Irosustat is a novel steroid sulfatase inhibitor for hormone-dependent cancer therapy. Its structure is a tricyclic coumarin-based sulfamate that undergoes desulfamoylation in aqueous solution, yielding the sulfamoyl-free derivative, 667-coumarin. The aim of the present work was to study the in vitro metabolism of irosustat, including its metabolic profile in liver microsomes and hepatocytes, the potential species differences, and the identification of the main metabolites and of the enzymes participating in its metabolism. Irosustat was extensively metabolized in vitro, showing similar metabolite profiles among rat, dog, monkey, and humans (both sexes). In liver microsomes, the dog was the species that metabolized irosustat most similarly to metabolism in humans. Marked differences were found between liver microsomes and hepatocytes, meaning that phase I and phase II enzymes contribute to irosustat metabolism. Various monohydroxylated metabolites of irosustat and of 667-coumarin were found in liver microsomes, which mostly involved hydroxylations at the C8, C10, and C12 positions in the cycloheptane ring moiety. 667-Coumarin was formed by degradation but also by non-NADPH-dependent enzymatic hydrolysis, probably catalyzed by microsomal steroid sulfatase. The main metabolites formed by hepatocytes were glucuronide and sulfate conjugates of 667-coumarin and of some of its monohydroxylated metabolites. The major cytochrome P450 enzymes involved in the transformation of irosustat were CYP2C8, CYP2C9, CYP3A4/5, and CYP2E1. Moreover, various phase II enzymes (UDP-glucuronosyltransferases and sulfotransferases) were capable of conjugating many of the metabolites of irosustat and 667-coumarin; however, the clinically relevant isoforms could not be elucidated.
Irosustat (also known as BN83495, 667 COUMATE, or STX64) is an irreversible steroid sulfatase (STS) inhibitor for steroid hormone-dependent cancer therapy and is currently under clinical development by the Ipsen Group. STS enzyme is widely distributed throughout the body and regulates the formation of estrone and dehydroepiandrosterone from their corresponding sulfate conjugates. Both compounds can be further converted to estradiol and androstenediol, respectively, which are described to promote tumor growth (Reed et al., 2005; Foster, 2008). Irosustat exhibits potent STS inhibition both in vitro, showing an IC50 value of 8 nM in placental microsome preparations (Woo et al., 2000) and in vivo in a MCF-7 xenograft breast cancer model (Foster et al., 2006). It also causes regression of estrone sulfate-stimulated nitrosomethylurea-induced mammary tumors in ovariectomized rats (Purohit et al., 2000). Moreover, irosustat was the first STS inhibitor to be tested in phase I clinical trials for the treatment of postmenopausal women with advanced metastatic hormone-dependent breast cancer, showing encouraging results (Stanway et al., 2006; Palmieri et al., 2011). In addition to breast cancer therapy, STS inhibitors may be useful for the treatment of other steroid hormone-dependent cancers such as prostate cancer (Selcer et al., 2002).
The irosustat structure is a tricyclic coumarin-based sulfamate (Fig. 1A). The presence of the sulfamoyl ester group is indispensable for its STS inhibitory activity and, in addition, confers to the irosustat molecule the ability to bind and to reversibly inhibit carbonic anhydrase II (Ho et al., 2003; Lloyd et al., 2005), an enzyme that is highly expressed in mammalian erythrocytes. Irosustat undergoes spontaneous desulfamoylation in aqueous solutions at nearly physiological pH (Ireson et al., 2003) leading to the formation of its major degradation derivative, 667-coumarin (structure shown in Fig. 1B), a process that is enhanced by increasing temperature. The binding to carbonic anhydrase II enzyme induces irosustat uptake and transport by red blood cells, while preventing it from degradation (Ireson et al., 2004).
During the early development phases of a new drug candidate, in vitro metabolism studies give essential information to help select the most suitable species for toxicological studies and to provide valuable foresight on metabolic pathways in humans. To this purpose, several in vitro test systems from different species and sexes, such as liver microsomes and hepatocytes, are commonly used in drug metabolism assessments. In the present work, the metabolic profile, the metabolite identification, and the potential species differences in the in vitro metabolism of irosustat were characterized using pooled liver microsome preparations from rats, dogs, monkeys, and humans and hepatocytes from rat, dog, and humans. Moreover, the enzymes participating in irosustat metabolism in humans were also investigated to help to predict possible drug-drug interactions.
Materials and Methods
Compounds irosustat [molecular weight of free base 309.3; purity by high-performance liquid chromatography (HPLC) 99.6%] and 667-coumarin (molecular weight of free base 230.3; purity by HPLC 99.1%) were synthesized by Panchim (Evry, France). [14C]Irosustat (specific activity 123 mCi/mmol; radiochemical purity by HPLC 98.7%) was custom-labeled by GE Healthcare (Little Chalfont, Buckinghamshire, UK). All reagents including all cytochrome P450 (P450)-specific inhibitors were purchased from Sigma-Aldrich (St. Louis, MO), except where indicated otherwise. Solvents used for HPLC analysis were of analytical or HPLC grade. Male and female Sprague-Dawley rat hepatocytes and liver microsomes and liver microsomes from male beagle dogs and from male and female cynomolgus monkeys were prepared in-house. Female human liver microsomes (HLM), female beagle dog liver microsomes, and the Reaction Phenotyping Kit version 7 (consisting of HLM from 16 separate donors) were purchased from XenoTech, LLC (Lenexa, KS). Fresh male beagle dog hepatocytes and cryopreserved female human hepatocytes were purchased from Biopredic (Rennes, France). Male HLM; commercial microsomes from Baculovirus-insect cell-expressed human P450s (Supersomes) CYP1A1, 1A2, 1B1, 2A6, 2B6, 2C8, 2C9*1, 2C19, 2D6*1, 2E1, 3A4, 3A5, and control; human UDP-glucuronosyltransferases (UGTs) 1A1, 1A3, 1A4, 1A6, 1A7, 1A8, 1A9, 1A10, 2B4, 2B7, 2B15, and 2B17; and human flavin-containing monooxygenase 3 (FMO3) were all purchased from BD Gentest (Woburn, MA). Human sulfotransferases (SULTs) 1A1*1 and 1A3 were purchased from Cypex (Dundee, Scotland, UK), and SULTs 1A1*2 and 1A2*1 were purchased from Invitrogen (Carlsbad, CA). All liver-derived samples were obtained with permission from the ethics committees of Ipsen Pharma S.A. and the other local manufacturers.
Preparation of Liver Microsomes.
Liver samples were excised and washed in isotonic saline. All subsequent steps were performed at 4°C. The liver was placed on crushed ice, minced with scissors, and then homogenized in 50 mM Tris-HCl buffer (pH 7.4) containing 154 mM KCl, using a Potter-Elvehjem homogenizer. The homogenate was centrifuged at 10,000g at 4°C for 20 min, and the resulting pellet was discarded. The supernatant was centrifuged at 100,000g at 4°C for 1 h (Lake, 1987), and the pellet (microsomes) was resuspended in 50 mM Tris-HCl buffer, pH 7.4, containing 154 mM KCl. The protein concentration was adjusted to 20 mg/ml, and the resulting microsomal suspensions were aliquoted and stored at −80°C. Microsomes were characterized by determination of total content of cytochrome P450 (Omura and Sato, 1964) and by determination of cytochrome c (P450) reductase (Lake, 1987), ethoxycoumarin O-deethylase (Peters et al., 1991), and testosterone 6β-hydroxylase activities (Kawano et al., 1987).
Hepatocyte Isolation and Culture.
Male and female Sprague-Dawley rats (180–220 g) were anesthetized with ketamine-medetomidine (10:0.1 mg). After loss of the righting reflex, an abdominal midline incision was made, and the portal vein was cannulated. Hepatocytes were further isolated by collagenase perfusion method adapted from Seglen (1976) and Gomez-Lechon et al. (1992). After wash in a suspension medium [50% Leibovitz L-15 medium and 50% Ham's F-12 nutrient mixture, containing 5% fetal calf serum, 0.94 mg/ml d-(+)-glucose, 2 mg/ml bovine serum albumin, 2 mM l-glutamine, 10 mM HNaCO3, 100 IU/ml penicillin, 100 μg/ml streptomycin, and 10−8 M insulin], the viability and cell density were measured by the trypan blue exclusion method using a hemocytometer. Finally, rat hepatocytes were plated in 24-well culture dishes coated with collagen substratum (cell density 25 × 104 viable cells/well). Fresh male beagle dog hepatocytes were purchased already plated in 24-well culture dishes coated with collagen substratum (cell density 30 × 104 viable cells/well). Female human cryopreserved hepatocytes were thawed according to the instructions provided by the supplier, and the viability and cell density were measured by the trypan blue exclusion method. Finally, human hepatocytes were plated in 24-well culture dishes coated with collagen substratum (cell density 0.4 × 106 viable cells/well).
Incubation of [14C]Irosustat with Liver Microsomes and Determination of Apparent Intrinsic Clearance.
Liver microsomes (1 mg/ml) from Sprague-Dawley rats, beagle dogs, cynomolgus monkeys, and humans (both sexes, separately) were incubated in duplicate with 25 μM [14C]irosustat at 37°C in 50 mM Tris-HCl buffer, pH 7.4, containing 5 mM MgCl2. Incubations were performed in the presence of a NADPH-generating system consisting of 4 mM d-glucose 6-phosphate, 2 IU/ml glucose-6-phosphate dehydrogenase and 1 mM β-NADP (0.5-ml final volume). The reactions were started by addition of β-NADP and were quenched at 0, 30, 60, and 120 min by addition of 1 volume of acetonitrile containing 10% acetic acid. Incubation mixtures without microsomes and without β-NADP incubated for 120 min were used as controls. The samples generated were either stored at −80°C until analysis or directly analyzed by HPLC after centrifugation at 20,000g at 4°C for 15 min and further dilution with HPLC mobile phase.
For each species and sex, the apparent intrinsic clearance (Clint, app) parameter was estimated using the t1/2 method, where the log percentage of substrate remaining versus time was plotted. For calculation, data from incubations up to 30 min were selected. Because 667-coumarin is formed by degradation during the incubations and both irosustat and 667-coumarin actually act as a substrate, the percentage remaining included both compounds, enabling the estimation of the P450-mediated Clint, app. The slope of the resulting linear regression (−k) was used to obtain in vitro t1/2 (in vitro t1/2 = 0.693/k). The in vitro t1/2 was further converted to Clint, app using the following equation: Clint, app = [(0.693/in vitro t1/2) · (milliliters of incubation per milligram of microsomes) · (45 mg of microsomes/g liver) · (X g of liver/kg b.wt.)], where X = 45 for rat, 25 for dog, 30 for monkey, and 20 for human (Bohnert and Gan, 2010).
Incubation of Unlabeled Irosustat with Hepatocytes.
Cultured hepatocytes from male and female Sprague-Dawley rats, male beagle dogs, and women, were incubated in duplicate with 50 μM unlabeled irosustat at 37°C in a humidified atmosphere containing 95% air and 5% CO2. The incubation volume was 0.5 ml, and the incubation time period was 3 h. The incubations were quenched by addition of 1 volume of acetonitrile containing 3% acetic acid. The samples generated were either stored at −80°C until analysis or directly analyzed by HPLC after centrifugation at 20,000g and 4°C for 15 min and dilution with HPLC mobile phase.
Incubation of Unlabeled Irosustat and 667-Coumarin with Rat Liver Microsomes and Rat Hepatocytes.
Liver microsomes (1 mg/ml) and hepatocytes from female Sprague-Dawley rats were incubated either with 50 μM irosustat or with 50 μM 667-coumarin for 120 min (liver microsomes) and 3 h (hepatocytes), under the same conditions as described previously.
Estimation of the Apparent Km and Vmax for Irosustat Metabolism in HLM.
Before assessment of the enzyme kinetics, the linearity of 14C-labeled irosustat transformation in HLM was evaluated in a range of increasing microsomal protein concentrations and incubation time periods. Linear metabolite formation was found up to approximately 1 mg of microsomal protein/ml and up to 40 min of incubation, respectively (data not shown). These incubation conditions were selected for the enzyme kinetics experiment. The microsomal incubations were conducted with final [14C]irosustat concentrations of 0, 5, 10, 25, 50, 80, 110, 150, and 250 μM (n = 4 for each concentration, except for 110 and 150 μM, for which n = 2). Reactions were quenched, and samples were analyzed with on-line radioactivity detection as described below. Apparent Km and Vmax were estimated after fitting the Michaelis-Menten equation V = Vmax S/(Km + S) to the data obtained, using WinNonlin (version 3.3; Pharsight, Mountain View, CA).
Determination of the Human P450 Enzymes Responsible for Irosustat Metabolism.
The main P450 enzymes responsible for the formation of phase I metabolites of irosustat were identified using the following three complementary approaches.
Correlation study with P450 isoform-specific activities in HLM.
The irosustat biotransformation rate (as metabolite formation) in incubations of the test compound with a panel of 16 individual batches of characterized HLM was correlated with the rate of 13 probe P450-specific reactions: 7-ethoxyresorufin O-dealkylation and phenacetin O-deethylation (for CYP1A2), coumarin 7-hydroxylation (for CYP2A6), S-mephenytoin N-demethylation and bupropion hydroxylation (for CYP2B6), paclitaxel 6α-hydroxylation (for CYP2C8), diclofenac 4′-hydroxylation (for CYP2C9), S-mephenytoin 4′-hydroxylation (for CYP2C19), dextromethorphan O-demethylation (for CYP2D6), chlorzoxazone 6-hydroxylation (for CYP2E1), testosterone 6β-hydroxylation and midazolam 1′-hydroxylation (for CYP3A4/5), and benzydamine N-oxidation (for FMO3). The incubations (n = 2) were performed with 50 μM unlabeled irosustat and 1 mg/ml microsomal protein concentration for 40 min, under the conditions described previously for incubations of irosustat with liver microsomes. The results of activity assays using specific cytochrome P450 substrates were taken from the data sheet supplied by the microsome manufacturer (Reaction Phenotyping Kit; XenoTech, LLC). The Pearson correlation coefficient (r) was estimated, and the correlation was accepted as significant when the probability level was lower than 0.01 (α = 0.01). NCSS 2001 (Intermountain Scientific/BioExpress, Kaysville, UT) was used for calculation.
Chemical inhibition of P450 enzymes in pooled HLM.
Incubations of 50 μM irosustat in the presence and absence of 10 specific P450 inhibitors (n = 2, except for the CYP2C19 inhibitors nootkatone and (+)-N-3-benzyl-nirvanol, n = 3) were performed using pooled HLM under the incubation conditions described above. The following chemical inhibitors and final concentrations were used (all working solutions were made in dimethyl sulfoxide): 1 μM furafylline (CYP1A2, mechanism-based inhibitor); 0.1 μM methoxsalen (CYP2A6, mechanism-based inhibitor); 750 μM orphenadrine (CYP2B6 reversible inhibitor); 10 μM quercetin (CYP2C8, reversible inhibitor); 2 μM sulfaphenazole (CYP2C9, reversible inhibitor); 100 μM nootkatone and 5 μM (+)-N-3-benzyl-nirvanol (CYP2C19, reversible inhibitors); 0.5 μM quinidine (CYP2D6, reversible inhibitor); 15 μM 4-methylpyrazole (CYP2E1, reversible inhibitor); and 0.5 μM ketoconazole (CYP3A4/3A5, reversible inhibitor). For the mechanism-based inhibitors furafylline and methoxsalen, HLM were preincubated for 15 min in the presence of inhibitor and the NADPH-generating system before addition of irosustat.
Incubation with cDNA-expressed human enzymes.
Unlabeled irosustat (50 μM) was incubated (n = 2) for 60 min with insect cell microsomes expressing individual human P450 enzymes (Supersomes, 100 pmol P450/ml): CYP1A1, 1A2, 1B1, 2A6, 2B6, 2C8, 2C9*1, 2C19, 2D6*1, 2E1, 3A4, and 3A5. Irosustat was also incubated with 500 μg/ml FMO3, control nonenzyme-expressing insect cell microsomes, and with CYP2B6 at shorter incubation times (10, 20, and 40 min). The incubations were performed at 37°C as described previously. The metabolism of 50 μM unlabeled 667-coumarin by the cDNA-expressed human enzymes was also investigated following the same incubation conditions.
Investigation of the Role of CYP1A2 in Irosustat Metabolism.
Native pooled HLM and CYP1A2-fortified HLM showing 2-fold increased CYP1A2 activity were used for these experiments. To prepare CYP1A2-fortified HLM, phenacetin O-deethylase activity was determined in both, CYP1A2 Supersomes and pooled HLM, using 500 μM phenacetin as substrate (concentration at Vmax). The data resulting from this test indicated that the concentration of recombinant human CYP1A2 necessary to increase the CYP1A2 activity in the HLM suspension 2-fold was 20 pmol/ml (data not shown). For the main experiment, 50 μM unlabeled irosustat was incubated with native pooled HLM (1 mg/ml) or CYP1A2-fortified HLM (1 mg/ml plus 20 pmol/ml CYP1A2) in the presence or absence of a mix of P450 chemical inhibitors (10 μM quercetin, 2 μM sulfaphenazole, 20 μM tranylcypromine, and 0.5 μM ketoconazole). In all cases, incubations were performed in the presence or absence of 1 μM furafylline. The incubations (n = 2) were performed for 40 min after a 15-min preincubation with or without furafylline, under the conditions described previously.
Determination of the Phase II Enzymes Responsible for the Metabolism of Irosustat.
Preliminary results indicated that cDNA-expressed human CYP1A2 is able to reproduce the metabolite profile of irosustat obtained in HLM. Therefore, unlabeled irosustat was incubated with CYP1A2 Supersomes (200 pmol P450/ml). After a 120-min incubation, UGT individual human enzymes (1 mg/ml final concentration, n = 2), UGT1A1, 1A3, 1A4, 1A6, 1A7, 1A8, 1A9, 1A10, 2B4, 2B7, 2B15, and 2B17, were added to the incubates together with 2 mM UDP-glucuronic acid, 0.025 mg/ml alamethicin, and MgCl2 supplementary solution (10 mM final concentration). The incubation volume was 400 μl, and the reactions were quenched after 120 min from UGT addition with 1 volume of acetonitrile containing 10% acetic acid.
As in the case of human UGTs, unlabeled irosustat was first incubated with 200 pmol/ml CYP1A2 Supersomes. After a 120-min incubation, SULT individual human enzymes (12.5 μg/ml final concentration, n = 2), SULT1A1*1, SULT1A1*2, SULT1A2*1, and SULT1A3, were added to the incubates together with 50 μM adenosine 3′-phosphate 5′-phosphosulfate, 10 mM dithiothreitol, and MgCl2 supplementary solution (5 mM final concentration). The incubation volume was 400 μl, and the reactions were quenched after 120 min from SULT addition with 1 volume of acetonitrile containing 10% acetic acid.
Formation of glucuronide and sulfate conjugates of 667-coumarin.
Unlabeled 667-coumarin was directly incubated (n = 2) with the human UGT and SULT enzymes, under the same conditions of incubation but without the preincubation step with CYP1A2 Supersomes.
[14C]Irosustat and its metabolites were separated by HPLC using a Waters 600 solvent delivery system equipped with a reverse-phase column (Sunfire C18, 150 × 4.6 mm, 5-μm particle size; Waters, Milford, MA). The mobile phase consisted of 50 mM ammonium formate, pH 5.0 (solvent A), and methanol (solvent B), and the flow rate was 1 ml/min. The initial mobile phase contained 25% solvent B and was maintained at this composition for 2 min. The percentage of solvent B was increased linearly up to 53% over the next 24 min and further increased to 70% in the following 7 min. The percentage of solvent B was rapidly changed to 100% in 0.1 min and maintained at this composition for 5 min. Finally, the percentage of solvent B was returned to 25% in the last 7 min. On-line radiochemical detection was performed using a LB508 radioflow detector equipped with a 150-μl solid scintillation cell (YG-150) from Berthold Technologies (Bad Wildbad, Germany). When nonradiolabeled irosustat was used, UV peak detection was performed using a Waters 486 variable wavelength or a Waters 2487 dual λ absorbance detector at 312 nm. Occasionally, the HPLC column was replaced by a Symmetry C18 column (250 × 4.6 mm, 5-μm particle size; Waters) or a Sunfire C18 column (100 × 4.6 mm, 3.5-μm particle size; Waters); the gradient times and the flow rate were adjusted to the column dimensions when necessary.
Mass Spectrometric Characterization.
Metabolites in representative incubate samples were selected for LC-MS characterization. Monitoring of metabolite ions was conducted using a ZQ2000 mass spectrometer (Waters) over the regions where HPLC peaks were present. The mass spectrometer operated with an electrospray ion source in positive ionization mode. Data were captured by means of full-scan mass spectra. The capillary and extractor voltages were set at 3 kV and 3 V, respectively, the radiofrequency lens voltage was set at 0.5 V, the source and desolvation temperatures were set at 120 and 250°C, respectively, and the desolvation and cone gas flow rates were 400 and 50 l/h, respectively. Two cone voltages (30 and 60 V) were used to determine the fragmentation pattern of the compounds. The spectra obtained in chromatograms from incubates with irosustat or 667-coumarin were compared with blank samples. Data were acquired by MassLynx 4.0 (Waters).
Phase I Metabolite Isolation.
The main phase I metabolites were produced by incubating 50 μM irosustat or 667-coumarin with rat liver microsomes (2 mg/ml protein concentration) over 120 min as described previously. Incubates were centrifuged, and the supernatants diluted with 50 mM ammonium formate, pH 5.0. The metabolites were purified by solid-phase extraction (SPE) using Sep-Pak Plus C-18 cartridges (Waters). SPE cartridges were activated with acetonitrile and further conditioned with 50 mM ammonium formate, pH 5.0. Then diluted supernatants (5 ml) were loaded onto each SPE cartridge, and a washing step was performed with 50 mM ammonium formate, pH 5.0. Metabolites were eluted using 2 ml of acetonitrile. The SPE fractions were pooled and dried under a stream of nitrogen, and the dry residues were reconstituted into 50 mM ammonium formate, pH 5.0, containing 17% acetonitrile. Metabolites were separated by HPLC with a Symmetry C18 column (250 × 4.6 mm, 5 μm; Waters) and purified by fraction collection of HPLC elutes. The isolated metabolite fractions were freeze-dried on a Christ Alpha 2-4 Lyophilizer (B. Braun-Biotech, Melsungen, Germany). The freeze-drying procedure was repeated three times.
NMR Analysis of Isolated Metabolites Lacking Sulfamoyl Group.
The characterization of the chemical structure of isolated 667-coumarin derivatives was performed by NMR (Serveis Cientifico-técnics, University of Barcelona, Barcelona, Spain). NMR experiments were performed on either a 600- or 800-MHz Avance NMR spectrometer fitted with TCI H-C/N-D-05 cryoprobes (Bruker, Wissembourg, France). Samples of the isolated metabolites were dissolved in dimethyl sulfoxide-d6, and NMR data sets were acquired at 25°C. The experiments performed were monodimensional 1H NMR spectra, two-dimensional homonuclear 1H-1H correlation experiments (correlation spectroscopy and total correlation spectroscopy), and two-dimensional heteronuclear 1H-13C correlation experiments (heteronuclear single quantum correlation and heteronuclear multiple bond correlation).
Identification of Irosustat Primary Metabolites (Containing Sulfamoyl Group).
Isolated irosustat primary metabolites were incubated in aqueous buffer at pH 7 to 8 at 37°C for 2.5 h to induce desulfamoylation. The resulting 667-coumarin derivatives where compared with the NMR-identified derivatives by HPLC coelution experiments.
Some of the main phase II metabolites of irosustat were tentatively identified by LC-MS as glucuronides of a number of oxidized derivatives of 667-coumarin. To assess the identity of the aglycones (actually phase I metabolites), the glucuronides were obtained by incubation of irosustat with hepatocytes and purified by fraction collection of HPLC elutes. The fractions containing each metabolite were pooled and dried under a stream of nitrogen. After dilution with 50 mM ammonium formate, pH 5.0, the specimens were freeze-dried. The conjugated metabolites were individually incubated with 3.75 mg/ml β-glucuronidase enzyme (from Helix pomatia, type H-1; Sigma-Aldrich) in 100 mM sodium acetate buffer, pH 5.0, at 37°C for 6 h. The incubations were quenched by addition of acetonitrile containing 10% acetic acid. All the samples generated were analyzed by HPLC, and the resulting phase I metabolites where compared with those already identified by HPLC coelution experiments.
In Vitro Metabolite Profile of [14C]Irosustat in Liver Microsomes from Different Species and Sexes.
The in vitro metabolic profile of [14C]irosustat in liver microsomes was compared among rats, dogs, monkeys, and humans (males and females separately). The relative percentage of peak area in the radio-HPLC profiles for each metabolite after 120-min incubations is presented in Table 1. When all species and sexes during the whole time course experiments were considered, up to 15 different labeled metabolites of [14C]irosustat were detected. The radio-HPLC metabolite profiles obtained were analogous to those obtained by UV detection at 312 nm. A representative HPLC-UV profile of a HLM incubate is shown in Fig. 2A.
The [14C]irosustat metabolites showing a percentage of radioactive peak area higher than 5% were considered as main metabolites. In liver microsomes from the different species under study, the main metabolites were M7, M8, M13, M14, M16, and 667-coumarin. They were found in all samples, except for M14, which was not detected in female monkey microsome incubates, appearing only in males. Besides these main metabolites, some minor metabolites accounting for relative percentages less than 4.4%, were also detected and showed some differences among species and/or between sexes, although these differences were not considered to be relevant.
Taking into account Clint, app parameters (Table 2), the highest transformation rate of [14C]irosustat was observed in monkey liver microsomes (faster in females than in males), followed by rat, dog, and human microsomes. The fastest metabolism rate in monkeys was associated with increased production of metabolites M7 and M8 compared with those of the other species (Table 1).
The in vitro metabolite profile of [14C]irosustat in dog liver microsomes was the closest to the one obtained in HLM both qualitatively and quantitatively, except for metabolite M11, which was detected in human but not in dog microsomes.
Results from control samples without microsomes demonstrated that irosustat is hydrolyzed to 667-coumarin under the incubation conditions (at 37°C and pH 7.4). However, the amount of 667-coumarin formed in these samples was lower than the amount formed in control samples containing microsomes without β-NADP (Table 2).
In Vitro Metabolite Profile of Irosustat in Hepatocytes from Different Species.
As shown in Fig. 2, the pattern of irosustat metabolism by hepatocytes was remarkably different from that obtained with liver microsomes. Five main metabolites of irosustat were detected in incubations with human hepatocytes: M1, M2, M12, M17, and 667-coumarin (Fig. 2B). Except for 667-coumarin, these metabolites were not formed in the microsome incubations, suggesting that most of them might be phase II metabolites. Figure 2, C and D, also shows the metabolite profile of irosustat in hepatocytes from rat and dog, respectively: six main peaks in addition to 667-coumarin (M2, M3, M7, M12, and M17) were detected in rat hepatocytes, and in dog the main metabolites detected were M12, M17, and 667-coumarin.
Determination of the Human P450 Enzymes Responsible for the Metabolism of Irosustat.
As step preceding the enzyme identification experiments, the substrate concentration-dependent transformation of [14C]irosustat was assessed in HLM. The kinetics showed a classic hyperbolic pattern (data not shown), and, therefore, the Michaelis-Menten kinetics equation was fitted to the metabolite formation data, giving a Km value of 43.4 μM and a Vmax value of 369 pmol/mg per min. From these results and former data on linearity of [14C]irosustat metabolism, the incubation conditions selected for the following phenotyping experiments were set as follows: 50 μM irosustat, 1 mg/ml microsomal protein concentration, and 40-min incubation time.
To identify the P450 enzymes responsible for the metabolism of irosustat, a combination of three experimental approaches was used: 1) correlation analysis of irosustat phase I metabolite formation rate with several P450 isoform-specific activities in a panel of 16 individual HLM; 2) assessment of the effect of chemical P450 isoform-specific inhibitors on irosustat metabolite formation in pooled HLM; and 3) formation of irosustat metabolites by cDNA-expressed P450 enzymes. Variable results were obtained for each approach. The correlation between specific P450 activities and irosustat metabolite formation seemed to be the most selective part of the study, and the results are shown in Table 3. Under our experimental conditions, the formation of all irosustat metabolites, except for P-36, correlated with the activity of one or more members of CYP2C family (i.e., CYP2C8, CYP2C9, and CYP2C19). For some of the metabolites, an additional correlation was found with CYP3A4/5 (M13, M14, and M16) and CYP2E1 (M7) activities. Metabolite P-36 did not correlate with any of the P450s tested. The results of the chemical inhibition phase (Table 4) showed that almost all inhibitors, except those for CYP1A2 (furafylline) and CYP2C19 [nootkatone and (+)-N-3-benzyl-nirvanol], relevantly decreased the formation of several irosustat metabolites. Table 5 shows the results obtained in incubations of cDNA-expressed P450 enzymes with irosustat. Recombinant CYP1A2 and CYP2C19 were able to form most of the irosustat metabolites produced in HLM incubations. CYP2A6 and FMO3 enzymes were not able to metabolize irosustat. Because CYP2B6 was able to form M16 and P-36 but not their respective sulfamoylated precursors M13 and M18, CYP2B6 was also incubated with irosustat for shorter incubation times, showing formation of M18 but not M13. 667-Coumarin was also incubated with the same cDNA-expressed P450 enzymes. The 667-coumarin metabolites M8, M16, and P-36 were formed by different enzymes. M8 was formed by the same enzymes as irosustat, M16 was formed by the same enzymes as irosustat plus CYP2A6, CYP2C8, and CYP2E1, and P-36 was formed by all P450s except for CYP2C8, CYP2C9, and CYP2E1.
Although all three approaches have advantages and disadvantages, a combination of them was required to identify which enzyme or enzymes could potentially be responsible for irosustat metabolism. The enzymes that showed positive results in the three approaches were CYP2C8, CYP2C9, CYP2E1, and CYP3A4/5, and they were identified as the main enzymes responsible for the phase I metabolism of irosustat.
Investigation of the Role of CYP1A2 in Irosustat Metabolism.
The effect of a combination of chemical inhibitors of the P450s involved in irosustat metabolism was assessed in native and CYP1A2-fortified HLM in the presence or absence of furafylline (CYP1A2 inhibitor). Table 6 shows the percentage of inhibition for each of the irosustat metabolites formed. Furafylline alone showed no inhibitory effect either in native or in CYP1A2-fortified HLM, except for a slight inhibition of M14 formation in the latter. The formation of all metabolites was remarkably inhibited by the combination of specific P450 inhibitors in both HLM models. Approximately the same inhibition percentages were found in irosustat incubations with native HLM when the mix of inhibitors was used either in the presence or absence of furafylline. However, in CYP1A2-fortified HLM, the inhibition of irosustat metabolism increased (for almost all metabolites) when furafylline was used together with the mix of P450-inhibitors.
Determination of the Phase II Enzymes Responsible for the Metabolism of Irosustat.
To identify the relative contribution of specific human liver enzymes responsible for the formation of the phase II metabolites of irosustat, incubations were performed using cDNA-expressed human UGT and SULT enzymes in the presence of the respective cofactors. The experimental approach consisted of incubating irosustat with recombinant CYP1A2 to induce the formation of most of the phase I in vitro metabolites (Table 5), followed by the addition of the different UGT and SULT enzymes and the required cofactors.
As shown in Table 7, up to nine peaks corresponding to putative glucuronides were produced by the different UGTs. These metabolites were as follows: M12 and M2 (already detected in hepatocyte incubations; Fig. 2) and G1, G2, G3, G4, M4, G7, and G8 (newly detected metabolites). M12 was also formed when 667-coumarin was incubated directly with all UGTs except for UGT1A4, which, in turn, was the unique UDP-glucuronosyltransferase unable to form any of the glucuronides.
As shown in Table 8, up to eight peaks corresponding to potential sulfate conjugates were found after incubation of irosustat phase I metabolites with different human SULTs (1A1*1, 1A1*2, 1A2*1, and 1A3), using the same experimental approach. The sulfate conjugates were named S2, S3, S4, S5, S6, and S7 (new peaks) and were formed in addition to M17 and M3, which had already been detected in hepatocyte incubations (Fig. 2). M17 formation was also confirmed when 667-coumarin was incubated directly with all the SULTs. Sulfotransferases SULT1A1*1, 1A1*2, and 1A2*1, but not SULT1A3, were capable of forming M17. SULT1A1*1 was able to produce all the sulfate conjugates identified and also in the largest amounts.
Preliminary Identification of Irosustat Main Metabolites by MS.
The MS spectra of irosustat and its main in vitro metabolites are shown in the supplemental data. The MS spectrum of irosustat was characterized by its protonated molecular ion of m/z 310 and by the formation of a main fragmentation product of m/z 231. This fragmentation product was consistent with the protonated molecular ion of the 667-coumarin molecule, which was produced after loss of the 3-O-sulfamoyl group in the MS ion source. In addition to 667-coumarin, the main in vitro irosustat metabolites were identified as monooxidized derivatives, and also glucuronide and sulfate conjugates.
Two groups of monooxidized metabolites were found in incubations of irosustat with microsomes: monooxidized irosustat derivatives (primary metabolites M7, M9, M13, and M14); and monooxidized 667-coumarin derivatives (secondary metabolites M8, M15, and M16). Differences in the fragmentation pattern of the metabolites at 60 V cone voltage (supplemental data) and, importantly, differences in their HPLC retention time (Fig. 2A) indicated that they are probably formed by oxidation in different atoms of the irosustat or 667-coumarin structure.
Three glucuronide metabolites could be identified by MS in hepatocyte incubations with irosustat: one corresponding to the 667-coumarin glucuronide (M12) and two monooxidized 667-coumarin glucuronides (M1 and M2).
One sulfate metabolite found in hepatocyte incubates was identified by MS as 667-coumarin sulfate and was named as M17.
In vitro incubations of 667-coumarin with liver microsomes and hepatocytes.
To help in the identification of irosustat metabolites, 667-coumarin and irosustat were incubated separately with both female rat liver microsomes and female rat hepatocytes at 37°C for 120 min and 3 h, respectively. The metabolite profiles obtained from 667-coumarin incubations were compared with those obtained with irosustat. As shown in Fig. 3, M8, M11, M15, M16, M12, M17, and P-36 were formed directly from 667-coumarin; conversely, the primary metabolites M7, M9, M13, and M14 were only formed when irosustat was used as substrate. These results supported the metabolite identification performed by MS.
Structure Elucidation of Irosustat Main Metabolites.
The main phase I metabolites of irosustat were purified as described under Materials and Methods. Those compounds, corresponding to putative monooxidized metabolites of irosustat, were converted to their respective 667-coumarin counterparts by incubation at neutral pH and 37°C temperature to force the desulfamoylation. The resulting oxidized 667-coumarin derivatives were further characterized by its HPLC retention time. The results showed that the irosustat oxidized metabolites or primary metabolites M7, M13, M14, and M18 were the sulfamate-containing precursors of the oxidized 667-coumarin derivatives M8, M16, M15, and P-36 (secondary metabolites), respectively.
The identification of the chemical structure of the monooxidized 667-coumarin derivatives (indeed, the position of the oxidized atoms) was performed by NMR, and the identification of their respective sulfamate-containing derivatives was automatically deduced from the data described previously (Fig. 4). The monodimensional 1H spectra of M8 and M16 indicated that no changes took place in the aromatic region with respect to 667-coumarin. The same conclusion was achieved for M15 after interpretation of the homonuclear 1H-1H correlation spectroscopy and total correlation spectroscopy and heteronuclear 1H-13C single quantum correlation and multiple bond correlation two-dimensional spectra. For all three cases, these spectra indicated that the hydroxyl group was located in the cycloheptane ring. The position of the hydroxyl group in the cycloheptane ring was determined from the study of the ring atoms in two-dimensional spectra. Thus, metabolites M8, M15, and M16 were hydroxylated at the C10, C12, and C8 positions in the cycloheptane ring of the 667-coumarin molecule, respectively. It is obvious that the same hydroxylation sites corresponded to their sulfamate-containing counterparts M7, M14, and M13. The structures described in Fig. 4 for these metabolites are compatible with the NMR data.
The glucuronidation and the sulfation of 667-coumarin may basically be produced through its unique hydroxyl group at the C3 position, yielding M12 (667-coumarin 3-O-glucuronide) and M17 (667-coumarin 3-O-sulfate), respectively (Fig. 4). Metabolites M1 and M2, which were preliminarily identified by mass spectrometry as glucuronides, were purified and treated with β-glucuronidase, leading to the formation of M11 and M16, respectively (Fig. 4).
The novel STS inhibitor irosustat was extensively metabolized in vitro in liver microsomes and hepatocytes. Its metabolite profile was similar among the species tested and between sexes. However, marked differences were found between both in vitro systems, meaning that phase I and phase II enzymes are involved in irosustat metabolism. Various monooxidized metabolites of irosustat (primary metabolites) and of its desulfamoylated derivative 667-coumarin (secondary metabolites) were formed in liver microsomes; therefore, oxidation was the predominant route of irosustat phase I metabolism. The most abundant monooxidized metabolites were identified by NMR, showing hydroxyl groups at different carbon atoms of the cycloheptane ring. As shown in Table 1 and Fig. 4, the main primary phase I metabolites were M7, M13, and M14 (10-hydroxy-, 8-hydroxy-, and 12-hydroxy-irosustat, respectively), and the corresponding secondary phase I metabolites were M8, M16, and M15 (10-hydroxy-, 8-hydroxy-, and 12-hydroxy-667-coumarin, respectively). On the other hand, the main metabolites in hepatocyte incubations were formed by phase II metabolic enzymes. This fact was further demonstrated by LC-MS analysis. Rat, dog, and human hepatocytes mainly converted irosustat to 667-coumarin and to 3-O-glucuronide and 3-O-sulfate conjugates of 667-coumarin (M12 and M17, respectively). In human hepatocytes, M1 and M2 derivatives were also formed and were identified as glucuronides of metabolites M11 and M16 (both 667-coumarin derivatives), respectively. However, the chemical structures of M1 and M2 were not fully determined. Because hepatocytes are a more physiologically significant in vitro model, a high relevance of phase II metabolism is anticipated in vivo.
The comparison of the microsomal metabolites of irosustat allowed us to detect metabolic similarities between animal species and humans. These results should be helpful in the selection of nonrodent species for toxicity evaluation. Qualitatively, all phase I human metabolites were formed by dog microsomes except for metabolite M11. Nevertheless, M11 was formed in rats, indicating that no unique human metabolites were formed in liver microsomes, and therefore no safety concerns are anticipated from the in vitro data. Quantitatively, the following rank order was established among species regarding irosustat Clint, app (Table 2): monkey ≫ rat > dog ≥ humans. As a conclusion, the dog was the species that showed the metabolic pattern closest to that of humans.
Under in vitro incubation conditions (at 37°C and pH 7.4), 667-coumarin was the major degradation product of irosustat formed by hydrolysis of the sulfamoyl ester group (Fig. 1). Desulfamoylation in aqueous solution probably occurs via the E1cB elimination reaction, assisted by the extended conjugation present in the coumarin motif (Lloyd et al., 2005). However, the results of the present work showed how incubations of irosustat with microsomes in the absence of cofactor produced increased amounts of 667-coumarin compared with incubations of irosustat in buffer alone (Table 2), demonstrating that 667-coumarin can be also formed by non-NADPH-dependent enzymatic hydrolysis. Therefore, 667-coumarin may be considered as a metabolite and not only as a degradation product. Moreover, 667-coumarin was previously described to be a product resulting after STS inhibition by irosustat (Woo et al., 2000). Because the STS enzyme is present in liver microsomes (Kauffman et al., 1998), STS is a probable candidate for the enzymatic formation of 667-coumarin.
Once the main metabolites of irosustat were identified and in consideration of the various phase I and phase II reactions involved, the enzymes capable of metabolizing irosustat were carefully evaluated. Figure 4 summarizes the in vitro metabolism pathways of irosustat including the enzymes considered to be responsible for its primary metabolism (those showing positive results in the correlation, inhibition, and recombinant enzyme approaches): M7 was formed by CYP2C9 and CYP2E1, M9 by CYP2C8, M13 by CYP2C8, CYP2C9, and CYP3A4/5, M14 by CYP3A4/5, and M18 by CYP2C9. Although the secondary metabolites M8, M11, M16, and P-36 were included in the phenotyping experiments, the enzymes involved in their formation should be considered as not fully elucidated because desulfamoylated derivatives can be formed by two sequential reactions: 1) irosustat hydroxylation and further hydrolysis of the sulfamoyl group or 2) loss of the sulfamoyl group followed by hydroxylation. These two ways were demonstrated in the present work: first, by converting hydroxylated irosustat metabolites into their respective 667-coumarin counterparts by incubation at neutral pH and at 37°C; and second, by direct incubation of 667-coumarin with rat microsomes and hepatocytes (Fig. 3) and with recombinant human P450s. Because of the structural similarities between irosustat and 667-coumarin, it is likely that the same enzymes could be involved in the transformation of both compounds. From the correlation and chemical inhibition tests (Tables 3 and 4), no relevant differences in phenotyping between the primary metabolites and the respective secondary metabolites were obtained, except for CYP2E1, which is related to M7 formation but not to M8. In the recombinant P450s approach, irosustat and 667-coumarin were transformed by almost the same enzymes. However, some main differences were found: 1) CYP2A6 was able to metabolize 667-coumarin but not irosustat; 2) CYP2C9 was able to metabolize irosustat but not 667-coumarin; and 3) CYP2B6 formed M16 from 667-coumarin but was not able to form M13 from irosustat. These different capabilities of CYP2A6, CYP2B6, and CYP2C9 to metabolize irosustat and 667-coumarin were considered as not relevant, given the high number of enzymes involved in the metabolism of both compounds.
Surprising results were obtained for CYP1A2 and CYP2C19 enzymes when the recombinant P450s approach was assessed. The incubation of irosustat with CYP1A2 Supersomes showed that this enzyme was able to produce a metabolite profile very similar to the one obtained by pooled HLM. However, the two specific CYP1A2 activities measured in the 16 individual HLM batches, 7-ethoxyresorufin O-dealkylation and phenacetin O-deethylation, did not correlate with the formation of any of the irosustat metabolites. In addition, the CYP1A2 mechanism-based inhibitor furafylline (1 μM) had no effect on the metabolism of irosustat in pooled HLM. The inhibition was also assessed using a 10-fold increased furafylline and 10-fold reduced irosustat concentration with similar results (data not shown). Furthermore, the extent of inhibition of furafylline on irosustat metabolism was studied comparatively in incubations with native HLM and CYP1A2-fortified HLM (Table 6). As expected, irosustat metabolism was markedly decreased in both HLM models in the presence of a combination of P450-specific inhibitors, confirming that irosustat is metabolized by diverse P450 isoforms. In contrast, furafylline (either alone or in combination with the mix of P450-inhibitors) showed a negligible effect in native HLM, whereas in CYP1A2-fortified HLM, it essentially enhanced the inhibition induced by the other P450-inhibitors, meaning that several P450s must be affected at the same time to produce any potential metabolic interaction outcome.
Concerning CYP2C19, the S-mephenytoin 4′-hydroxylation activity in 15 individual HLM batches (one was considered as an outlier) correlated with the formation of some of the irosustat metabolites in HLM. In addition, the incubation of irosustat with recombinant CYP2C19 also produced several irosustat metabolites. However, the two specific CYP2C19 inhibitors used, nootkatone and (+)-N-3-benzyl-nirvanol (Tassaneeyakul et al., 2000; Suzuki et al., 2002), did not inhibit the formation of any irosustat metabolite in the pooled HLM model.
Overall, CYP1A2 and CYP2C19 enzymes were considered as not relevant in irosustat metabolism when other P450 enzymes are present and active, although a minor role cannot be completely discarded (i.e., a weak contribution of CYP1A2 in the case of overexpression or induction). From the phenotyping results, we may conclude that, even if coadministered drugs inhibit one of the identified drug-metabolizing enzymes, the pharmacokinetics of irosustat is unlikely to be noticeably affected because of probable metabolic compensatory mechanisms produced by other P450 isoforms.
When irosustat phase I metabolites were incubated with different cDNA-expressed UGTs and SULTs, up to nine different glucuronides and eight sulfates were produced, respectively. Although the actual role of each phase II enzyme was not fully clarified, phase II metabolism is a secondary pathway in irosustat metabolism and can only occur after phase I transformation of the parent compound, or after formation of 667-coumarin or any of its hydroxylated metabolites. Therefore, although highly involved in the overall drug disposition process, phase II enzymes would neither influence irosustat clearance nor be the target enzymes involved in possible drug-drug interaction processes, unless the biological activity of any of the phase I metabolites was demonstrated.
To summarize, multiple irosustat metabolic pathways including desulfamoylation, primary and secondary metabolism, and phase I and phase II reactions were involved in the metabolism of irosustat in liver microsomes and hepatocytes from several species. CYP2C8, CYP2C9, CYP3A4/5, and CYP2E1 were identified as the main enzymes responsible for the primary transformation of irosustat. Although a number of recombinant phase II enzymes were tested with positive results, their actual clinical relevance could not be clarified. Glucuronide and sulfate conjugation reactions would be secondary to phase I transformation of irosustat or 667-coumarin formation, and, therefore, they should not play an important role in irosustat clearance. The in vitro studies presented in this work have also been useful in identifying potential metabolites of irosustat and in obtaining valuable information for planning and interpreting future toxicology and in vivo metabolism of irosustat.
Participated in research design: Ventura and Solà.
Conducted experiments: Ventura and Solà.
Contributed new reagents or analytic tools: Ventura and Solà.
Performed data analysis: Ventura.
Wrote or contributed to the writing of the manuscript: Ventura, Solà, Peraire, and Obach.
Other: Celma contributed to the NMR analysis interpretation.
We thank M. Victor and C. Maté for their technical assistance, T. Ali for his expert review, and the Serveis Cientifico-Técnics, University of Barcelona, for performing the NMR analysis.
This work was sponsored by the Ipsen Group.
Article, publication date, and citation information can be found at http://dmd.aspetjournals.org.
- steroid sulfatase
- high-performance liquid chromatography
- cytochrome P450
- human liver microsomes
- flavin-containing monooxygenase
- Clint, app
- apparent intrinsic clearance
- liquid chromatography
- mass spectrometry
- solid-phase extraction.
- Received January 21, 2011.
- Accepted April 4, 2011.
- Copyright © 2011 by The American Society for Pharmacology and Experimental Therapeutics