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Departments of Pharmacokinetics, Dynamics & Metabolism (H.-Z.B.,W.F.P., E.Y.W., B.V.S.) and Clinical Research (S.R.R., M.A.A.), Pfizer Global Research and Development, San Diego, California
(Received January 9, 2004; accepted April 6, 2004)
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Abstract |
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, 2002; Lalezari et al., 2003). Highly active antiretroviral therapy that combines at least three anti-HIV drugs has become the standard care of HIV disease, leading to dramatic falls in mortality and morbidity associated with acquired immune deficiency syndrome (Mocroft et al., 1998; Palella et al., 1998). However, these regimens become increasingly ineffective with time due to the high rates of replication, mutation, and selection of HIV (Katzenstein et al., 1992; Ho et al., 1995; Wei et al., 1995). To effectively treat antiretroviral-experienced patients, new drugs with activity against the emerging drug-resistant viruses are required.
Capravirine (AG1549 or S-1153; Fig. 1) represents a new NNRTI agent (De Clercq, 2001, 2002; Fujiwara et al., 1998, 1999; Ohkawa et al., 1998; Ren et al., 2000). Capravirine inhibits replication of HIV-1 strains that are resistant to nucleoside/nucleotide reverse transcriptase inhibitors and other NNRTIs, and is more potent than nevirapine and delavirdine of this class (Fujiwara et al., 1998). In particular, capravirine is active against the single K103N mutation of RT that renders HIV-1 resistant to each of the currently marketed NNRTIs (De Clercq, 2001) and requires at least two RT mutations to become resistant (Fujiwara et al., 1998). A definitive in vitro metabolism study (H.-Z. Bu, manuscript in preparation) suggests that cytochrome P450 3A4 (CYP3A4) is the predominant enzyme responsible for the metabolic clearance of capravirine. A number of metabolites of capravirine were identified in rat liver microsomal incubations, with sulfoxidation, N-oxidation, and sulfonation as the major biotransformation pathways and hydroxylation on the isopropyl moiety as the secondary pathway (Ohkawa et al., 1998). The objective of the present study was to characterize the disposition of capravirine and its metabolites in humans after a single oral dose of [14C]capravirine alone (group A) or after multiple oral doses of ritonavir, a potent CYP3A4 inhibitor (Kumar et al., 1996), followed by a single oral dose of [14C]capravirine (group B). Since capravirine will most likely be used in combination with one or more protease inhibitors to treat HIV-1 infection in antiretroviral-experienced patients, determining the disposition of capravirine in the presence of a potent CYP3A4 inhibitor such as ritonavir is beneficial. Qualitative and quantitative metabolite profiles of capravirine in humans were achieved using HPLC coupled in-line with a radiochemical detector and an ion-trap mass spectrometer (LC-RAM-MS).
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Materials and Methods |
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Human Studies. The study protocol was approved by an independent institutional review board. All subjects (healthy male volunteers) understood the procedures and agreed to participate in the study by giving written informed consent (Declaration of Helsinki).
Five group A subjects were administered on day 1 a single oral dose of capravirine (1400 mg) as a solution of [14C]capravirine (50 µCi, 700 mg; 100 mg/ml in PEG-400) and one capravirine tablet (700 mg). Five group B subjects were administered ritonavir (100-mg capsule) orally once daily for 14 days beginning on day -7. One single oral dose of capravirine (1400 mg) was coadministered with ritonavir on day 1 as a solution of [14C]capravirine (50 µCi, 700 mg; 100 mg/ml in PEG-400) and one capravirine tablet (700 mg). Blood samples were collected at predose and at 0.25, 0.5, 1, 1.5, 2, 2.5, 3, 4, 6, 8, 12, 24, 36, 48, 72, 96, 120, 144, and 168 h after [14C]capravirine dosing. Urine samples were collected at predose and at 0 to 4, 4 to 8, 8 to 12, 12 to 24, 24 to 36, 36 to 48, 48 to 72, 72 to 96, 96 to 120, 120 to 144, and 144 to 168 h after [14C]capravirine dosing. Fecal samples were collected and pooled at 24-h intervals for 7 days after [14C]capravirine dosing. All samples were stored at -20° C before analysis.
Radioanalysis. Aliquots of blood (150 µl) were transferred into tared cones and pads, weighed before combustion using a PerkinElmer Model A307 sampler oxidizer (PerkinElmer Life and Analytical Sciences, Boston, MA). The resulting 14CO2 was trapped in PerkinElmer Carbo-Sorb and mixed with PerkinElmer Permafluor E+ scintillation fluid. Aliquots of plasma (75 µl) and urine (0.3 ml) samples were transferred into tared scintillation vials, weighed, and mixed with 15 ml of Ready-Safe scintillation cocktail (Beckman Coulter, Fullerton, CA). Fecal samples were homogenized with water [20% (w/w) feces/water). Aliquots of fecal homogenates (0.3 ml) were transferred into tared cones and pads and then dried for 24 to 48 h before combustion using a PerkinElmer Model A307 sampler oxidizer. The resulting 14CO2 was trapped in PerkinElmer Carbo-Sorb and mixed with PerkinElmer Permafluor E+ scintillation fluid. All radioassays were performed in triplicate. Radioactivity was measured for 2 min using a Beckman liquid scintillation counter (Beckman Coulter). Data were automatically corrected for counting efficiency using an external standardization technique and an instrument-stored quench curve generated from a series of sealed quench standards.
Sample Preparation for Metabolite Profiling and Structure Elucidation. Plasma. For each of the group A subjects (6–10), an equal volume (1.43 ml) of each plasma sample collected at 1, 1.5, 2, 2.5, 3, 4, and 6 h postdosing was taken to generate a plasma pool (10 ml). For each of the group B subjects (1–5), an equal volume (1.11 ml) of each plasma sample collected at 1, 1.5, 2, 2.5, 3, 4, 6, 8, and 12 h postdosing of [14C]capravirine was taken to generate a plasma pool (10 ml). Each pooled sample was precipitated by the addition of 3 volumes of acetonitrile while vortexing vigorously. After centrifugation, the supernatant was removed and retained. The pellet was re-extracted as above, and the supernatants were combined and transferred into glass tubes for evaporation to dryness under N2 at 45° C. The residues were reconstituted in 140 µl of 30:70 (v/v) methanol/20 mM ammonium acetate (pH 4), and the solutions were transferred into autosampler vials for analysis. The injection volume was 100 µl.
Urine. Urine samples collected at 0 to 4, 4 to 8, 8 to 12, and 12 to 24 h postdosing of [14C]capravirine were pooled on a percent weight basis to generate a urine pool (10 ml) for each subject. Each pooled urine sample was diluted with an equal volume of acetonitrile, vortexed, and centrifuged. The supernatant was transferred into glass tubes for evaporation to dryness under N2 at 45° C. The residues were reconstituted in 0.2 ml of 30:70 (v/v) methanol/20 mM ammonium acetate (pH 4), and the solutions were transferred into autosampler vials for analysis. The injection volume was 100 µl. When more urine samples were needed for structure elucidation, the above procedure would be repeated for selected subjects.
Feces. Individual fecal homogenates with sufficient radioactivity were selected for metabolite profiling. Aliquots (4 ml) of the selected fecal homogenates were extracted with 3 volumes of acetonitrile while vortexing vigorously. After centrifugation, the supernatants were removed and retained. The pellets were re-extracted as above, and the supernatants were combined and transferred into glass tubes for evaporation to dryness under N2 at 45° C. The residues were reconstituted in 0.2 ml of 30:70 (v/v) methanol/20 mM ammonium acetate (pH 4), and the solutions were transferred into autosampler vials for analysis. The injection volume was 100 µl.
Metabolite Profiling. Metabolite profiling was conducted on an Agilent 1100 chromatograph (Agilent Technologies, Palo Alto, CA) coupled in-line with an IN/US ß-RAM radiochemical detector (IN/US Systems, Tampa, FL) and a Finnigan LCQ-Deca ion-trap mass spectrometer (Thermo Finnigan, San Jose, CA). Separation was performed using a Phenomenex (Torrance, CA) Aqua C18 column (150 x 4.6 mm, 5 µm) at a flow rate of 1.0 ml/min. The effluent was split to allow 20% to the mass spectrometer via the supplied electrospray ionization (ESI) source and 80% to mix with the ULTIMA FLO-M scintillation cocktail (at 2.4 ml/min; PerkinElmer Life and Analytical Sciences), and then to flow through the ß-RAM detector. A mobile phase gradient of (A) 20 mM ammonium acetate (pH 4) and (B) methanol was programmed as follows: initiated with 100% A for 10 min, changed to 60% A from 10 to 30 min, changed to 55% A from 30 to 35 min, held at 55% A from 35 to 60 min, changed to 40% A from 60 to 70 min, changed to 10% A from 70 to 80 min, held at 10% A from 80 to 90 min, changed to 100% A from 90 to 92 min, and held at 100% A from 92 to 100 min for the system to be equilibrated. Note that all the above gradient changes were linear. Major operating parameters for the ion-trap ESI-MS method are shown as follows: positive ion mode with a spray voltage of 4.5 kV, capillary temperature of 275° C, sheath gas flow rate of 80 (arbitrary), and an auxiliary gas flow rate of 20 (arbitrary). Full-scan mass spectra were acquired over a mass range of m/z 100 to1000. Win-Flow V1.5 (IN/US Systems) and Xcalibur V1.2 (Thermo Finnigan) were used to control the ß-RAM detector and the LC-MS system, respectively, for data acquisition and processing.
Metabolite Identification. Rapid determination of N-oxides. A molecular ion [M + H]+ containing an N-oxide moiety usually generates a source-induced fragment ion [M + H - O]+ under atmospheric pressure chemical ionization (APCI) conditions (Ramanathan et al., 2000; Tong et al., 2001). This methodology was used to identify N-oxide metabolites of capravirine in humans. The same HPLC method as above was used with the ion-trap mass spectrometer coupled to the supplied APCI source. Note that the HPLC effluent was exclusively passed through the APCI source. Major operating parameters for the APCI-MS method are indicated as follows: positive ion mode with a discharge current of 4.0 µA, capillary temperature of 175° C, vaporizer temperature of 300° C, sheath gas flow rate of 80 (arbitrary), and an auxiliary gas flow rate of 20 (arbitrary). Full-scan mass spectra were acquired over the mass range of m/z 100 to 1000.
Metabolite identification. Helium at a constant pressure of 40 psi was used as the damping and collision gas for all ion-trap LC-ESI-MSn (n = 2–4) experiments. Precursor isolation window, activation amplitude, activation Q, and activation time were set at 1 amu, 40%, 0.25 ms, and 30 ms, respectively. All other conditions were the same as described under Metabolite Profiling.
Incubation of urine samples with ß-glucuronidase. A 0- to 24-h pooled urine sample (10 ml) from group A (2 ml from each subject) was mixed with 10 ml of acetonitrile. After centrifugation, the supernatant was transferred into glass tubes for evaporation to dryness under N2 at 45° C. The residue was reconstituted with 0.4 ml of 100 mM sodium acetate buffer (pH 5). After mixing and centrifugation, the supernatant was split into two equivalent aliquots in two 1.5-ml microcentrifuge tubes. One milligram of ß-glucuronidase was added to one tube and no ß-glucuronidase was added to the other tube that served as control. The two samples were incubated overnight at 37° C in a shaking water bath. The two incubates (100 µl) were injected for LC-RAM-MS analysis to aid in the structure elucidation of glucuronides of capravirine.
Quantification of Capravirine in Plasma. The concentration of capravirine in plasma was determined using a reversed-phase LC-tandem MS method. Capravirine was isolated from plasma by ethyl acetate/hexane liquid-liquid extraction. This method was validated for the analysis of capravirine in 100-µl heparinized human plasma samples over a concentration range of 50 to 10,000 ng/ml with acceptable accuracy and precision.
Pharmacokinetic Analysis. Plasma concentration-time data were analyzed by a noncompartmental pharmacokinetic method using WinNonlin Version 4.0.1 (Pharsight, Mountain View, CA). The maximum plasma concentration (Cmax) and the time at which Cmax was achieved (tmax) were taken directly from the concentration data. The area under the plasma concentration-time curve (AUClast) through the time (tlast) of the last quantifiable concentration (Clast) was estimated by linear trapezoidal approximation. AUC extrapolated to infinity (AUC
) was estimated by adding AUClast and the ratio of Clast/
, where is the plasma terminal elimination rate constant estimated by linear regression analysis of the terminal slope of log plasma concentration-time curve. Apparent terminal elimination half-life (t1/2) was calculated as 0.693/.
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Results |
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The mean total recovery of radioactivity was 86.3% for group A and 79.0% for group B. Radioactivity was generally eliminated equally to urine and feces for groups A and B with approximately 44% and 42% of the dose, respectively, recovered in urine, and 42% and 37% of the dose, respectively, recovered in feces (Table 2).
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Metabolite Profiling. Radiochromatographic peaks were assigned component numbers (e.g., component 9 as C9) in order of retention time (Table 3). However, some component numbers were cross-referenced to the corresponding metabolite numbers (e.g., metabolite 7 as M7) assigned in a previous in vitro study (Ohkawa et al., 1998) (e.g., C9 corresponding to M7 was expressed as C9/M7). The characteristic chlorine isotopic ion clusters [35Cl35Cl/35Cl37Cl/37Cl37Cl = 9/6/1 in relative abundance (McLafferty and Ture
ek, 1993)] coupled with the high sensitivity of the ion-trap mass spectrometer led to the observation of over 50 metabolites of capravirine, of which 19 metabolites were radiochemically quantifiable (Table 3). In this study, both radiochemical profiling (based on radiochromatographic peak areas of radiolabeled drug-derived components) and MS profiling (based on ion chromatographic peak areas of nonradiolabeled drug-derived components) were achieved. The highly sensitive MS method provided additional useful information for the comparison in metabolic profiles between the two treatment groups, especially for the metabolites unquantifiable by radiochemical detection.
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No useful radiochemical profiles were obtained from group A plasma due to insufficient radioactivity (Table 3; Fig. 3A). In group B plasma, unchanged capravirine was the major radiochemical component and C19/M4, C20/M5, C21, and C26/M2 represented the major circulating metabolites (Table 3; Fig. 3B). Based on the MS plasma profiles, the levels of all multioxygenated metabolites, the monooxygenated metabolite C23/M3, the sulfone, and the trioxygenated glucuronide C6 decreased, and the levels of the parent drug, the three other monooxygenated metabolites C19/M4, C20/M5, and C26/M2, and the two other glucuronides C7 and C21 increased, following the coadministration of low-dose ritonavir (Table 3).
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Eight urinary metabolites in group A subjects were radiochemically quantifiable: C4, C6, C9/M7, C11 (the most abundant for this group), C15/M6, C21, C22, and C26/M2; and nine urinary metabolites in group B subjects were radiochemically quantifiable: C6, C7, C12, C14, C19/M4, C20/M5, C21, C23/M3, and C26/M2 (Table 3; Fig. 4). In both groups, unchanged capravirine in urine was observable by MS detection only. Based on the MS urinary profiles, the levels of most multioxygenated metabolites, the sulfone, and the trioxygenated glucuronide C6 decreased, and the levels of all four monooxygenated metabolites C19/M4, C20/M5, C23/M3, and C26/M2, two dioxygenated metabolites C12 and C14, two other glucuronides C7 and C21, and the sulfate C13 increased, following the coadministration of low-dose ritonavir (Table 3).
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Since feces were not available for some subjects at some time intervals, along with the radioactivity of some fecal samples being insufficient, fecal metabolite profiling was performed only for selected samples. For the profiled fecal samples, only the metabolite profiles of the 24- to 48-h fecal homogenates, which were available from most of the subjects and had the highest mean recovery of radioactivity, are presented here (Table 3; Fig. 5). In group A subjects, quantifiable fecal metabolites included C3, C9/M7 (the most abundant for this group), C16, C18, C19/M4, C20/M5, and C23/M3. In group B subjects, the dominant fecal metabolite was C13 (
17% of the dose), which was poorly chromatographed. Other quantifiable fecal metabolites for group B were C3, C9/M7, C19/M4, C20/M5, C21, C23/M3, and C26/M2. The amount of unchanged capravirine in feces was low (0.4% of the dose for group A and 1.4% of the dose for group B). Based on the MS fecal profiles, the levels of most multioxygenated metabolites, the sulfone and the monooxygenated metabolite C23/M3 decreased, and the levels of the three other monooxygenated metabolites, C19/M4, C20/M5, and C26/M2, the dioxygenated metabolite C12, and the sulfate increased, after the coadministration of low-dose ritonavir (Table 3). In summary, coadministration of low-dose ritonavir (a potent CYP3A4 inhibitor) drastically reduced the levels of sequential oxygenated metabolites, and markedly elevated the levels of the parent drug and primary oxygenated metabolites overall in plasma, urine, and feces.
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Metabolite Identification. Capravirine underwent extensive oxidative biotransformation in humans exclusively via individual or combined pathways of sulfoxidation, pyridinyl N-oxidation, and hydroxylation at the primary and tertiary carbons of the isopropyl group (Fig. 1; Table 4), each of which was generally determined based on the characteristic mass spectrometric fragmentation feature(s). In addition, multistage product ion mass spectra of selected capravirine metabolites were generated on the monoisotopic precursor/product ions containing either 35Cl35Cl or 37Cl37Cl when ion intensity was sufficient. The comparison of 35Cl35Cl-containing product ion mass spectra to corresponding 37Cl37Cl-containing ion spectra facilitated the interpretation of collision-induced dissociation pathways and the assignment of metabolite structures. Note that metabolite identification was performed only for the 19 radiochemically quantifiable metabolites and the capravirine sulfone, which was minor in abundance but an important precursor metabolite for sequential metabolism (Table 3).
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Under LC-MSn (n = 2–4) conditions, neutral losses of 48 amu (SO) and 64 amu (SO2) were indicative of sulfoxidation and sulfonation, respectively, of capravirine. Five metabolites, C3, C4, C9/M7, C15/M6, and C23/M3, involved sulfoxidation, and six metabolites, C6, C10, C11, C16, C18, and C22, involved sulfonation, as identified using this approach (Table 4). Based on the characteristic neutral loss of O (16 amu) under APCI-MS conditions, eight metabolites involving pyridinyl N-oxidation were determined: C4, C10, C11, C12, C14, C15/M6, C22, and C26/M2 (Table 4).
It is usually difficult to determine regiochemical positions of hydroxylation by mass spectrometry. However, it was straightforward to distinguish hydroxylation at the primary carbon from that at the tertiary carbon of the isopropyl moiety of capravirine simply, based on relative abundance in the loss of water under MSn (n = 2–4) conditions. As shown in Fig. 6, C19/M4 formed via the tertiary hydroxylation had the loss of water as the predominant fragmentation pathway (i.e., m/z 467 
449 and m/z 406 388). In striking contrast, C20/M5 formed via the primary hydroxylation had the loss of water as the minor fragmentation pathway. This observation was also true for other hydroxylated metabolites of capravirine.
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In general, all multioxygenated metabolites were identified by the simultaneous use of the above specific features. As an example, the MS2 product ion mass spectrum of C11 is illustrated in Fig. 7. The characteristic neutral loss of 64 amu (m/z 454 390) was suggestive of a sulfone, and the predominant loss of water (m/z 515 497 and m/z 454 436) indicated hydroxylation at the tertiary carbon of the isopropyl group of capravirine. In addition to the finding that C11 involved N-oxidation, based on the neutral loss of 16 amu under APCI-MS conditions, the specific fragment ion at m/z 169 (Fig. 7) was also indicative of N-oxidation of capravirine. The structure of the small fragment ion was proposed based on an accurate mass measurement (data not shown), which involved the transfer of the carbamate group. In fact, MS2, MS3, and MS4 ion-trap mass spectrometry was performed for all of the 19 identified metabolites of capravirine, and only the metabolites involving N-oxidation exhibited the specific fragment ion at m/z 169 in their MS2 (neither MS3 nor MS4) product ion mass spectra (Table 4). Metabolites C12, C14, C19/M4, C20/M5, C22, C23/M3, and C26/M2 had authentic standards available, the structures of which were confirmed by similarity of chromatographic retention times and multistage product ion mass spectra (in vivo metabolites versus authentic standards; data not shown).
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Four phase II metabolites of capravirine were also identified in the study: C6, C7, C13, and C21. C13 exhibited [M + H]+ at m/z 547 (= 451 + 16 + 80), consistent with a sulfate of a monooxygenated metabolite of capravirine. As a result of thermal lability under the ESI source conditions, C13 underwent a significant neutral loss of SO3 (80 amu), characteristic of a sulfate, and generated a fragment ion at m/z 467 (= 451 + 16), consistent with a monooxygenated product of capravirine. The product ion mass spectrum of the source-induced ion at m/z 467 (data not shown) suggested that the fragment ion represented the protonated molecular ion of C20/M5. Therefore, C13 was proposed as a sulfate product of C20/M5.
Incubation of human urine samples with ß-glucuronidase was conducted to identify glucuronides of capravirine. C21 with [M + H]+ at m/z 627 (= 451 + 176) was shown to be a glucuronide of capravirine by comparing ion chromatograms of the incubations in the absence versus presence of ß-glucuronidase (data not shown). Likewise, C6 with [M + H]+ at m/z 675 [= 451 + (3 x 16) + 176] was demonstrated to be a glucuronide of C18. C7 with [M + H]+ at m/z 659 [= 451 + (2 x 16) + 176] was a glucuronide of a dioxygenated derivative of capravirine. However, due to the fact that C7 was minor in human urine and that several dioxygenated metabolites of capravirine coexisted, it was not possible to assign a precursor metabolite for the formation of C7 (data not shown).
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Discussion |
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, and t1/2 were increased 3.5-, 9.3-, and 4.6-fold, respectively, for unchanged capravirine and 2.3-, 4.8-, and 2.3-fold, respectively, for total radioactivity in the presence of ritonavir, which is consistent with metabolic inhibition by ritonavir. This is an example of beneficial (or positive) drug-drug interaction (Kumar et al., 1999; Jin et al., 2003), in which the parent drug provides a majority of antiviral activity and a longer duration of higher exposure in plasma may lead to better virologic outcomes. A definitive in vitro study using human liver microsomes has shown that the metabolism of capravirine is mediated predominantly by CYP3A4 (>90%), with minor contribution by CYP2C8 and possibly 2C9 and 2C19 (<10%) and no contribution by the flavin-containing monooxygenases (H.-Z. Bu, manuscript in preparation). Specifically, all monooxygenations (C19/M4, C20/M5, C23/M3, and C26/M2) are mediated primarily by CYP3A4 with minor contribution by the CYP2C isoforms, and all sequential oxygenations (C3, C4, C9/M7, C11, C12, C15/M6, C18, C22, and the capravirine sulfone) appear to be mediated exclusively by CYP3A4. These results indicate that the elevated plasma (also urinary and fecal) levels of the three monooxygenated metabolites (C19/M4, C20/M5, and C26/M2) in the ritonavir-pretreated group are likely attributed to: 1) a significant decrease in the sequential metabolism of the primary metabolites, and 2) a relative increase in the CYP2C-dependent oxygenation of capravirine to the primary metabolites. The elevated plasma level of the glucuronide C21 in the ritonavir-pretreated group is possibly due to the increased level of the parent drug, the precursor for the glucuronidation. The dramatically increased urinary and fecal levels of the sulfate C13 in the ritonavir-pretreated group is consistent with the significantly elevated level of C20, the precursor metabolite for the sulfation. It was expected that the low-dose ritonavir would only exert a partial inhibition of CYP3A4 activity and, therefore, CYP3A4 would still be the primary isozyme responsible for the extensive oxygenations of capravirine in the ritonavir-pretreated group. In addition, the three major monooxygenated metabolites, C19/M4, C20/M5, and C26/M2, in plasma have been tested for anti-HIV activity in vitro. Each of the three shows significantly less activity compared with capravirine (J. P. Graham, unpublished data).
Capravirine is biotransformed to a large number of metabolites (greater than 50) in humans, with up to 15 phase I metabolites and 4 phase II metabolites characterized, and all others observed at trace levels. Formation of all the identified metabolites except C21 (a glucuronide of capravirine) involves oxygenations (mono-, di-, tri-, or tetra-). Oxygenation sites are restricted to the sulfur atom (for sulfoxidation or sulfonation), the pyridinyl nitrogen atom (for N-oxidation), and the isopropyl group (for mono- or dihydroxylation) (Fig. 8). Among the 15 phase I metabolites, 9, 8, 7, and 5 metabolites involve S-oxidation (5 for sulfoxidation and 4 for sulfonation), N-oxidation, tertiary hydroxylation, and primary hydroxylation, respectively, indicating that the first three reactions constitute the primary routes and the last one the secondary route in the oxidative metabolism of capravirine. These data show that capravirine has a great tendency to undergo sequential biotransformation reactions, resulting in the numerous multioxygenated metabolites.
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The chemical structures of capravirine metabolites in humans were proposed based on complementary APCI-MS and ESI-MSn (n = 2–4) mass spectral data as well as ß-glucuronidase incubation for glucuronides. Although the structure elucidation involved various isomeric oxygenated metabolites (4 mono-, 4 di-, 4 tri-, and 2 tetraoxygenated), it was still achievable to distinguish those isomers using the aforementioned strategies. In addition to the characteristic features of neutral losses, other specific product ions for a specific metabolite were also used for the identification of the metabolite. The ion-trap mass spectrometer can perform multistage collision-induced dissociation fragmentations of any precursor ions of interest if the ion current is sufficient. This unique function makes the tool very powerful in distinguishing those isomeric metabolites.
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Acknowledgments |
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Footnotes |
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Address correspondence to: Dr. Hai-Zhi Bu, Department of Pharmacokinetics, Dynamics and Metabolism, Pfizer Global Research and Development, San Diego, CA 92121. E-mail: haizhi.bu{at}pfizer.com
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References |
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ek F (1993) Interpretation of Mass Spectra. University Science Books, Mill Valley, CA.
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