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Research ArticleArticle

In Vitro Characterization of the Human Liver Microsomal Kinetics and Reaction Phenotyping of Olanzapine Metabolism

Porntipa Korprasertthaworn, Thomas M. Polasek, Michael J. Sorich, Andrew J. McLachlan, John O. Miners, Geoffrey T. Tucker and Andrew Rowland
Drug Metabolism and Disposition November 2015, 43 (11) 1806-1814; DOI: https://doi.org/10.1124/dmd.115.064790

Abstract

Olanzapine (OLZ) is an atypical antipsychotic used in the treatment of schizophrenia and related psychoses. The metabolism of OLZ is complex and incompletely characterized. This study aimed to elucidate the enzymes and pathways involved in the metabolism of OLZ and to determine the kinetics of OLZ oxidation and glucuronidation by human liver microsomes, recombinant cytochrome P450 (rP450) enzymes, and recombinant UDP-glucuronosyltransferase (rUGT) enzymes. An ultra-performance liquid chromatography–mass spectrometry method was developed and validated to quantify OLZ, its four oxidative metabolites (N-desmethyl-OLZ, 2-hydroxymethyl-OLZ, 7-hydroxy-OLZ, and OLZ-N-oxide), and two N-glucuronides (OLZ-10-N-glucuronide and OLZ-4′-N-glucuronide). Consistent with previous reports, UGT1A4, CYP1A2, and flavin-containing monooxygenase 3 play major roles in catalyzing the formation of OLZ-10-N-glucuronide, 7-hydroxy-OLZ, and OLZ-N-oxide, respectively. In addition, a previously uncharacterized major contribution of CYP2C8 to OLZ-N-demethylation was demonstrated. The kinetics of OLZ metabolite formation (Km and Vmax) by human liver microsomes, rP450 enzymes, and rUGT enzymes were characterized in the presence of bovine serum albumin [2% (w/v)]. Consistent with the known effect of bovine serum albumin on CYP1A2, CYP2C8, and UGT1A4 activities, Km values reported here are lower than previously reported values for OLZ metabolic pathways. In addition to CYP1A2-mediated OLZ-N-demethylation, these results suggest that other P450 enzymes, particularly CYP2C8, contribute significantly to oxidative OLZ metabolism through catalysis of OLZ-N-demethylation.

Introduction

Olanzapine (OLZ) is an atypical antipsychotic indicated for the treatment of schizophrenia and related psychoses (Rosenheck et al., 2003). Although OLZ is effective for the acute management of psychosis, up to 80% of patients discontinue treatment within 5 years due to either a lack of long-term efficacy or drug-related adverse events (Schwenger et al., 2011). In this regard, plasma OLZ and metabolite (primarily N-desmethyl-OLZ) concentrations have been identified as important markers of both clinical response to OLZ and the occurrence of adverse effects such as weight gain, hyperinsulinemia, and hyperlipidemia (Perry et al., 2001; Mauri et al., 2007; Nozawa et al., 2008). Accordingly, therapeutic drug monitoring for OLZ is recommended in a number of clinical guidelines, and a therapeutic concentration range of 20–80 µg/l has been established (Baumann et al., 2004).

OLZ is cleared primarily via hepatic metabolism, with <10% of the drug excreted unchanged in urine. The metabolism of OLZ is complex, involving multiple pathways and drug-metabolizing enzyme families (Fig. 1). Consistent with their roles in the metabolism of a myriad of drugs (Miners and Mackenzie, 1991; Rowland et al., 2013; Zanger and Schwab, 2013), available evidence indicates that members of the cytochrome P450 (P450) and UDP-glucuronosyltransferase (UGT) superfamilies are extensively involved in the oxidative and conjugative metabolism of OLZ, respectively, with a minor contribution from the flavin-containing monooxygenase (FMO) enzymes (Ring et al., 1996; Linnet, 2002). The major primary metabolites of OLZ formed in humans are N-desmethyl-OLZ, OLZ-10-N-glucuronide, 2-hydroxymethyl-OLZ, and OLZ-N-oxide (Kassahun et al., 1997). Although not detected in vivo, formation of 7-hydroxy-OLZ has been reported in vitro with human liver microsomes (HLMs) as the enzyme source (Ring et al., 1996). Despite the established relationship between plasma N-desmethyl-OLZ concentration and clinical outcomes, there has been only one study that has evaluated the oxidative pathways of OLZ in a systematic manner (Ring et al., 1996). Current data suggest that CYP1A2, CYP2C9, CYP2D6, CYP2E1, CYP3A4, and FMO3 contribute to the oxidative metabolism of OLZ. CYP1A2 has been identified as the major enzyme responsible for the formation of N-desmethyl-OLZ and 7-hydroxy-OLZ, whereas CYP2D6 has been identified as the major enzyme responsible for 2-hydroxymethyl-OLZ formation (Ring et al., 1996). However, the roles of additional hepatic drug-metabolizing P450 enzymes, particularly CYP2B6, CYP2C8, and CYP2C19, in the oxidative metabolism of OLZ remain poorly characterized. Notably, when considering data presented by Ring et al. (1996), the identity of P450 enzymes involved in OLZ oxidative metabolism was primarily assigned using HLMs as the enzyme source and on the basis of surrogate markers (i.e., correlations with immunoquantified enzyme levels and enzyme-selective catalytic activities).

Fig. 1.
Fig. 1.

Schematic of OLZ metabolism investigated in vitro by this study. A previous study identified OLZ-10-N-glucuronide as a major metabolite of OLZ in urine (Kassahun et al., 1997). Gluc, glucuronide.

Given the purported major role of CYP1A2 in the formation of N-desmethyl-OLZ, a number of studies have attempted to correlate CYP1A2 phenotype and genotype with plasma OLZ and metabolite concentrations (Hägg et al., 2001; Carrillo et al., 2003; Shirley et al., 2003; Nozawa et al., 2008; Ghotbi et al., 2010; Laika et al., 2010; Skogh et al., 2011; Söderberg et al., 2013a). Although studies have demonstrated moderate correlations with end points such as OLZ oral clearance (Carrillo et al., 2003; Shirley et al., 2003), these reports have consistently demonstrated a lack of correlation between CYP1A2 phenotype or genotype and plasma OLZ metabolite concentrations. Taken together, these data support the hypothesis that other P450 enzymes play a clinically important role in the metabolism of OLZ, particularly the formation of N-desmethyl-OLZ. Given the importance of N-desmethyl-OLZ concentrations as a marker of clinical outcomes, clarification of the identity of the P450 enzymes involved in this OLZ metabolic pathway is warranted.

This study sought to reevaluate the metabolic pathways involved in OLZ clearance using ultra-performance liquid chromatography (UPLC)–mass spectrometry (MS) for metabolite quantification and accounting for recently identified potential confounding factors associated with in vitro drug kinetic studies, notably nonspecific binding of substrate (McLure et al., 2000; Rowland et al., 2007, 2008a; Wattanachai et al., 2011, 2012) and inhibition of selected P450 and UGT enzymes by long-chain unsaturated fatty acids released from membranes during the course of an incubation (Tsoutsikos et al., 2004; Yao et al., 2006; Rowland et al., 2008 a,b). Data confirmed the role of FMO3 in the formation of OLZ-N-oxide but excluded a significant role of this enzyme in the formation of other OLZ metabolites. Kinetic analyses were performed to characterize the P450-catalyzed OLZ oxidative pathways and OLZ glucuronidation pathways using both HLMs and recombinant enzymes. Kinetic data demonstrate that in addition to CYP1A2, other P450 enzymes, most notably CYP2C8, catalyze the formation of N-desmethyl-OLZ at a comparable rate.

Materials and Methods

Materials.

2-Hydroxymethyl-OLZ, 2-hydroxymethyl-OLZ-d3, N-desmethyl-OLZ, N-desmethyl-OLZ-d8, OLZ, OLZ-d3, OLZ-d3-N-oxide, and OLZ-N-oxide were purchased from Toronto Research Chemicals (North York, ON, Canada); ticlopidine was from LKT Laboratories (St. Paul, MN); montelukast was from Cayman Chemical (Ann Arbor, MI); and bovine serum albumin (BSA) (essentially fatty acid free), CYP3cide, d-glucose-6-phosphate disodium salt hydrate, diethyldithiocarbamic acid, furafylline, glucose-6-phosphate dehydrogenase, hecogenin, NADP, quinidine, sulfaphenazole, ticlopidine, tranylcypromine, troleandomycin, and UDP-diphosphoglucuronic acid (UDP-GlcUA) were from Sigma-Aldrich (St. Louis, MO). Human UGT2B4, UGT2B7, UGT2B10, UGT2B15, UGT2B17, and FMO3 Supersomes were purchased from BD Biosciences (Woburn, MA). All other reagents and solvents were of analytical reagent grade or higher.

Preparation of HLMs and Expression of Recombinant P450 and UGT Enzymes.

Human livers (H7, H10, H12, H13, and H40) were obtained from the human liver bank of the Flinders University School of Medicine Department of Clinical Pharmacology. Approval was obtained from the Flinders Medical Centre Research Ethics Committee for the use of human liver tissue in xenobiotic metabolism studies. Microsomes were prepared by differential centrifugation, as described by Bowalgaha et al. (2005).

Recombinant CYP1A1, CYP1A2, CYP2B6, CYP2C8, CYP2C9, CYP2E1, CYP3A4, and NADPH P450 oxidoreductase were coexpressed in Escherichia coli according to the general procedure of Boye et al. (2004). The P450/NADPH P450 oxidoreductase ratios were close to unity. cDNAs encoding UGT1A1, UGT1A3, UGT1A4, UGT1A6, UGT1A7, UGT1A8, UGT1A9, and UGT1A10 were stably expressed in the human embryonic kidney 293T (HEK293T) cell line, as described previously (Uchaipichat et al., 2004). Activity of these recombinant UDP-glucuronosyltransferases (rUGTs) (except UGT1A4) was confirmed using the nonselective substrate 4-methylumbelliferone (Uchaipichat et al., 2004). The activity of UGT1A4 was confirmed using lamotrigine as substrate (Rowland et al., 2006).

OLZ Oxidative Metabolism Assay.

Incubations, in a total volume of 200 µl, contained phosphate buffer (1 mM, pH 7.4), HLMs (0.5 mg/ml), and OLZ (5–400 μM). After a 5-minute preincubation, reactions were initiated by the addition of an NADPH-generating system (1 mM NADP, 10 mM glucose-6-phosphate, 2 IU/ml glucose-6-phosphate dehydrogenase, and 5 mM MgCl2). Incubations were performed at 37°C in a shaking water bath for 60 minutes. Reactions were terminated by the addition of ice-cold methanol containing 4% acetic acid (200 µl) and cooling on ice. Samples were subsequently centrifuged at 5000 × g for 10 minutes at 10°C, and a 150-µl aliquot of the resultant supernatant fraction was diluted in 350 µl mobile phase containing deuterated internal standards (OLZ-d3, N-desmethyl-OLZ-d8, 2-hydroxymethyl-OLZ-d3, and OLZ-d3-N-oxide; 25 nM). A 7.5-µl aliquot was injected directly onto the UPLC column.

For reactions performed using recombinant cytochrome P450 (rP450) enzymes, incubation mixtures contained P450 protein expressed in E. coli membranes in place of HLM protein. Screening experiments were initially performed to assess the capacity of individually expressed rP450 to catalyze the oxidative metabolism of OLZ. These experiments were performed at an rP450 concentration of 25 pmol P450/ml and OLZ concentrations of 50 and 300 µM, with samples incubated for 2 hours at 37°C in a shaking water bath. Incubation conditions for subsequent kinetic studies with rP450 were as described above for HLMs with the following amendments: rP450 content (10 pmol P450/ml), OLZ (5–1000 µM), and incubation time (60 and 10 minutes for rCYP1A2 and rCYP2C8, respectively). BSA [2% (w/v)] was included in all experiments (HLM and rP450) performed to assess the kinetics of CYP1A2- and CYP2C8-catalyzed reactions. Control samples were incubated in the absence of an NADPH-generating system and/or protein. Rates of product formation were optimized for linearity with respect to protein concentration and incubation time.

OLZ Glucuronidation Assay.

Incubations, in a total volume of 200 µl, contained phosphate buffer (1 mM, pH 7.4), MgCl2 (4 mM), HLMs (0.25 mg/ml), and OLZ (50–1500 µM). HLMs were fully activated by the addition of the pore-forming polypeptide alamethicin (50 µg/mg protein) with incubation on ice for 30 minutes (Boase and Miners, 2002). After a 5-minute preincubation, reactions were initiated by the addition of UDP-GlcUA (5 mM). Incubations were performed at 37°C in a shaking water bath for 45 minutes. Reactions were terminated by the addition of ice-cold methanol containing 4% acetic acid (200 µl) and cooling on ice. Samples were subsequently centrifuged at 5000 × g for 10 minutes at 10°C, and a 150-µl aliquot of the resultant supernatant fraction was diluted in 350 µl mobile phase containing a deuterated OLZ-d3 internal standard (25 nM). A 7.5-µl aliquot was injected directly onto the UPLC column.

For reactions performed using human rUGTs, incubation mixtures contained UGT1A enzymes expressed in HEK293T cells (as lysate) or UGT2B enzymes expressed in Supersomes. Screening experiments were initially performed to assess the capacity of individually expressed rUGT to glucuronidate OLZ. These experiments were performed at an rUGT concentration of 1 mg/ml and OLZ concentrations of 350 and 1500 µM, with samples incubated for 2 hours at 37°C in a shaking water bath. Incubation conditions for subsequent kinetic studies with rUGT were as described above for HLMs with the following amendments: rUGT content (0.5 mg/ml) and incubation time 30 minutes. For the experiments performed in the presence of BSA, reactions were terminated by the addition of ice-cold methanol containing 4% acetic acid (400 µl) and cooling on ice. Control samples were incubated in the absence of UDP-GlcUA and/or protein. The rate of product formation was optimized for linearity with respect to protein concentration and incubation time.

Inhibition Experiments.

The effects of P450 enzyme–selective inhibitors on the formation of the oxidative metabolites of OLZ were investigated using pooled HLMs as the enzyme source. Furafylline (10 μM), tranylcypromine (1 μM), ticlopidine (3 μM), montelukast (0.2 μM), sulfaphenazole (2.5 μM), quinidine (1 μM), diethyldithiocarbamic acid (10 μM), and CYP3cide (1.5 μM) were used as selective inhibitors of CYP1A2, CYP2A6, CYP2B6/CYP2C19, CYP2C8, CYP2C9, CYP2D6, CYP2E1, and CYP3A4, respectively, whereas hecogenin (10 μM) was used as a selective UGT1A4 inhibitor. Pooled HLMs (0.5 mg/ml) were incubated with 50 μM OLZ in the absence and presence of the selective inhibitors. When required, inhibitors were dissolved in either 50% acetonitrile (CYP3cide) or methanol (furafylline, montelukast, and sulfaphenazole) and added to incubations such that the final solvent concentration was either 0.5% or 1% (w/v). Control samples were prepared without inhibitors but with an equivalent concentration of solvent. Since furafylline, ticlopidine, diethyldithiocarbamic acid, and CYP3cide are mechanism-based inhibitors, pooled HLM proteins were preincubated with these compounds in the presence of the NADPH-generating system for 30 minutes prior to the addition of OLZ. Reaction mixtures were incubated at 37°C for 1 hour in a shaking water bath and then terminated and further prepared for analysis as described in the oxidative metabolism assay. The percent inhibition of metabolite formation by enzyme-specific inhibitors was calculated by comparing activity with control activity (without inhibitor) (n = 4).

Separation of Olanzapine Metabolites.

Chromatographic separations were performed on a Waters Acquity BEH C18 analytical column (100 mm × 2.1 mm, 1.7 µm; Waters Corporation, Milford, MA) using a Waters Acquity UPLC system. The column temperature was maintained at 40°C, whereas the sample compartment was maintained at 15°C. Analytes were separated by gradient elution at a flow rate of 0.2 ml/min. Initial conditions were 87% ammonium formate (10 mM, pH 3.0; mobile phase A) and 13% acetonitrile (mobile phase B). The proportion of mobile phase B was increased linearly to 70% over 7 minutes and held for 0.5 minutes. The total run time, including reconditioning of the column to initial conditions, was 9 minutes. Retention times of 2-hydroxymethyl-OLZ, 7-hydroxy-OLZ, N-desmethyl-OLZ, OLZ-N-oxide, OLZ-10-N-glucuronide (two peaks), and OLZ-4′-N-glucuronide under these conditions were 1.7, 2.2, 3.4, 3.8, 1.4, 1.8, and 3.5 minutes, respectively (Fig. 2).

Fig. 2.
Fig. 2.

Extracted ion chromatograms for OLZ oxidative and glucuronide metabolites formed by HLMs. The oxidation assay was performed using HLMs (0.5 mg/ml) and 50 μM OLZ, whereas the glucuronidation assay contained HLMs (1 mg/ml) and 1 mM OLZ. Both reactions were performed at 37°C for 1 hour. (A) Peaks at m/z 489.18. (B) Peak at m/z 299.14. (C) Peaks at m/z 329.15. Peak 1, OLZ-10-N-glucuronide isomer 1; peak 2, OLZ-10-N-glucuronide isomer 2; peak 3, OLZ-4′-N-glucuronide; peak 4, N-desmethyl-OLZ; peak 5, 2-hydroxymethyl-OLZ; peak 6, 7-hydroxy-OLZ; and peak 7, OLZ-N-oxide.

Quantification of Olanzapine Metabolites.

Column elutant was monitored by MS, performed on a Waters Q-ToF Premier quadrupole, orthogonal acceleration time-of-flight tandem mass spectrometer operated in positive ion mode with electrospray ionization. Instrument acquisition settings are described in Table 1. Time-of-flight data were collected in wide-pass (MS) mode, with the resolving quadrupole acquiring data between mass-to-charge (m/z) ratios of 100 and 650. Selected ion data were extracted at precursor m/z ratios of 299.14 (N-desmethyl-OLZ), 329.15 (2-hydroxymethyl-OLZ, 7-hydroxy-OLZ, and OLZ-N-oxide), and 489.18 (OLZ-10-N-glucuronide and OLZ-4′-N-glucuronide). Resulting pseudo–multiple reaction monitoring spectra were analyzed using Waters QuanLynx software (Waters Corporation, Sydney, Australia). Quantification of analytes in incubation samples was accomplished by comparison of peak areas with those of authentic standards for each metabolite (25–500 nM), with the exception of 7-hydroxy-OLZ and OLZ-10-N-glucuronide, which were measured by comparing the peaks with those of the respective external standards 2-hydroxymethyl-OLZ and OLZ.

View this table:
TABLE 1

Mass spectrometer instrument settings

Binding of OLZ to BSA, HLMs, rP450 Enzymes, and rUGTs.

Binding of OLZ to HLMs, rP450 enzymes, rUGTs, and BSA in each enzyme source was measured by equilibrium dialysis according to the method of McLure et al. (2000). The experiments were performed using a Dianorm equilibrium dialysis apparatus consisting of Teflon dialysis cells (capacity of 1.2 ml/side) separated by a Sigma-Aldrich dialysis membrane (molecular mass cut off of 12 kDa). One side of the dialysis cell was loaded with 1 ml OLZ (10–100 µM) in phosphate buffer (1 mM, pH 7.4). The other compartment was loaded with 1 ml HLMs, rP450 enzymes, rUGTs, or a combination of BSA with each enzyme source in phosphate buffer (1 mM, pH 7.4). The dialysis cell assembly was immersed in a water bath maintained at 37°C and rotated at 12 rpm for 4 hours. Control experiments were also performed with phosphate buffer or BSA on both sides of the dialysis cells at low and high concentrations of OLZ to ensure that equilibrium was attained. A 200-µl aliquot was collected from each compartment, treated with ice-cold methanol containing 4% acetic acid (400 µl), and cooled on ice. Samples were subsequently centrifuged at 5000 × g for 10 minutes at 10°C, and a 5-µl aliquot of the supernatant fraction was analyzed by high-performance liquid chromatography.

Quantification of OLZ Binding.

Separation of OLZ was achieved on an Agilent Zorbax Eclipse XDB-C8 column (4.6 × 150 mm, 5 µm; Agilent Technologies, Santa Clara, CA) using a mobile phase comprising 90% 10 mM ammonium formate (pH 3.0) and 10% acetonitrile, at a flow rate of 1 ml/min. The column eluent was monitored at 257 nm. The retention time of OLZ was 3.1 minutes. OLZ concentrations in dialysis samples were determined by comparison of peak areas with those of an OLZ standard (concentration range of 10–100 µM).

Kinetic Analysis.

Data points represent the mean of duplicate (<10% variance) or quadruplicate measurements. Kinetic constants for OLZ oxidative metabolism and glucuronidation were obtained by fitting Michaelis–Menten, substrate inhibition, or Hill equations (eqs. 1–3) to the experimental data using EnzFitter software (version 2.0; Biosoft, Cambridge, UK). Goodness of fit was assessed by comparison of the F statistic, coefficient of determination (r2), S.E. of the parameter fit, and 95% confidence intervals.

The Michaelis–Menten equation is as follows:Embedded Image(1)where v is the rate of reaction, Vmax is the maximum velocity, Km is the Michaelis–Menten constant (substrate concentration at 0.5 Vmax), and [S] is the substrate concentration.

The Hill equation, which describes positive or negative cooperative kinetics, is as follows:Embedded Image(2)where S50 is the substrate concentration resulting in 50% of Vmax (analogous to Km in the previous equation) and n is the Hill coefficient.

The substrate inhibition equation is as follows:Embedded Image(3)where Ksi is the constant describing the substrate inhibition reaction.

Results

Binding of OLZ to HLMs, rP450 Enzymes, rUGTs, and BSA.

It has been reported that the in vitro activities of certain P450 (particularly CYP1A2, CYP2C8, and CYP2C9) and UGT (particularly UGT1A9 and UGT2B7) enzymes are inhibited by long-chain unsaturated fatty acids released from the microsomal membrane during incubations. This inhibition largely manifests as an increase in Km values (with occasional effects on Vmax) and hence intrinsic clearance for the added substrate, which results in an underprediction of the in vitro intrinsic clearance of the substrate (Tsoutsikos et al., 2004; Rowland et al., 2007, 2008a,b; Wattanachai et al., 2011, 2012). In addition, it has been shown that the inhibitory effect of fatty acids can be overcome by sequestration with BSA (Rowland et al., 2007, 2008a; Wattanachai et al., 2012). Thus, BSA [2% (w/v)] was included in incubations for the kinetic experiments performed in this study. However, since the effect of BSA on the kinetics of UGT1A4-catalyzed glucuronidation is less well characterized, the kinetics of OLZ-10-N-glucuronidation were determined in the absence and presence of albumin.

Accordingly, the binding of OLZ to HLMs, E. coli membrane (used to express rP450 enzymes), HEK293T cell lysate (used to express rUGT enzymes), and BSA [2% (w/v)] was determined by equilibrium dialysis. The extent of OLZ binding was calculated as the concentration of drug in the buffer compartment divided by the concentration of drug in the protein compartment and expressed as a fraction unbound in the incubation mixture. The binding of OLZ to HLMs, E. coli membrane, and HEK293T cell lysate was negligible (<15%) across the OLZ concentration range investigated (10–100 µM). However, OLZ did bind significantly to mixtures of HLMs and BSA with a mean ± S.D. fraction unbound in the incubation mixture value of 0.69 ± 0.01. Thus, the concentration of OLZ present in reaction mixtures was corrected for binding in the calculation of kinetic parameters for experiments performed in the presence of BSA.

Identification of P450 Enzymes Mediating the Oxidative Metabolism of OLZ.

The capacity of recombinantly expressed CYP1A1, CYP1A2, CYP2B6, CYP2C8, CYP2C9, CYP2E1, and CYP3A4 to form oxidative metabolites of OLZ was assessed at OLZ concentrations of 50 and 300 μM. At the higher concentration of OLZ (Fig. 3), N-desmethyl- OLZ and 2-hydroxymethyl-OLZ were formed to the greatest extent by rCYP2C8 (730 and 60 pmol/min per nmol P450, respectively), whereas 7-hydroxy-OLZ was formed mainly by rCYP1A2 (130 pmol/min per nmol P450). In addition, rCYP1A2, rCYP2B6, rCYP2C9, and rCYP3A4 formed 2-hydroxymethyl-OLZ, albeit to a lesser extent (<50 pmol/min per nmol P450). OLZ-N-oxide was primarily formed by FMO3 (483 pmol/min per mg; data not shown) with minor contributions by CYP2C8 and CYP3A4 (200 and 120 pmol/min per nmol P450, respectively).

Fig. 3.
Fig. 3.

In vitro formation of OLZ oxidative metabolites by E. coli–expressed P450. Formation of N-desmethyl-OLZ, 2-hydroxymethyl-OLZ, 7-hydroxy-OLZ, and OLZ-N-oxide by rP450 enzymes (25 pmol P450/ml) incubated with 300 μM OLZ at 37°C for 2 hours. Data are shown as the means of duplicate determinations from a single experiment.

Kinetics of OLZ Oxidative Metabolism by HLMs and rP450 Enzymes.

Four oxidative metabolites formed by incubations of OLZ with HLMs were identified by UPLC-MS. These were 2-hydroxymethyl-OLZ, 7-hydroxy-OLZ, N-desmethyl-OLZ, and OLZ-N-oxide (retention times of 1.7, 2.2, 3.4, and 3.8 minutes, respectively; Fig. 2, peaks 4–7). The kinetics of OLZ-N-demethylation and OLZ-7-hydroxylation were best described by the Michaelis–Menten equation, whereas OLZ-2-hydroxylation exhibited negative cooperativity (n ranging from 0.6 to 0.9). OLZ-N-oxide formation followed Michaelis–Menten kinetics in four of five livers; in the remaining liver (H12), the formation of this metabolite was best described by the Hill equation (positive cooperativity, n = 1.5). Kinetic parameters for oxidative OLZ metabolism by HLMs are summarized in Table 2 and Eadie–Hofstee plots are shown in Fig. 4, A–C.

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TABLE 2

Derived kinetic parameters for OLZ oxidative pathways by HLMs in the presence of BSA [2% (w/v)]

Data are the mean ± S.D. for microsomes from five individual livers.

Fig. 4.
Fig. 4.

Representative Eadie–Hofstee plots for OLZ oxidative metabolism. (A) OLZ-N-demethylation by HLM_H7. (B) OLZ-2-hydroxylation by HLM_H7. (C) OLZ-7-hydroxylation by HLM_H7. (D) OLZ-N-demethylation by rCYP1A2. (E) OLZ-N-demethylation by rCYP2C8. (F) OLZ-7-hydroxylation by rCYP1A2, in the presence of BSA. Points are experimentally determined values (means of duplicate and quadruplicate measurements at each concentration for HLM_H7 and rP450 enzymes, respectively), whereas curves are from model fitting.

Kinetic parameters for the formation of the oxidative metabolites of OLZ by rP450 enzymes over the substrate concentration range of 5–1000 μM are shown in Table 3, and the Eadie–Hofstee plots are shown in Fig. 4, D–F. OLZ-N-demethylation by rCYP1A2 was best described by the Michaelis–Menten equation, whereas rCYP1A2-catalyzed OLZ-7-hydroxylation was best described by the Hill equation (negative cooperativity, n = 0.7). Kinetic data for OLZ-N-demethylation by rCYP2C8 were best described by the Michaelis–Menten equation.

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TABLE 3

Derived kinetic parameters for OLZ oxidative metabolism by rP450 enzymes in the presence of BSA [2% (w/v)]

Data are mean ± S.D. from quadruplicate experiments.

Identification of UGT Enzymes Mediating the Glucuronidation of OLZ.

The capacity of human UGT1A1, UGT1A3, UGT1A4, UGT1A6, UGT1A7, UGT1A8, UGT1A9, UGT1A10, UGT2B4, UGT2B7, UGT2B10, UGT2B15, and UGT2B17 to catalyze the glucuronidation of OLZ was evaluated at substrate concentrations of 350 and 1500 µM. Only UGT1A4 converted OLZ to OLZ-10-N-glucuronide and OLZ-4′-N-glucuronide (Fig. 2; peaks 1–3). This result differs from a previous in vitro study, which demonstrated that when overexpressed in HEK293 cells at an unusually high final incubation concentration of 30 mg/ml, UGT2B10 glucuronidated OLZ, with intrinsic clearances for OLZ-10-N-glucuronide and OLZ-4′-N-glucuronide of 5 and 7 nl/min per mg, respectively (Erickson-Ridout et al., 2011). Screening experiments in this study using Supersome-expressed UGT2B10 (1 mg/ml; 2-hour incubation) demonstrated no activity. Thus, subsequent kinetic studies were performed only with rUGT1A4.

OLZ Glucuronidation by HLMs and rUGT1A4.

Previous studies characterizing OLZ metabolism in vitro have demonstrated the formation of three compounds with m/z ratios consistent with that of OLZ-N-glucuronides (489.18). These compounds have been identified as two isomers of OLZ-10-N-glucuronide (Fig. 2; peaks 1 and 2) and OLZ-4′-N-glucuronide (Fig. 2; peak 3) (Kassahun et al., 1998; Erickson-Ridout et al., 2011). In this study, limited formation of the OLZ-4′-N-glucuronide (<5% that of OLZ-10-N-glucuronide) was only detected in incubations performed at an increased HLM concentration of 1 mg/ml (a positive control for screening of UGT activities). As such, no kinetic analysis was performed to characterize the formation of this metabolite. Because of the lack of commercial availability of an authentic OLZ-10-N-glucuronide standard, chromatographic peaks were assigned to OLZ-10-N-glucuronide on the basis of mass spectral characteristics. OLZ-10-N-glucuronide was characterized by [M-H+] peaks at m/z 313.16 [after the loss of the glucuronic moiety; molecular mass = 176 g/mol] and m/z 432.15 [after the loss of a methyl piperazine (CH2 = CH-NH-CH3) moiety; molecular mass = 57 g/mol]. The fragmentation pattern observed for OLZ-10-N-glucuronide was consistent with previous reports for this compound (Kassahun et al., 1998; Erickson-Ridout et al., 2011). Representative kinetic plots for OLZ-10-N-glucuronide formation by HLMs (liver H40), in the absence and presence of BSA, are shown in Fig. 5A and mean kinetic parameters are summarized in Table 4. OLZ-10-N-glucuronidation by HLMs in the absence and presence of BSA was best described by the substrate inhibition equation. In the absence of BSA, the mean ± S.D. Km and Vmax values for OLZ 10-N-glucuronidation by HLMs were 634 ± 287 µM and 443 ± 176 pmol/min per mg, respectively. The addition of BSA did not alter kinetic parameters for HLM-catalyzed OLZ 10-N-glucuronidation (Km 511 ± 245 µM and Vmax 484 ± 170 pmol/min per mg), although substantial interliver variability was observed and may have masked any small effect. For experiments performed with rUGT1A4, where variability is limited to experimental error, the addition of BSA decreased the Km for OLZ 10-N-glucuronidation from 341 µM to 183 µM, with minor effects on Vmax and Ksi.

Fig. 5.
Fig. 5.

Eadie–Hofstee plots for OLZ 10-N-glucuronidation. (A and B) OLZ 10-N-glucuronidation by HLM_H40 (A) and OLZ 10-N-glucuronidation by rUGT1A4 (B), in the presence (filled triangles) and absence (open diamonds) of BSA. Points are experimentally determined values (means of quadruplicate measurements at each concentration); lines are model fits.

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TABLE 4

Derived kinetic parameters for OLZ-10-N-glucuronidation by HLMs and rUGT1A4 in the absence and presence of albumin

For HLMs, data are mean ± S.D. from quadruplicate measurements for HLMs from each of five individual livers. For UGT1A4, data are mean ± S.D. from quadruplicate measurements.

As observed in previous studies and incubations performed with HLMs, rUGT1A4 generated three chromatographic peaks identified as two isomers of OLZ-10-N-glucuronide and OLZ-4′-N-glucuronide. However, the low rate of OLZ-4′-N-glucuronide formation precluded kinetic analysis. As found with HLMs, experimental data for rUGT1A4 were best described by the substrate inhibition equation (Table 3). An Eadie–Hofstee plot for rCYP1A4-catalyzed OLZ glucuronidation is shown in Fig. 5B. Km and Vmax values for OLZ-10-N-glucuronidation by rUGT1A4 were lower than those for OLZ-10-N-glucuronidation by HLMs by 36%–64%.

Inhibition Studies in Pooled HLMs.

The effects of CYP3cide, diethyldithiocarbamic acid, furafylline, hecogenin, montelukast, quinidine, sulfaphenazole, ticlopidine, and tranylcypromine on the formation of OLZ oxidation products by pooled HLMs were determined. The formation of N-desmethyl-OLZ was inhibited by 56%, 27%, and 35% by furafylline, montelukast, and CYP3cide, respectively. The formation of 7-hydroxy-OLZ was markedly inhibited (84%) by furafylline, whereas the effects of other P450 enzyme–selective inhibitors were minor (<20% inhibition), except for CYP3cide, which inhibited the formation of 7-hydroxy-OLZ by 24%.

The formation of 2-hydroxymethyl-OLZ was inhibited by furafylline, tranylcypromine, montelukast, and CYP3cide by approximately 20%–30%. CYP3cide also inhibited OLZ-N-oxide formation to a moderate extent (27%), whereas other P450 enzyme–selective inhibitors had no effect on the formation of this metabolite. Tranylcypromine, ticlopidine, sulfaphenazole, diethyldithiocarbamate, and quinidine had only a minor (<20% inhibition) or no effect on the formation of the OLZ oxidative metabolites. Human liver microsomal OLZ-10-N-glucuronide formation was inhibited >80% by hecogenin.

Discussion

This study reports, for the first time, the major involvement of additional P450 enzymes, most notably CYP2C8, in OLZ-N-demethylation. This finding is of considerable clinical importance, given the strong correlations between plasma N-desmethyl-OLZ concentration, clinical response to OLZ, and the occurrence of adverse effects (Hägg et al., 2001; Carrillo et al., 2003; Shirley et al., 2003; Nozawa et al., 2008; Ghotbi et al., 2010; Laika et al., 2010; Skogh et al., 2011; Söderberg et al., 2013a). Variability in CYP2C8 protein expression and catalytic activity is well documented (Sonnichsen et al., 1995; Baldwin et al., 1999; Totah and Rettie, 2005; Naraharisetti et al., 2010). It is likely that the contribution of CYP2C8 and other P450 enzymes to N-desmethyl-OLZ formation may explain the poor correlation between CYP1A2 phenotype or genotype and plasma concentration of this metabolite (Hägg et al., 2001; Carrillo et al., 2003; Shirley et al., 2003). Indeed, given the comparable involvement of multiple enzymes in the formation of this metabolite, it is unlikely that consideration of any single metabolic pathway will provide a robust predictive marker of clinical outcomes with OLZ, and that a more complex model accounting for multiple covariates is required.

The sole previous study considering the P450 enzymes responsible for OLZ metabolism demonstrated that OLZ-N-demethylation is primarily catalyzed by CYP1A2, with minor contributions from CYP3A4 and CYP2D6 (Ring et al., 1996). FMO3 was reported to be the primary enzyme involved in the formation of OLZ-N-oxide, with minor contributions from CYP1A2, CYP2D6, CYP2C9, CYP2E1, and CYP3A4 (Ring et al., 1996). However, the initial reaction phenotyping for OLZ oxidative metabolism was assigned on the basis of correlations of the rate of OLZ metabolite formation with immunoquantified P450 (CYP1A2, CYP2A6, CYP2C9, CYP2C19, CYP2D6, CYP2E1, and CYP3A) and FMO3 levels in HLMs and (where possible) with the catalytic activity of form-selective probe substrates for these enzymes. Notably, confirmation of enzyme involvement using cDNA-expressed enzymes was only reported for a limited panel of enzymes (CYP1A2, CYP2A6, CYP2C9, CYP2D6, CYP2E1, and CYP3A4). Ring et al. (1996) did postulate the possible involvement of CYP2C8 in the formation of OLZ metabolites on the basis of correlation of the rates of OLZ metabolite formation with the immunoquantified level of this enzyme. However, the significance of the role of CYP2C8 in terms of OLZ metabolite formation was excluded on the basis of poor correlations; correlation coefficients (r) ranged from 0.19 for 2-hydroxymethyl-OLZ formation to 0.40 for N-desmethyl-OLZ formation. The approach used to assess the immunoquantified level of P450 was not reported by Ring et al. (1996). However, given the significant similarities in amino acid sequence within the CYP2C family (CYP2C8, CYP2C9, and CYP2C19), it is plausible that antibody cross-reactivity may have confounded this result. In addition, it should be noted that although correlations of the rate of N-desmethyl-OLZ formation with catalytic activities for CYP1A2-mediated ethoxyresorufin-O-deethylase and caffeine-3-demethylase (form-selective catalytic activities) were significant (r = 0.64 and 0.66, respectively), correlation of the rate of N-desmethyl-OLZ formation with the immunoquantified level of microsomal CYP1A2 (r = 0.35) was not significant and in fact was poorer than the correlation observed for the immunoquantified level of CYP2C8. Because of the lack of a selective CYP2C8 probe at the time of the previous study, correlation of the rate of OLZ metabolite formation with a form-selective catalytic activity for CYP2C8 was not undertaken (Ring et al., 1996). In our study, OLZ-N-demethylations by rCYP2C8 and rCYP1A2 were the major oxidative pathways; scaling of kinetic data for relative liver P450 abundance demonstrate that these enzymes account for 28% and 26% of the total in vitro metabolism of OLZ, respectively. The role of FMO3 in the oxidative metabolism of OLZ (primarily OLZ-N-oxide formation) was confirmed. However, given the limited role of OLZ-N-oxide in determining OLZ plasma concentration and clinical outcomes (Söderberg et al., 2013b), as well as the recent in vitro characterization of this pathway (Cashman et al., 2008), subsequent kinetic studies were not performed.

Previous in vitro studies found that OLZ-10-N-glucuronide is formed by both UGT1A4 (Linnet, 2002) and UGT2B10 (Erickson-Ridout et al., 2011); the latter enzyme was also reported to be primarily responsible for the formation of OLZ-4′-N-glucuronide (a minor metabolite). However, the catalytic activity of rUGT2B10 was approximately 28-fold lower than that of rUGT1A4. Given the comparable levels of mRNA expression of UGT1A4 and UGT2B10 in human liver (1110 and 1004 copies per 109 copies of 18S rRNA, respectively) (Court et al., 2012), it is likely that the activity of UGT1A4 is more important in determining the formation of OLZ-N-glucuronides in vivo. Consistent with this finding, UGT1A4 was observed to be the major enzyme involved in OLZ-10-N-glucuronide and OLZ-4′-N-glucuronide formation. These data are similarly consistent with the report of Kato et al. (2013), where despite referring to OLZ as a UGT2B10 substrate, the observed activity for UGT2B10 was <5% that of UGT1A4 and was too low to facilitate kinetic analysis. In our study, the low rate of OLZ-4′-N-glucuronide formation by rUGT1A4 similarly precluded kinetic analysis of this pathway; this observation is also consistent with the previously reported low intrinsic clearance for UGT1A4-mediated OLZ-4′-N-glucuronidation of 10 ± 4 nl/min per μg (Erickson-Ridout et al., 2011). In contrast with the previous study, UGT2B10 did not catalyze OLZ-N-glucuronidation. Notably, the previously reported capacity of UGT2B10 to catalyze OLZ-N-glucuronidation was only observed in an experiment performed at an unusually high recombinant enzyme concentration of 30 mg/ml (Erickson-Ridout et al., 2011). Consistent with the major role of UGT1A4 in the formation of OLZ-10-N-glucuronides, inhibition experiments with hecogenin demonstrated >80% inhibition of OLZ-10-N-glucuronide formation.

It has been demonstrated that endogenous long-chain unsaturated fatty acids, such as arachidonic acid, linoleic acid, and oleic acid, are released from biologic membranes during in vitro incubations (Rowland et al., 2008b) and can inhibit the catalytic activities of multiple P450 and UGT enzymes, including CYP1A2, CYP2C8, and UGT1A4 (Rowland et al., 2006; Wattanachai et al., 2011, 2012). This artifact of in vitro kinetic experiments typically manifests as a decrease in “apparent” enzyme affinity (increase in Km) for the added substrate and subsequent underprediction of enzyme activity for the affected pathway. Inclusion of BSA in incubation samples restores the catalytic activity of affected enzymes by sequestering these fatty acids, thereby preventing their interaction with drug-metabolizing enzymes (Rowland et al., 2008a). A concentration effect relationship for albumin has been demonstrated; an added BSA concentration of 0.5% (5 mg/ml) causes a consistent >80% maximal decrease in Km, whereas the addition of 2% BSA (20 mg/ml) has been demonstrated to cause a maximal decrease in Km across multiple drug-metabolizing enzymes. Notably, addition of BSA at a concentration <0.5% (most commonly 0.1%) has been associated with substantial variability in effect. In this study, HLMs, E.coli membranes expressing rP450 enzymes and HEK293T cells expressing rUGT were used as enzyme sources in kinetic experiments. Given the major roles of CYP1A2, CYP2C8, and UGT1A4 in the formation of OLZ metabolites, and limited binding of OLZ to BSA, kinetic parameters were determined in the presence of BSA (2%). Consistent with the effect of BSA the activity of these enzymes, the kinetics of OLZ metabolism demonstrated greater affinity (lower Km) compared with previous findings (Ring et al., 1996). Because the “albumin effect” has been demonstrated for both CYP1A2 and CYP2C8 (Wattanachai et al., 2011, 2012), studies were not performed in the absence of BSA for these enzymes. By contrast, given the comparatively limited data concerning the effect of BSA on UGT1A4, studies with this enzyme were performed in the presence and absence of BSA (2%).

In conclusion, this study demonstrates that multiple P450 enzymes are involved in the metabolism of OLZ, particularly with regard to the formation of N-desmethyl-OLZ. This is a critical finding, given the importance of N-desmethyl-OLZ as a marker of clinical response and adverse effects associated with the administration of OLZ. Indeed, data presented here raise the possibility that inter- and intraindividual variability in the activities of CYP2C8 and CYP3A4 may additionally contribute to the therapeutic efficacy and tolerability of OLZ in humans.

Acknowledgments

The authors thank Dr. Benjamin Lewis for assistance with rP450 enzyme expression and Drs. Vidya Perera and Jason Roberts for contributions to study conception and design.

Authorship Contributions

Participated in research design: Korprasertthaworn, Polasek, McLachlan, Miners, Rowland.

Conducted experiments: Korprasertthaworn, Rowland.

Performed data analysis: Korprasertthaworn, Miners, Rowland.

Wrote or contributed to the writing of the manuscript: Korprasertthaworn, Polasek, Sorich, McLachlan, Miners, Tucker, Rowland.

Footnotes

    • Received April 29, 2015.
    • Accepted August 31, 2015.

  • This research was supported by Simcyp Limited (a Certera company) [Simcyp Grant and Partnership Scheme].

  • dx.doi.org/10.1124/dmd.115.064790.

Abbreviations

BSA
bovine serum albumin
FMO
flavin-containing monooxygenase
HEK293T
human embryonic kidney 293T
HLM
human liver microsome
MS
mass spectrometry
m/z
mass-to-charge
OLZ
olanzapine
P450
cytochrome P450
rP450
recombinant cytochrome P450
rUGT
recombinant UDP-glucuronosyltransferase
UDP-GlcUA
UDP-diphosphoglucuronic acid
UGT
UDP-glucuronosyltransferase
UPLC
ultra-performance liquid chromatography

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In Vitro Characterization of Olanzapine Metabolism

Porntipa Korprasertthaworn, Thomas M. Polasek, Michael J. Sorich, Andrew J. McLachlan, John O. Miners, Geoffrey T. Tucker and Andrew Rowland
Drug Metabolism and Disposition November 1, 2015, 43 (11) 1806-1814; DOI: https://doi.org/10.1124/dmd.115.064790
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