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Published in final edited form as: Toxicol In Vitro. 2019 Oct 16;62:104669. doi: 10.1016/j.tiv.2019.104669

Apoptosis contributes to the cytotoxicity induced by amodiaquine and its major metabolite N-desethylamodiaquine in hepatic cells☆

Yangshun Tang 1, Qiangen Wu 1, Frederick A Beland 1, Si Chen 1, Jia-Long Fang 1,*
PMCID: PMC12376074  NIHMSID: NIHMS2105837  PMID: 31629065

Abstract

Amodiaquine (ADQ), an antimalarial drug used in endemic areas, has been reported to be associated with liver toxicity; however, the mechanism underlying its hepatoxicity remains unclear. In this study, we examined the cytotoxicity of ADQ and its major metabolite N-desethylamodiaquine (NADQ) and the effect of cytochrome P450 (CYP)-mediated metabolism on ADQ-induced cytotoxicity. After a 48-h treatment, ADQ and NADQ caused cytotoxicity and induced apoptosis in HepG2 cells; NADQ was slightly more toxic than ADQ. ADQ treatment decreased the levels of anti-apoptotic Bcl-2 family proteins, which was accompanied by an increase in the levels of pro-apoptotic Bcl-2 family proteins, indicating that ADQ-induced apoptosis was mediated by the Bcl-2 family. NADQ treatment markedly increased the phosphorylation of JNK, extracellular signal-regulated kinase (ERK1/2), and p38, indicating that NADQ-induced apoptosis was mediated by MAPK signaling pathways. Metabolic studies using microsomes obtained from HepG2 cell lines overexpressing human CYPs demonstrated that CYP1A1, 2C8, and 3A4 were the major enzymes that metabolized ADQ to NADQ and that CYP1A2, 1B1, 2C19, and 3A5 also metabolized ADQ, but to a lesser extent. The cytotoxicity of ADQ was increased in CYP2C8 and 3A4 overexpressing HepG2 cells compared to HepG2/CYP vector cells, confirming that NADQ was more toxic than ADQ. Moreover, treatment of CYP2C8 and 3A4 overexpressing HepG2 cells with ADQ increased the phosphorylation of JNK, ERK1/2, and p38, but not the expression of Bcl-2 family proteins. Our findings indicate that ADQ and its major metabolite NADQ induce apoptosis, which is mediated by members of the Bcl-2 family and the activation of MAPK signaling pathways, respectively.

Keywords: Amodiaquine, Cytotoxicity, Cytochrome P450, Apoptosis, Metabolism

1. Introduction

Malaria is a major, potentially fatal, disease in developing countries that leads to approximately 2 million deaths every year (Snow et al., 2005). Amodiaquine (ADQ), an antimalarial drug, has been widely used in endemic areas for > 50 years, especially for the treatment against chloroquine-resistant isolates of Plasmodium falciparum (Watkins et al., 1984). Although ADQ has high efficacy compared to other 4-amino-quinoline antimalarial drugs, the risk of hepatotoxicity and idiosyncratic agranulocytosis diminishes its therapeutic advantages (Adjei et al., 2009; Larrey et al., 1986; Neftel et al., 1986). These potential side effects of ADQ have led to the withdrawal of ADQ in several countries and the prohibition of its prophylactic use. However, the drug is still widely used in Africa and Asia where chloroquine resistance has increased.

The clinical use of ADQ is associated with hepatotoxicity, and this adverse effect has been attributed to the bioactivation of the drug to a quinone imine metabolite (Jewell et al., 1995). Oxidative stress has been suggested to play a role in the hepatotoxicity induced by ADQ due to the induction of redox cycling by the quinone imine metabolite (Park et al., 1994; Tafazoli and O’Brien, 2009). ADQ also caused hepatotoxicity in a GSH-depleted mouse model, which suggests that hepatic GSH levels and covalent binding may be related to hepatic injury (Shimizu et al., 2009). However, the exact mechanism of the hepatotoxicity remains to be elucidated. Understanding the mechanism of the hepatoxicity caused by ADQ may help characterize the risk factors and aid in the protection of susceptible patients against ADQ toxicity. Thus, it is important to evaluate the cytotoxicity induced by ADQ as well as understanding the underlying cellular mechanisms.

After oral administration, ADQ is rapidly absorbed and extensively metabolized to its major metabolite N-desethylamodiaquine (NADQ) (Laurent et al., 1993). Other minor metabolites include N-bisdesethylamodiaquine and 2-hydroxydesethylamodiaquine (Churchill et al., 1985; Mount et al., 1986). Both ADQ and NADQ have antimalarial activity, with the activity of ADQ being about three-fold more potent than that of NADQ. NADQ has a 20- to 50-fold longer half-life (t1/2) than ADQ; as such, the area-under-curve of NADQ is approximately 100- to 240-fold higher than ADQ (Laurent et al., 1993; Scarsi et al., 2014). Therefore, the antimalarial activity of ADQ may be mainly determined by NADQ (Churchill et al., 1985).

The main enzyme responsible for the conversion of ADQ to NADQ is cytochrome P450 (CYP) 2C8 (Li et al., 2002). It has been reported that CYP1A1 and CYP1B1 convert ADQ and NADQ to the aldehyde metabolite 5-[(7-chloro-4-quinolinyl)amino]2-hydroxybenzaldehyde (AADQ) (Johansson et al., 2009). CYP3A4, 2C9, and 2D6 have also been shown to be involved in the metabolism of ADQ (Zhang et al., 2017). The administration of ketoconazole (a CYP3A4 inhibitor) to rats reduced the biliary excretion of ADQ by 50% and decreased the amount of drug irreversibly bound to liver proteins (Jewell et al., 1995). This indicates that CYP enzymes played a role in the bioactivation of ADQ to a reactive metabolite that conjugates with glutathione and binds to proteins. As the expression levels and activities of hepatic human CYPs vary highly among different populations (Achour et al., 2014; Zanger et al., 2014), the overexpression of bioactivating human CYPs may be an important risk factor determining the susceptibility to the cytotoxicity induced by ADQ. Although the P450-dependent bioactivation of ADQ by human liver microsomes has been reported (Zhang et al., 2017), the impact of specific human CYPs on the cytotoxicity of ADQ has not been established.

In this study, we examined the cytotoxicity of ADQ and its major metabolite NADQ in human hepatoma HepG2 cells. In addition, we investigated the metabolism of ADQ by human CYPs. The metabolic activity of each CYP isoform toward ADQ was examined using mass spectrometry and the effect of human CYP-mediated metabolism of ADQ on its cytotoxicity in HepG2 cells was evaluated.

2. Materials and methods

2.1. Chemicals and reagents

ADQ, NADQ, [diethyl-d10]ADQ, and [ethyl-d5]NADQ were purchased from Toronto Research Chemicals (North York, Ontario, Canada). Dimethyl sulfoxide (DMSO), Williams’ Medium E, thiazolyl blue tetrazolium bromide (MTT), ketoconazole, and rifampicin were obtained from Sigma-Aldrich (St. Louis, MO). Methanol, penicillin–-streptomycin solution, and 2.5% trypsin were purchased from Thermo Fisher Scientific, Inc. (Pittsburgh, PA). Fetal bovine serum (FBS) was acquired from Atlanta Biologicals (Lawrenceville, GA).

2.2. Antibodies

Rabbit monoclonal antibodies against caspase-3, cleaved caspase-3, Bcl-2, Mcl-1, Bax, JNK, phospho-JNK (Thr 183/Thr 185), ERK1/2, phospho-ERK1/2 (Thr 202/Tyr 204), p38, phospho-p38 (Thr 180/Tyr 182), and anti-rabbit IgG HRP-linked antibody were purchased from Cell Signaling Technology (Danvers, MA). Mouse monoclonal antibodies against human CYP3A4 and β-actin, and anti-mouse IgG HRP-linked antibody were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Goat polyclonal antibody against human CYP2C8 was purchased from Cell Signaling Technology (Danvers, MA).

2.3. Cell culture

The human hepatoma cell line HepG2 was obtained from the American Type Culture Collection (ATCC, Manassas, VA). HepG2 cells overexpressing human CYP isoforms 1A1, 1A2, 1B1, 2A6, 2A7, 2A13, 2B6, 2C8, 2C9, 2C18, 2C19, 2D6, 2E1, 3A4, 3A5, 3A7, 4A11, and 4B1 were established previously (Wu et al., 2017). The cells were cultured in Williams’s Medium E supplemented with 10% FBS and penicillin–-streptomycin at 37 °C in a humidified atmosphere containing 95% air and 5% CO2. The cells were seeded at a concentration of 2.5 × 105 cells/ml in a volume of 100 μl/well in 96-well plates; 1 × 105 cells/ml in a volume of 1 ml/well in 24-well plates; and 0.5 × 106 cells/ml in a volume of 2 ml/well in 6-well plates. Cells were cultured for 24 h prior to treatment with the indicated concentrations of ADQ, NADQ, or the vehicle (0.1% methanol) for 48 h.

2.4. MTT cell viability assay

An MTT reduction assay was used to measure the cell viability. Briefly, cells were cultured and treated with ADQ or NADQ in 24-well plates. After 48 h, the cells were incubated for 4 h with fresh culture medium containing 1 mg/ml MTT. The resulting formazan was dissolved in DMSO, and the absorbance at 540 nm was determined with a Synergy 2 Microplate Reader (BioTek, Winooski, VT). The IC50 values were obtained from the cell growth curves using GraphPad Prism 6.0.

2.5. Lactate dehydrogenase (LDH) assay

Cytotoxicity was measured with an LDH assay. Briefly, after treatment with ADQ or NADQ in 96-well plates for 48 h, 6 μl of cell-free supernatant from each well was transferred into a new clear 96-well plate. Then, 10 μl of 10% Triton X-100 was added to the treated cells. Following a 1-h lysis, 10 μl of lysates was transferred into empty wells of the clear 96-well plate containing the corresponding supernatant. Then, 230 μl of reaction buffer (203.3 mM NaCl, 81.3 mM Tris, 0.2 mM NADH, and 1.7 mM monosodium pyruvate, pH 7.2) was added to the wells containing the supernatants or cell lysates. Absorption was measured at 340 nm for 5 min at 1-min intervals using a BioTek Synergy 2 Microplate Reader. LDH release was calculated by the equation: percent cytotoxicity = 100 × (decrease in supernatant absorption/decrease in cell lysate absorption).

2.6. Caspase-3/7 activity measurement

The caspase-3/7 activity was examined using a Caspase-Glo® 3/7 assay system (Promega, Madison, WI). The induction of caspase-3/7 activity was calculated by comparing the luminescence of the treated cells to that of the vehicle control.

2.7. Western blot analysis

After treatment, cells were trypsinized and washed twice in PBS. Approximately 107 cells were lysed in RIPA buffer with a complete protease inhibitor cocktail (Roche Diagnostics, Basel, Switzerland) for 30 min on ice and centrifuged at 12,000g for 20 min at 4 °C. The supernatants were collected, and total protein concentrations were measured using BCA protein assay kits (Pierce, Rockford, IL). The tissue lysates (20–60 μg total protein) were subjected to electrophoresis in a 10% SDS-PAGE gel. The resolved proteins were electrophoretically transferred onto a PVDF membrane (Bio-Rad, Hercules, CA). Both electrophoresis and blotting were performed with a Bio-Rad Mini-PROTEAN® 3 electrophoresis system. Blots were blocked with 5% milk in PBS/Tween-20 and incubated with specific primary antibodies against caspase-3 (1:1000), Bcl-2 (1:1000), Mcl-1 (1:1000), Bax (1:1000), JNK (1:1000), phospho-JNK (Thr 183/Thr 185) (1:1000), ERK1/2 (1:1000), phospho-ERK1/2 (Thr 202/Tyr 204) (1:1000), p38 (1:1000), phospho-p38 (Thr 180/Tyr 182) (1:1000), CYP2C8 (1:1000), CYP3A4 (1:1000), or β-actin (1:2000), followed by a secondary antibody conjugated to horseradish peroxidase. The blots were visualized by chemiluminescence using an ECL detection kit (Millipore Corporation, Billerica, MA) and quantified with a FluorChem R System (ProteinSimple, San Jose, CA). β-Actin was used as the loading control. All primary antibodies were incubated with the same membrane after consecutive stripping using Restore Western Blot Stripping Buffer (Pierce).

2.8. Preparation of microsomes from HepG2 cells and HepG2/CYP-overexpressing cell lines

For each cell line, 2 × 108 cells were harvested, suspended in 6 ml of ice-cold microsome preparation buffer (250 mM sucrose, 10 mM Tris, pH 7.4), and sonicated three times for 10 s each. The homogenate was centrifuged at 10,000g for 10 min at 4 °C to remove debris and large organelles, and then the supernatant was centrifuged at 100,000g for 60 min at 4 °C. The pellet was resuspended in 1 ml of microsome preparation buffer, and aliquots were stored at −80 °C. The protein concentration of the microsomal preparation was determined using a Pierce BCA protein assay kit.

2.9. Metabolism of ADQ by microsomes isolated from HepG2/CYP-overexpressing cells using HPLC-MS/MS spectrometry

Metabolism of ADQ by microsomes isolated from HepG2/CYP-overexpressing cells was assayed in a 125-μl final reaction volume by incubating microsomes (1.0 mg of microsomal protein) with 0.8 mM ADQ, 4 mM MgCl2, 2 mM NADPH, and 100 mM potassium phosphate buffer (pH 7.2) at 37 °C for 30 min with gentle shaking. Negative controls for the assay included incubations conducted with microsomes from parent HepG2 cells and HepG2/vector cells, as well as incubations conducted in the absence of ADQ or NADPH. At the end of the incubation, 10 μl from the reactions was collected and added into 490 μl of ice-cold acetonitrile containing 200 ng/ml [diethyl-d10]ADQ, which was used as internal standard for ADQ, and 200 ng/ml [ethyl-d5]NADQ, which was used as internal standard for NADQ and AADQ. The samples were vortexed briefly, centrifuged at 14,000 rpm for 5 min at room temperature, and the supernatant was analyzed by HPLC-mass spectrometry, using a Shimadzu Prominence HPLC coupled with an AB Sciex 3200 QTRAP mass spectrometer (Foster City, CA). Ten μl of the final sample was injected onto a Waters Atlantis T3 C18 column (4.6 × 150 mm, 5 μm particle size) at 40 °C using a gradient between solvent A (10 mM ammonium acetate containing 0.1% formic acid) and solvent B (acetonitrile containing 0.1% formic acid) at a flow rate of 0.5 ml/min. The gradient started with 25% solvent B for 0.5 min, followed by a linear gradient of 25 to 90% solvent B in 9.5 min, then returning solvent B to 25% in 0.5 min, and maintained for 5 min to equilibrate the column. The eluate was monitored with a QTRAP mass spectrometer in the positive electrospray mode using multiple reaction monitoring of the following transitions: [M + H]+ m/z 356.1 to 283.1 for ADQ, m/z 366.2 to 283.1 for [diethyl-d10]ADQ, m/z 328.1 to 283.1 for NADQ, m/z 333.0 to 283.1 for [ethyl-d5]NADQ, and m/z 299.0 to 269.1 for AADQ. The experimental parameters were optimized as follow: curtain gas 40, ion spray voltage 4000 V, temperature 600 °C, ion source gas 1 80, ion source gas 2 80, CAD gas medium, declustering potential 60, entrance potential 6.5, collision energy 30 V, collision cell entrance potential 40, and collision cell exit potential 4. Analytes were quantified using linear regression, with calibration curves ranging from 1.6 to 1000 ng/ml in Analyst 1.6 software. The enzymatic activities of CYPs are expressed as pmol metabolite/mg protein/min.

2.10. ADQ metabolites in HepG2 cells overexpressing CYP2C8 and 3A4

After 48 h of exposure to ADQ at the IC50, the culture medium from HepG2/CYP vector cells and HepG2 cells overexpressing CYP2C8 and 3A4 was collected and the cells were harvested. Cell lysates were obtained by adding 50 μl Nanopure water to cell pellet, followed by three cycles of freezing and thawing. The protein concentration of the lysate was determined using a Pierce BCA protein assay kit. The cell culture medium or cell lysate was then diluted with four volumes of ice-cold acetonitrile containing 200 ng/ml of the internal standards ADQ-d10 and NADQ-d5, vortexed, and centrifuged at 14,000 rpm for 5 min at room temperature. The supernatant was analyzed by HPLC–MS/MS mass spectrometry as described above.

2.11. Statistical analyses

Data are presented as the mean ± standard deviation of three independent experiments. Statistical analyses were performed using GraphPad Prism 6.0. Two-way ANOVA (analysis of variance) with Dunnett’s post hoc test was used to compare the treatments of ADQ and NADQ at the indicated concentrations in HepG2 cells. One-way ANOVA with Tukey’s post hoc test was used to compare the HepG2/CYP Vector cell line and the CYP overexpressing HepG2 cell lines. Comparison between CYP overexpressing HepG2 cell lines with and without the CYP-specific inhibitor was made by Student’s t-test. The results were considered significant at p < .05.

3. Results

3.1. ADQ and its major metabolite NADQ caused cytotoxicity

To assess whether ADQ and NADQ caused cytotoxicity in HepG2 cells, cells were treated with ADQ or NADQ at concentrations ranging from 1.56 to 50 μM for 48 h. An MTT assay showed that ADQ and NADQ induced cytotoxicity. The values of IC50 of ADQ and NADQ were 17.4 and 15.0 μM, respectively (Table 1), indicating that ADQ is slightly less cytotoxic to HepG2 cells than its major metabolite NADQ.

Table 1.

The half inhibitory concentration (IC50) of ADQ and NADQ on cell growth.a

IC50 (μM) 95% confidence intervals (μM)
ADQ 17.4* 16.8–18.0
NADQ 15.0 14.1–15.9
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a

HepG2 cells were incubated with various concentrations of ADQ or NADQ (1.56–50 μM) for 48 h. The IC50 values were obtained from the cell growth curves, using GraphPad Prism 6.0. Data are presented as the mean IC50 of three independent experiments and 95% confidence intervals of the IC50.

*

Significantly (p < .05) different from cells exposed to NADQ.

To evaluate whether the decrease in cell viability was due to necrotic cell death, an LDH release assay was used. LDH release was increased in a concentration-dependent manner following the ADQ and NADQ treatment (Fig. 1a). NADQ treatment at the concentrations above the 0.5 IC50 caused > 4-fold release of LDH than the vehicle control. ADQ treatment at the IC50 induced approximate a 3-fold greater release of LDH than the vehicle control. These results indicated that the cytotoxicity induced by ADQ and NADQ is due partly to necrosis and that NADQ led to more necrotic cell death than ADQ.

Fig. 1.

Fig. 1.

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ADQ and NADQ induced cell death and apoptosis in HepG2 cells. HepG2 cells were treated with ADQ and NADQ at the indicated concentrations for 48 h. The cells were then analyzed for (a) LDH release and (b) caspase-3/7 activity. (c) Immunoblotting for the cleaved caspase-3 and β-actin using whole-cell lysates. The intensity of each band was quantified, and the relative protein levels were calculated using β-actin as the internal reference. (d) Relative level of the cleaved caspase-3. Data are presented as the mean ± standard deviation. *Significantly (p < .05) different from the vehicle control. ^Significant (p < .05) difference between ADQ and NADQ treatment. #Significant (p < .05) concentration-related trend.

To determine whether apoptosis was induced after treatment with ADQ or NADQ, cells were treated with the 0.1 IC50, 0.5 IC50, and IC50 of ADQ or NADQ for 48 h, and apoptosis was analyzed. As shown in Fig. 1b, exposure of HepG2 cells to ADQ and NADQ increased the activity of caspase-3/7 in a concentration-dependent manner. At the IC50, NADQ produced greater caspase-3/7 activity than ADQ. An increase in the expression level of cleaved caspase-3 was also observed following the treatment of ADQ and NADQ (Fig. 1c and d). These results showed that ADQ and NADQ induced apoptosis in HepG2 cells, with the extent being greater with NADQ.

3.2. Bcl-2 family proteins contributed to the apoptosis induced by ADQ

Apoptosis is a highly regulated form of programmed cell death in response to predeath signaling pathways. Several signaling pathways have been identified that are involved in the initiation and amplification of apoptosis, and most of them are caspase-dependent pathways (Xu and Shi, 2007). To determine which pathways are involved in the apoptosis induced by ADQ and NADQ, the expression levels of the Bcl-2 family proteins, which control a key step in the apoptosis pathway by governing permeabilization of the mitochondrial outer membrane (Shamas-Din et al., 2013), were measured. Treatment of ADQ for 48 h decreased the expression levels of the anti-apoptotic members Bcl-2 and Mcl-1 and increased the expression level of the pro-apoptotic member Bax, whereas no change was observed after the treatment of NADQ (Fig. 2). These data indicated that the Bcl-2 family-mediated apoptotic pathway was involved in the apoptosis induced by ADQ.

Fig. 2.

Fig. 2.

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Involvement of the Bcl-2 family proteins and the MAPK signaling pathways in apoptosis induced by ADQ and NADQ in HepG2 cells. Western blotting of Bcl-2, Mcl-1, Bax, and the phosphorylation of p38, JNK, ERK1/2, and β-actin using whole-cell lysates from HepG2 cells treated with ADQ and NADQ at the concentrations indicated for 48 h. The intensity of each band was quantified, and the relative protein levels were calculated using β-actin as the internal reference. (a) Immunoblotting for the proteins. Relative levels of the proteins in HepG2 cells treated with (b) ADQ and (c) NADQ. Data are presented as the mean ± standard deviation. *Significantly (p < .05) different from the vehicle control.

3.3. Involvement of MAPK pathways in the apoptosis induced by NADQ

Mitogen-activated protein kinase (MAPK) pathways have been shown to be involved in the apoptotic process (Chang and Karin, 2001). MAPKs has been reported to respond to chemical and physical stresses (Pearson et al., 2001), thereby controlling cell survival and cell death. Mammals express three major groups of MAPKs: extracellular signal-regulated kinase (ERK1/2), Jun amino-terminal kinases (JNK), and p38 proteins (Raman et al., 2007). ERK1/2 stimulates DNA synthesis and modulates cell proliferation and cell survival, whereas JNK and p38 are linked to the induction of apoptotic cell death (Cai et al., 2006; Lu and Xu, 2006). To investigate whether ADQ or NADQ induced apoptosis through the MAPK signaling pathways, the expression levels of p38, JNK, and ERK1/2 were measured. As shown in Fig. 2, treatment of NADQ for 48 h markedly increased the phosphorylation of p38, JNK, and ERK1/2, whereas no change was observed after the treatment of ADQ. These data indicate that the MAPK signaling pathways are involved in the apoptosis induced by NADQ.

3.4. The effect of CYP-mediated metabolism on the ADQ-induced cytotoxicity

As human CYPs can be important risk factors in determining the susceptibility to the cytotoxicity induced by ADQ, we investigated the ability of 18 human CYPs to metabolize ADQ. ADQ was incubated with microsomes isolated from HepG2/CYP-overexpressing cells, and its major metabolite NADQ and a minor metabolite AADQ were quantified by HPLC-MS/MS mass spectrometry. As shown in Table 2, fourteen CYPs (CYP1A1, 1A2, 1B1, 2A6, 2A13, 2B6, 2C8, 2C18, 2C19, 2D6, 2E1, 3A4, 3A5, and 3A7) metabolized ADQ to NADQ, with CYP1A1, 2C8, and 3A4 having the greatest metabolic activity. Among the CYPs, five (CYP1A1, 1B1, 2D6, 3A4, and 3A5) metabolized ADQ to NADQ and AADQ, with CYP1A1 and 1B1 having the greatest metabolic activity.

Table 2.

Metabolism of ADQ by individual human CYPs.a

pmol metabolite/mg microsomal protein/min
NADQ AADQ
HepG2 0.74 ± 0.05 0.32 ± 0.04
CYP vector 1.91 ± 0.01 0.24 ± 0.08
CYP1A1 58.9 ± 1.90* 54.9 ± 2.10*
CYP1A2 11.2 ± 0.42* 0.54 ± 0.01
CYP1B1 15.4 ± 0.32* 13.0 ± 0.71*
CYP2A6 3.66 ± 0.28* 0.21 ± 0.02
CYP2A7 0.26 ± 0.08 0.19 ± 0.01
CYP2A13 3.73 ± 0.05* 0.17 ± 0.01
CYP2B6 2.71 ± 0.19* 0.23 ± 0.04
CYP2C8 106 ± 6.90* 0.27 ± 0.02
CYP2C9 0.68 ± 0.04 0.25 ± 0.03
CYP2C18 2.33 ± 0.13* 0.24 ± 0.02
CYP2C19 11.9 ± 0.32* 0.17 ± 0.01
CYP2D6 7.76 ± 0.37* 0.39 ± 0.04*
CYP2E1 3.47 ± 0.11* 0.19 ± 0.02
CYP3A4 29.6 ± 1.90* 1.05 ± 0.02*
CYP3A5 14.5 ± 0.75* 0.79 ± 0.07*
CYP3A7 2.96 ± 0.12* 0.20 ± 0.03
CYP4A11 0.57 ± 0.13 0.16 ± 0.05
CYP4B1 0.98 ± 0.13 0.15 ± 0.02
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a

Metabolism of ADQ by microsomes isolated from HepG2 cells overexpressing human CYPs was assayed in a 125 μl final reaction volume by incubating microsomes (1.0 mg of microsomal protein) with 0.8 mM ADQ, 4 mM MgCl2, 2 mM NADPH, and 100 mM potassium phosphate buffer (pH 7.2) at 37 °C for 30 min with gentle shaking. The metabolites were analyzed using HPLC-MS/MS as described in Materials and Methods. Data are presented as the mean ± standard deviation.

*

Significantly (p < .05) different from the CYP vector cell line.

To investigate the effect of CYP-mediated metabolism of ADQ on its cytotoxicity, the cytotoxicity of ADQ in HepG2/CYP vector cells and HepG2 cells overexpressing CYP1A1, 1A2, 1B1, 2C8, 2C19, 3A4, and 3A5 was examined. After treatment for 48 h, the IC50 values of ADQ was increased in HepG2 cells overexpressing CYP1A1 and 1B1 and decreased in HepG2 cells overexpressing CYP2C8 and 3A4 when compared to the CYP vector cell line (Table 3). LDH release was elevated in HepG2 cells overexpressing CYP2C8 and 3A4 at the IC50 when compared to the CYP vector cell line (Fig. 3a). As shown in Fig. 3b, the activity of caspase-3/7 increased at the 0.5 IC50 and IC50 in HepG2 cells overexpressing CYP2C8 and 3A4. These results indicate that exposing of HepG2 cells overexpressing CYP2C8 and 3A4 to ADQ caused an increase in cytotoxicity.

Table 3.

IC50 values of ADQ in HepG2 cell lines overexpressing human CYPs.a

IC50 (μM) 95% confidence intervals (μM)
CYP vector 17.1 15.7–18.6
CYP1A1 26.4* 25.6–27.3
CYP1A2 18.7 17.2–20.2
CYP1B1 25.7* 24.0–27.6
CYP2C8 11.7* 10.1–13.4
CYP2C19 17.3 13.0–23.1
CYP3A4 14.0* 13.0–15.1
CYP3A5 17.1 15.4–18.9
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a

HepG2 cell lines overexpressing human CYPs were incubated with various concentrations of ADQ (1.56–50 μM) for 48 h. The IC50 values were obtained from the cell growth curves, using GraphPad Prism 6.0. Data are presented as the mean IC50 of three independent experiments and 95% confidence intervals of the IC50.

*

Significantly (p < .05) different from CYP vector cells.

Fig. 3.

Fig. 3.

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ADQ induces cell death in HepG2 cells overexpressing CYP2C8 and 3A4. Cells were treated with ADQ at the concentrations indicated for 48 h. The cells were analyzed for (a) LDH release and (b) caspase-3/7 activity. Data are presented as the mean ± standard deviation. *Significantly (p < .05) different from the CYP vector cells. #Significant (p < .05) concentration-related trend.

To evaluate whether the major metabolite NADQ was formed in HepG2 cells overexpressing CYP2C8 and 3A4 upon treatment with ADQ, the levels of NADQ in the culture media and the cells were measured by HPLC-MS/MS spectrometry. Exposing HepG2 cells overexpressing CYP2C8 and 3A4 to ADQ at the IC50 for 48 h led to the formation of NADQ in both the culture media and the cells (Fig. 4). These data demonstrate that ADQ is metabolized to NADQ in HepG2 cells overexpressing CYP2C8 and 3A4 and that the metabolism increases the cytotoxicity of ADQ.

Fig. 4.

Fig. 4.

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Metabolism of ADQ to NADQ in HepG2 cells overexpressing CYP2C8 and 3A4. Cells were treated with ADQ at the IC50 for 48 h. The levels of NADQ in (a) the culture media and (b) the cell lysate were analyzed by HPLC-tandem mass spectrometry. The relative levels of NADQ (fold of CYP vector cells) are presented as the mean ± standard deviation. *Significantly (p < .05) different from the CYP vector cell line.

3.5. MAPK pathway involved in the apoptosis induced by ADQ in CYP2C8 and 3A4 overexpressing cells

To investigate which apoptosis signaling pathway is involved in the apoptosis induced by ADQ in HepG2 cells overexpressing CYP2C8 and 3A4, the protein expression levels of the Bcl-2 family and MAPKs were examined. As shown in Fig. 5, treatment with ADQ for 48 h did not alter the expression levels of Bcl-2 or Mcl-1 or Bax in HepG2 cells overexpressing CYP2C8 and 3A4, while the phosphorylation of p38, JNK, and ERK1/2 was markedly increased. These results indicate that exposing HepG2 cells overexpressing CYP2C8 and 3A4 to ADQ mediated the MAPK signaling pathways.

Fig. 5.

Fig. 5.

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Involvement of MAPK signaling pathways in apoptosis induced by ADQ in HepG2 cells overexpressing CYP2C8 and 3A4. Western blotting of Bcl-2, Mcl-1, Bax, and the phosphorylation of p38, JNK, ERK1/2, and β-actin using whole-cell lysates from HepG2 cells overexpressing CYP2C8 and 3A4 treated with ADQ at the concentrations indicated for 48 h. The intensity of each band was quantified, and the relative protein levels were calculated using β-actin as the internal reference. (a) Immunoblotting for the proteins. Relative levels of the proteins in HepG2 cells overexpressing (b) CYP2C8 and (c) CYP3A4. Data are presented as the mean ± standard deviation. *Significantly (p < .05) different from the vehicle control.

3.6. Effect of CYP2C8 and 3A4 on the cytotoxicity of ADQ

To verify further the effect of CYP2C8 and 3A4 on the cytotoxicity of ADQ, the CYP2C8 inhibitor rifampicin and CYP3A4 inhibitor ketoconazole (Kajosaari et al., 2005) were co-incubated with ADQ. The CYP inhibitors at the concentration of 30 μM were not toxic to HepG2 cells overexpressing CYP2C8 and 3A4, and markedly reduced the expression levels of CYP2C8 and 3A4 (Fig. 6a and b).

Fig. 6.

Fig. 6.

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Effect of CYP2C8 and 3A4 on the cytotoxicity of ADQ. HepG2 cells overexpressing CYP2C8 and 3A4 were co-incubated with ADQ at the concentrations indicated with or without a CYP inhibitor (30 μM rifampicin or ketoconazole) for 48 h. The cells were then analyzed for the expression levels of CYP2C8 and 3A4 by (a) Western blotting, (b) relative levels of CYP2C8 and 3A4, (c, d) LDH release, and (e, f) caspase-3/7 activity. β-Actin was the loading control. Data are presented as the mean ± standard deviation. *Significantly (p < .05) different from the vehicle control. ^Significantly (p < .05) different from cells treated with the CYP inhibitor. #Significant (p < .05) concentration-related trend.

Following a 48-h co-treatment of ADQ with the CYP inhibitors, the IC50 values of ADQ in HepG2 cells overexpressing CYP2C8 and 3A4 were increased to the same value as in HepG2/CYP vector cells (Table 4). Moreover, co-treatment of ADQ with rifampicin and ketoconazole in HepG2 cells overexpressing CYP2C8 and 3A4 reversed the elevated necrotic cell death (Fig. 6c and d) and the activities of caspase3/7 (Fig. 6e and f) induced by ADQ. In addition, co-treatment of ADQ with rifampicin and ketoconazole in HepG2 cells overexpressing CYP2C8 and 3A4 decreased the expression levels of anti-apoptotic members Bcl-2 and Mcl-1 and increased the expression level of pro-apoptotic member Bax, but did not alter the phosphorylation of p38, JNK, or ERK1/2 (Fig. 7).

Table 4.

IC50 values of ADQ in HepG2 cell lines overexpressing CYP2C8 and 3A4.a

IC50 (μM) 95% confidence intervals (μM)
CYP Vector 18.9 16.3–22.0
CYP2C8 11.6 9.8–13.7
CYP2C8 + Rifampicin 17.9* 16.2–19.7
CYP3A4 14.8 13.0–16.9
CYP3A4 + Ketoconazole 19.6* 17.2–22.4
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a

HepG2 cell lines overexpressing CYP2C8 and 3A4 were incubated with various concentrations of ADQ (3.13–50 μM) in the presence or the absence of the CYP inhibitor (30 μM rifampicin or ketoconazole) for 48 h. The IC50 values were obtained from the cell growth curves, using GraphPad Prism 6.0. Data are presented as the mean IC50 of three independent experiments and 95% confidence intervals of the IC50.

*

Significantly (p < .05) different from cells treated without the CYP inhibitor.

Fig. 7.

Fig. 7.

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Effect of CYP2C8 and 3A4 on the apoptosis induced by ADQ. Western blotting of Bcl-2, Mcl-1, Bax, and the phosphorylation of p38, JNK, ERK1/2, and β-actin using whole-cell lysates from HepG2 cells overexpressing CYP2C8 and 3A4 treated with ADQ at the concentrations indicated for 48 h. The intensity of each band was quantified, and the relative protein levels were calculated using β-actin as the internal reference. (a) Immunoblotting for the proteins. Relative levels of the proteins in HepG2 cells overexpressing (c) CYP2C8 and (d) CYP3A4. Data are presented as the mean ± standard deviation. *Significantly (p < .05) different from the vehicle control.

4. Discussion

This study investigated the cytotoxicity of ADQ and its major metabolite NADQ and characterized the effect of CYP-mediated metabolism on ADQ-induced cytotoxicity by using CYP overexpressing HepG2 cell lines. After treating HepG2 cells for 48 h, ADQ and NADQ caused cytotoxicity and induced apoptosis, with NADQ being slightly more toxic than ADQ. ADQ activated the Bcl-2 family-mediated apoptotic pathway, whereas NADQ activated MAPK signaling pathways. Metabolic studies demonstrated that 14 human CYPs metabolized ADQ to NADQ, with CYP1A1, 2C8, and 3A4 being the major enzymes. Among the seven human CYPs with ADQ metabolic activity > 10 pmol metabolite/mg microsomal protein/min, the IC50 values of ADQ were decreased in HepG2 cells overexpressing CYP2C8 and 3A4. The overexpression of CYP2C8 and CYP3A4 in HepG2 cells increased the cytotoxicity induced by ADQ, caused no change in Bcl-2 family members, and activated the phosphorylation of p38, JNK, and ERK1/2.

The activation of caspase-3 initiates apoptotic damage and is an essential event during apoptosis, making it a main indicator for the analysis of apoptotic cells (Degterev et al., 2003). Thus, we measured the caspase-3/7 activity to investigate the induction of apoptosis by ADQ and NADQ. Treatment of ADQ and NADQ for 48 h increased the activity and the levels of the cleaved form of caspase-3 (Fig. 1b–d). The Bcl-2 protein family plays a vital role in the apoptotic pathway that senses cellular stress, oligomerizes in and permeabilizes the mitochondrial outer membrane, and activates the final effector caspases of apoptosis (Rodriguez et al., 2011; Shamas-Din et al., 2013). Treatment with ADQ but not NADQ increased the level of pro-apoptotic Bax and decreased the levels of anti-apoptotic Bcl-2 and Mcl-1, suggesting that the Bcl-2 family is responsible for the apoptosis induced by ADQ (Fig. 2a and b). MAPKs participate in signal transduction pathways that control intracellular events in response to extracellular chemical and physical stimulations (Cargnello and Roux, 2011). MAPK family members ERK1/2, JNK, and p38 are involved in the apoptotic pathway (Deng et al., 2003; Pearson et al., 2001). In this study, the role of MAPK signaling pathways was investigated in the apoptosis induced by ADQ and NADQ. Our findings indicated that NADQ, but not ADQ, activated ERK1/2, JNK, and p38 pathways (Fig. 2a and c). Similar results were observed in HepG2 cells overexpressing CYP2C8 and 3A4, further confirming the pathways involved in the apoptosis induced by NADQ, the major ADQ metabolite (Figs. 5 and 7). Although it has been reported that glutathione depletion and oxidative stress may be risk factors for the cytotoxicity induced by ADQ (Heidari et al., 2014; Shimizu et al., 2009; Tafazoli and O’Brien, 2009), our study demonstrates that the Bcl-2 family mediates the apoptosis induced by ADQ and that MAPK signaling pathways are involved in the apoptosis induced by NADQ.

CYP-dependent bioactivation of ADQ by human liver microsomes has been reported previously (Jurva et al., 2008; Zhang et al., 2017); nonetheless, the specific role of CYPs in the cytotoxicity induced by ADQ has not been examined. In this study, by using microsomes prepared from HepG2 cells overexpressing human CYPs, 14 human CYPs, including previous reported CYP2C8, 1A1, and 3A4, were able to mediate the conversion of ADQ to NADQ (Table 2). By using MTT and LDH release assays, our data indicated that the cytotoxicity of ADQ was increased in HepG2 cells overexpressing CYP2C8 and 3A4 and decreased in HepG2 cells overexpressing CYP1A1 and 1B1 (Table 3). The increase in cytotoxicity observed with CYP2C8 and 3A4 may be due to the metabolism of ADQ to NADQ, which is slightly more toxic than ADQ. The decrease in toxicity observed with CYP1A1 and 1B1 may be a consequence of the conversion of NADQ to AADQ. Since the Cmax of NADQ in vivo is > 20-fold higher than that of ADQ and the t1/2 of NADQ is much longer than ADQ (Lai et al., 2009), NADQ may be responsible for the adverse effects of ADQ in humans.

The largest contribution to bioactivation of ADQ in liver has been attributed to CYP2C8 and 3A4 (Zhang et al., 2017). Our study confirmed that CYP2C8 and 3A4 are involved in ADQ-induced cytotoxicity because cells with higher levels of CYP2C8 and 3A4 were sensitive to ADQ and co-treatment of ADQ with inhibitors of CYP2C8 and 3A4 significantly decreased the cytotoxicity induced by ADQ. A meta-analysis study by Achour et al. (Achour et al., 2014) has shown that there are variable levels of hepatic CYPs (up to 10-fold differences) and a significant positive correlation between the abundance of CYP2C8 and 3A4. Thus, individuals with high expression levels of hepatic CYP2C8 and/or 3A4 may have an increased risk of the adverse effects from ADQ.

Acknowledgements

Yangshun Tang was supported by an appointment to the Postgraduate Research Program in the Division of Biochemical Toxicology at the National Center for Toxicological Research administered by Oak Ridge Institute for Science Education through an inter-agency agreement between the U.S. Department of Energy and the U.S. Food and Drug Administration.

Footnotes

☆

The views presented in this article do not necessarily reflect those of the U.S. Food and Drug Administration.

Declaration of competing interest

The authors declare that there are no conflicts of interest.

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