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Metabolism and Disposition of a -Aminobutyric Acid Type A Receptor Partial Agonist in Humans

Departments of Pharmacokinetics, Pharmacodynamics, and Metabolism (C.L.S., M.G., A.D.V.), Clinical Pharmacology (K.V.), and Clinical Research Operations (A.H.B.), Pfizer Global Research and Development, Groton/New London Laboratories, Pfizer Inc., Groton, Connecticut

(Received November 14, 2007; accepted January 3, 2008)

	Abstract

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The metabolism and disposition of N-[3-fluoro-4-[2-(propylamino)ethoxy]phenyl]-4,5,6,7-tetrahydro-4-oxo-1H-indole-3-carboxamide (1), a potent subtype-selective partial agonist at the -aminobutyric acid type A receptor complex, were elucidated in humans following a p.o. dose of N-[3-fluoro-4-[2-(propylamino)ethoxy]phenyl]-4,5,6,7-tetrahydro-4-oxo-1H-[3-¹⁴C]indole-3-carboxamide monomethane-sulfonate ([¹⁴C]1). Overall, 1 was well tolerated, with approximately twice as much radioactivity excreted in feces (64.8 ± 13.3%) as in urine (28.4 ± 8.8%). Across subjects, the oral clearance of 1 was composed of both renal (10%) and metabolic (90%) components, with the biotransformation of 1 happening predominately via oxidative deamination to either 2-fluoro-4-[(4-oxo-4,5,6,7-tetrahydro-1H-indole-3-carbonyl)-amino]-phenoxy acetic acid (2) or 4-oxo-4,5,6,7-tetrahydro-1H-indole-3-carboxylic acid [3-fluoro-4-(2-hydroxy-ethoxy)-phenyl]-amide (3) and minimally by aliphatic hydroxylation and carbamate formation. Active renal secretion of 1 was observed as its unbound renal clearance was 6-fold greater than the glomerular filtration rate. Experiments using human hepatic in vitro systems were undertaken to better understand the enzyme(s) involved in the clinically observed oxidative biotransformation pathways. N-Dealkylation of 1, the principal metabolic route observed in vivo, was found to be predominately monoamine oxidase-B-mediated with the resulting putative aldehyde intermediate undergoing subsequent oxidation to 2 or reduction to 3.

Benzodiazepines exert their efficacy in a full-agonist fashion by allosterically modulating the -aminobutyric acid type A (GABA_A) receptor complex (Olsen and Tobin, 1990; Drexler et al., 2006), which potentiates GABA-mediated neuronal inhibition (Sieghart, 1992; Mohler et al., 2002). Unlike full agonists, partial agonists of the GABA_A receptor, particularly those that are subtype-selective (Low et al., 2000; Mohler et al., 2002; Griebel et al., 2003), may afford concurrently the desired anxiolytic effects of a benzodiazepine while minimizing or eliminating its unwanted side effects, resulting in a superior safety profile for the general and chronic treatment of GAD (Lader, 1994).

N-[3-Fluoro-4-[2-(propylamino)ethoxy]phenyl]-4,5,6,7-tetrahydro-4-oxo-1H-indole-3-carboxamide (1), a potent subtype-selective partial agonist at the GABA_A receptor complex, elicits anxiolytic effects in rats without the side effects of benzodiazepines (Pfizer Inc., unpublished internal data). The study reported herein was undertaken to ascertain definitively the metabolic and excretory pathways of 1 in humans following a single p.o. dose of N-[3-fluoro-4-[2-(propylamino)ethoxy]phenyl]-4,5,6,7-tetrahydro-4-oxo-1H-[3-¹⁴C]indole-3-carboxamide monomethanesulfonate ([¹⁴C]1) (Fig. 1). Determination of the clearance routes, metabolites, and pharmacokinetics of 1 in the clinic has provided deeper insight into its overall human disposition, and confirmed rats and monkeys as appropriate toxicological species (Shaffer et al., 2005).

View larger version (7K): [in this window] [in a new window]   FIG. 1. Chemical structure of protonated [14C]1 (m/z 374, *, 14C) and its CID fragmentation pathway.

	Materials and Methods

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Chemicals and Reagents. Compounds 1, 2, and 3 were provided by Neurogen Corp. (Branford, CT); [¹⁴C]1 (50.7 mCi/mmol, 99.0% radiochemical purity) was made by the Radiochemical Synthesis Group at PGRD, Pfizer Inc. (Groton, CT). The chemical purity of all the synthetic compounds was >99%. Human liver microsomes (HLMs) [17.6 mg of protein/ml, 0.32 nmol of cytochrome P450 (P450)/mg protein] pooled from 53 individual donors and human liver mitochondria (20 mg of protein/ml) were purchased from Xeno-Tech LLC (Lenexa, KS). Cryopreserved human hepatocytes (from three donors) were procured from In Vitro Technologies Inc. (Baltimore, MD). Recombinant human monoamine oxidase (MAO)-A (20 mg of protein/ml) and MAO-B (20 mg of protein/ml) membranes were provided by Professor Dale E. Edmondson under contract with Pfizer Inc. Chemicals and solvents of reagent or high-performance liquid chromatography (HPLC) grade were supplied by Aldrich Fine Chemical Co. (Milwaukee, WI), Fisher Scientific (Pittsburgh, PA), and the J. T. Baker Chemical Co. (Phillipsburg, NJ). TruCount liquid scintillation mixture was purchased from IN/US Systems, Inc. (Tampa, FL). All the excreta and plasma were collected gravimetrically and stored at –20°C until analysis.

Dosing of Human Volunteers and Collection of Samples. The study was a nonrandomized, open-label, single-dose study to investigate the absorption, metabolism, and excretion of 1 in humans. The study population was composed of six healthy, nonsmoking adult white male volunteers between the ages of 18 and 64 years and weighing 61 to 91 kg. The study was conducted in compliance with an Institutional Review Board/Independent Ethics Committee, informed consent regulations, and International Conference on Harmonization/Good Clinical Practice Guidelines. The Institutional Review Board approved both the study protocol and informed consent documents before drug shipment. After being informed of the design, purpose, and potential risks of the study, written informed consent was required from each subject before his enrollment at the Pharmaceutical Product Development Phase I Clinic (Austin, TX), where the subjects were kept under continuous medical surveillance from 24 h predose to 264 h postdose. Subjects were required to fast 8 h before and 4 h after dosing. Each subject was administered 200 mg (approximately 91 µCi) of [¹⁴C]1 dissolved in sterile H₂O (100 ml), which was swallowed directly from an amber glass dosing bottle with a 30-cc-wide mouth, followed by more sterile H₂O (two 70-ml bottle rinses).

From each subject, urine was collected predose and from 0 to 12 and 12 to 24 h during day 1, and in 24-h intervals from 24 to 240 h postdose, and feces were collected predose and as passed at 24- or 48-h intervals for 11 days postdose. Blood samples sufficient to provide 5 ml of plasma were collected into sodium heparinized tubes via arm venipuncture predose and at 0.5, 1, 2, 4, 8, 12, 24, 48, 72, 96, 120, 144, 168, 192, 240, 264, 288, and 312 h postdose for the pharmacokinetic evaluation of 1 and total radioactivity; having been released from the Clinic on day 11 (264 h postdose), subjects returned to the Clinic for the 288 and 312 h postdose blood draws. Blood samples sufficient to provide 20 ml of plasma were collected similarly at 1, 4, 8, 12, and 24 h postdose for the profiling of metabolites of 1. Control plasma was harvested from predose blood samples.

Determination of Radioactivity within Biological Matrices. Triplicate gravimetric aliquots (0.1 g) of each sample of urine and plasma were mixed with TruCount scintillation mixture (7 ml) and counted for 4 min in a model LS 6500 liquid scintillation counter (Beckman Coulter, Fullerton, CA). Fecal samples were homogenized with distilled H₂O (approximately 20% w/w, feces/H₂O) using a Stomacher homogenizer (Seward, Worthing, West Sussex, UK). Triplicate gravimetric aliquots (0.5–0.7 g) of fecal homogenate were transferred into tared cones and pads, weighed, dried for a minimum of 24 h at ambient temperature and combusted by a Packard Instruments Model A0387 sample oxidizer (Packard BioScience, Co., Meriden, CT). Combustion efficiency using a ¹⁴C standard was determined daily before the combustion of study samples, and the measured radioactivity content in feces was adjusted using daily combustion efficiency values. The resulting [¹⁴C]CO₂ was trapped in Monophase-S (PerkinElmer Life and Analytical Sciences, Boston, MA), mixed in Perma Fluor E scintillation fluid (Packard BioScience, Co.), and quantified over 10 min in a Packard Instruments model 2300 liquid scintillation counter (Packard BioScience, Co.). Scintillation counter data were automatically corrected for counting efficiency using an external standardization technique and an instrument-stored quench curve generated from a series of sealed quench standards.

Quantitative Analysis of 1 in Plasma. Plasma concentrations of 1 for each subject were determined using a validated liquid chromatography/tandem mass spectrometry (LC/MS/MS) assay (Venkatakrishnan et al., 2007) at MDS Pharma Services (Lincoln, NE). The dynamic range of the assay was 1 to 1000 ng/ml for 1.

Calculations. Pharmacokinetic parameters were calculated for each subject by noncompartmental analyses using WinNonlin version 3.2 (Pharsight Corp., Mountain View, CA). Values used to determine the pharmacokinetic parameters of total radioactivity were calculated by converting the liquid scintillation counting (LSC)-generated raw data to concentrations (ng-Eq/ml) using the specific activity (0.17 mCi/mmol) of administered [¹⁴C]1. The area under the plasma concentration-time curve (AUC)_0-tlast was calculated using the linear trapezoidal method; elimination rate constant (k_el) was determined by linear regression of the log concentration versus time data during the terminal phase; half-life (t_1/2) was calculated as (ln2)/k_el, and AUC_0- was calculated as the sum of AUC_0-tlast and AUC_tlast-, which was determined by dividing the plasma concentration at t_last by k_el. Both maximal plasma concentration (C_max) and its time of first occurrence (T_max) were taken directly from the concentration versus time data. Means and S.D. were calculated when half or greater of the values exceeded the lower limit of quantification (LLOQ) for 1 (1.0 ng/ml) or total radioactivity (72.7 ng-Eq/ml). A value of 0 was used when a measured concentration was below its LLOQ. For each subject, oral plasma clearance (CL_p/F) of 1 was calculated by dividing the dose by respective plasma AUC_0-, and renal clearance (CL_R) of 1 was calculated by dividing the amount of 1 excreted unchanged in urine over 48 h (Ae_0–48), as determined by the radioprofiling of pooled individual urine (see below), by its plasma AUC_0–48; both clearance values were then subject weight-normalized to afford units of milliliter per minute per kilogram. The use of urinary excretion data collected over 48 h postdose for CL_R calculations is justified as being approximately 5 times the estimated t_1/2 of 1 in this study.

Sample Preparation for Metabolite Profiling and Identification. At each step during the sample preparation of all the biological matrices, total radioactivity levels were determined by LSC for recovery calculations. Following preparation, all the samples were analyzed as described below by LC/MS/MS with radiometric detection. Predose and blank samples served as controls for determining background radioactivity and endogenous, non–drug-related ions observed within respective matrices or their extracts by LC/MS/MS.

Urine. Urine samples from each subject collected from 0 to 48 h postdose representing >95% of total urine radioactivity were pooled proportional to the amount of urine in each sampling period for analysis by LC/MS/MS with radiometric detection. Pooled urine samples (13–30 ml) were concentrated by an N₂ stream at 37°C, reconstituted in 1% isopropanol in 2 mM ammonium acetate, pH 4.3 (0.5 ml, solvent A), and centrifuged [832 relative centrifugal force (rcf) for 10 min] to afford the analytical sample, which retained 90% of the radioactivity contained within the pooled sample before concentration.

Feces. Fecal homogenates from each subject were pooled based on sample time intervals representing >90% of total fecal radioactivity and were pooled for analysis as described above for urine. Pooled fecal homogenates (7–10 g) were diluted with H₂O (1 ml/g homogenate) and agitated at 37°C overnight in a shaking water bath. The softened stool samples were diluted with MeCN (2 ml/g homogenate), vortex-mixed, and centrifuged (832 rcf for 10 min), and the resulting supernatant was transferred to a clean vial. The remaining fecal pellets were extracted an additional one or two times with 33% H₂O in MeCN (9 ml) to ensure that >90% of radioactivity, as determined by LSC analysis of the combined supernatants, from each pooled fecal sample was extracted. The combined supernatants were concentrated by an N₂ stream at 37°C and reconstituted in solvent A (2 ml) to yield the analytical sample, which retained 90% of the radioactivity contained within the pooled sample before concentration.

Plasma. Plasma from blood samples drawn from each individual at 1, 4, 8, 12, and 24 h postdose was used for circulatory metabolite profiling and identification; these time points accounted for 100% of the total radioactivity AUC because for all the subjects the last quantifiable time point for radioactivity was 24 h postdose. Plasma samples were pooled according to the method of Hamilton et al. (1981), i.e., 2, 3.5, 4, 8, and 6 ml, respectively, of plasma from each time point were combined. To remove dissolved proteins, pooled plasma samples (23.5 ml) were diluted with MeCN (47 ml), vortex-mixed, and centrifuged (3661 rcf for 10 min), and the resulting supernatants, which contained >90% of the radioactivity from each pooled sample, were isolated. Each supernatant was concentrated to near dryness at 35°C under N₂ and reconstituted in solvent A (250 µl) to provide the analytical sample.

Metabolite Profiling and Identification. Samples were analyzed by an LC/MS/MS system comprising a PE Sciex API-3000 tandem quadrupole mass spectrometer with a turbo ion spray interface (PerkinElmer Life and Analytical Sciences), two Shimadzu LC-10A HPLC pumps (Shimadzu USA, Columbia, MD), and a CTC PAL autosampler (LEAP Technologies, Carrboro, NC) in series with a β-RAM radiometric detector (IN/US Systems, Inc.) containing a liquid scintillant cell (250 µl). Analytes within sample aliquots (25–100 µl) were eluted on a Column Engineering Monitor C₁₈ analytical column (5 µm, 4.6 x 150 mm) at 1 ml/min with solvent A and MeCN (solvent B). The following two-step gradient was used: 0 to 45 min, 5 to 60% solvent B in solvent A; 45 to 50 min, 60 to 80% B in A. Following the elution of 1 and its metabolites, the column was washed with 80% B in A for 2 min and then returned over 3 min to 5% B in A, where it remained for 10 min before the next injection. For each matrix, >94% of the radioactivity injected onto the column eluted during the first 40 min of the gradient program. HPLC effluent was split 1:9 between the mass spectrometer and the radiometric flow detector; liquid scintillation mixture flowed at 3 ml/min to the radiometric detector. Mass spectral data were collected using positive ionization in full, precursor ion, neutral loss, product ion, and multiple-reaction monitoring (MRM) scanning modes. Instrument settings and potentials were adjusted to provide optimal data in each mode. Masschrom version 1.1.1 (PerkinElmer Life and Analytical Sciences) and Winflow version 1.4 (IN/US Systems, Inc.) software were used for the acquisition and processing of mass spectral and radiochromatographic data, respectively.

Because the radioactivity in all the reconstituted plasma samples was too low for quantification by radiometric flow detection, HPLC effluent was isolated in 30-s intervals by a Gilson FC 204 fraction collector (Gilson, Inc., Middleton, WI), and each respective fraction was mixed with scintillation fluid (7 ml) and subjected to LSC for 5 min. Individual plasma radiochromatograms were generated from respective liquid scintillation data using Microsoft Excel (Microsoft Office 2000 9.0.4402 SR-1; Microsoft, Redmond, WA) and paired with their respective MRM chromatograms.

In Vitro Incubations with [¹⁴C]1. All the samples from incubations using [¹⁴C]1 were analyzed by LC/MS/MS with radiometric detection system as described previously; inline radiodetection quantified 1 and its metabolites, whose identification was confirmed by previously observed liquid chromatography retention times (LC t_Rs) and collision-induced dissociation (CID) spectra.

HLMs. To determine NADPH-dependent metabolites, incubations (2.5 ml) were first performed in duplicate with NADPH (3.3 µmol) in 10-ml Erlenmeyer flasks open to air at 37°C in a shaking water bath. Each incubation contained HLMs (1.5 mg of protein/ml 0.1 M KH₂PO₄ buffer, pH 7.4), MgCl₂ (7.5 µmol), and [¹⁴C]1 (50 nmol, 5.1 nCi/nmol). Sample aliquots (1 ml) were removed by micropipette at 0 and 60 min after NADPH addition, quenched with MeCN (5 ml), centrifuged (1509 rcf for 5 min), and the resulting supernatant concentrated and reconstituted in solvent A (0.5 ml) for LC/MS/MS analysis. To determine non–NADPH-dependent metabolites of 1, incubations were performed as described above but lacked NADPH, and sample aliquots were taken at 0 and 4 h after addition of [¹⁴C]1. For HLM incubations with metabolites 2-fluoro-4-[(4-oxo-4,5,6,7-tetrahydro-1H-indole-3-carbonyl)-amino]-phenoxy acetic acid (2) and 4-oxo-4,5,6,7-tetrahydro-1H-indole-3-carboxylic acid [3-fluoro-4-(2-hydroxy-ethoxy)-phenyl]-amide (3), 1-h incubations (±NADPH) were conducted in duplicate.

Human Hepatocytes. Incubations (2.5 ml), which were gassed every hour with 5% CO₂ in O₂, were performed in duplicate in capped 10-ml Erlenmeyer flasks at 37°C in a shaking water bath. Each incubation contained thawed cryopreserved human hepatocytes (2 x 10⁶ viable cells/ml bicarbonate-based Williams' E medium with 10% fetal bovine serum) and [¹⁴C]1 (50 nmol, 5.1 nCi/nmol). A sample aliquot (0.5 ml) was removed by micropipette just after addition of 1 and quenched with MeCN (1 ml). After 4 h of reaction, the incubation was terminated with MeCN (4 ml). Quenched samples were vortex-mixed and centrifuged (1509 rcf for 5 min), and the resulting supernatant was evaporated under N₂ at 35°C, and the residue was reconstituted in solvent A (0.1–0.5 ml) for LC/MS/MS analysis.

Human Liver Mitochondria. Incubations (2 ml) were performed in duplicate using the apparatus employed for HLMs. Each incubation contained human liver mitochondria (1 mg of protein/ml 0.1 M KH₂PO₄ buffer, pH 7.4) and [¹⁴C]1 (40 nmol, 5.1 nCi/nmol). Incubations were terminated by the addition of MeCN (4 ml) 3 h after substrate addition, and the duplicate incubations were combined to optimize analytically the total amount of radioactivity within ultimate sample aliquots. Quenched incubations were processed as described earlier to afford the analytical supernatant for LC/MS/MS analysis.

In Vitro Studies Investigating the MAO-Mediated Conversion of 1 to 2 and 3. All the samples from incubations using nonradiolabeled 1 were analyzed by an LC/MS/MS system consisting of a TSQ Quantum triple quadrupole mass spectrometer with an electrospray ionization source (Thermo Fisher Scientific, Inc., Waltham, MA), a Series 1100 HPLC pump with membrane degasser, and an autosampler (Agilent Technologies, Inc., Santa Clara, CA). Analytes within sample aliquots (50 µl) were eluted on a Phenomenex Luna C₁₈ analytical column (5 µm, 2.0 x 50 mm) at 0.5 ml/min with 5 mM ammonium formate, pH 3.0 (solvent C), and solvent B using the following gradient: 0 to 2 min, 25% B in C (effluent diverted to waste); 2 to 4 min, 25 to 90% B in C; 4 to 6 min, 90% B in C. On elution of 1, 2, and 3, the column was returned over 0.5 min to 25% B in C, where it remained for 2.5 min before the next injection. Instrument settings and potentials were adjusted to provide optimal data. Mass spectral data were collected using positive ionization in MRM scanning mode monitoring m/z 374.0 162.0 (1, LC t_R = 2.6 min), m/z 347.0 162.0 (2, LC t_R = 4.6 min), and m/z 333.0 162.0 (3, LC t_R = 4.8 min); monitored ions were windowed by ±0.7 atomic mass units. Quantification of 1, 2, and 3 was accomplished using standard curves ranging from 10 to 1000 nM. LCQuan (Thermo Fisher Scientific, Inc.) software was used for the acquisition and processing of mass spectral data.

Recombinant Human MAO-A and MAO-B Membranes. To determine more precisely the MAO-A and/or MAO-B contributions to, and kinetics for, the metabolism of 1 to 2 and 3, incubations (1 ml) were performed in duplicate in 1.4-ml polypropylene microtubes open to air at 37°C in a shaking water bath. Each incubation contained either MAO-A or MAO-B membranes (10 µg of protein/ml 50 mM KH₂PO₄ buffer, pH 7.4) and 1 (20 nmol). Sample aliquots (100 µl) were removed by micropipette at 0, 10, 20, 30, 40, 50, 60, and 70 min after addition of 1, quenched with 10% formic acid in MeCN (500 µl) containing an internal standard, vortex-mixed, and then filtered through a Millipore mixed cellulose ester membrane (0.45 µm). Filtrates were evaporated to dryness under an N₂ stream at 37°C, and the resulting residues were reconstituted in 10% MeCN in H₂O (100 µl) and analyzed by LC/MS/MS as described previously.

	Results

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Clinical Observations. Six healthy, adult male volunteers were enrolled as planned. All the subjects completed the study and were evaluable for adverse events, safety laboratory tests, and pharmacokinetics. There were no serious adverse events, withdrawals resulting from adverse events, or deaths associated with this study. Following administration of 1, the only reported adverse event was diarrhea in one subject. There were no vital signs, electrocardiogram, or safety laboratory test findings of potential clinical concern. Overall, 1 was well tolerated.

Excretion of Total Radioactivity. Mean overall recovery of excreted drug-derived material (Table 1) was 93.2 ± 14.3%, with approximately half as much radioactivity in urine (28.4 ± 8.8%) as in feces (64.8 ± 13.3%). Although 94% of administered radioactivity was recovered from five of the subjects over the study period, collected excreta accounted for only 64.5% of the dose given to Subject 4, an obvious outlier within the study group. Interestingly, urinary recovery for Subject 4 (22.4%) was consistent, whereas fecal recovery (42.1%) was inconsistent with that of the other five subjects (29.6 ± 9.3% and 69.4 ± 8.2%, respectively). This discrepancy is manifested in a larger S.D. in the mean fecal and total recoveries including Subject 4; mean urinary recovery S.D. is essentially unaffected by Subject 4. Similarities in both urinary recovery and pharmacokinetics (for both 1 and total ¹⁴C) for Subject 4 relative to all the other subjects suggest this volunteer's much lower overall recovery was most likely caused by deficient fecal collection rather than partial dose ingestion. The excretion rate of total drug-related material was moderate in all the subjects; on average, >80% of the recovered radioactivity was excreted within 4 days postdose.

Pharmacokinetics of 1 and Total Radioactivity. Raw data for determining the pharmacokinetic parameters of 1 and total radioactivity were acquired using a validated LC/MS/MS assay and LSC, respectively. In all the subjects, concentrations of 1 and total radioactivity were below their individual LLOQ within plasma sampled beyond 48 and 24 h postdose, respectively, defining these time points as respective t_last values. For 1 and total radioactivity, mean plasma concentrations versus time are plotted in Fig. 2, and mean pharmacokinetic parameters are listed in Table 2.

View larger version (10K): [in this window] [in a new window]   FIG. 2. Semilogarithmic (top) and linear (bottom) plots of mean plasma concentrations of 1 () and total radioactivity () in humans.

Structural Rationalization of 1 and Its Metabolites. Compound 1 had a protonated molecular ion of m/z 374 and an LC t_R of approximately 19.8 min; its CID product ion spectrum contained fragment ions with m/z 271, 213, 193, 180, 162, and 86 (Fig. 1; Table 3). Precursor ion scanning of diagnostic fragment ions m/z 162 and 213 determined whether metabolites of 1 were modified on either its 4-oxotetrahydroindole or aniline moiety, respectively. A summary of all the metabolite LC/MS/MS data is found in Table 3.

The identification of a metabolite as a fully characterized synthetic standard (i.e., 2 or 3) was determined by the compounds' indistinguishable CID spectra and LC t_R. Metabolites M1, M2, M3, M4, M5, and M7, for which authentic standards did not exist, were each assigned a tentative structure based on their respective m/z, precursor ion scan response, and CID spectrum. Unknown metabolites U1 and U2 were not assigned even a speculative structure because neither ionized within the MS in either positive or negative ion mode, which precluded generation of any mass spectral data. Because of extremely limited remaining plasma quantities following circulatory radioprofiling and the inability to form U1 or U2 in vitro, no further analyses were conducted for the structure elucidation of either metabolite.

Quantitative Profile of [¹⁴C]1 and Its Metabolites in Excreta and Plasma. Unchanged 1 and metabolite 2 were the only compounds observed in the urine of all the subjects, whereas metabolites U1, U2, M1, M2, M4, and M5 were detected in at least half of the six volunteers (Table 4). On average, 9.9 ± 2.6% of administered 1 was renally excreted (Table 4). Unchanged 1, M1, 2, M4, and M5 were detected in the feces of all the subjects, whereas 3 and M2 were observed for five and two subjects, respectively (Table 5). Regardless of subject, 2 was the predominate metabolite in both excretory matrices. In plasma, 1, U1, M2, 2, M4, and 3 were observed in all the subjects; U2 and M7 were observed in five (Table 6). On average, 1 comprised 19.1 ± 4.2% of total circulatory radioactivity in humans, consistent with the mean pharmacokinetic-derived AUC_0–24 ratios for 1 and total radioactivity of 30.9 ± 6.0%.

In Vitro Metabolism of 1. Following identification of the human in vivo metabolites of 1, experiments using human liver-derived in vitro systems and [¹⁴C]1 were first undertaken to identify the hepatic subcellular fraction(s) responsible for the oxidative deamination of 1, the most clinically relevant metabolic clearance pathway identified in the human mass balance study with [¹⁴C]1. The use of [¹⁴C]1 in the initial leg of these studies permitted both the direct quantification of 1 and its metabolites without the need for analytical calibration curves, as well as the detection of nonionizable metabolites U1 and U2 (if formed in vitro). Although the enzymology underlying the conversion of 1 to 2 and 3 was of greatest interest based on the human in vivo metabolite profiles from a quantitative perspective, the general enzyme identification for all the metabolic pathways was undertaken.

For studies using [¹⁴C]1 (Table 7), HLMs converted approximately 39% of 1 to M1, 2, 3, and M5 in the presence of NADPH; in the absence of NADPH, HLMs transformed approximately 49% of 1 to 2 and 3 only. These data suggested that the HLM-mediated formation of M1 and M5 was NADPH-dependent, whereas that of 2 and 3 was predominately not NADPH-dependent. A combination of the oxidative deamination of 1, ultimately affording 2 and 3, being a largely NADPH-independent metabolic clearance pathway in HLMs and reports (Jarrott and Iversen, 1968; Unzeta et al., 1983; Pfizer Inc., unpublished internal data) that liver microsomes are often contaminated with MAO suggested MAO was the culprit of the N-dealkylation of 1. In human hepatocytes, 1 formed 2 and 3 only following approximately 60% consumption of 1, once again suggesting the role of MAO in the oxidative deamination of 1. Next, human liver mitochondria, the hepatic organelle containing the vast majority of MAO (Testa, 1995b), metabolized 1 to 2 and 3 only (Table 7), further validating 1 as an MAO substrate. In no tested human hepatic-derived in vitro systems using [¹⁴C]1 were U1 or U2 observed.

Based on the in vitro metabolism studies conducted with [¹⁴C]1, recombinant human MAO-A and MAO-B membranes were used to determine more precisely the MAO-A and/or MAO-B contributions to and preliminary rates of the oxidation of 1 to 2 and 3 (Table 8). In the MAO-A membrane system, 1 was converted minimally to 2 and 3 at equivalent rates (approximately 0.56 pmol of metabolite/min/mg). In the MAO-B membrane system, 1 was also metabolized to both 2 (3.1 pmol of 2/min/mg) and 3 (31.8 pmol of 3/min/mg), but at formation rates 5- to 60-fold greater, respectively, than those observed in the MAO-A system (Table 8).

In Vitro Metabolism of 2 and 3. Incubation of 2 and 3 with HLMs for purely qualitative purposes afforded M4 and M5, respectively, in an NADPH-dependent manner; no other metabolites were observed for either 2 or 3.

	Discussion

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The metabolism and disposition of 1 were quantitatively characterized in humans following a single p.o. dose of [¹⁴C]1 (200 mg, 91 µCi), which was well tolerated in all six healthy volunteers. Mean overall recovery of administered radioactivity was 93.2 ± 14.3%, with approximately twice as much in feces (64.8 ± 13.3%) as in urine (28.4 ± 8.8%). The excretion rate of total drug-related material was moderate in humans, with >80% of the recovered radioactivity excreted within 4 days postdose. A combination of short T_max for both 1 and total ¹⁴C, sizeable amounts of urinary radioactivity, and the predominance of non–microflora-generated metabolites within feces suggested 1 was rapidly and substantially (76% of dose) absorbed orally in humans. Across subjects, the oral clearance of 1 was composed chiefly of metabolic components with only minimal renal clearance (approximately 10%). The precise metabolic contribution to oral clearance is unknown because noteworthy fecal amounts (approximately 14% of dose) of 1 suggested a possible biliary component to its oral clearance. However, the size of this contribution was undeterminable because it was unknown how much of 1 (if any) detected in feces was actually absorbed following ingestion. Biliary clearance of 1 was minimal in both rats and monkeys (Shaffer et al., 2005).

Approximately 10% of administered 1 was excreted unchanged in urine, equal to the pharmacokinetically derived contribution of renal clearance (CL_R) to apparent oral clearance (CL_p/F) (Table 3). Active renal secretion of 1 was observed as its unbound CL_R was 6-fold greater than the glomerular filtration rate (GFR) (Table 3), consistent with the identification of 1 as a multidrug resistance 1 P-glycoprotein substrate (Venkatakrishnan et al., 2007) and the established contribution of this transporter to active renal secretion of its substrates (Lee and Kim, 2004). A similar extent of active renal secretion was observed in monkeys (6x GFR) but not rats (2x GFR) (Shaffer et al., 2005).

Overall, qualitatively and quantitatively similar metabolite profiles in all biological matrices were observed across subjects (Tables 4, 5, 6). A proposed schematic overview of the human metabolism of 1 is presented in Fig. 3. Based on the identification of all the urinary and circulatory metabolites, other than U1 and U2, human biotransformation of 1 occurs predominately via N-dealkylation, which has been shown in vitro to be mediated predominately by MAO-B, with only minor contributions by MAO-A or P450(s). The resulting putative aldehyde undergoes one of two facile fates: oxidation to 2 or reduction to 3. In vitro experiments suggest 2 and 3 each undergo P450-catalyzed hydroxylation to M4 and M5, respectively. The enzymes suspected in the conversion of the aldehyde intermediate to 2 and 3 are aldehyde dehydrogenase (ALDH) and alcohol dehydrogenase (ADH), respectively, both of which are found in hepatocytes and hepatic microsomes, cytosol, and mitochondria (Testa, 1995a) and are consistent with the presented observations. Interestingly, HLMs may have greater intrinsic reductive (versus oxidative) capacity of the putative aldehyde intermediate because greater quantities of 3 relative to 2 were observed in HLMs (±NADPH) than in either hepatocytes or liver mitochondria (Table 7). However, the observation of considerable quantities of 2 and 3 in HLM incubations lacking NADPH, which itself and its oxidized form (NADP⁺) could act as respective cofactors for ALDH and ADH within NADPH-containing HLM incubations, does suggest that the formation mechanisms of 2 and 3 from their common aldehyde precursor in cofactor-free HLMs are unresolved redox quandaries. Conversely, 3-fold greater quantities of 2 versus 3 observed in both hepatocytes and liver mitochondria (Table 7) were more indicative of the clinical excretory metabolic profiles, suggesting these in vitro systems may contain most appropriately the necessary cofactors for both ALDH (NAD⁺) and ADH (NADH), facilitating conversion of the intermediate aldehyde to 2 (and possibly the conversion of 3 to 2 as well).

View larger version (8K): [in this window] [in a new window]   FIG. 3. An overview of the proposed metabolic pathways of 1 in humans. Bold numerical assignments designate observed metabolites; the bracketed structure is a putative metabolite intermediate. [O], enzyme-mediated oxidation; H2, enzyme-mediated reduction.

The hypothesized enzymatic conversion of 3 to 2 via the reversible ADH pathway may play a role in the human metabolism of 1. This suspicion has been preliminarily confirmed by qualitative studies conducted in human liver cytosol in which 3 was converted to 2 in an NAD⁺-dependent manner. Specifically, although 3 comprises the same amount (approximately 13%) of total circulatory radioactivity as 2, 2 and its monooxygenated metabolite M4 account for approximately 53% of excreted drug-related material, whereas 3 and its metabolite M5 comprise only approximately 5%. These data when viewed in context of the in vitro findings suggest that 3 may be converted to 2 via the reversible ADH pathway (and subsequent ALDH oxidation) in vivo, which could result in the majority of drug-related material being excreted in the form of 2 and M4, although a contribution of potential differences in volumes of distribution of these metabolites to quantitative differences between excretory and circulatory metabolite profiles cannot be assessed. This phenomenon was also observed in monkeys but not rats (Shaffer et al., 2005).

Two very minor human metabolic pathways for 1 are its aliphatic hydroxylation to regioisomers M1 and M2 and direct conjugation to carbamate M7. As opined previously (Shaffer et al., 2005), the site of 4-oxotetrahydroindole hydroxylation is believed to be the same in M2, M4, and M5 based on CID spectra interpretation, although this was never determined unequivocally. Because of the inconsequential contribution of these two hydroxylated metabolites to the overall human metabolism of 1, the P450 isoform(s) responsible for their formation, as well as that of the monohydroxylated metabolites of 2 and 3, were not determined. An in-depth discussion of M7 formation has also been already undertaken (Shaffer et al., 2005). It is hypothesized that M7 may form by direct nucleophilic attack of dissolved CO₂ and/or carbonic acid by 1 in circulation; plasma is the lone matrix in which 1 may optimally interact with physiological carbonic acid-dissolved CO₂ pool equilibriums affecting M7 formation. In humans, total bicarbonate concentrations in plasma tend to be approximately 2-fold greater than that in tissues (Davis et al., 1993), with a fraction (0.5 mM) of plasma CO₂ in carbamate bonds with plasma proteins (White et al., 1968). Furthermore, human venous blood carries approximately a 2-fold greater amount of CO₂ within carbamino bonds than arterial blood (White et al., 1968). Hence, the conditions may be prime for M7 formation in venal plasma. Whether the formation of M7 is enzyme-mediated remains to be determined, although M7 was never observed in any human in vitro incubations with [¹⁴C]1.

Structure elucidation of U1 and U2 was elusive. Although these two metabolites each comprised on average approximately 9% of total plasma radioactivity, both were only detected in urine at <1% of administered 1. These data, coupled with the very short reverse-phase LC t_Rsof U1 and U2, suggest these metabolites are highly polar with very low volumes of distribution, which would rationalize them both contributing notably to total circulatory radioactivity and undergoing renal excretion while insignificantly comprising total urinary drug-related material. Such highly polar compounds would not be expected to permeate readily cellular constituents in vivo, making them of little concern from a safety perspective (Smith and Obach, 2005). It is conceivable that these unidentified, un-ionizable metabolites are some type(s) of conjugate(s) that revert(s) to 1 or one of its various metabolites before excretion. Metabolites U1 and U2 were never observed in any human liver-derived in vitro system using [¹⁴C]1.

Overall, the human metabolism and disposition data suggest both rats and monkeys were appropriate preclinical safety species. Particularly, monkeys most accurately projected the metabolism and disposition of 1 in humans as these species' excretion profiles, metabolite profiles, active renal clearance, and pharmacokinetics were essentially identical (Shaffer et al., 2005).

	Acknowledgments

 
We thank Neurogen Corporation for their contribution to the collaborative development of 1 with Pfizer Inc. and for supplying 1, 2, and 3. We also thank Dr. Klaas Schildknegt of the Radiosynthesis Group at PGRD, Groton, CT for the synthesis and purification of [¹⁴C]1, Beth Obach for conducting excreta and plasma radioanalysis, and Dr. Robert Berman for medical monitoring.

	Footnotes

 
Article, publication date, and citation information can be found at http://dmd.aspetjournals.org.

doi:10.1124/dmd.107.019760.

ABBREVIATIONS: GAD, generalized anxiety disorder; GABA_A, -aminobutyric acid type A; 1, N-[3-fluoro-4-[2-(propylamino)ethoxy]phenyl]-4,5,6,7-tetrahydro-4-oxo-1H-indole-3-carboxamide monomethanesulfonate; [¹⁴C]1, N-[3-fluoro-4-[2-(propylamino)ethoxy]phenyl]-4,5,6,7-tetrahydro-4-oxo-1H-[3-¹⁴C]indole-3-carboxamide monomethanesulfonate; HLM, human liver microsome; P450, cytochrome P450; MAO, monoamine oxidase; HPLC, high-performance liquid chromatography; LC/MS/MS, liquid chromatography/tandem mass spectrometry; LSC, liquid scintillation counting; AUC, area under the plasma concentration-time curve; k_el, elimination rate constant; LLOQ, lower limit of quantification; rcf, relative centrifugal force; MRM, multiple-reaction monitoring; LC t_R, liquid chromatography retention time; CID, collision-induced dissociation; 2, 2-fluoro-4-[(4-oxo-4,5,6,7-tetrahydro-1H-indole-3-carbonyl)-amino]-phenoxy acetic acid; 3, 4-oxo-4,5,6,7-tetrahydro-1H-indole-3-carboxylic acid [3-fluoro-4-(2-hydroxy-ethoxy)-phenyl]-amide; GFR, glomerular filtration rate; ALDH, aldehyde dehydrogenase; ADH, alcohol dehydrogenase.

¹ Current affiliation: Novartis Institutes for BioMedical Research, Department of Metabolism and Pharmacokinetics, Cambridge, MA.

² Current affiliation: Millennium Pharmaceuticals Inc., Department of Clinical Pharmacology, Cambridge, MA.

Address correspondence to: Dr. Christopher L. Shaffer, Pharmacokinetics, Dynamics and Metabolism, Pfizer Global Research and Development, Groton/New London Laboratories, Pfizer Inc., Eastern Point Road, MS 8220-4186, Groton, CT 06340. E-mail: christopher.l.shaffer{at}pfizer.com

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