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Drug Metab Dispos. Apr 2011; 39(4): 627–635.
PMCID: PMC3063720

In Vitro Metabolism of 17-(Dimethylaminoethylamino)-17-demethoxygeldanamycin in Human Liver Microsomes

Abstract

The objective of this study was to investigate the oxidative metabolism pathways of 17-(dimethylaminoethylamino)-17-demethoxygeldanamycin (17-DMAG), a geldanamycin (GA) derivative and 90-kDa heat shock protein inhibitor. In vitro metabolic profiles of 17-DMAG were examined by using pooled human liver microsomes (HLMs) and recombinant CYP450 isozymes in the presence or absence of reduced GSH. In addition to 17-DMAG hydroquinone and 19-glutathionyl 17-DMAG, several oxidative metabolites of 17-DMAG were detected and characterized by liquid chromatography-tandem mass spectrometry. Different from previously reported primary biotransformations of GA and GA derivatives, 17-DMAG was not metabolized primarily through the reduction of benzoquinone and GSH conjugation in HLMs. In contrast, the primary biotransformations of 17-DMAG in HLMs were hydroxylation and demethylation on its side chains. The most abundant metabolite was produced by demethylation from the methoxyl at position 12. The reaction phenotyping study showed that CYP3A4 and 3A5 were the major cytochrome P450 isozymes involved in the oxidative metabolism of 17-DMAG, whereas CYP2C8, 2D6, 2A6, 2C19, and 1A2 made minor contributions to the formation of metabolites. On the basis of the identified metabolite profiles, the biotransformation pathways for 17-DMAG in HLMs were proposed.

Introduction

The 90-kDa heat shock protein (Hsp90) is a molecular chaperone to mediate the folding, activation, and assembly of many oncogenic client proteins, which stimulate cancer cell growth (McIlwrath et al., 1996). Geldanamycin (GA) is an Hsp90 inhibitor that binds to Hsp90 and disrupts the interaction between Hsp90 and its client proteins (An et al., 1997). This disruption depletes the oncogenic proteins and results in antitumor activity. To develop potent antitumor agents, a number of GA derivatives have been synthesized and characterized biologically. Among GA derivatives, 17-(allylamino)-17-demethoxygeldanamycin (17-AAG) and 17-(dimethylaminoethylamino)-17-demethoxygeldanamycin (17-DMAG) have been introduced into clinical trials (Glaze et al., 2005).

Both GA and 17-AAG are known to undergo extensive metabolism (Egorin et al., 1998; Musser et al., 2003; Guo et al., 2005, 2006; Lang et al., 2007). Although GA and 17-AAG are structurally similar (see Fig. 1), their metabolite profiles in liver microsomes are different (Lang et al., 2007). GA is primarily (40–73%) reduced into geldanamycin hydroquinone (GAH2) (Lang et al., 2007; Guo et al., 2008). During exposure to oxygen, GAH2 slowly reverts to GA. In the presence of reduced GSH, more than 50% of GA is rapidly converted into 19-glutathionyl geldanamycin hydroquinone (Cysyk et al., 2006; Lang et al., 2007). No significant amount of oxidative metabolites of GA in the incubations with human liver microsomes (HLMs) has been detected (Lang et al., 2007). The metabolic pathways of 17-AAG in liver microsomes are controversial. Guo et al. (2008) reported that quinone/hydroquinone conversion was the primary metabolism mode of 17-AAG and 17-DMAG in microsomal preparation. In the presence of reduced GSH, 15% of 17-AAG was conjugated with GSH after incubation in liver microsomes for 24 h. However, Lang et al. (2007) observed that only 2% of 17-AAG was reduced into hydroquinone in HLMs, and no significant amount of 19-GSH conjugate of 17-AAG was detected in HLMs in the presence of 5 mM GSH. Furthermore, they found that, different from GA, 17-AAG in HLMs primarily underwent oxidative metabolism on the 17-allylamino side chain to form 17-aminogeldanamycin (17-AG) (see Fig. 1) and 17-(2′,3′-dihydroxypropylamino)-geldanamycin, which was consistent with a previous study (Egorin et al., 1998).

Fig. 1.
Structures of GA, 17-AAG, 17-DMAG, and 17-AG.

17-DMAG is much more metabolically stable than 17-AAG because of the limited oxidative metabolism on 17-dimethylaminoethylamino side chain (Glaze et al., 2005). Compared with 17-AAG, 17-DMAG exhibits a longer terminal half-life of 16 to 19 h (Hwang et al., 2006; Moreno-Farre et al., 2006) (4 h for 17-AAG) and a lower total clearance of 7.4 to 17.7 l/h (Hwang et al., 2006; Moreno-Farre et al., 2006) (36 l/h for 17-AAG) in humans. Although the preclinical (Egorin et al., 2002) and clinical (Glaze et al., 2005; Goetz et al., 2005) pharmacokinetics of 17-DMAG have been investigated, to our knowledge, the biotransformation information of 17-DMAG is still limited and controversial. Reduction of quinone was proposed to be the primary metabolism of 17-DMAG in liver microsomes, and 17-DMAG was observed to undergo more rapid GSH conjugation than 17-AAG (Guo et al., 2008). However, these findings cannot explain the less in vivo metabolism of 17-DMAG than that of 17-AAG in animals and humans (Musser et al., 2003; Hwang et al., 2006).

Biotransformation of GA and its derivatives is related to their antitumor activity and toxicity. For example, the reduction of benzoquinone ansamycins into hydroquinone ansamycins enhanced Hsp90 inhibition (Guo et al., 2006; Lang et al., 2007), whereas GSH conjugation of benzoquinone ansamycins was correlated with their hepatic toxicity (Guo et al., 2008). Hence, it is important to elucidate the major biotransformation pathways of 17-DMAG in liver microsomes for discovery of more stable, potent, and less toxic GA analogs.

In this study, we investigated the biotransformation pathways of 17-DMAG in HLMs and especially focused on quinone-hydroquinone conversion and GSH conjugation. The relative percentages of major metabolites of 17-DMAG were estimated by normalizing their peak areas. The major metabolites in incubations were tentatively characterized using liquid chromatography-tandem mass spectrometry (LC-MS/MS), and the cytochrome P450 (P450) enzymes responsible for the formation of the metabolites were identified. On the basis of the identified metabolite profiles, the biotransformation pathways for 17-DMAG in HLMs were proposed.

Materials and Methods

Materials.

GA, 17-AAG, and 17-DMAG were purchased from LC Laboratories (Woburn, MA). Reduced GSH, reduced β-NADPH, MgCl2, 0.1 M phosphate buffer, formic acid α-naphthoflavone, tranylcypromine, quercetin, sulfaphenazole, ticlopidine, diethyldithiocarbamate, quinidine, and ketoconazole were supplied by Sigma-Aldrich (St. Louis, MO). MI-63 was obtained from Prof. S. Wang (Department of Medicinal Chemistry, University of Michigan, Ann Arbor, MI). High-performance liquid chromatography (HPLC)-grade acetonitrile was purchased from Thermo Fisher Scientific (Waltham, MA). HPLC water was purified using a MilliQ water system (Millipore Corporation, Billerica, MA). Pooled HLMs (20 mg/ml), purified Escherichia coli-expressed recombinant human P450 enzymes coexpressed with human cytochrome b5 (1 nmol/vial), and E. coli-expressed control microsomes were obtained from XenoTech, LLC (Lenexa, KS).

Metabolic Stability Assay.

GA, 17-AAG, or 17-DMAG was incubated with HLMs in the absence or presence of reduced GSH at 37°C. The enzymes were activated by reduced β-NADPH. The incubation solution was diluted with 0.1 M phosphate buffer (containing MgCl2) to 0.6 ml. The final concentrations of drug, microsomes, β-NADPH, phosphate buffer, and MgCl2 were 5 μM, 1 mg/ml, 1 mM, 0.1 M, and 3.3 mM, respectively. For GSH conjugation assays, the final concentration of reduce GSH was 5 mM (Lang et al., 2007). An aliquot of 40 μl of mixture was collected at 1, 5, 10, 15, 30, 45, 60, and 120 min, and then proteins were precipitated with 120 μl of ice-cold acetonitrile containing an internal standard MI-63 (100 ng/ml). The samples were centrifuged at 14,000 rpm × 5 min, and 10 μl of supernatant was injected into LC-MS. To determine the apparent Km values for the formation of 17-AG from 17-AAG, various concentrations of 17-AAG (1–200 μM) were incubated in 0.25 mg/ml HLMs for 15 min in triplicate. Likewise, to determine the Km values for the formation of M1 and M3 from 17-DMAG, various concentrations of 17-AAG (0.5–100 μM) were incubated in 0.5 mg/ml HLMs for 30 min in triplicate. The formation rates of the metabolites were fitted with the Michaelis-Menten equation using WinNonlin (Pharsight, Mountain View, CA).

Incubations with HLMs for Metabolite Identification.

17-DMAG was incubated with HLMs in 0.6 ml of phosphate buffer (0.1 M) at 37°C. The final concentrations of microsomes, β-NADPH, phosphate buffer, and MgCl2 were the same as described in stability assay. The initial concentration of 17-DMAG was 10 μM. Two different negative controls were prepared simultaneously by using boiled microsomes (100°C for 5 min) or spiking 10 μM 17-DMAG after protein precipitation. Both samples and negative controls were incubated for 2 h, and the reaction was terminated with 1.2 ml of ice-cold acetonitrile to precipitate proteins. In addition, to evaluate GSH conjugation, 17-DMAG (10 μM) was incubated with reduced GSH (5 mM) and β-NADPH (1 mM) in 0.1 M phosphate buffer (containing MgCl2) in the presence or absence of HLMs (1 mg/ml) at 37°C for 2 h. After protein precipitation, samples and negative controls were centrifuged at 14,000 rpm × 5 min for LC-MS analysis.

LC-MS/MS.

An Agilent 1200 HPLC system (Agilent Technologies, Santa Clara, CA) was used for separation. The processed samples were injected on a Zobarx SB-C18 column (2.1 × 50 mm, 3.5 μm) (Agilent Technologies). Mobile phase A, consisting of 0.1% formic acid and 10 mM ammonium formate in water, and mobile phase B, consisting of 0.1% formic acid in acetonitrile, were used for a linear gradient elution as follows: 10 to 90% B in 10 min, hold 90% B for 3 min, return to 10% B in 0.1 min, and hold 10% B for 3.9 min to equilibrate the column. The flow rate was 0.4 ml/min. Mass spectrometric detection was performed on a QTRAP 3200 mass spectrometer (Applied Biosystems/MDS Sciex, Foster City, CA) equipped with an electrospray ionization source. 17-DMAG was detected under positive ionization mode, whereas 17-AAG and GA were detected under negative ionization mode. The multiple reaction monitoring (MRM) ion transition was m/z 617 → 58 for 17-DMAG, m/z 559 → 516 for GA, and m/z 584 → 541 for 17-AAG (Smith et al., 2004). Ionization source temperature was set at 650°C. Curtain gas, gas 1, and gas 2 were set at 30, 50, and 50 arbitrary units, respectively. Ion spray voltage was set at 5500 V for 17-DMAG and −4500 V for GA and 17-AAG. A neutral loss of 43 Da was observed, likely because of the loss of N-methylenemethanamine from 17 side chain. Hence, the MRM ion transitions at m/z 561 → 518, 864 → 821, and 866 → 823 were used to detect GAH2, 19-glutathionyl GA, and 19-glutathionyl geldanamycin hydroquinone, respectively. MRM ion transitions at m/z 586 → 543, 889 → 846, and 891 → 848 were used to detect 17-(allylamino)-17-demethoxygeldanamycin hydroquinone (17-AAGH2), 19-glutathionyl 17-(allylamino)-17-demethoxygeldanamycin (17-AAG-SG), and 19-glutathionyl 17-(allylamino)-17-demethoxygeldanamycin hydroquinone, respectively. Meanwhile, the tertiary amine of 17-DMAG facilitated its protonation and resulted in a high MS sensitivity under positive ionization. As a result, MRM ion transitions at m/z 619 → 58, 922 → 58, and 924 → 58 were used to detect 17-(dimethylaminoethylamino)-17-demethoxygeldanamycin hydroquinone (17-DMAGH2), 19-glutathionyl 17-(dimethylaminoethylamino)-17-demethoxy geldanamycin (17-DMAG-SG), and 19-glutathionyl 17-(dimethylaminoethylamino)-17-demethoxygeldanamycin hydroquinone, respectively.

Screening and Characterization of Metabolites.

GA, 17-AAG, and 17-DMAG were infused into a mass spectrometer to obtain their MS, MS2, and MS3 spectra. On the basis of the similarities and difference among their mass spectra, the structures of fragment ions of protonated 17-DMAG were proposed tentatively. MRM ion transitions 617 → 58, 617.3 → 159, and 617 → 524 were used to generate 240 additional MRM ion transitions for metabolites screening using Metabolite ID software (Applied Biosystems), including 40 common biotransformation processes. To detect all the metabolites, full scan and precursor scan also were conducted. Only the components detected in the sample and absent in all the control samples were regarded as possible metabolites. To characterize the possible metabolites, both the sample and controls were injected on the LC-MS for EPI and MS3 scans to obtain their MS2 and MS3 spectra. On the basis of the MS2, MS3 spectra of the metabolites, and the proposed structures of 17-DMAG fragment ions, the metabolites of 17-DMAG were characterized.

Inhibition of 17-DMAG Oxidative Metabolism by Selective P450 Inhibitors.

The incubation mixtures, containing 10 μl of pooled HLMs (20 mg protein/ml), were preincubated in 0.1 M phosphate buffer (pH 7.4) in the presence of reduced β-NADPH (1 mM), MgCl2 (3.3 mM), and chemical inhibitor for 3 min at 37°C in a shaking water bath. Parallel control incubations were conducted in the absence of chemical inhibitors. The reactions were initiated by adding 17-DMAG (final concentration, 1 μM). The reaction mixtures (final volume 0.2 ml) were incubated for 20 min at 37°C. The P450 isoform-selective inhibitors used were α-naphthoflavone for 1A2 (10 μM), tranylcypromine for 2A6 (1 μM), ticlopidine (10 μM) for 2B6 and 2C19, quercetin (1 μM) for 2C8, sulfaphenazole (1 μM) for 2C9, quinidine (1 μM) for 2D6, diethyldithiocarbamate (10 μM) for 2E1, and ketoconazole (1 μM) for 3A (Kim et al., 2003; Lee et al., 2006). In positive control experiments, 10 μM α-naphthoflavone inhibited the formation of acetaminophen from phenacetin (20 μM); 1 μM tranylcypromine inhibited the formation of 7-hydroxycoumarin from coumarin (5 μM); 10 μM ticlopidine inhibited the formation of hydroxybupropion from bupropion (50 μM) and formation of hydroxyomprazole from omprazole (50 μM); 1 μM quercetin inhibited the formation of 6α-hydroxypaclitaxel from paclitaxel (10 μM); 1 μM sulfaphenazole inhibited hydroxytolbutamide formation from tolbutamide (100 μM); 1 μM quinidine inhibited dextrorphan formation from dextromethorphan (5 μM); 10 μM diethyldithiocarbamate inhibited the formation of hydroxychlorzoxazone from chlorzoxazone (50 M); and 1 μM ketoconazole inhibited 1′-hydroxymidazolam formation from midazolam (5 μM) (Lee et al., 2006). The reactions were terminated by adding 400 μl of ice-cold acetonitrile containing 100 ng/ml MI-63. The samples were vortexed for 30 s and centrifuged at 14,000 rpm for 5 min. The supernatant was analyzed by LC-MS/MS to monitor formation rate of 17-DMAG metabolites. The MRM ion transitions for determination of M1, M2, M3, M4, and M6 were 633 → 175, 633 → 322, 603 → 510, 603 → 524, and 665 → 157, respectively. The percentages of inhibition were calculated by the ratio of the amounts of metabolites formed with and without the specific inhibitor.

Incubation with Recombinant CYP450 Enzymes.

The incubation mixtures (200 μl), containing 20 pmol of recombinant CYP450 enzyme, 0.2 mg of reduced β-NADPH, and 3.3 mM MgCl2, were preincubated in 0.1 M phosphate buffer (pH 7.4) at 37°C in a shaking water bath. E. coli-expressed control microsomes without cDNA of human P450 were used as a negative control. The reactions were initiated by adding 17-DMAG (final concentration 1 μM). After 30-min incubation, the reactions were terminated by adding 400 μl of ice-cold acetonitrile containing 100 ng/ml MI-63. LC-MS/MS was used to monitor formation rate of 17-DMAG metabolites. The formation rates of the metabolites in each CYP450 isozyme were expressed as the percentages relative to their formation rates in CYP3A4.

Results

Relative Percentages of Metabolites in Incubations.

The apparent Km values for the formation of 17-AG from 17-AAG and M1 and M3 from 17-DMAG in HLMs were determined as 29.1, 6.61, and 7.83 μM, respectively, which were higher than the initial concentrations of 17-AAG and 17-DMAG (5 μM) in stability assays. The relative percentages of GA and 17-AAG and their corresponding hydroquinone and 19-glutathionyl conjugates determined in stability assays were shown in Fig. 2. The values were expressed as percentages normalized by peak area ratio of GA or 17-AAG and internal standard at 1 min of incubation. GA exhibited high metabolic stability in HLMs in the absence of reduced GSH (Fig. 2A). Seventy-three percent of GA was not metabolized after 2 h of incubation in 1 mg/ml HLMs. A total of 6.4 to 9.3% of GAH2 was obtained during the course of incubation. A similar amount of GAH2 was also detected in the incubation with boiled HLMs (negative control), suggesting that the formation of GAH2 was not dependent on CYP450 enzymes. In contrast, the presence of 5 mM reduced GSH resulted in the rapid metabolism of GA during the first 5 min of incubation. Only 75.3 and 46.6% of GA remained after 5- and 120-min incubation, respectively. Consistently, the reduced GSH resulted in a quick onset of the formation of GAH2 in HLMs, and 85.7% of GAH2 formed 1 min after the addition of 5 mM reduced GSH. In addition, the formation of GAH2 was also observed in the incubation of GA and GSH in the absence of HLMs, and the peak area ratio of GAH2 and GA was 1.93 after 2 h of incubation. No significant amount of GSH conjugates were detected in all the above incubations using negative MRM ion transitions m/z 864 → 821 and 866 → 823.

Fig. 2.
Relative percentages of GA and GAH2 (A); 17-GA and 17-GAH2 (B); and 17-DMAG (C) and their metabolites in the incubations with HLMs in the presence or absence of 5 mM reduced GSH. The values were expressed as percentages normalized by peak area ratio of ...

In contrast to GA, 17-AAG was extensively metabolized in 1 mg/ml HLMs and totally disappeared 45 min after the beginning of incubation (Fig. 2B). Less than 1% of 17-AAGH2 was detected during the first 30 min of incubation in HLMs, and no 17-AAGH2 was detected after incubation for 1 h. The presence of reduced GSH did not significantly change the metabolism rate of 17-AAG in HLMs but resulted in a quicker onset of 17-AAGH2 formation. Furthermore, 3.04% of 17-AAGH2 was obtained when 5 μM 17-AAG was incubated with 5 mM reduced GSH for 2 h in the absence of HLMs. GSH conjugates of 17-AAG were not detected over the course of incubations using the negative MRM ion transitions m/z 889 → 846 and 891 → 848.

Figure 2C showed the relative percentages of 17-DMAG and its metabolites during the course of incubation. The characterization of 17-DMAG metabolites M1 to M7 is discussed in the following section. Compared with 17-AAG, 17-DMAG exhibited improved metabolic stability. Twenty-four percent of 17-DMAG remained in HLMs after 2 h of incubation. Only a trace amount (0.24%) of 17-DMAGH2 was obtained after incubation for 2 h. Reduced GSH showed no effect on the metabolism of 17-DMAG and formation of 17-DMAGH2 in HLMs. The maximum percentage (0.33%) of 17-DMAG-SG during the incubation was detected at 60 min, whereas 19-glutathionyl 17-(dimethylaminoethylamino)-17-demethoxygeldanamycin hydroquinone was not detected during the incubation. In addition, 1.16% of 17-DMAG-SG was obtained when 17-DMAG was incubated with reduced GSH for 2 h in the absence of HLMs.

Identification of 17-DMAG Metabolites.

To assign the major fragment ions of protonated 17-DMAG, the MS/MS and MS3 spectra of GA, 17-AAG, and 17-DMAG were obtained. Figure 3A showed the MS/MS spectrum of protonated 17-DMAG. The product ions at m/z 58 and 72 of 17-DMAG, which were not detected in the MS/MS spectra of GA and 17-AAG, were assigned to the N,N-dimethylethanamine and trimethylamine moieties of 17-dimethylaminoethylamino side chain (Fig. 3B). Both product ions at m/z 187 and 159 were detected in MS/MS spectra of GA, 17-AAG, and 17DMAG. On the basis of the accurate mass measurement, the product ion at m/z 187 was assigned to C1–10 as shown in Fig. 3B (Lang et al., 2007). On the MS3 spectra, the ion at m/z 187 was further fragmented to generate ions at m/z 159, 131, and 117, and their structures were tentatively proposed in Fig. 3B.

Fig. 3.
A, MS/MS spectrum and MS3 spectrum (m/z 617 → 187) of protonated 17-DMAG. B, the proposed fragmentation pathways of protonated 17-DMAG.

In addition to 17-DMAGH2 and 17-DMAG-SG, 12 metabolites of 17-DMAG were detected in the HLM incubation but not in negative controls by MS full scan and MRM scan. Three metabolites had a protonated molecular ion (Mr) of 665, five had an Mr of 633, one had an Mr of 627, one had an Mr of 615, and two had an Mr of 603. The major metabolites were the two compounds with an Mr of 603, five compounds with an Mr of 633, and one compound with an Mr of 665. The metabolites with Mr of 603 and 633 were also observed previously in the bile of rats administered with 17-DMAG, although their structures were not identified (Egorin et al., 2002).

Five components were detected in the LC-MS/MS chromatogram (m/z 633) of HLM incubation extract but not the negative controls (Fig. 4A). The Mr (m/z 633) of each metabolites showed a mass shift of 16 Da compared with protonated 17-DMAG, indicating the addition of one oxygen. The MS/MS spectrum (Fig. 4B) of the component eluted at 8.21 min (M1) showed the product ions at m/z 58 and 72, suggesting that the oxidation did not occur at the 17 side chain. The product ions of M1 at m/z 203, 175, 147, and 133 exhibited a mass shift of 16 Da compared with the corresponding product ions of 17-DMAG, indicating that one hydroxyl group generated on C1–10. Furthermore, MS3 spectrum of protonated M1 (m/z 617 → 203) showed no neutral loss of water from the ion at m/z 203, indicating that there was no hydrogen on the carbon next to the hydroxylated carbon. Hence, the hydroxyl of M1 was tentatively assigned to C23. Product ions at m/z 187 and 159 were detected in the MS/MS spectrum (Fig. 4C) of the component eluted at 9.04 min (M2), suggesting that the hydroxylation did not occur at C1–10 moiety. The detection of a product ion at m/z 70 instead of 72 suggested the oxidation on N,N-dimethylethanamine, whereas the presence of a product ion at m/z 58 indicated that the trimethylamine moiety was intact. Hence, the hydroxyl was tentatively assigned to the α-carbon. The product ion at m/z 70 was proposed to be generated from α-hydroxylated 17-DMAG by cleaving the bond between nitrogen and α-carbon, followed by dehydration. Similar α-hydroxylation on a 17 side chain also was observed in the metabolism of 17-AAG in HLMs (Lang et al., 2007).

Fig. 4.
A, LC-MS/MS chromatogram (m/z 633) of HLM incubation extract. Five metabolites of 17-DMAG formed by the addition of one oxygen; B, MS/MS spectrum of the metabolite eluted at 8.21 min (M1); C, MS/MS spectrum of the metabolite eluted at 9.04 min (M2).

The metabolite eluted at 7.14 min (Fig. 4A) showed a similar profile of product ions as that of M2, except the presence of the metabolite's product ion was at m/z 72 instead of m/z 70. The major product ions of this metabolite were ions at m/z 159 and 187 instead of ions at m/z 322 and 304. The metabolite was probably formed by the hydroxylation of C25 or the nitrogen adjacent to C17. The metabolites eluted at 6.71 and 7.53 min exhibited the same MS/MS pattern, and the major product ions were ions at m/z 572, 540, 306, 274, 246, 192, and 177, suggesting the oxidation might occur at C1–10 moiety, C22, C24, or two methoxyl groups. The components eluted at 9.65 and 10.14 min might not be metabolites because they were also detected in the negative controls. A complete separation of the peaks at 7.53 and 7.80 min on the chromatogram was not obtained. The structure of the metabolite at 7.80 min was not identifiable because of the low intensities of its product ions.

Two metabolites with the identical Mr at m/z 603 (parent − 14) were eluted at 7.57 min (M3) and 7.13 min (M4). Both metabolites were likely formed by demethylation (Fig. 5A). The two metabolites had similar MS/MS fragmentation pattern, except for the detection of product ions of M3 at m/z 571 and 510 and product ions of M4 at m/z 585 and 524 (Fig. 5, B and D). The ions at m/z 571 and 510 were derived from protonated M3 by the loss of methanol and subsequent loss of carbamic acid from C7. Compared with the loss of methanol from C12, the loss of methanol from C6 position was more energetically favored because it would increase the amount of conjugation in the ansamycin ring. Hence, the product ion of M3 at m/z 571 was generated by the loss of methanol from C6, and M3 was produced by the demethylation from the methoxyl at C12 of 17-DMAG (Fig. 5C). Likewise, the product ions of M4 at m/z 585 and 524 were generated by the loss of water from C6 and subsequent loss of carbamic acid from C7 (Fig. 5D). M4 was generated by the demethylation from the methoxyl at C6. The component eluted at 8.08 min was also detected in the negative controls, indicating that it was not a metabolite of 17-DMAG.

Fig. 5.
A, LC-MS/MS chromatogram (m/z 603) of HLM incubation extract. Two metabolites of 17-DMAG formed by demethylation; B, MS/MS spectrum and MS3 spectrum (m/z 603 → 306) of the metabolite eluted at 7.57 min (M3); C, the proposed formation and fragmentation ...

One metabolite eluted at 8.4 min (M5) exhibited a Mr at m/z 615 (parent − 2), indicating a loss of H2O from hydroxylized 17-DMAG (Fig. 6A). The detection of m/z 187 suggested that the metabolism did not occur at C1–10 position. The ions at m/z 72 and 58 were not detected in the MS/MS spectrum of M5, suggesting that the metabolism was likely to occur on N,N-dimethylethanamine moiety. The base product ion peak of protonated M5 at m/z 304 was also detected in the MS/MS spectrum of M2 (Fig. 4C). Hence, M5 is proposed to be derived from M2 by the loss of H2O, which resulted in the formation of a carbon-nitrogen double bond and increased amount of conjugation in the system (Fig. 6A).

Fig. 6.
MS/MS spectrum of the metabolite eluted at 8.40 min (M5) (A) and the metabolite eluted at 6.80 min (M6) (B).

It is worth noting that the product ion at m/z 306 was detected in the MS/MS spectra of M1 (Fig. 4B), M3 (Fig. 5B), and M4 (Fig. 5C). Its counterpart ion at m/z 322 in MS/MS spectrum of M2 (Fig. 4C) showed a mass shift of 16 Da. Another counterpart ion at m/z 304 in MS/MS of M5 (Fig. 6A) showed a mass shift of 2 Da. Hence, the ion at m/z 306 was possibly generated by the loss of methanol (M1 and M4) or H2O (M3) from C12 and subsequent cleavage of the amide bond in the ring and the bond between C10 and C11 (Fig. 4C). MS3 spectrum of M3 (m/z 633 → 306) showed that the ion at m/z 306 generated its fragment ions at m/z 274, 246, 229, and 218 (Fig. 5B) by the loss or sequential losses of CH3OH, CO, and NH3 (Fig. 5C). The product ion of M2 at m/z 322 (Fig. 4C) and the product ion of M5 at m/z 304 (Fig. 6A) were produced through similar fragmentation.

The metabolite eluted at 6.8 min (M6) showed an Mr at m/z 665 (parent + 48), indicating the addition of the three oxygen to 17-DMAG. The detection of ion at m/z 338 (Fig. 6B), a counterpart of the ion at m/z 306 of M3 and M4 (a mass shift of 32 Da), indicated that two oxygen atoms were added to C11–20 moiety. The presence of the ion at m/z 58 suggested that two N-methyl groups were not oxidized. Two hydroxyl groups were tentatively assigned to the α-carbon of 17 side chain and C25, respectively. The product ions of M6 at m/z 185 and 157 (Fig. 6B) exhibited a mass shift of 2 Da from their counterpart ions of 17-DMAG at m/z 187 and 159, respectively, indicating a third hydroxyl group located at C1–10 moiety of M6, and this hydroxyl was ready to be deprived from protonated M6 through dehydration. Hence, the third hydroxyl was tentatively assigned to C24 because the formation of a double bond between C24 and C10 would increase the amount of conjugation and stabilize the product ions of M6. The relative amounts of the six metabolites M1 to M6 in HLM incubation during 1 to 120 min were shown in Fig. 2C. The relative amounts of 17-DMAG metabolites at 60 min were M3 (2.19%) > M1 (1.48%) > M6 (1.40%) > M2 (1.38%) > M4 (0.98%) > M5 (0.52%) > 17-DMAGH2 (0.19%).

One minor metabolite eluted at 9.2 min exhibited an Mr at m/z 627, a mass shift of 10 Da from the counterpart ion of 17-DMAG. The major product ion of this metabolite included ions at m/z 566, 538, and 331. The product ion at m/z 566 was derived from the Mr by the loss of carbamic acid. A subsequent neutral loss of 28 Da instead of 32 Da (the loss of methanol C12 or C6) from m/z 566 was detected and gave rise to m/z 538, suggesting the possible loss of an ethylene or carbon monoxide. Because of the low concentration of this metabolite in HLM incubation, the MS3 spectra of its product ions were not obtainable. Additional study is required to definitively identify the structure of the metabolite.

Inhibition of 17-DMAG Oxidative Metabolism by Selective P450 Inhibitors.

Identification of the P450 isozymes responsible for oxidative metabolism of 17-DMAG was performed using both P450-selective inhibitors. The results of experiments using specific P450 inhibitors to prevent the formation of 17-DMAG metabolites in HLMs were summarized in Fig. 7. The formation of M1, M2, M3, M4, and M6 in HLMs was inhibited dramatically by a CYP3A inhibitor. Ketoconazole (1 μM) resulted in 75 to 88% inhibition on the formation of the five metabolites. The other inhibitors did not cause remarkable inhibition on the formation of metabolites in HLMs. Results from positive control experiments confirmed that the incubations were performed under optimum conditions.

Fig. 7.
Effect of selective CYP450 inhibitors on the oxidative metabolism of 17-DMAG by HLMs.

Oxidative Metabolism of 17-DMAG by Recombinant P450 Isoenzymes.

17-DMAG (1 μM) was incubated with recombinant human P450 isozymes, and the relative formation rates (percentage of that of CYP3A4) of M1, M2, M3, M4, and M6 were shown in Fig. 8. Among the CYP450 isoforms tested, CYP3A4 produced the highest formation rates of M1, M2, M3, M4, and M6. CYP3A5 showed a lower activity for the metabolism of 17-DMAG relative to CYP3A4. CYP2C8, 2D6, 2A6, 2C19, and 1A2 produced M6 at moderate rates compared with CYP3A4 and 3A5. Other CYP450 enzymes did not metabolize 17-DMAG at an appreciable rate. The incubation of 17-DMAG with E. coli-expressed control microsomes did not show the formation of the metabolites.

Fig. 8.
Relative formation rates of M1, M2, M3, M4, and M6 by recombinant human CYP450 isozymes. The values were expressed as percentages normalized by the formation rate in CYP3A4 incubation.

Discussion

The molecular modeling analysis showed that the reduction of quinone into hydroquinone facilitated the formation of two more hydrogen bonds between GAH2 and Hsp90, resulting in a tighter binding between GAH2 and Hsp90 (Lang et al., 2007). GSH plays an important role in the detoxification of reactive drugs and metabolites formed by hepatic drug-metabolizing enzymes (Satoh, 1995). GA and its analogs were observed to react chemically (i.e., nonenzymatically) with GSH to form GSH conjugates at 19-position, and the GSH conjugation could be important for the toxicity of GA and its analogs (Cysyk et al., 2006). One objective of this study was to detect and compare the quinone-hydroquinone conversion and GSH conjugation among GA, 17-AAG, and 17-DMAG in HLM incubations. Previous study (Lang et al., 2007) and our preliminary study showed that only trace amounts of hydroquinone metabolites and 19-glutathionyl metabolites of 17-AAG or 17-DMAG formed in the HLM incubation. To generate enough metabolites, a substrate concentration of 5 μM and a HLM concentration of 1 mg/ml were selected for metabolic stability assays. The substrate concentration was lower than the Km values for the formation of major metabolites of 17-AAG and 17-DMAG. To evaluate the formation of GSH conjugates of GA derivatives in HLMs, GSH in reduced form was added to the HLM incubations to reach a final concentration of 5 mM as described previously (Lang et al., 2007; Guo et al., 2008). It was also reported (Guo et al., 2008) that less than 15% of 17-AAG-SG formed even after 24 h of incubation in the presence of 5 mM reduced GSH, indicating a slow GSH conjugation rate. Hence, the total incubation time of metabolic stability assays was chosen as 2 h to acquire as high an amount of GSH conjugates as possible while maintaining reasonable enzyme activity.

A total of 6.4 to 9.3% of GAH2 was detected in HLM incubation in the absence of GSH, much lower than the amount (40–73%) detected in a published report (Lang et al., 2007). The inconsistency might be partially caused by oxygen exposure during sample preparation as some GAH2 was oxidized into GA. The reduced GSH stimulated the reduction of GA, and 78 to 88% of GAH2 was detected, which was consistent with the amount of GAH2 obtained in HLM incubation under hypoxic conditions (Lang et al., 2007). The reduction of GA was not dependent on CYP450 enzymes because the incubation of GA with GSH in the absence of HLMs produced more GAH2 than the incubation in the presence of HLMs. In contrast, reduced GSH did not enhance the formation of 17-AAGH2 and 17-DMAGH2 in HLMs. Less than 1% of 17-AAGH2 and 17-DMAGH2 formed in HLM incubations. This result agreed well with a previous study (Lang et al., 2007) in which only 2% of 17-AAGH2 were detected after incubation in HLMs for 60 min. The reason for different formation rates of GAH2 and 17-AAGH2 in HLMs has been discussed: a 17-allylamino group of 17-AAG is a stronger electron-donating substituent than the 17-methoxy of GA, leading to a more negative shift in one-electron redox potential for 17-AAG. As a result, 17-AAG is not as easily reduced as GA (Lang et al., 2007). Likewise, 17-dimethylaminoethylamino is also a stronger electron-donating substituent than the 17-methoxy, resulting in the slower formation of 17-DMAGH2 than GAH2. With the tight binding between GAH2 and Hsp90, the tendency to get reduced might result in a higher nonselective toxicity of GA than that of 17-AAG and 17-DMAG.

GA, 17-AAG, and 17-DMAG were reported to react chemically with GSH to form GSH conjugates at 19-position (Cysyk et al., 2006). However, in the current study, only 0.33% of 17-DMAG-SG was formed in the incubation of 17-DMAG with GSH in the presence of HLMs, and 1.16% of 17-DMAG-SG was formed in the absence of HLMs. The presence of proteins in the incubation might decrease GSH conjugation. Consistent with previous in vitro and in vivo studies (Egorin et al., 1998; Lang et al., 2007), no significant amount of GSH conjugates of 17-AAG was detected. Likewise, the GSH conjugates of GA were not detected in the presence and absence of HLMs.

Quinone/hydroquinone cycling was proposed to be the primary metabolism pathway of GA and its analogs (Guo et al., 2008). NADPH-dependent redox cycling rates of GA analogs were determined by measuring the oxygen consumption rates: 17-DMAG > GA > 17-AAG; however, the GSH conjugation rates are as follows: GA > 17-DMAG > 17-AAG. However, these findings were inconsistent with the rapid in vitro and in vivo metabolism of 17-AAG (Egorin et al., 2001; Musser et al., 2003).

Our study showed that benzoquinone reduction and GSH conjugation were not the primary metabolism pathways of 17-DMAG in HLMs. This observation was supported by in vivo metabolism data of 17-DMAG in rats (Egorin et al., 2002). In the study, 11 compounds with UV spectra similar to that of 17-DMAG were detected in rat bile after intravenous administration of 17-DMAG. Although the structures of the 11 compounds were not identified, the metabolism did not involve the alterations to the benzoquinone moiety because their UV spectra were not altered compared with 17-DMAG. Among the 11 metabolites, 4 had Mr of 633 and 2 had Mr of 603, which were also detected in our HLM incubations.

The results of metabolite identification revealed that the biotransformation of 17-DMAG by CYP450 enzymes differed from that of 17-AAG. The extensive oxidative metabolism of 17-AAG on the 17-allylamino side chain resulted in the formation of 17-AG as well as epoxide and diol metabolites (Egorin et al., 1998). The hydroxylation on C22 and demethylation from methoxy moieties were detected as the minor biotransformation pathways of 17-AAG in HLMs (Lang et al., 2007). In contrast, the hydroxylation on C23 (M1) and demethylation from methoxy moieties (M3 and M4) were the primary biotransformation of 17-DMAG in HLMs (Fig. 8). Similar to 17-AAG, the hydroxylation of the adjacent carbon to the nitrogen generated a stable carbinolamine intermediate (M2). Subsequently, M2 underwent the loss of H2O and formed an imine metabolite (M5). In general, carbinolamines are known as unstable intermediates in N-demethylation. Most carbinolamines tend to form imine intermediates by the loss of water or decompose into dealkylated amine by the loss of aldehyde or ketone. However, stable carbinolamines and imines exist in aqueous solutions under some specific circumstances. For example, carbinolamines and imines can be stabilized by adjacent electron-withdrawing groups (Upthagrove and Nelson, 2001). Furthermore, a stable carbinolamine, hydroxymethylpentamethylmelamine, was detected as a metabolite in mouse plasma (Abikhalil et al., 1986) and rat liver microsomes (Ames et al., 1983) because the adjacent substituents delocalize the lone pair nitrogen electrons and stabilize the carbinolamine. In addition, stable carbinolamines have been isolated or detected as metabolites to a number of drugs including benzamides (Huizing et al., 1980), carbamates (Dorough and Casida, 1964), procarbazine (Weinkam and Shiba, 1978), N-methylcarbazole (Ebner et al., 1991), medazepam (Schwartz and Kolis, 1972), verapamil (Walles et al., 2002), pyrimidinyl-pyridopyrazines (Prakash and Soliman, 1997), and ecabapide (Fujimaki et al., 1995). Quinone is known as an electron-withdrawing group by resonance (Hiramatsu et al., 1983). The presence of the quinone adjacent to the nitrogen in M2, M5, and M6 of 17-DMAG allows the delocalization of the lone pair nitrogen electrons and stabilizes the metabolites.

17-AAG was a known substrate of CYP3A4 (Lang et al., 2007). In the current study, the reaction phenotyping experiments using recombinant human P450s and selective P450 chemical inhibitors revealed that CYP3A4 and CYP3A5 were the major P450 isozymes responsible for the oxidative metabolism of 17-DMAG in HLMs. In addition, CYP2C8, 2D6, 2A6, 2C19, and 1A2 were observed to contribute to the formation of M6. On the basis of the identified metabolite profiles, the in vitro oxidative biotransformation pathways for 17-DMAG in HLMs were proposed (Fig. 9).

Fig. 9.
Summary of in vitro oxidative metabolism pathways of 17-DMAD in HLMs.

In conclusion, this study tentatively characterized six oxidative metabolites of 17-DMAG in HLMs and revealed that the primary biotransformations of 17-DMAG in HLMs were hydroxylation and demethylation on its side chains. CYP3A4 and CYP3A5 were identified as the major P450 isozymes responsible for the oxidative metabolism of 17-DMAG in HLMs. The metabolic data generated in the current study have important implications for the discovery of more metabolically stable and less toxic GA analogs for the treatment of solid tumors.

This work was supported in part by the National Institutes of Health National Cancer Institute [Grants R01-CA120023, R21-CA143474]; and the University of Michigan Cancer Center Research [Grant Munn].

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

doi:10.1124/dmd.110.036418.

ABBREVIATIONS:

Hsp90
90-kDa heat shock protein
17-AAG
17-(allylamino)-17-demethoxygeldanamycin
17-AAG-SG
19-glutathionyl 17-(allylamino)-17-demethoxygeldanamycin
17-AAGH2
17-(allylamino)-17-demethoxygeldanamycin hydroquinone
17-AG
17-aminogeldanamycin
17-DMAG
17-(dimethylaminoethylamino)-17-demethoxy geldanamycin
17-DMAG-SG
19-glutathionyl 17-(dimethylaminoethylamino)-17-demethoxy geldanamycin
17-DMAGH2
17-(dimethylaminoethylamino)-17-demethoxygeldanamycin hydroquinone
EPI
enhanced product ion
GA
geldanamycin
GAH2
geldanamycin hydroquinone
HLM
human liver microsome
HPLC
high-performance liquid chromatography
LC-MS/MS
liquid chromatography-tandem mass spectrometry
Mr
protonated molecular ion
MRM
multiple reaction monitoring
MS
mass spectrometry
P450
cytochrome P450.

Authorship Contributions

Participated in research design: Wang.

Conducted experiments: Sun.

Contributed new reagents or analytic tools: Wang.

Performed data analysis: Sun, Zheng, and Zou.

Wrote or contributed to the writing of the manuscript: Zheng and Zou.

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