasedc
sensitivity
AKR1C1
AKR1C2
AKR1C4
DHT
kcat (min−1
Km (␮M)
kcat/Km (min−1 M−1
Estrone
2-methoxyestrone
18.52
8.33
740
170
25-fold
50-fold
)
0.74 0.03
4.2 0.5
1.98 0.12
2.9 0.2
3.08 0.15
<0.2
a
Underivatized analyte.
PFB derivative.
Negative ion APCI of underivatized sample compared with ECAPCI of PFB deriva-
)
)
)
)
1.8 × 105
6.8 × 105
>1.5 × 107
b
c
DHTG
kcat (min−1
Km (␮M)
kcat/Km (min−1 M−1
tive.
)
0.53 0.03
16.1 2.4
3.3 × 104
0.24 0.03
4.1 1.2
3.28 0.08
<0.2
5.9 × 104
>1.6 × 107
were available which showed identical RT and negative molec-
ular ions. Assignment of catalytic efficiencies to these reactions
was again performed by conducting fluorimetric assays which
measured the consumption of NADPH, Table 1. These assays
revealed that the reduction of TibS occurred with similar catalytic
efficiencies to those reported for the reduction of tibolone by each
AKR1C isoform [12].
DHTS
kcat (min−1
Km (␮M)
kcat/Km (min−1 M−1
)
0.75 0.05
5.2 0.8
1.0 0.1
4.2 0.5
2.4 × 105
3.07 0.07
<0.2
1.4 × 105
>1.5 × 107
Tib
kcat (min−1
Km (␮M)
)
0.88 0.06
0.76 0.13
1.2 × 106
–
12.7 3.7
0.87 0.34
1.4 × 107
1.83 0.06
1.02 0.07
1.8 × 106
–
kcat/Km (min−1 M−1
Ki (␮M)
0.7 0.29
3.3. Analysis of estrogens by LC–ECAPCI/MS
TibS
kcat (min−1
Km (␮M)
)
1.9 0.3
2.5 0.28
7.6 × 105
–
12.1 2.7
1.2 0.5
1.0 × 107
2.1 0.6
1.20 0.03
0.5 0.1
2.4 × 106
–
Increased risk to breast cancer is associated with life-time
exposure to estrogens. Some estrogen metabolites are thought to
be genotoxic e.g., 4-hydroxy-17␤-estradiol (4-OHE2) while oth-
ers are thought to be non-genotoxic and anti-proliferative e.g.,
2-hydroxy-17␤-estradiol (2-OHE2) [20–23]. The balance between
these competing metabolic pathways may be an important deter-
minant of breast cancer risk, yet the immunochemical methods
available for quantifying endogenous E2 and its metabolites in
plasma have proved to be controversial [4–6]. Stable isotope
LC–MS-based methods can potentially resolve problems in estro-
gen quantification through the rigorous application of stable
isotope dilution methodology. Unfortunately, underivatized estro-
gens are relatively insensitive to conventional LC–ESI/MS-based
methods. Therefore, we developed LC–ECAPCI/MS methodology,
coupled with the use of PFB derivatives in order to substantially
improve the sensitivity of detection for plasma estrogens. In this
method, the nitrogen sheath gas is bombarded with electrons from
the coronal discharge to yield a nitrogen radical cation and a low
energy thermal electron [10] (Fig. 4). An E1 molecule modified to
have a high collision cross section through formation of electron
capturing PFB-ether derivative (Fig. 5) will then undergo highly
efficient dissociative electron capture. This results in the formation
of an intense M-PFB negative ion through the loss of a PFB radical.
Similar results have been obtained with other electron capturing
groups such as p-nitrobenzyl (PNB, Fig. 5) [24]. MRM analysis of the
transition m/z 269 → 145 which monitors the fragmentation of the
C and D rings upon CID provides the necessary rigor for unequivocal
identification of E1 [10]. The limit of detection (LOD) is estimated as
740 attomole and 140 attomole for E1 and 2-methoxy-E1, respec-
tively (Table 2). Thus, ECAPCI/MS was found to increase sensitivity
over traditional negative ion APCI by 25–50-fold.
kcat/Km (min−1 M−1
Ki (␮M)
)
Steady state parameters determined by fluorimetric assay. Values are given as
means SD (n > 2). Values for AKR1C3 catalyzed reactions were not determined due
to low turnover.
The ability of AKR1C1–AKR1C4 isoforms to reduce TibS
was therefore examined. Incubation with recombinant
AKR1C1–AKR1C4 followed by product profiling by LC–ESI/MS
in the negative ion mode showed that AKR1C1–AKR1C3 all
reduced TibS to 3␤-OH-TibS (RT = 9.8 min). By contrast AKR1C4
reduced TibS to a mixture of the 3␣- and 3␤-hydroxy isomers
(RT = 7.0 min and 9.9 min, respectively) (Fig. 3). Identification of the
products was unambiguous since authentic synthetic standards
3.4. Analysis of plasma estrogens by stable isotope dilution
LC–ECAPCI/MS
The specificity of the LC–ECAPCI/MS method was further
increased by the use of stable isotope labeled analogs as inter-
nal standards for each analyte and the use of normal-phase LC
for the separation of isomeric estrogens. The stable isotope dilu-
tion LC–ECAPCI/MS method was validated for the simultaneous
detection of E2 and seven of its plasma metabolites: 2-methoxy-E2,
4-methoxy-E2, 16␣-hydroxy-E2, E1, 2-methoxy-E1, 4-methoxy-
E1, and 16␣-hydroxy-E1 (Table 3). Typical chromatograms for male
plasma samples to which authentic estrogens had been were added
are shown for E2 and its metabolites (Fig. 6) and for E1 and its
Fig. 3. LC–MS analysis of the reduction of TibS catalyzed by human AKR1C isoforms.
(A) The ion chromatogram (m/z 200–400) of a mixture of authentic standards of TibS,
3␣-OH-TibS, and 3␤-OH-TibS; (B–F) Corresponding ion chromatograms of reaction
samples containing no enzyme (B) and AKR1C1–AKR1C4 (C–F). Samples were pre-
pared as described in Section 2. Reproduced with permission from the American
Society of Biochemists and Molecular Biologists.

T.M. Penning et al. / Journal of Steroid Biochemistry & Molecular Biology 121 (2010) 546–555
551
Fig. 4. Proposed mechanism for ECAPCI/MS. Reproduced with permission from Analytical Chemistry.
Fig. 5. Estrone derivatives that provide increased LC–MS sensitivity compared with underivatized estrone.

552
T.M. Penning et al. / Journal of Steroid Biochemistry & Molecular Biology 121 (2010) 546–555
Fig. 7. Stable isotope dilution LC–ECAPCI/MRM/MS analysis of E1 and metabolites.
Fig. 6. Stable isotope dilution LC–ECAPCI/MRM/MS analysis of E2 and metabolites.
basis of the assay is to monitor the formation or consumption
of NAD(P)H linked to the oxidation or reduction of a hydroxy or
ketosteroid. Assay validation is usually performed in discontin-
uous assays which rely upon separating substrate from product
by TLC and identifying unknowns by co-migration with authentic
synthetic standards. Often the TLC systems do not have sufficient
resolving power to separate all mixtures of stereoisomers of
interest and product identify is based on reactions with struc-
turally related steroid substrates. However, bacterial [26–28] and
mammalian HSDs [12,29] are also not always positional or stereo-
specific increasing the number of products possible. The problem
of product identity can be exacerbated if steroid conjugates are
examined as substrates since conjugates do not migrate from the
origin of normal-phase TLC plates.
metabolites (Fig. 7). Linear standard curves were obtained in the
range 5–2000 pg/mL plasma for E2 and E1 (Fig. 8) as well as the
other six E2 metabolites (data not shown). Replicate determina-
tions (n = 5) of quality control samples were conducted one day at
the LLOQ (5 pg/mL). They were conducted on three separate days
for the LQC (10 pg/mL), MQC (50 pg/mL), and HQC (500 pg/mL) on
three separate days. The intra- and inter-day accuracy of all qual-
ity control samples was within 100 15% of theoretical with a
precision (coefficient of variation) better than 15%. The method
can also be further modified by the inclusion of a hydrolysis
step prior to extraction, where the plasma is treated with ␤-
glucuronidase/arylsulfatase step to release the free steroids.
4. Discussion
We have used LC–MS methods to resolve the products of reac-
tions catalyzed by steroid hormone transforming AKRs [15,18].
We show that LC–ESI/MS in the negative ion mode provides suf-
ficient resolving power to generate umambiguous product profiles
for the reduction of steroid conjugates of DHT and the synthetic
steroid tibolone catalyzed by AKR1C isoforms. These methods
Hydroxysteroid dehydrogenases (HSDs) belong to two
gene superfamilies (AKRs) and the short-chain dehydroge-
nase/reductases [25]. Historically many substrate specificity
studies have been performed with these enzymes in which the
Table 3
LC–ECAPCI/MRM/MS analysis of estradiol and its metabolites.
Estrogen
[M−PFB] parent ion (m/z)
Product ion (m/z)
Collision energy (eV)
Retention time (min)
Estradiol (E2)
271
274
301
304
301
304
287
291
269
273
299
302
299
302
287
291
183
185
286
286
286
286
171
173
145
147
284
284
284
284
171
173
43
43
28
28
28
28
40
40
45
45
28
28
28
28
40
40
17.4
17.4
17.7
17.7
18.5
18.5
22.6
22.7
6.95
6.96
7.39
7.40
8.37
8.44
22.6
22.7
[2H3]-estradiol (E2-D3)
2-Methoxy-E2 (2-MeO-E2)
[2H3]-2-methoxy-E2 (2-MeO-E2-D3)
4-Methoxy-E2 (4-MeO-E2)
[2H3]-4-methoxy-E2 (4-MeO-E2-D3)
16␣-Hydroxy-estradiol (16␣-OH-E2)
[2H4]-16␣-hydroxy-estradiol (16␣-OH-E2-D4)
Estrone (E1)
[2H4]-estrone (E1-D4)
4-Methoxy-E1 (4-MeO-E1)
[2H3]-4-methoxy-E1 (4-MeO-E1-D3)
2-Methoxy-E1 (2-MeO-E1)
[2H3]-2-methoxy-E1 (2-MeO-E1-D3)
16␣-Hydroxy-estrone (16␣-OH-E1)
[2H4]-16␣-hydroxy-estrone (16␣-OH-E1-D4)

T.M. Penning et al. / Journal of Steroid Biochemistry & Molecular Biology 121 (2010) 546–555
553
analyzed estrogen metabolites in the plasma of post-menopausal
women where basal levels are in the low pg/mL range [10].
Stable isotope dilution LC–MS methods can provide the optimal
specificity for estrogen analysis because internal standards with
identical physicochemical properties to the relevant analytes are
carried through the entire analysis procedure. This makes it possi-
ble to compensate for losses that occur through all steps. Equally
important is the ability of the stable isotope analogs to act as car-
riers and prevent non-specific losses through binding to glassware
or other surfaces when the corresponding endogenous analytes
are present in only trace amounts. Furthermore, the endogenous
analyte has to meet three strict criteria in order to satisfy the con-
straints of the analytical method—it must have the same parent
ion, product ion as the authentic material, and the relative reten-
tion time to the internal standard should be identical from run
to run. Ideally, [13C]-estrogen analogs should be used as internal
standards but they have only recently become available and so
it has been necessary to used deuterium labeled analogs, which
elute slightly ahead of the protium forms under reversed phase
conditions or slightly after the protium forms under normal-phase
conditions. Therefore, differential suppression or enhancement of
ionization could still affect the quality of the analytical data. For-
tunately, [13C]-analogs of many estrogens have become available
recently from Cambridge Isotope Laboratories (Andover, MA), so it
should be possible to introduce another level of rigor by employ-
ing these standards in the future instead of the deuterium labeled
versions.
Endogenous estrogens are not effectively ionized using con-
ventional ESI or APCI methodology. Therefore, it is necessary to
enhance their ionization characteristics by derivatization and three
approaches have been described. The first approach uses con-
ventional derivatization coupled with LC–ESI/MS. This approach
was exemplified by studies of the Ziegler and co-authors [30],
Yamashita et al. [31], and Xu and Spink [32] groups in which a dan-
syl (D), picolinoyl (P), or pyridyl-3-sulfonyl (PS) group is attached to
the 3-hydroxy moiety of the estrogen (Fig. 5). The second approach
involves the preparation of pre-ionized (quaternized) derivatives,
so that ionization is not required in the ESI source of the mass
spectrometer. This approach was exemplified by studies of the
Chen and co-authors [33], Higashi and co-authors [34], and Adamec
and co-authors [35] groups in which an N-methyl-2-pyridyl (MP),
1-(2,4-dinitro-5-fluorphenyl-4,4,-dimethylpiperazinyl (MPPZ), or
a N-methyl-nicotinyl (NMN) group is attached to the 3-hydroxy
moiety of the estrogen (Fig. 5). The third approach, which we
have pioneered, involves the preparation of an electron captur-
ing PFB derivative of the estrogen (Fig. 5) coupled with the use
of ECAPCI/MS. The Higashi group [24] has also explored the util-
ity of ECAPC/MS for estrogen analysis by using different electron
capturing derivatives such as PNB (Fig. 5).
The three derivatization strategies described are all capa-
ble of quantifying plasma estrogens in the low pg/mL range.
Therefore, in the future, we and others will be able to rigor-
ously establish the precise levels of individual estrogens that are
present in the plasma of post-menopausal women. The ratio of 4-
methoxy-estrogens to 2-methoxy-estrogens provides an indirect
measurement of the catechol estrogens 4-hydroxy-E2/4-hydroxy-
E1 and 2-hydroxy-E2/2-hydroxy-E1. This is potentially important
since the 4-hydroxylated catechols are considered to be geno-
toxic estrogens while the 2-hydroxy catechols are metabolized
very rapidly to 2-methoxy-estrogens that are considered to be
anti-proliferative and protective against mammary carcinogenesis
[20–23].
Fig. 8. Standard curves for E1 and E2 in the range of 5–2000 pg/mL. Linear regression
lines were obtained with r2 values of 0.998 or better.
also highlight the danger of using one reaction to determine the
stereochemical course of another. Thus AKR1C2 will produce the
3␤-hydroxysteroid product from DHTG but the 3␣-hydroxysteroid
product from DHTS. Knowing the product profiles has enabled
us to assign kinetic constants (Km, kcat and kcat/Km) to each of
the reactions with confidence. For AKR1C4 catalytic efficiency for
the reduction of DHT, DHTG and DHTS are unaltered by the pres-
ence of the conjugate group raising the possibility that conjugation
reactions (Phase 2 reactions) could precede reduction reactions
(Phase 1 reactions). In liver we proposed that the conjugation
pathway occurs first for DHT since 17␤-hydroxy-5␣-androstane-
3␣-glucuronide would be the expected metabolite if reduction
occurred first but this product is not detected in the plasma or
urine. Instead the 3␣-OH-DHTG is observed which would be con-
sistent with reduction occurring after conjugation, i.e., the Phase 2
enzyme reaction occurs first [15].
For AKR1C2 and AKR1C4 we show that the catalytic efficiencies
for tibolone and TibS are similar again raising the issue that reduc-
tion of the 3-ketosteroid could follow 17␤-sulfation. If tibolone
conjugation precedes reduction in any tissue, the estrogenic 3␣-
and 3␤-hydroxy-tibolones would not form since they would be
present as their 17␤-sulfates. The difference in the balance between
reduction of tibolone or sulfation of tibolone could contribute to the
selective tissue estrogenic effects of the drug in post-menopausal
women.
We have also exploited the power of stable-isotope dilution
LC–ECAPCI/MS to reliably quantify plasma E1 and E2 and their
downstream metabolites. The use of an electron capturing deriva-
tive such as PFB increases sensitivity by approximately 50-fold
compared with conventional LC–MS methodologies employing
underivatized estrogens. This level of sensitivity is required to
Hormone dependent malignancies also synthesize steroid hor-
mones in the tumor to drive tumor growth [36,37] and this explains
the effectiveness of aromatase inhibitors in treating breast cancer
in post-menopausal women and the potential use of 5␣-reductase

554
T.M. Penning et al. / Journal of Steroid Biochemistry & Molecular Biology 121 (2010) 546–555
inhibitors in treating prostate cancer [38–42]. Considerable effort
has been made in diagnosing these tumors in terms of the presence
of nuclear receptors, real-time-PCR to measure transcript levels of
steroidogenic enzymes, and immunohistochemical approaches to
measure both receptor and enzyme levels. A characteristic property
less developed which is a pre-requisite to complete the molecular
pathology of these tumors are robust methods to measure intra-
tumoral levels of estrogens and androgens. The methods we have
developed offer promise to measure steroid metabolomes within
prostate and breast tumor biopsy samples and the information
gleaned could be used to determine treatment paradigm.
The regulation of ligand occupancy of nuclear receptors is often
governed by pairs of HSDs. For example regulation of the ER is gov-
erned by type 1 17␤-HSD (which reduces E1 to E2) and by type
2/4 17␤-HSD (which oxidizes E2 to E1) [43–46], while regulation
of the AR is governed by AKR1C2 (which reduces 5␣-DHT to 3␣-
diol) and by HSD17B6 (which oxidizes 3␣-diol to DHT) [47,48].
Thus a component of measuring intra-tumoral levels of steroid
hormones is to distinguish between the intracrine formation of
ketosteroids from hydroxysteroids. The LC–ECAPCI-MS method
requires derivatization to an electron capturing group. This can
be accomplished using the pentafuorobenzyl bromide or pentaflu-
orobenzylcarboxymethoxime derivative, for hydroxysteroids and
ketosteroids, respectively. The derivatization with pentafluoroben-
zylcarboxymethoxime was developed originally for EC negative
chemical ionization gas chromatography–MS of ketosteroids Alter-
natively, conventional derivatives such as N-hydroxy-oximes [49]
or pre-ionized derivatives such as those formed by the Girard T
reagent [50] could be employed to improve sensitivity of ketos-
teroid detection by ESI/MS or APCI/MS [51,52]. The next phase of
our work will be to implement such methods.
[10] G. Singh, A. Gutierrez, K. Xu, I.A. Blair, Liquid chromatography/electron cap-
ture atmospheric pressure chemical ionization/mass spectrometry: analysis of
pentafluorobenzyl derivatives of biomolecules and drugs in the attomole range,
Anal. Chem. 72 (14) (2000) 3007–3013.
[11] M.E. Burczynski, R.G. Harvey, T.M. Penning, Expression and characteri-
zation of four recombinant human dihydrodiol dehydrogenase isoforms:
oxidation of trans-7,8-dihydroxy-7,8-dihydrobenzo[a]pyrene to the activated
o-quinone metabolite benzo[a]pyrene-7,8-dione, Biochemistry 37 (32) (1998)
6781–6790.
[12] S. Steckelbroeck, Y. Jin, S. Gopishetty, B. Oyesanmi, T.M. Penning, Human
cytosolic 3␣-hydroxysteroid dehydrogenases of the aldo–keto reductase
superfamily display significant 3␤-hydroxysteroid dehydrogenase activity:
implications for steroid hormone metabolism and action, J. Biol. Chem. 279
(11) (2003) 10784–10795.
[13] S. Steckelbroeck, Y. Jin, B. Oyesanmi, H.J. Kloosterboer, T.M. Penning, Tibolone
is metabolized by the 3␣/3␤-hydroxysteroid dehydrogenase activities of the
four human isozymes of the aldo–keto reductase 1C subfamily: inversion of
stereospecificity with a ꢁ5(10)-3-ketosteroid, Mol. Pharmacol. 66 (6) (2004)
1702–1711.
[14] K. Ramana, B.L. Dixit, S. Srivastava, G.K. Balendiran, S.K. Sivastava, A. Bhat-
nagar, Selective recognition of glutathiolated aldehydes by aldose reductase,
Biochemistry 39 (40) (2000) 12172–12180.
[15] Y. Jin, L. Duan, S.H. Lee, H.J. Kloosterboer, I.A. Blair, T.M. Penning, Human
cytosolic hydroxysteroid dehydrogenases of the aldo–keto reductase super-
family catalyze reduction of conjugated steroids: implications for phase I and
phase II steroid hormone metabolism, J. Biol. Chem. 284 (15) (2009) 10013–
10022.
[16] P. Albertazzi, R. di Micco, E. Zanardi, Tibolone: a review, Maturitas 30 (3) (1998)
295–305.
[17] R. Moore, Livial: a review of clinical studies, Br. J. Obstet. Gynaecol. 106 (Suppl.
19) (1999) 1–21.
[18] S. Stecklebroeck, B. Oyesanmi, Y. Jin, S.-H. Lee, H.J. Kloosterboer, T.M. Penning,
Tibolone metabolism in human liver is catalyzed by 3␣/3␤-hydroxysteroid
dehydrogenase activities of the four isoforms of the aldo–keto reduc-
tase (AKR)1C subfamily, J. Pharmacol. Exp. Therap. 316 (3) (2006) 1300–
1309.
[19] R.M. Vos, S.F. Krebbers, C.H. Verhoeven, L.P. Delbressine, The in vivo human
metabolism of tibolone, Drug Metab. Dispos. 30 (2) (2002) 106–112.
[20] J.Q. Chen, T.R. Brown, J.D. Yager, Mechanisms of hormone carcinogenesis: evo-
lution of views, role of mitochondria, Adv. Exp. Med. Biol. 630 (2008) 1–18.
[21] A.H. Eliassen, S.A. Missmer, S.S. Tworoger, S.E. Hankinson, Circulating 2-
hydroxy- and 16␣-hydroxy estrone levels and risk of breast cancer among
postmenopausal women, Cancer Epidemiol. Biomarkers Prev. 17 (8) (2008)
2029–2035.
Acknowledgements
[22] J. Russo, M. Hasan Lareef, G. Balogh, S. Guo, I.H. Russo, Estrogen and its metabo-
lites are carcinogenic agents in human breast epithelial cells, J. Steroid Biochem.
Mol. Biol. 87 (1) (2003) 1–25.
[23] R. Santen, E. Cavalieri, E. Rogan, J. Russo, J. Guttenplan, J. Ingle, W. Yue, Estrogen
mediation of breast tumor formation involves estrogen receptor-dependent,
as well as independent, genotoxic effects, Ann. N.Y. Acad. Sci. 1155 (2009)
132–140.
This work was supported by the following research grants:
P30ES013587 and R01CA90744 from the National Institutes of
Health and a Prostate Cancer Foundation Challenge Grant (TMP),
R01CA091016 (IAB), and a Pilot-Project Grant awarded with
P30ES13587 (YJ).
[24] T. Higashi, N. Takayama, T. Nishio, E. Taniguchi, K. Shimada, Procedure for
increasing the detection responses of estrogens in LC–MS based on introduction
of a nitrobenzene moiety followed by electron capture atmospheric pressure
chemical ionization, Anal. Bioanal. Chem. 386 (3) (2006) 658–665.
[25] T.M. Penning, Molecular endocrinology of hydroxysteroid dehydrogenases,
Endocr. Rev. 18 (3) (1997) 281–305.
[26] J.H. Abalain, S. Di Stefano, M.L. Abalain-Colloc, H.H. Floch, Cloning, sequencing
and expression of Pseudomonas testosteroni gene encoding 3␣-hydroxysteroid
dehydrogenase, J. Steroid Biochem. Mol. Biol. 55 (2) (1995) 233–238.
[27] D. Ghosh, Z. Wawrzak, C.M. Weeks, W.L. Duax, M. Erman, The refined three-
dimensional structure of 3␣,20␤-hydroxysteroid dehydrogenase and possible
roles of the residues conserved in short-chain dehydrogenases, Structure 15
(10) (1994) 629–640.
References
[1] V. Moal, E. Mathieu, P. Reynier, Y. Malthièry, Y. Gallois, Low serum testosterone
assayed by liquid chromatography–tandem mass spectrometry. Comparison
with five immunoassay techniques, Clin. Chim. Acta 386 (1–2) (2007) 12–19.
[2] L.M. Thienpont, K. Van Uytfanghe, S. Blincko, C.S. Ramsay, H. Xie, R.C. Doss, B.G.
Keevil, L.J. Owen, A.L. Rockwood, M.M. Kushnir, K.Y. Chun, D.W. Chandler, H.P.
Field, P.M. Sluss, State-of-the-art of serum testosterone measurement by iso-
tope dilution–liquid chromatography–tandem mass spectrometry, Clin. Chem.
54 (8) (2008) 1290–1297.
[3] C. Wang, C.H. Catlin, L.M. Demers, B. Starcevic, R.S. Swerdloff, Measurement
of total serum testosterone in adult men: comparison of current laboratory
methods versus liquid chromatography–tandem mass spectrometry, J. Clin.
Endocrinol. Metab. 89 (6) (2004) 534–543.
[4] R. Giese, Measurement of endogenous estrogens: analytical challenges and
recent advances, J. Chromatogr. A. 1000 (1–2) (2003) 401–412.
[5] F. Stanczyk, J.S. Lee, R.J. Santen, Standardization of steroid hormone assays:
why, how and when? Cancer Epidemiol. Biomarkers Prev. 16 (9) (2007)
1713–1719.
[6] X. Xu, J.M. Roman, H.J. Issaq, L.K. Keefre, T.D. Veenstra, R.G. Ziegler, Quantitative
measurement of endogenous estrogens and estrogen metabolites in human
serum by liquid chromatography–tandem mass spectrometry, Anal. Chem. 79
(20) (2007) 7813–7821.
[7] H. Adlercreutz, M.J. Tikkanen, D.H. Hunneman, Mass fragmentographic deter-
mination of eleven estrogens in the body fluids of pregnant and nonpregnant
subjects, J. Steroid Biochem. 5 (3) (1974) 211–217.
[8] H. Adlercreutz, P. Kiuru, S. Rasku, K. Wähälä, T. Fotsis, An isotope dilution gas
chromatographic–mass spectrometric method for the simultaneous assay of
estrogens and phytoestrogens in urine, J. Steroid Biochem. Mol. Biol. 92 (5)
(2004) 399–411.
[9] R.J. Santen, L. Demers, S. Ohorodnik, J. Settlage, P. Langecker, D. Blanchett, P.E.
Goss, S. Wang, Superiority of gas chromatography/tandem mass spectrometry
assay (GC/MS/MS) for estradiol for monitoring of aromatase inhibitory therapy,
Steroids 72 (8) (2007) 666–671.
[28] R. Strickler, D.F. Covey, B. Tobias, Study of 3␣,20 ␤-hydroxysteroid dehydro-
genase with an enzyme-generated affinity alkylator: dual enzyme activity at a
single active site, Biochemistry 19 (22) (1980) 4950–4954.
[29] T.M. Penning, M.E. Burczynski, J.M. Jez, C.-F. Hung, H.-K. Lin, H. Ma, M. Moore,
N. Palackal, K. Ratnam, Human 3␣-hydroxysteroid dehydrogenase isoforms
(AKR1C1–AKR1C4) of the aldo–keto reductase superfamily: functional plas-
ticity and tissue distribution reveals roles in the inactivation and formation of
male and female sex hormones, Biochem. J. 351 (Pt 1) (2000) 67–77.
[30] X. Xu, T.D. Veenstra, S.D. Fox, J.M. Roman, H.J. Issaq, R. Falk, J.E. Saavedra, L.K.
Keefer, R.G. Ziegler, Measuring fifteen endogenous estrogens simultaneously
in human urine by high-performance liquid chromatography–mass spectrom-
etry, Anal. Chem. 77 (20) (2005) 6646–6654.
[31] K. Yamashita, M. Okuyama, Y. Watanabe, S. Honma, S. Kobayashi, M.
Numazawa, Highly sensitive determination of estrone and estradiol in human
serum by liquid chromatography–electrospray ionization-tandem mass spec-
trometry, Steroids 72 (11–12) (2007) 819–827.
[32] L. Xu, D.C. Spink, Analysis of steroidal estrogens as pyridine-3-sulfonyl deriva-
tives by liquid chromatography–electrospray tandem mass spectrometry, Anal.
Biochem. 375 (1) (2008) 105–114.
[33] Y.H. Lin, C.Y. Chen, G.S. Wang, Analysis of steroid estrogens in water using liquid
chromatography/tandem mass spectrometry with chemical derivatizations,
Rapid Commun. Mass Spectrom. 21 (13) (2007) 1973–1983.

T.M. Penning et al. / Journal of Steroid Biochemistry & Molecular Biology 121 (2010) 546–555
555
[34] T. Nishio, T. Higashi, A. Funaishi, J. Tanaka, K. Shimada, Development and appli-
cation of electrospray-active derivatization reagents for hydroxysteroids, J.
Pharm. Biomed. Anal. 44 (3) (2007) 786–795.
[35] W.C. Yang, F.E. Regnier, D. Sliva, J. Adamec, Stable isotope-coded quaternization
for comparative quantification of estrogen metabolites by high-performance
liquid chromatography–electrospray ionization mass spectrometry, J. Chro-
matogr. B: Analyt. Technol. Biomed. Life Sci. 870 (2) (2008) 233–240.
[36] F. Labrie, A. Belanger, J. Simard, Intracrinology. Autonomy and freedom of
peripheral tissues, Ann. Endocrinol. 56 (1) (1995) 23–29.
[37] F. Labrie, V. Luu-The, S.X. Lin, J. Simard, C. Labrie, M. El-Alfy, G. Pelletier, A.
Belanger, Intracrinology: role of the family of 17␤-hydroxysteroid dehydro-
genases in human physiology and disease, J. Mol. Endocrinol. 25 (1) (2000)
1–16.
[44] S. Andersson, 17␤-Hydroxysteroid dehydrogenase: isozymes and mutations, J.
Endocrinol. 146 (2) (1995) 197–200.
[45] S. Andersson, W.M. Geissler, S. Patel, L. Wu, The molecular biology of androgenic
17␤-hydroxysteroid dehydrogenases, J. Steroid Biochem. Mol. Biol. 53 (1–6)
(1995) 37–39.
[46] F. Labrie, V. Luu-The, S.-X. Lin, C. Labrie, J. Simard, R. Breton, A. Belanger, The key
role of 17␤-hydroxysteroid dehydrogenases in sex steroid biology, Steroids 62
(1) (1997) 148–158.
[47] D.R. Bauman, S. Steckelbroeck, M.V. Williams, D.M. Peehl, T.M. Penning, Iden-
tification of the major oxidative 3␣-hydroxysteroid dehydrogenase in human
prostate that converts 5␣-androstane-3␣,17␤-diol to 5␣-dihydrotestosterone:
a potential therapeutic target for androgen dependent disease, Mol. Endocrinol.
20 (2) (2006) 444–458.
[38] A. Brodie, Q. Liu, Y. Liu, B. Long, Aromatase inhibitors and their antitumor effects
in model systems, Endocr. Relat. Cancer 6 (2) (1999) 205–210.
[39] A. Brodie, W.M. Garrett, J.R. Hendrickson, C.H. Tsai-Morris, P.A. Marcotte, C.H.
Robinson, Inactivation of aromatase in vitro by 4-hydroxy-4-androstene-3,17-
dione and 4-acetoxy-4-androstene-3,17-dione and sustained effects, Steroids
38 (6) (1981) 693–702.
[40] H.G. Bull, M. Garcia-Calvo, S. Andersson, W.E. Baginsky, H.K. Chan, D.E.
Ellsworth, R.R. Miller, R.A. Stearns, R. Bakshi, G.H. Rasmusson, R.L. Tolmna,
R.W. Myers, J.W. Kozarich, G.S. Harris, Mechanism-based inhibition of human
steroid 5␣-reductase by finasteride: enzyme-catalyzed formation of NADP+-
dihydrofinasteride, a potent bisubstrate analog, J. Am. Chem. Soc. 118 (10)
(1996) 2359–2365.
[48] T.M. Penning, D. Bauman, Y. Jin, T.L. Rizner, Identification of the molecular
switch that regulates access of 5␣-DHT to the androgen receptor, Mol. Cell
Endocrinol. 265–266 (2007) 77–82.
[49] S.-H. Lee, I.A. Blair, Characterization of 4-oxo-2-nonenal as a novel product of
lipid peroxidation, Chem. Res. Toxicol. 13 (2000) 698–702.
[50] W.J. Griffiths, S. Liu, G. Alvelius, J. Sjovall, Derivatization for the charac-
terisation of neutral oxosteroids by electrospray and matrix-assisted laser
desorption/ionisation tandem mass spectrometry: the Girard P derivative,
Rapid Commun. Mass Spectrom. 17 (9) (2003) 924–935.
[51] T. Higashi, N. Takayama, M. Kyutoku, K. Shimada, E. Koh, M. Namiki, Liquid
chromatography–mass spectrometric assay of androstenediol in prostatic tis-
sue: influence of androgen deprivation therapy on its level, Steroids 71 (11–12)
(2006) 1007–1013.
[41] G.J. Gormley, 5␣-Reductase inhibitors in prostate cancer, Endocr. Relat. Cancer
3 (1) (1996) 57–67.
[52] T. Higashi, A. Yamauchi, K. Shimada, E. Koh, A. Mizokami, M. Namiki,
Determination of prostatic androgens in 10 mg of tissue using liquid
chromatography–tandem mass spectrometry with charged derivatization,
Anal. Bioanal. Chem. 382 (4) (2005) 1035–1043.
[42] W.R. Miller, Aromatase Inhibitors, Endocr. Relat. Cancer 3 (1) (1996) 65–79.
[43] J. Adamski, T. Normand, F. Leenders, D. Monte, A. Begue, D. Stehelin, P.W. Jung-
blut, Y. de, Launoit, Molecular cloning of a novel widely expressed human
80 kDa 17␤-hydroxysteroid dehydrogenase IV, Biochem. J. 311 (Pt. 2) (1995)
437–443.