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DMD # 70185
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Species Differences in the Oxidative Desulfuration of a Thiouracil-Based
Irreversible Myeloperoxidase Inactivator by Flavin-Containing
Monooxygenase Enzymes
Heather Eng, Raman Sharma, Angela Wolford, Li Di, Roger B. Ruggeri, Leonard Buckbinder,
Edward L. Conn, Deepak K. Dalvie, and Amit S. Kalgutkar
Pharmacokinetics, Pharmacodynamics, and Metabolism Department, Pfizer Inc., Groton, CT
(H.E., R.S., A.W., L.D.), La Jolla, CA (D.K.D.) and Cambridge MA (A.S.K.); Worldwide
Medicinal Chemistry (E.L.C., R.B.R.); Cardiovascular and Metabolic Research Unit,
Cambridge, MA (L.B.)
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Running Title: Oxidative desulfuration by flavin-containing monooxygenases
Address correspondence to: Amit S. Kalgutkar, Pharmacokinetics, Dynamics, and
Metabolism-New Chemical Entities, Pfizer Worldwide Research and Development, 610 Main
Street, Cambridge, MA 02139, USA. Tel: +(617)-551-3336. E-mail: amit.kalgutkar@pfizer.com
Text Pages (including references): 30
Tables: 1
Figures: 8
References: 59
Abstract: 252
Introduction: 471
Discussion: 1550
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Abbreviations used are: 1, 6-(2,4-dimethoxyphenyl)-1-(2-hydroxyethyl)-2-thioxo-2,3-
dihydropyrimidin-4(1H)-one; AUC(0-∞), area under the plasma concentration–time curve from
zero to infinity; CID, collision-induced dissociation; CLint, intrinsic clearance; CLp, plasma
clearance; CLrenal, renal clearance; DMSO, dimethyl sulfoxide; FMO, flavin-containing
monooxygenase; GSH, reduced glutathione; H2O2, hydrogen peroxide; LC-MS/MS, liquid
chromatography tandem mass spectrometry; KM, Michaelis-Menten constant; M1, 5-(2,4-
dimethoxyphenyl)-2,3-dihydro-7H-oxazolo[3,2-a]pyrimidin-7-one; MPO, myeloperoxidase;
NADPH, reduced nicotinamide adenine dinucleotide phosphate; P450, cytochrome P450; PTU,
propylthiouracil; SAR, structure-activity relationship; t1/2, half-life; tR, retention time; TPO,
thyroid peroxidase; Vdss, steady state distribution volume; Vmax,maximum rate of oxidative
desulfurization.
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Abstract:
N1-Substituted-6-arylthiouracils (represented by 6-(2,4-dimethoxyphenyl)-1-(2-hydroxyethyl)-2-
thioxo-2,3-dihydropyrimidin-4(1H)-one (1)) represent a novel class of selective irreversible
inhibitors of human myeloperoxidase. The present account represents a summary of our in vitro
studies on the facile oxidative desulfuration in 1 to a cyclic ether metabolite M1 in NADPH-
supplemented rat (t1/2 (mean±standard deviation)=8.6±0.4 min) and dog liver microsomes
(t1/2=11.2±0.4 min), but not in human liver microsomes (t1/2 >120 min). The in vitro metabolic
instability also manifested in moderate-to-high plasma clearances of the parent compound in rats
and dogs with significant concentrations of M1 detected in circulation. Mild heat deactivation of
liver microsomes or co-incubation with the flavin-containing monooxygenase (FMO) inhibitor
imipramine significantly diminished M1 formation. In contrast, oxidative metabolism of 1 to M1
was not inhibited by the pan cytochrome P450 inactivator 1-aminobenzotriazole. Incubations
with recombinant FMO isoforms (FMO1, FMO3, and FMO5) revealed that FMO1 principally
catalyzed the conversion of 1 to M1. FMO1 is not expressed in adult human liver, which
rationalizes the species difference in oxidative desulfuration. Oxidation by FMO1 followed
Michaelis-Menten kinetics with KM, Vmax, and CLint values of 209 μM, 20.4 nmoL/min/mg
protein, and 82.7 μL/min/mg protein, respectively. Addition of excess glutathione essentially
eliminated the conversion of 1 to M1 in NADPH-supplemented rat and dog liver microsomes,
which suggested that the initial FMO1-mediated S-oxygenation of 1 yields a sulfenic acid
intermediate capable of redox cycling to the parent compound in a glutathione-dependent fashion
or undergoing further oxidation to a more electrophilic sulfinic acid species that is trapped
intramolecularly by the pendant alcohol motif in 1.
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Introduction
We recently reported structure-activity relationship (SAR) studies on N1-substituted-6-aryl-2-
thiouracil derivatives as irreversible, mechanism-based inactivators of the hemoprotein
myeloperoxidase (MPO, EC 1.11.2.2) with a high degree of selectivity for MPO relative to
peroxidases such as thyroid peroxidase (TPO) and cytochrome P450 (P450) enzymes (Ruggeri et
al., 2015). The thiouracil analogs behave as suicide substrates of MPO and covalently adduct to
the heme prosthetic group through an oxidized sulfur species (presumably a thiyl radical)
generated during catalysis (Ruggeri et al., 2015; Tidén et al., 2011). The antithyroid drug
propylthiouracil (PTU, Figure 1), which irreversibly inhibits MPO and TPO in a non-selective
fashion (Lee et al., 1990; Ruggeri et al., 2015), was used as a starting point in our SAR work to
identify selective MPO inhibitors. Introduction of polar N1 substituents and replacement of the
C6 propyl group in PTU with electron-rich aromatic functionalities resulted in significant
improvements in MPO inhibitory activity (inferred from inactivation kinetics parameters (kinact
and KI) and partition ratio) and virtually abolished TPO inhibition with the resultant compounds.
Concern over the risk of immune-mediated toxicity (e.g., agranulocytosis and hepatotoxicity)
associated with chronic PTU treatment (Cooper, 2005; Futcher and Massie, 1950; Ichiki et al.,
1998) via oxidative bioactivation of the thiouracil motif to protein- and thiol-reactive
intermediates (Jiang et al., 1994; Lee et al., 1988, Lee et al., 1990; Waldhauser and Uetrecht,
1991) was principally mitigated by tethering pendant nucleophilic groups to the N1-substituent,
which could potentially quench reactive species in an intramolecular fashion. Out of this
exercise emerged the lead compound 6-(2,4-dimethoxyphenyl)-1-(2-hydroxyethyl)-2-thioxo-2,3-
dihydropyrimidin-4(1H)-one (1, Figure 1) with significant improvements noted in MPO
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inactivation potency and selectivity relative to PTU. Intramolecular trapping of reactive species
was demonstrated by reacting 1 with excess hydrogen peroxide (H2O2) (Kalm, 1961; Kitamura,
1934), which quantitatively converted 1 to the pharmacologically inactive cyclic ether 5-(2,4-
dimethoxyphenyl)-2,3-dihydro-7H-oxazolo[3,2-a]pyrimidin-7-one (M1, Figure 1), presumably
via an unstable oxidized sulfur intermediate (Ruggeri et al., 2015). Importantly, no thiol
conjugates of 1 were generated upon addition of reduced glutathione (GSH) to the H2O2 and
MPO/H2O2 incubations. Moreover, compound 1 was resistant towards metabolic turnover in
reduced nicotinamide adenine dinucleotide phosphate (NADPH)-supplemented human liver
microsomes (half-lives (t1/2) > 120 min) and cryopreserved human hepatocytes (t1/2 > 240 min),
which was generally consistent with its physicochemical properties (molecular weight = 308,
lipophilicity (logD7.4 = 1.1), and topological polar surface area = 71.03 Å2). In contrast, a high
metabolic turnover of 1 was noted in NADPH-supplemented rat and dog liver microsomes,
which translated in moderate to high plasma clearance (CLp) in these preclinical species. In vitro
mechanistic studies were initiated to rationalize the species difference in metabolism and
revealed a facile conversion of 1 principally to the cyclic ether metabolite M1 by rat and dog
liver flavin-containing monooxygenase (FMO) 1, which is not expressed in adult human liver.
The collective findings from these studies are reported, herein.
Materials and Methods
Materials. The synthesis of compound 1 (chemical purity > 99% by HPLC and NMR) has been
previously reported (Ruggeri et al., 2015). The preparation of the M1 metabolite is described in
the supplemental section. NADPH, 1-aminobenzotriazole, reduced GSH, and imipramine were
purchased from Sigma-Aldrich (St. Louis, MO). Pooled male Wistar-Han rat and male and
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female human liver microsomes (pool of 126 livers, age 8-10 weeks) were purchased from BD
Gentest (Woburn, MA), male beagle dog liver microsomes (pool of 14 livers, age 1-4 years) and
pooled male and female human kidney microsomes from XenoTech (Kansas City, KS).
Recombinant human FMO1, FMO3, and FMO5 supersomes were purchased from Corning
(Oneonta, New York).
Liver and Kidney Microsomal Stability. Stock solutions of 1 were prepared in dimethyl
sulfoxide (DMSO) then diluted with methanol and acetonitrile. The final concentration of
solvents in the incubation mixture were 0.025% DMSO, 0.5% methanol, and 0.475%
acetonitrile. Assessment of t1/2 in liver or kidney microsomes was determined in triplicate with
microsomes (1 mg/mL microsomal protein for rat, dog, and human) in 0.1�M potassium
phosphate buffer (pH 7.4) containing 3.3 mM MgCl2 at 37�°C. The reaction mixture was
prewarmed with 1.3 mM NADPH at 37�°C for 5�min before initiating the reaction with the
addition of 1 (1 µM). Aliquots of the reaction mixture at 0.25, 5.0, 10, 20, 30, 40 and 60�min
were added to acetonitrile containing 0.1% formic acid and an internal standard terfenadine (2
ng/mL). The samples were centrifuged prior to dilution of supernatant with an equal volume of
water containing 0.1% formic acid and liquid chromatography/tandem mass spectrometry (LC-
MS/MS) analysis of the disappearance of 1 and the formation of cyclized metabolite M1. For
control experiments, NADPH was omitted from these incubations. A parallel incubation of liver
microsomes from rat and dog containing compound 1 (1 μM), GSH (5 mM) and NADPH (1.3
mM) was conducted in order to evaluate redox cycling of the initial S-oxygenation product of 1
formed during the process of enzymatic oxidation. To test the involvement of the thermally
unstable FMO in the oxidation, rat and dog liver microsomes were preincubated at 50 °C for 5
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min in the absence of NADPH co-factor and then cooled on ice followed by incubations
described above. For the purposes of metabolite identification studies, the concentration of 1 in
the microsomal incubations was raised to 10�µM. The duration of the incubation time was
60�min. Separate incubations of 1 (10 μM) were also conducted in human liver microsomes
containing NADPH (1.3 mM) and glutathione (1 mM) for the purposes of trapping potentially
reactive species arising from the oxidative metabolism of 1. To determine the effects of P450 or
FMO inhibition on the metabolic conversion of 1 to M1, 1-aminobenzotriazole (1 mM final
concentration) [P450 inactivator] or imipramine (250 μM final concentration) [FMO competitive
inhibitor] were preincubated with rat and dog liver microsomes in the presence of NADPH for
20 or 2 min, respectively, prior to initiation of the reaction with 1.
Incubations of 1 with Recombinant FMO Isoforms. Potassium phosphate buffer (0.1 M, pH
7.4), magnesium chloride (3.3 mM), NADPH (1.3 mM), and recombinant FMO (0.5 mg/mL)
were combined and pre-warmed at 37 °C for 2 min. To initiate reaction, 1 was added at a final
concentration of 1 µM (final 0.025% DMSO, 0.5% methanol, and 0.475% acetonitrile). At each
time point (0.25, 5.0, 10, 20, 30, 40, and 60 min) a 50 µL aliquot of reaction mixture was
transferred to 200 µL acetonitrile containing 0.1% formic acid and internal standard terfenadine
(2 ng/mL). After centrifugation at 2000 g, equal volumes of supernatant and water containing
0.1% formic acid were mixed, and the disappearance of 1 and formation of M1 was examined by
LC-MS/MS.
To evaluate linearity of product (i.e., M1) formation by human FMO1 and FMO3 isoforms,
recombinant FMO1or FMO3 supersomes (0.1-1 mg/mL) and NADPH (3.3 mM) were
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preincubated in 0.1 M phosphate buffer pH 7.4 for 2 minutes at 37 °C. Reactions (5000 μL)
were initiated by the addition of compound 1 (1 μM), and were allowed to continue at 37 °C for
0.25 to 60 min. Aliquots (50 µL) of reactions were quenched with 200 µL of acetonitrile
containing 0.1% formic acid and internal standard terfenadine (2 ng/mL), and after centrifugation
at 2000 g, supernatants were combined with an equal volume of water containing 0.1% formic
acid, and the formation of M1 was examined by LC-MS/MS.
For determination of the Michaelis-Menten constant KM, maximum rate of oxidative
desulfurization Vmax, and intrinsic clearance (CLint, Vmax/KM) for 1, incubations were repeated at a
single protein concentration and time point determined to be in the linear range of metabolite
formation (0.1 mg/mL FMO1, 1 mg/mL FMO3, 60 min), containing 12 concentrations of 1 (1-
300 µM). Kinetic parameters were obtained for FMO1 using the Michaelis-Menten nonlinear
regression in GraphPad PRISM (La Jolla, CA). As the FMO3 reaction was not saturated within
the range of substrate concentrations tested, the slope of formation rate of M1 versus substrate
concentration (CLint) was calculated using linear regression.
Animal Pharmacokinetics. Dog experiments were conducted in our AAALAC-accredited
facilities and were reviewed and approved by Pfizer Institutional Animal Care and Use
Committee. Rat studies were done at BioDuro, Pharmaceutical Product Development Inc.
(Shanghai, PRC); animal care and in vivo procedures were conducted according to guidelines
from the BioDuro Institutional Animal Care and Use Committee. Jugular vein cannulated male
Wistar-Han rats (~250 g), obtained from Vital River (Beijing, China), and male Beagle dogs (~
8-11 kg) were used for these studies. Rats were provided ad libitum access to water and food.
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Dogs were fasted overnight and fed 4 hr after dosing. Compound 1 was administered
intravenously (i.v.) in 5% DMSO/95% of 30% 2-hydroxypropyl-β-cyclodextrin or 25% 2-
hydroxypropyl-β-cyclodextrin/75% 100 mM Tris buffer (pH 8.0) via the tail vein of rats (n=3) or
saphenous vein (dogs, n=3) at a dose of 1.0 mg/kg in a dosing volume of 1 (rat) or 0.5 (dog)
mL/kg. Serial blood samples were collected before dosing and 0.033 (rat only), 0.083, 0.25, 0.5,
1, 2, 4, 7, and 24 h after dosing. Blood samples from the pharmacokinetic studies were
centrifuged to generate plasma. All plasma samples were kept frozen until analysis. Urine
samples (0–7.0 and 7.0–24 h) were also collected after i.v. administration. Aliquots of plasma or
urine (10-50 μl) were transferred to 96-well blocks and methanol/acetonitrile (1:1, v/v, 200 μl)
containing an internal standard was added to each well. Supernatant was diluted 20-fold with
methanol/water (1:1, v/v) containing 0.1% formic acid. Samples were then analyzed by LC-
MS/MS and concentrations of 1 and M1 in plasma and urine were determined by interpolation
from a standard curve. Range of quantitation was 1-2000 ng/mL for rat (linear R2 0.991 1 and
linear R2 0.996 M1) and 1-5000 ng/mL for dog (linear R2 0.995 1 and quadratic R2 0.978 M1).
Determination of Pharmacokinetic Parameters. Pharmacokinetic parameters were
determined using noncompartmental analysis (Watson v.7.4, Thermo Scientific, Waltham, MA).
The area under the plasma concentration-time curve from t = 0 to 24 h (AUC0-24) and t = 0 to
infinity (AUC0-∞) was estimated using the linear trapezoidal rule and CLp was calculated as the
intravenous dose divided by AUC0–∞i.v.. The terminal rate constant (kel) was calculated by a
linear regression of the log-linear concentration-time curve, and the terminal elimination t1/2 was
calculated as 0.693 divided by kel. Apparent steady state distribution volume (Vdss) was
determined as the i.v. dose divided by the product of AUC0–∞ and kel. Percentage of unchanged 1
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excreted in urine over 24 h was calculated using the following equation: amount (in mg) of 1 in
urine over the 24 h interval post dose/actual amount of the dose of 1 administered (mg) x 100%.
The renal clearance (CLrenal) was derived as the ratio of amount (in mg) of 1 in urine over the 24
h interval post dose/AUC0-24.
LC-MS/MS Analysis for Quantitation of 1 and M1. Concentrations of analytes from in vitro
and in vivo studies were determined on a Sciex 5500 LC-MS/MS triple quadrupole mass
spectrometer (Sciex, Framingham, MA). Analytes were chromatographically separated using
Agilent 1290 (Santa Clara, CA) or Shimadzu LC-20AD (Shimadzu Scientific Instruments, MD)
pumps. A CTC PAL autosampler was programmed to inject 1 or 10 μL on a Phenomenex
Kinetex C18 30 x 3 mm HPLC (Phenomenex, Torrance, CA) or Mac Mod Halo C18 50 x 2.1
mm UPLC column (Mac Mod Analytical, Chadds Ford, PA) using a mobile phase consisting of
water containing 0.1% (v/v) formic acid (solvent A) and acetonitrile containing 0.1% formic
(solvent B) at a flow rate of 0.5 mL/min. Compounds 1 and M1 were detected using
electrospray ionization (positive ion mode) in the multiple reaction monitoring mode monitoring
for mass-to-charge (m/z) transition 309.1 → 164.2 or 291.1 and 275.1 →189.1, respectively.
Compound 1 and M1 standards were fit by least-squares regression of their areas to a weighted
linear equation, from which the unknown concentrations were calculated. The dynamic range of
the assay was 1.0-2000 ng/mL. Assay performance was monitored by the inclusion of quality
control samples with acceptance criteria of ± 30% target values.
Bioanalytical Methodology for Metabolite Identification. Qualitative assessment of the
metabolism of 1 was conducted using a Thermo Finnegan Surveyor photodiode array plus
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detector, Thermo Acela pump and a Thermo Acela Autosampler (Thermo Scientific, West Palm
Beach, FL). The monitoring wavelength (λ) was 280 nm. Chromatography was performed on a
Phenomenex Hydro RP C18 (4.6 mm x 150 mm, 3.5 μm) column. The mobile phase composed
of 5 mM ammonium formate buffer with 0.1% formic acid (pH=3) (solvent A) and acetonitrile
(solvent B) at a flow rate of 1 mL/min. The binary gradient was as follows: solvent A to solvent
B ratio was held at 95:5 (v/v) for 3 min and then adjusted to 55:45 (v/v) from 0 to 35 min, 30:70
(v/v) from 35 to 45 min, and 5:95 (v/v) from 45 to 52 min where it was held for 3 min and then
returned to 95:5 (v/v) for 6 min before next analytical run. Identification of the metabolites was
performed on a Thermo Orbitrap mass spectrometer operating in positive ion electrospray mode.
The spray potential was 4 V and heated capillary was at 275 °C. Xcalibur software version 2.0
was used to control the HPLC-MS system. Product ion spectra were acquired at a normalized
collision energy of 65 eV with an isolation width of 2 amu. Metabolites from liver microsomes
were identified in the full-scan mode (from m/z 100 to 850) by comparing t = 0 samples with t =
60 min samples or through comparison with synthetic standard(s), and structural information was
generated from collision-induced dissociation (CID) spectra of protonated molecular ions.
Results
Microsomal stability. To examine microsomal stability, compound 1 was incubated in rat, dog,
and human liver microsomes or human kidney microsomes in the presence and absence of
NADPH co-factor; periodically, aliquots of the incubation mixture were examined for the
depletion of 1 (Supplemental Figure 1). The t1/2 (mean ± standard deviation) for depletion of 1 in
NADPH-supplemented rat, dog, and human liver microsomes was 8.6 ± 0.4, 11.2 ± 0.4 min, and
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> 120 min, respectively. No substrate depletion (t1/2 > 120 min) was noted in rat, dog, and
human incubations that lacked NADPH co-factor or liver microsomes. Figure 2 depicts the
extracted ion chromatograms of incubation mixtures of 1 in NADPH-supplemented liver
microsomes from rat, dog, and human. The major metabolite M1 was detected in rat and dog
liver microsomes in a NADPH-dependent fashion. The MS2/MS3 spectra of compound 1
(retention time (tR) = 11.25 min, exact mass (M+H)+ = 309.0904) and M1 (tR = 9.61 min, exact
mass (M+H)+ = 275.1026 ) are shown in Supplemental Figures 2 and 3, respectively.
Theoretical exact masses for the proposed fragment ion structures in the CID spectrum of 1 and
M1 were consistent with the observed accurate masses (< 2 ppm difference). The tR and mass
spectrum of M1 was identical to the one discerned with an authentic standard, which was
chemically synthesized via S-methylation of 1 to the corresponding thioether derivative 2
followed by peroxide-mediated oxidative desulfuration/intramolecular cyclization presumably
via an electrophilic sulfoxide intermediate (see supplemental section for detailed synthetic
protocol). Metabolites M2 (tR = 9.19 min) and M3 (tR = 8.75 min) were rat-specific metabolites
with an identical exact mass (295.0747 (M+H)+) and CID spectra (see Supplemental Figure 4 for
a representative CID spectra of M3), implying that these metabolites were isomeric phenols
derived from cytochrome P450-mediated O-demethylations in 1. Consistent with the metabolic
stability results, M1–M3 were only detected in trace quantities in human liver microsomal
incubations of 1 in the presence of NADPH. Compound 1 (1 μM) appeared to be stable (t1/2 >
120 min) towards metabolic turnover in NADPH-supplemented human kidney microsomes, with
minimal amount of M1 (80 nM) formed during the course of the 60 min incubation.
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Compound 1 was devoid of reactive metabolite formation in NADPH-supplemented rat, dog,
and human liver microsomes as inferred from the lack of GSH conjugates formed when GSH (5
mM) was included in the microsomal incubations (data not shown). Incidentally, inclusion of
excess GSH significantly attenuated oxidative desulfuration of 1 (to M1) in liver microsomal
incubations from rat and dog (Supplemental Figure 5). In the presence of GSH, 1 was virtually
resistant to metabolic turnover in NADPH-supplemented dog liver microsomes (t1/2 (- GSH) =
11.2 ± 0.4 min; t1/2 (+ GSH) > 120 min). In contrast, the impact of GSH on the overall metabolic
stability of 1 in NADPH-supplemented rat liver microsomes was less severe (t1/2 (- GSH) = 8.6 ±
0.4; t1/2 (+ GSH) 49.7 ± 1.7 min). Qualitative examination of metabolite formation in NADPH-
and GSH-supplemented rat liver microsomal incubations of 1 (10 μM) revealed that the
formation of the O-demethylated metabolites M2 and M3 was not impacted in the presence of
the thiol nucleophile, which was in contrast to the complete disappearance of M1 (Figure 3).
Identification of enzymes responsible for oxidative desulfuration of 1 to M1. Incubations
of 1 (1 μM) in NADPH-supplemented rat or dog liver microsomes, which had been subjected to
heat treatment (50 °C) for 5 min in the absence of NADPH co-factor induced metabolic
resistance in 1 virtually abrogated the formation of M1 as shown in a representative plot of an
incubation mixture of 1 in heat-inactivated rat liver microsomes (Figure 4), implying a potential
role for a FMO isoform(s) in oxidative desulfuration.
Incubations of 1 (1 μM) in 0.5 mg/mL human recombinant FMO1, FMO3, and FMO5 in the
presence of NADPH revealed that FMO1 was principally responsible for the formation of M1
with a minor contribution from FMO3 (Figure 5). Contribution of FMO5 towards M1 formation
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was insignificant in this analysis; as such enzyme kinetics experiments were not pursued. The
reactions with FMO1 and FMO3 were linear as a function of incubation time (5–60 min) and
protein concentration up to 1.0 mg/mL of microsomal protein (results not shown). The effects of
substrate concentrations on oxidative desulfuration of 1 by recombinant human FMO1 and
FMO3 were investigated, and the results are shown in Figure 6. Oxidation of 1 to M1 by FMO1
followed Michaelis– Menten kinetics (Figure 6, panel A), with Km, Vmax, and CLint ± standard
error values of 209 ± 12 μM, 20.4 ± 0.6 nmoL/min/mg protein, and 97.7 μL/min/mg protein,
respectively. In the case of FMO3, the Michaelis–Menten plot (see Figure 6, panel B) showed
that conversion of 1 to M1 was linear up to the highest substrate concentration of 300 μM,
indicating that the apparent KM value was > 300 μM. Therefore, KM and Vmax values for FMO3
could not be determined. The corresponding CLint value calculated from the slope of the
Michaelis–Menten plot was 0.54 μL/min/mg protein.
Consistent with these observations, conversion of 1 to M1 in NADPH-supplemented rat and
dog liver microsomes was strongly inhibited upon co-incubation with imipramine (250 μM), a
selective inhibitor of FMO1 (Dixit and Roche, 1984; Lee et al., 2009; Yamazaki et al., 2014), as
reflected from changes in t1/2 from 8.6 to ~ 90 min (rats) and 11 to 81 min (dogs). In contrast, the
effects of the non-selective P450 inactivator, 1-aminobenzotriazole (1 mM) (Boily et al., 2015;
Caldwell et al., 2005; Parrish et al., 2015; Strelevitz et al., 2006), on oxidative desulfuration was
less severe (rat: t1/2 from 8.6 to 25 min, dog: t1/2 from 11.0 to 12 min (Figure 7).
Intravenous pharmacokinetics of 1 after single doses to rats and dogs. The pharmacokinetic
parameters describing the disposition of 1 and M1 after administration of 1 to Wistar-Han rats
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and Beagle dogs are shown in Table 1 and Supplemental Figure 6. Compound 1 demonstrated
moderate to high CLp (rat CLp = 73 ± 13 mL/min/kg; dog CLp = 12 ± 3 mL/min/kg), and a
moderate Vdss (rat Vdss = 1.4 ± 0.5 L/kg; dog Vdss = 1.3 ± 0.3 L/kg) resulting in terminal
elimination t1/2 values of 0.4 ± 0.2 and 5.3 ± 0.8 h, respectively, in rats and dogs. M1 was also
detected in circulation after i.v. administration of 1 to rats and dogs. The corresponding AUC0-∞
values of 1 and M1 in rats were 233 ± 37 and 49.8 ± 11.2 ng.h/mL, respectively, whereas, the
corresponding AUC0-∞ values of 1 and M1 in dogs were 1450 ± 310 and 769 ± 136 ng.h/mL,
respectively. M1 had a slightly longer elimination t1/2 (relative to 1) in rats and dogs. Renal
excretion of unchanged 1 (< 1% in rats and ~ 1.1% in dogs) and unchanged M1 (~ 4.7% in rats
and ~ 14.2% in dogs) was relatively low.
Discussion
Concerns over the liberation of indiscriminate electrophilic species during MPO-mediated
oxidation of N1-substituted-6-aryl-2-thiouracils were minimized by ensuring a high partition
ratio for MPO inactivation and by tethering nucleophilic functional groups in proximity of the
thiouracil sulfur. Our medicinal chemistry strategy was also weighted towards the design of
compounds in the lower range of lipophilicity (logD < 1.5, topological polar surface area < 100
Å2) to minimize the potential for oxidative metabolism/bioactivation of the thiouracil motif in
human liver. Apart from peroxidases, enzymatic bioactivation of thioureas and related analogs
(e.g., thiones, thiocarbamides, etc.) to electrophilic intermediates by mammalian P450 and/or
FMO isoforms can also lead to toxicity (Decker and Doerge, 1992; Henderson et al., 2004; Ji et
al., 2007; Neal and Halpert, 1982; Onderwater et al., 1999; 2004; Poulsen et al., 1979; Smith and
Crespi, 2002). For instance, cases of clinical hepato- and/or nephrotoxicity noted with the
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antithyroid drug methimazole (Martinez-Lopez et al., 1962) and the antiparasitic agent
thiabendazole (Manivel et al., 1987) have been causally linked with their metabolism to the
proximal toxicants N-methylthiourea and thioformamide, respectively, via an initial P450-
catalyzed oxidative ring scission of the 2-mercaptobenzimidazole and thiazole motifs present in
these drugs. S-Oxidation of the N-methylthiourea and thioformamide metabolites to reactive
metabolites by FMO enzymes is believed to represent the key step resulting in toxicity (Mizutani
et al., 1993; 2000).
Consistent with our design philosophy, the N1-substituted-6-aryl-2-thiouracil class of MPO
inhibitors (represented in the present study by compound 1) were stable towards metabolism in
NADPH-supplemented human liver microsomes and/or cryopreserved human hepatocytes
(Ruggeri et al., 2015), and were latent to the formation of reactive species as judged from the
absence of GSH conjugates in human recombinant MPO and human liver microsomes
supplemented with an excess of the thiol nucleophile. Compound 1 was also devoid of
reversible and time-dependent inhibitory effects against major human cytochrome P450 enzymes
(Pfizer data on file), which made it an attractive candidate for advancement in preclinical toxicity
studies. In contrast with the metabolic resistance in human hepatic tissue, compound 1 was
converted to cyclic ether M1 in NADPH-supplemented rat and dog liver microsomes. Heat
inactivation which abolishes FMO activity while preserving P450 activity (Ziegler, 1980)
provided circumstantial evidence for the involvement of an FMO isoform(s) in the formation of
M1. Consistent with this initial finding, the non-selective P450 inactivator 1-aminobenzotriazole
had little effect on the conversion of 1 to M1 in rat and dog liver microsomes. In contrast, the
FMO1 competitive inhibitor imipramine dramatically reduced the oxidative desulfuration in rat
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and dog liver microsomes, respectively, implying that the conversion of 1 to M1 in rat and dog
liver microsomes is facilitated primarily by FMO1 rather than P450s. The possibility of
imipramine’s inhibitory effects occurring through inhibition of rat CYP isoforms (Murray and
Field, 1992; Masubuchi et al., 1995) can be ruled out on the basis of the results obtained with 1-
aminobenzotriazole.
Mammalian FMOs (E.C.1.14.12.8) comprise a group of flavin adenine dinucleotide-
containing enzymes that utilize NADPH and molecular oxygen to generate a 4α-
hydroperoxyflavin intermediate, which mediates the two-electron oxidation of soft, highly
polarizable nucleophilic heteroatom (nitrogen, sulfur, and phosphorus)-containing xenobiotics
(Cashman, 1995; Cashman et al., 1995, Hines et al., 1994; Krueger and Williams, 2005; Phillips
and Shephard, 2008; Ziegler, 2002). Our findings on the facile decomposition of 1 to M1 in the
presence of H2O2 and FMO1 are consistent with the notion that FMO1 will generally oxygenate
any nucleophilic heteroatom-containing compound that can be oxidized by H2O2 and/or peracids
(Bruice et al., 1983). To date, five distinct forms of FMO (i.e., FMO1–5) have been identified
(Hernandez et al., 2004; Lawton et al., 1994). Examinations of adult human liver mRNA
indicate high FMO3 (and FMO5) expression but low FMO1 expression (Dolfin et al., 1996;
Hines, 2006; Koukouritaki et al., 2002; Koukouritaki and Hines, 2005; Shimizu et al., 2011;
Zhang and Cashman, 2006; Chen et al., 2016). In contrast, rat livers have been shown to express
high levels of FMO1 protein (Cherrington et al., 1998; Itoh et al., 1993; Lattard et al., 2002a;
Yamazaki et al., 2014). Expression of FMO1 and FMO3 in dog liver has also been reported with
84–89% amino acid sequence identity to the corresponding orthologs from rat and human
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(Lattard et al., 2002b; Ripp et al., 1999; Stevens et al., 2003). As such, dog FMO1 and dog
FMO3 exhibit only 56% identities in primary amino acid sequence (Lattard et al., 2002b).
To identify the specific FMO isoform responsible for the oxidative desulfuration of 1, studies
were conducted using recombinant human FMO isoforms. FMO3 and FMO5 were used since
they represent isoforms most abundant in human liver and FMO1 because it is the ortholog of the
form most abundant in adult rat and dog liver. The formation of M1 was principally mediated by
recombinant FMO1 with little to no contribution from FMO3 or FMO5 (apparent CLint for M1
formation by FMO1 was 154-fold higher than that by FMO3), which is consistent with the
inhibitory effects of imipramine on oxidative desulfuration of 1 in rat and dog liver microsomes.
Overall, these results suggest that liver microsomal FMO1 could contribute to the relatively high
FMO-mediated oxidative desulfuration of 1 in rat and dog liver microsomes and that lower
expression of FMO1 in human livers is a major determinant of oxidation potential in livers from
preclinical species and humans. The in vitro metabolic instability of 1 also manifested in
moderate to high CLp in rats and dogs with significant circulating M1 concentrations measured
in both species, which provided an in vivo context for the in vitro findings, especially when
considering that renal excretion of 1 in unchanged form was negligible in rats and dogs.
Additional case studies on species differences in FMO-mediated metabolism have also appeared
in the literature, which strengthen our observations on the selective nature of FMO1-mediated
oxidative desulfuration in 1. For instance, a recent report from Liu et al., (2013) demonstrated
that quinuclidine ring N-oxidation in a selective α7 neuronal acetylcholine receptor agonist
ABT-107 occurred primarily in liver microsomes from rat and dog (but not in human), and was
also principally mediated by FMO1. Because mRNA expression levels for FMO1 are higher in
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human kidney (relative to liver) (Dolphin et al., 1996; Zhang and Cashman, 2006; Chen et al.,
2016), the oxidative desulfuration of 1 was also examined in NADPH-supplemented human
kidney microsomes. Compared with recombinant FMO1, 1 was relatively stable in kidney
microsomes with minimal amount of M1 (~ 80 nM) formed in a NADPH-dependent fashion.
Because of the unknown amount of FMO in the commercial preparation of the human kidney
microsomes, no rate comparisons can be made between recombinant and microsomal
preparations at the present time.
Mammalian FMOs typically display high activity toward S-oxidation in thioureas, thiones,
and thiocarbamides. The initial oxygenation of the sulfur atom produces the electrophilic
sulfenic acid (R-SOH) species that is capable of reacting with nucleophiles, including GSH
(Decker and Doerge, 1992; Henderson et al., 2004; Kim and Ziegler, 2000; Krieter et al., 1984;
Neal and Halpert, 1982; Onderwater et al., 1999; Poulsen et al., 1979; Smith and Crespi, 2002).
The sulfenic acid derivatives can undergo redox cycling in the presence of GSH coupled with the
oxidation of GSH to GSSG. The sulfenic acid metabolite can also undergo a second oxidation
by FMO to the unstable sulfinic acid (R-SO2H), which is more reactive than the sulfenic acid
metabolite and can damage the cell directly or alkylate proteins (Ji et al., 2007; Onderwater et al.,
1999; 2004). Our investigations on the oxidative desulfuration of 1 to M1 (Figure 8) largely
parallel the mechanistic insights noted in the literature. For example, addition of excess GSH
essentially eliminated the conversion of 1 to M1 in NADPH-supplemented rat and dog liver
microsomes suggesting that the initial FMO1-mediated S-oxygenation of 1 leads to the
corresponding sulfenic acid derivative 3 that undergoes redox cycling to the parent compound 1
in a GSH-dependent fashion (presumably via oxidation of GSH to GSSG). A second oxidation
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of the sulfenic acid derivative 3 yields the more electrophilic sulfinic acid species 4 that is
trapped intramolecularly (perhaps in a diffusion-controlled fashion) by the pendant alcohol motif
on the N1-substitutent in 1.
In conclusion, our studies underscore one of the limitations of rat and dog as surrogates of
adult human FMO-dependent drug metabolism studies and the conclusions from previous animal
studies that lack significant amounts of liver FMO3 (i.e., rats and dogs) may need to be
reconsidered. From a drug discovery perspective, our findings provide a cautionary note against
the use of allometric scaling of clearance from animals to human without a thorough knowledge
of the overall disposition/metabolic elimination mechanism of the molecule(s) under
consideration.
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Authorship Contributions
Participated in research design: Heather, Eng, Raman Sharma, Angela Wolford, and
Amit S. Kalgutkar
Conducted in vitro experiments: Heather Eng, Raman Sharma, and Edward L. Conn
Contributed new reagents or analytic tools: Edward L. Conn, and Roger B. Ruggeri
Performed data analysis: Heather Eng, Raman Sharma, Angela Wolford, Deepak Dalvie,
and Amit S. Kalgutkar
Wrote or contributed to the writing of the manuscript: Heather Eng, Deepak K. Dalvie,
Leonard Buckbinder, Roger B. Ruggeri, Li Di, and Amit S. Kalgutkar
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Declaration of interest
H.E., R.S., A.W., L.D., R.B.R., L.B. E.L.C., D.D. and A.S.K. are employees of, and/or hold
stock in, Pfizer Inc.
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Figure Legends
Figure 1. Oxidative desulfuration of the N1-substituted-6-arylthiouracil 1 by MPO or H2O2.
Figure 2. HPLC-UV (λ = 280 nm) chromatogram of an incubation mixture of 1 (10 μM) in
NADPH-supplemented rat (panel A), dog (panel B), and human (panel C) liver microsomes.
Figure 3. HPLC-UV chromatogram of an incubation mixture of 1 (10 μM) in NADPH-
supplemented rat liver microsomes in the absence or presence of GSH.
Figure 4. Oxidative desulfuration of 1 (1 μM, ●) to M1 (□) in NADPH-supplemented rat liver
microsomes in the absence (panel A) or presence of heat inactivation (5 min, 50 °C) (panel B).
Symbols depict means and error bars for standard deviations.
Figure 5. Oxidative desulfuration of 1 (1 μM) to M1 in human recombinant FMO1, FMO3, and
FMO5. Incubations were conducted using 0.5 mg/mL Supersomes in the presence of NADPH
(1.3 mM) for 5 min at 37 °C. Symbols depict means and error bars for standard deviations.
Figure 6. Kinetics of oxidative desulfuration of 1 to M1 by human recombinant FMO1 (panel A,
0.1 mg/mL microsomal protein) and FMO3 (panel B, 1 mg/mL). Incubations were conducted in
the presence of NADPH (1.3 mM) at 37 °C for 60 min in triplicate. Symbols depict means and
error bars for standard deviations.
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Figure 7. Oxidative desulfuration of 1 (1 μM) to M1 in NADPH-supplemented rat (panel A) and
dog (panel B) liver microsomes in the absence or presence of FMO1 and CYP inhibitors,
imipramine (250 μM) and 1-aminobenzotriazole (1000 μM), respectively. Symbols depict means
and error bars for standard deviations.
Figure 8. Proposed mechanism of oxidative desulfuration of 1 to M1 in NADPH-supplemented
rat and dog liver microsomes.
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Tables
TABLE 1
Mean pharmacokinetic parameters of compound 1 and cyclic ether metabolite M1 in after i.v. (1
mg/kg) administration of 1 to Wistar-Han rats and Beagle dogs.
ai.v. administration of 1 in 5% DMSO/95% of 30% 2-hydroxypropyl-β-cyclodextrin. bconcentrations of M1were estimated in animals administered with 1 via the i.v. route. ci.v. administration of 1 as a solution in 25% 2-hydroxypropyl-β-cyclodextrin/ Tris buffer (100 mM) (pH = 8.0). N.A. not applicable. Pharmacokinetic parameters are expressed as mean ± standard deviation.
Compound Species Dose (mg/kg)
CLp (mL/min/kg)
Vdss (L/kg)
t1/2 (h)
AUC(0-∞) (ng.h/mL)
1 Ratsa 1.0 (n=3)
73 ± 13 1.4 ± 0.5 0.4 ± 0.2 233 ± 37
M1b N.A. N.A. 0.8 ± 0.3 49.8 ± 11.2
1 Dogsc 1.0 (n=3)
12 ± 3.0 1.3 ± 0.3 5.3 ± 0.8 1450 ± 310
M1b N.A. N.A. 6.3 ± 0.8 769 ± 136
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Figure 2
6 7 8 9 10 11 12 13 140
20
60
100 9.61
9.19 11.25
8.75
6 7 8 9 10 11 12 13 140
20
60
100 9.61
11.25
6 7 8 9 10 11 12 13 14Time (min)
020
60
100 11.26Rel
ativ
e Ab
sorb
ance
1
M1
M2
M3
1
1
M1
M1M3 M2
A
B
C
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6 7 8 9 10 11 12 13 14Time (min)
020
60
100
Rel
ativ
e Ab
sorb
ance
11.25
9.19
8.75
6 7 8 9 10 11 12 13 140
20
60
100 9.61
9.19 11.25
8.75
1
1
M2
M3
M2
M3
M1
+ GSH
- GSH
Figure 3
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Con
cent
ratio
n (n
M)
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0 20 40 600
200
400
600
800
1000
Time (min)
M1 FMO1
FMO3FMO5
Figure 5
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A B
Figure 6T
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this version.D
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