Genotoxic and oxidative stress effects of 2-amino-9H-pyrido[2,3-b]indole in human hepatoma G2 (HepG2) and human lung alveolar epithelial (A549) cells.

http://informahealthcare.com/txm ISSN: 1537-6516 (print), 1537-6524 (electronic) Toxicol Mech Methods, Early Online: 1–11 ! 2015 Informa Healthcare USA, Inc. DOI: 10.3109/15376516.2015.1025345

RESEARCH ARTICLE

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Genotoxic and oxidative stress effects of 2-amino-9H-pyrido[2,3-b]indole in human hepatoma G2 (HepG2) and human lung alveolar epithelial (A549) cells Ting-Ting Zhang, Ge Zhao, Xiang Li, Fu-Wei Xie, Hui-Min Liu, and Jian-Ping Xie Key Laboratory of Tobacco Chemistry, Zhengzhou Tobacco Research Institute of CNTC, Zhengzhou, China

Abstract

Keywords

2-Amino-9H-pyrido[2,3-b]indole (AaC), which is present in high quantities in cigarette smoke and also in fried food, has been reported to be a probable human carcinogen. However, few studies have reported on the genotoxicity and oxidative stress induced by AaC. This study investigated the genotoxic effects of AaC in human hepatoma G2 (HepG2) and human lung alveolar epithelial (A549) cells using the comet assay. Significant increases in DNA fragment migration indicated that AaC causes serious DNA damage in HepG2 and A549 cells. The role of oxidative stress in the mechanism of AaC-induced genotoxicity was clarified by measuring the level of intracellular reactive oxygen species (ROS), the GSH/GSSG ratio and the formation of 8-hydroxydeoxyguanosine (8-OHdG), a marker of oxidative DNA damage. The results showed that the levels of ROS and 8-OHdG increased, whereas the GSH/GSSG ratio decreased. The concentration of 8-OHdG was positively related to DNA damage. Taken together, these results indicate that AaC can induce genotoxicity and oxidative stress and that AaC likely exerts genotoxicity in HepG2 and A549 cells through ROS-induced oxidative DNA damage. This is the first report to describe AaC-induced genotoxic and oxidative stress in HepG2 and A549 cells.

A549 cells, 2-amino-9H-pyrido[2,3-b]indole (AaC), genotoxic effects, HepG2 cells, oxidative stress

Introduction 2-Amino-9H-pyrido[2,3-b]indole (AaC) is a mutagenic and carcinogenic heterocyclic aromatic amine (HAA) that is present in fried or broiled food, cigarette smoke and diesel exhaust particles (Manabe et al., 1991a; Matsumoto et al., 1981; Robbana-Barnat et al., 1996; Turesky et al., 2005). This chemical was classified as a Group 2B carcinogen by a working group of the International Agency for Research on Cancer (IARC, 1993). Up to 258 ng AaC per cigarette was detected in mainstream tobacco smoke (Nauwelaers et al., 2011), a much greater amount than other HAAs such as 2-amino-1-methyl-6-phenylimidazo[4,5-b]pyridine (PhIP), present at 11–23 ng/cigarette (Manabe et al., 1991b); 3-amino-1,4-dimethyl-5H-pyrido[4,3-b]indole (Trp-P-1) and 3-amino-1-methyl-5H-pyrido[4,3-b]indole (Trp-P-2), present at 1–4 ng/cigarette (Zhao et al., 2014) and 2-amino-3methylimidazo[4,5-f]quinoline (IQ), present at less than 1 ng/cigarette (Yamashita et al., 1986). The difference in the levels of AaC in the urine is significantly different between smokers and non-smokers (Fu et al., 2014). Thus, cigarette Address for correspondence: Fu-Wei Xie, Key Laboratory of Tobacco Chemistry, Zhengzhou Tobacco Research Institute of CNTC, No. 2 Fengyang Street, High-Tech Zone, Zhengzhou 450001, China. Tel: +8637167672502. Fax: +8637167672055. E-mail: [email protected]

History Received 19 November 2014 Revised 16 February 2015 Accepted 1 March 2015 Published online 23 March 2015

smoking leads to significant exposure to AaC. However, the previous literature has mainly been focused on the toxicity of IQ-type HAAs, which are related to cooked meats (Dumont et al., 2010; Pezdirc et al., 2013; Sasaki et al., 1997). In contrast, fewer studies have investigated the genotoxic and carcinogenic properties of AaC. Thus, it is important to investigate the toxic effects of AaC. Oral administration of AaC has been reported to induce lymphomas and liver tumors in CDF1mice (Ohgaki et al., 1984), mutations in the colons of Big BlueÔ mice (Zhang et al., 1996) and DNA adduct formation in the livers of Sprague–Dawley rats (Snyderwine et al., 1998). AaC has also been reported to be mutagenic in a number of bacteria (Nagao et al., 1983; Pfau et al., 1999) and mammalian systems (Yoshimi et al., 1988) including Chinese hamster lung cells (IARC, 1986), and AaC induces chromosomal damage in Chinese hamster ovary cells and human lymphoblastoid (MCL-5) cells (IARC, 1986; Pfau et al., 1999). Many other HAAs such as IQ, PHIP and Trp-P-2 have been reported to induce oxidative stress, which may be a component of the toxicological mechanisms of the genotoxic effects of these compounds through increasing the production of free radicals and reactive oxygen species (ROS) and/or decreasing antioxidant defense mechanisms (Kato et al., 1996; KurzawaZegota et al., 2012; Trevisan et al., 2001; Wakata et al., 1985).

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Abbreviations: HepG2, human hepatoma G2 A549, human lung alveolar epithelial HAAs, heterocyclic aromatic amines AaC, 2-amino-9H-pyrido[2,3-b]indole IQ, 2-amino-3-methylimidazo[4,5-f]quinoline PhIP, 2-amino-1-methyl-6-phenylimidazo[4,5-b]pyridine Trp-P-1,3-amino-1,4-dimethyl-5H-pyrido[4,3-b]indole Trp-P-2,3-amino-1-methyl-5H-pyrido[4,3-b]indole ROS, reactive oxygen species 8-OHdG, 8-hydroxydeoxyguanosine GSH, reduced glutathione GSSG, oxidized glutathione dG, 20 -deoxyguanosine DCFH-DA, 20 ,70 -dichlorofluorescein diacetate DNase, Deoxyribonuclease PBS, phosphate-buffered saline MLC-5, human lymphoblastoid [13C115N2]-8-OH-G, [13C115N2]-8-hydroxydeoxy-guanosine OTM, Olive tail moment TM, tail moment D1-dG, 10 -deoxyguanosine monohydrate-10 -d TNB, 5-thio-2-nitrobenzoic acid DTNB, 50 ,50 -dithiobis-2-nitrobenzoic acid

However, the oxidative stress induced by AaC and its role in the functional mechanism of the genotoxicity of AaC remain unclear. In this study, human hepatoma G2 (HepG2) and human lung alveolar epithelial (A549) cells were used as in vitro models to investigate the genotoxic and oxidative stress effects of AaC and the possible mechanisms of those effects. HepG2 cells retain many characteristics of hepatocytes and could reflect the metabolism of xenobiotics in the human body better than other metabolically incompetent cells (Knasmu¨ller et al., 1998; Mersch-Sundermann et al., 2004; Pinti et al., 2002). They have been shown to be a suitable system for genotoxicity testing (Dearfield et al., 1983; Knasmuller et al., 2004). A549 cells are type II alveolar cells of the pulmonary epithelium, and have been widely used to assess the adverse effects of inhaled chemicals in toxicological and pharmaceutical studies (Castell et al., 2005). A549 and HepG2 cells have been used to investigate the genotoxicity of some xenobiotics in many studies (Jiang et al., 2007; Peng et al., 2013; Zou et al., 2009). As AaC is produced in high quantities in cigarette smoke and is believed to exert its genotoxicity through its metabolites N-acetoxyAaC and N-sulfonyloxy-AaC derivatives (King et al., 1999, 2000), A549 and HepG2 cells are considered excellent models to investigate the toxicity of AaC. An alkaline comet assay was applied to assess the genotoxicity of AaC. Simultaneously, the levels of intracellular reactive oxygen species (ROS) were detected by the 20 ,70 -dichlorofluorescein diacetate (DCFH-DA) assay. 8-Hydroxydeoxy-deoxyguanosine (8-OHdG), a reliable marker for ROS-induced oxidative DNA damage, was analyzed by HPLC-MS/MS. The intracellular GSH/GSSG

Toxicol Mech Methods, Early Online: 1–11

ratio, a known traditional marker of oxidative stress (Balendiran et al., 2004), was detected using a GSH/GSSG ratio assay kit.

Materials and methods Materials AaC, 8-OHdG, [13C115N2]-8-hydroxydeoxy-guanosine 13 15 ([ C1 N2]-8-OH-G), 20 -deoxyguanosine (dG) and 0 1 -Deoxyguanosine Monohydrate-10 -d (D1-dG) were obtained from Toronto Research Chemicals (Toronto, ON, Canada); Desferrioxamine mesylate salt and DCFH-DA were purchased from Sigma (St. Louis, MO); Dimethyl sulfoxide (DMSO) was obtained from Amresco (Solon, OH); All compounds were 498% pure. HPLC grade acetic acid, methanol and acetonitrile were purchased from J.T. Baker (Phillipsburg, NJ); Bovine pancreas deoxyribonuclease (DNase) and alkaline phosphatase were obtained from New England BioLabs (Ipswich, MA); The Blood and Cultured Cell DNA Extraction Kit was obtained from Epigentek (Farmingdale, NY); The GSH/GSSG Ratio Assay Kit was purchased from Sciencell (San Diego, CA); Low melting point agarose, normal melting point agarose and acridine orange were purchased from Solarbio (Beijing, China). Cell culture and treatment HepG2 (American Type Culture Collection (ATCC): HB-8065) and A549 cells (ATCC: CCL-185) were obtained from the Cell Bank of the Chinese Academy of Sciences (Shanghai, China) and cultured in RPMI 1640 nutrient mixture (Solarbio, Beijing, China) containing 15 or 10% fetal bovine serum (Gibco, Grand Island, NY), respectively, and cultured at 5% CO2/95% air at 37 C. According to experimental needs, cells were seeded on 6- and 96-well plates and 60-mm dishes (Corning, NY) and allowed to grow for 24 h (to approximately 80–90% confluence). The cells were then treated with vehicle (serum-free medium containing 0.1% DMSO) or different concentrations of AaC. Cytotoxicity measurements (neutral red assay) HepG2 and A549 cells were seeded on 96-well plates at a density of 1.5 104 cells/well. After 24 h incubation, cells were exposed to 10–100 mg/ml AaC or 0.1% DMSO for 24 h. At the end of the exposure, cells were washed twice with phosphate-buffered saline (PBS) and stained with 50 mg/ml neutral red solution for 3 h at 37 C, and then rapidly washed with a 1% formaldehyde solution and destained with 50% ethanol/1% acetic acid. The absorbance was measured at 540 nm with a microplate reader (Molecular Devices Spectra Max M5, Sunnyvale, CA). Comet assay The comet assay was carried out according to the protocol of Olive et al. (1992) with minor modifications. Briefly, 5 105 cells/well were seeded on six-well plates and incubated for 24 h, followed by treatment with different concentrations of AaC (5–20 mg/ml for HepG2 cells and 5–30 mg/ml for A549cells) or 0.1% DMSO (for control cells) for 24 h. At the end of the treatment, cells were washed with PBS and


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suspended in PBS at a concentration of 5 105 cells/ml. Cultures with greater than 80% cell viability were used to determine DNA damage. Twenty microliters cell suspensions were mixed with 180 ml of low melting point agarose (0.8%, w/v in PBS) and loaded onto a microscope slide pre-coated with 200 ml normal melting point agarose (1%, w/v in PBS). The slides were covered with a coverslip, and the agarose was solidified in a refrigerator at 4 C for 20 min. Thereafter, the coverslips were removed and the slides were immersed in cold fresh lysing solution (2.5 M NaCl, 100 mM Na2EDTA, 10 mM Tris, 1% sodium sarcosine, 1% triton X-100 and 10% DMSO, pH 10) for 2 h at 4 C. Subsequently, the slides were treated with cold fresh alkaline solution (1 mM Na2EDTA and 300 mM NaOH, pH 13) for 20 min to allow DNA unwinding, followed by electrophoresis at 20 V and 200 mA for another 20 min. After electrophoresis, the slides were bathed three times for 5 min in neutralizing solution (0.4 M Tris-HCl, pH 7.5). Then, the DNA was stained with 50 ml acridine orange solution (100 mg/ml) just prior to analysis. Finally, the slides were imaged by a fluorescence microscope (Olympus BX-51, Omachi, Japan) equipped with a blue excitation filter. At least 100 randomly selected cells were analyzed per sample using the Comet Assay Software Project CAPS-1.2.2 program (University of Wroclaw, Poland). Tail DNA (%), Olive tail moment (OTM), tail moment (TM) and tail length were used as the indicators of DNA damage. TM and OTM were calculated according to Olive et al. (Fenech, 2000; Olive et al., 1992). Measurement of intracellular ROS 20 ,70 -Dichlorofluorescein diacetate (DCFH-DA) is known as a highly selective fluorescent probe to detect the level of intracellular ROS (LeBel et al., 1992; Myhre et al., 2003). Briefly, HepG2 and A549 cells were seeded on 96-well plates at 1.5 104 cells/well and incubated for 24 h. Then, cells were loaded with 10 mM DCFH-DA for 1 h at 37 C in the dark. After washing away excess probe, the cells were treated with different concentrations of AaC (5–20 mg/ml for HepG2 cells and 5–30 mg/ml for A549 cells) or 0.1% DMSO (for control cells) for 2 h, and then washed three times with ice-cold PBS. The relative fluorescence intensity of the wells was measured by a microplate fluorescence reader (MD Spectra Max M5, Sunnyvale, CA) with an excitation wavelength of 488 nm and an emission wavelength of 525 nm. Measurement of the intracellular GSH/GSSG ratio Total cellular glutathione (GSH + GSSG) and oxidized glutathione (GSSG) were determined using a GSH/GSSG ratio Assay Kit. Briefly, HepG2 and A549 cells were seeded on six-well plates at 5 105 cells/well and incubated for 24 h. Then, cells were treated with different concentrations of AaC (5–20 mg/ml for HepG2 cells and 5–30 mg/ml for A549 cells), with 0.1% DMSO treatment as a control. The medium was removed after 2 h. Cells were washed twice with cold PBS and scraped into a micro centrifuge tube. Then, 40 ml 5% 5-sulfosalicylic acid was added to the cell pellet and vortexed. Samples were kept on ice for 5 min and then centrifuged at 10 000 rpm for 10 min at 4 C to remove insoluble material. Total glutathione in the supernatant was determined by

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measuring the increase in absorbance due to the formation of 5-thio-2-nitrobenzoic acid (TNB) from 50 ,50 -dithiobis-2nitrobenzoic acid (DTNB) in the presence of NADPH and glutathione reductase at 412 nm. The amount of GSSG was measured following derivatization of reduced glutathione (GSH) to remove it from the reaction. Then, the assay was performed as described for total glutathione. A standard curve with GSSG in the range of 0.195–6.25 mM was prepared prior to measurement of the samples. The GSH/GSSG ratio was calculated according to the manufacturer’s instructions. Measurement of intracellular 8-OHdG HepG2 and A549 cells were seeded on 60-mm dishes at a density of 1 106 cells/dish. After incubation for 24 h at 37 C, the cells were treated with different concentrations of AaC (5–20 mg/ml for HepG2 cells and 5–30 mg/ml for A549 cells) or 0.1% DMSO (for control cells) for 24 h. Then, the cells were washed twice with PBS and scraped into a microcentrifuge tube. DNA was isolated from the cell pellets using the Blood and Cultured Cell DNA extraction kit according to the manufacturer’s instructions. The amount of DNA was determined by UV spectroscopy, assuming that 50 mg/ml DNA exhibits an absorbance of 1.0 at 260 nm (Nauwelaers et al., 2011). DNA hydrolysis and purification were performed as described by Mangal et al. (2009) with some modifications. Deferoxamine was added to DNA elution solutions (approximately 200 ml containing 6 mg DNA) up to 0.1 mM. Then, the enzymatic hydrolysis was initiated by the addition of 20 U of DNase and 20 ml of 10 DNase buffer, followed by incubation for 2 h at 37 C. Thereafter, 20 ml of 10 alkaline phosphatase buffer was added together with 10 U of alkaline phosphatase, and the incubation continued at 37 C for 2 h. Finally, 10 ml of the sample was spiked with D1-dG (10 ng) and HPLC-grade water (1 ml). Then, the solution was filtered through a 0.22-mm filter (ANPEL, Shanghai, China). At the same time, a portion of the sample (150 ml) was spiked with internal standard [13C115N2]-8-OH-G (1 ng) and fractionated by SPE on a Bond Elut C18 cartridge (Agilent, Lake Forest, CA). The cartridge was washed with 2 ml water and eluted with 1 ml methanol. The collected fraction was dried under gentle nitrogen gas flow and reconstituted to a final volume of 100 ml. The HPLC-MS/MS analysis was conducted using a 1290 Infinity LC coupled with a 6490 Triple Quad mass spectrometer (Agilent, Palo Alto, CA). Chromatographic separation was performed using a Poroshell 120 SB C18 column (3.0 150 mm, 2.7 mm) at 30 C with an injection volume of 5 ml. The mobile phases were 0.1% acetic acid (A) and acetonitrile (B) and the flow rate was 0.4 ml/min. The gradient elution was performed as follows: 0 min, 5% B; 3 min, 10% B; 6–10 min, 100% B; equilibration time: 3 min. The mass spectrometer parameters were as follows: ionization mode, ESI+; detection mode, multiple reaction monitoring (MRM); dry gas (N2) flow at 12 L/min and 250 C; nebulizer, 40 psi; sheath gas (N2), 400 C, 12 l/min; capillary voltage: 4000 V; fragmental voltage, 380 V; collision gas, N2. Optimal MRM conditions and collision energies of

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Figure 1. Cytotoxic effects of AaC in HepG2 and A549 cells. Cells were exposed to 0.1% DMSO (control) or AaC for 24 h. Results are expressed as % of the value found in control cells, which arbitrarily set at 100%. Data are means ± SD of three independent experiments. *p50.05 and **p50.01 compared with control cells. aStatistical difference from A549 cells exposure at the same concentration, p50.05.

Table 1. Effect of AaC on DNA damage in HepG2 cells measured by the comet assay. AaC (mg/ml)

Tail length (mm)

Tail DNA (%)

Olive tail moment

Tail moment

0 5 10 15 20

37.60 ± 11.41 91.25 ± 12.35** 130.50 ± 20.94** 181.56 ± 25.76** 259.3 ± 31.73**

5.50 ± 1.36 11.89 ± 2.00** 19.29 ± 2.33** 27.61 ± 4.03** 33.36 ± 3.09**

4.07 ± 1.60 13.06 ± 1.97** 22.58 ± 4.59** 35.23 ± 9.5** 52.87 ± 9.28**

1.08 ± 0.95 13.04 ± 3.16** 25.40 ± 6.59** 50.70 ± 13.39** 83.53 ± 15.19**

DNA damage measured in a comet assay in HepG2 cells exposed to AaC (5–20 mg/ml) and 0.1% DMSO (for control cells) for 24 h. The data represent the mean value ± SD determined from three independent experiments. **p50.01 compared with control cells.

target compounds were as follows: 8-OhdG: m/z 284.2 ! 168.1 at 20 eV; dG: m/z 268.1 ! 152.1 at 11 eV; [13C115N2]8-OH-G: m/z 303.2 ! 171 at 12 eV; and D1-dG:m/z 269.1 ! 152.1 at 11 eV. The amounts of 8-OHdG and dG were estimated using calibration curves constructed by adding a fixed amount of [13C115N2]-8-OH-G (10 ng) and D1-dG (10 ng) internal standards to various amounts of 8-OHdG (0.01–2 ng/ml) and dG (1–200 ng/ml) standard solutions. The calibration curves were fitted to a straight line (area of response of the target compounds/internal standard versus the amount of the target compounds/internal standard) using ordinary leastsquares with equal weightings. The coefficient of determination (R2) values of the slopes exceeded 0.999. Statistical analysis All of the data were expressed as the mean ± standard deviation (SD). The statistical significance of differences between vehicle control and treated cells was analyzed with an ANOVA following Dunnett’s multiple comparison test. The difference in the viability of the two cell lines were compared using Student’s t-test. p Values 50.05 were considered statistically significant. IC50 values for AaC

were calculated with Regression following Probit. The Relationship among indicators was analyzed with Linear Fit using Origin 8.5 software (OriginLab Corporation, Northampton, MA).

Results Assessing the cytotoxicity of AaC to HepG2 cells and A549 cells by the NR incorporation assay Cell viability was estimated after a 24 h exposure to various concentrations of AaC (10–100 mg/ml). Figure 1 reveals a cell viability dose-response for AaC concentrations from 20 to 100 mg/ml and 30 to 100 mg/ml in HepG2 and A549 cells, respectively. The viability of A549 cells was higher than that of HepG2 cells when exposed to the same concentrations of AaC. AaC did not decrease the cell viability by more than 20% at concentrations up to 20 mg/ml in HepG2 cells and 30 mg/ml in A549 cells. The IC50 values were (58.2 ± 2.4) and (36.5 ± 3.01) mg/ml for A549 cells and HepG2 cells, respectively. These results indicated that HepG2 cells were more sensitive to AaC-induced cytotoxic effects than A549 cells. Based on these results, comet assays, ROS, GSH/GSSG ratio and 8-OHdG measurements were performed at concentrations


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Table 2. Effect of AaC on DNA damage in A549 cells measured by the comet assay. AaC (mg/ml)

Tail length (mm)

Tail DNA (%)

Olive tail moment

Tail moment

0 5 10 20 30

38.30 ± 11.41 66.32 ± 14.53 118.67 ± 16.00** 164.89 ± 19.56** 241.1 ± 22.11**

4.52 ± 1.58 7.61 ± 1.85 17.53 ± 3.22** 24.08 ± 4.68** 31.23 ± 2.56**

3.44 ± 1.42 8.13 ± 2.05 19.62 ± 5.23** 31.38 ± 6.88** 47.99 ± 11.92**

1.50 ± 0.92 5.16 ± 1.85 21.17 ± 6.33** 40.08 ± 10.70** 75.42 ± 9.19**


DNA damage measured in the comet assay in A549 cells exposed to AaC (5–30 mg/ml) and 0.1% DMSO (for control cells) for 24 h. Data represent the mean values ± SD determined in three independent experiments. **p50.01 compared with the control cells.

Figure 2. Effect of AaC on ROS generation in HepG2 and A549 cells evaluated using the DCFH-DA assay. (A) Effect of AaC on ROS generation in HepG2 cells. (B) Effect of AaC on ROS generation in A549 cells. Data represent the mean value ± SD of three independent experiments. *p50.05 and **p50.01 compared with the control cells.

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Figure 3. Effects of AaC on intracellular GSH/GSSG ratio in HepG2 and A549 cells. (A) GSH/GSSG ratio in HepG2 cells. (B) GSH/GSSG ratio in A549 cells. Data represented the mean value ± SD obtained with three independent experiments. *p50.05 and **p50.01 compared with control cells.

ranging from 5 to 20 mg/ml for HepG2 cells and 5 to 30 mg/ml for A549 cells, respectively. DNA damage measured by the comet assay DNA damage was measured in HepG2 and A549 cells by the comet assay under alkaline conditions (pH ¼ 13). Tables 1 and 2 show that AaC induced dose-dependent increases in Tail DNA content (%), OTM, TM and Tail Length at concentrations of 5–20 and 10–30 mg/ml in HepG2 cells and

A549 cells, respectively, compared with control cells. These data demonstrate that AaC induced DNA damage in HepG2 and A549 cells, and induced significant DNA migration at lower concentrations in HepG2 cells than in A549 cells. Intracellular ROS The generation of ROS in HepG2 and A549 cells treated with AaC or 0.1% DMSO (control cells) was measured by the DCFH-DA assay. As shown in Figure 2, AaC induced a


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Figure 4. AaC-induced oxidative DNA damage in HepG2 and A549 cells. (A) 8-OHdG formation in HepG2 cells treated with different concentrations of AaC or 0.1% DMSO for 24 h. (B) 8-OHdG formation in A549 cells treated with different concentrations of AaC or 0.1% DMSO for 24 h. Results are presented as the amount of 8-OHdG per 106 DNA bases (8-OHdG/ 106dG). The data represent the mean ± SD of three independent experiments. *p50.05 and **p50.01 compared with control cells.

significant increase in the intracellular generation of ROS at concentrations of 10–20 and 20–30 mg/ml in HepG2 cells and A549 cells, respectively. These results demonstrate that AaC affects ROS generation in HepG2 and A549 cells. Intracellular GSH/GSSG ratio The intracellular GSH/GSSG ratio was measured in A549 and HepG2 cells to examine the involvement of reduced glutathione (GSH) in the oxidative stress induced by AaC. As seen in Figure 3, after 2 h treatment with AaC, the intracellular GSH/ GSSG ratio was significantly decreased in both HepG2 and A549 treated at concentrations greater than 10 mg/ml compared with their respective controls (Figure 3). These data suggest that AaC induced either the consumption of GSH or the accumulation of GSSG.

Intracellular 8-OHdG 8-OHdG, a DNA adduct induced by oxygen free radicals, has been widely used as a marker for oxidative DNA damage (Loft et al., 1992, 1994; Toraason et al., 1999). The induction of intracellular 8-OHdG formation by AaC was examined by HPLC-MS/MS, and Figure 4 presents the results as the ratio of 8-OHdG to dG (8-OHdG/106dG). These results show that 8-OHdG/106dG increased in a dose-dependent manner over the range of 5–20 mg/ml in HepG2 cells and from 10 to 30 mg/ml in A549 cells.

Discussion A series of genotoxic studies have been performed to evaluate the risk of AaC to human health (Pfau et al., 1999; Robbana-Barnat et al., 1996; Yoshida & Matsumoto,


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Figure 5. The relationships among the levels of 8-OHdG, ROS and GSH/GSSG ratio in HepG2 and A549 cells. (A) The relationships among the levels of 8-OHdG, ROS and GSH/GSSG ratio in HepG2 cells. (B) The relationships among the levels of 8-OHdG, ROS and GSH/GSSG ratio in A549 cells. The data represent the results of three independent experiments.

1980; Yoshimi et al., 1988). However, few studies have addressed oxidative stress effects of AaC. In this study, we used HepG2 and A549 cells as in vitro models to investigate the potential genotoxic effects of AaC and discussed the role of oxidative stress in the mechanisms of those effects. At concentrations above 5 and 10 mg/ml, AaC induced a dose-dependent increase in DNA migration in HepG2 and A549 cells, respectively (Tables 1 and 2). These results clearly show that AaC can induce genotoxicity in HepG2 and A549 cells. It was consistent with that AaC caused dose-dependent induction of micronuclei in HepG2 cells (Majer et al., 2004). Moreover, these findings confirm the results of previous studies that AaC induced unscheduled DNA repair in hepatocytes from mice, rats and hamsters (Yoshimi et al., 1988). In the comet assay, AaC caused a significant increase in DNA migration in MCL-5 cells (Pfau et al., 1999). Considering all these results, AaC can clearly cause DNA damage. AaC induced significant DNA migration at lower concentrations in HepG2 cells than in A549 cells. It was likely due in part to the greater metabolic activity of these cells, as some

studies have shown that AaC exerts genotoxicity through its metabolites (King et al., 1999, 2000). Increased ROS production and/or a decrease in antioxidant defense can lead to oxidative stress (Trevisan et al., 2001). ROS could attack other cellular components such as lipids, leaving behind reactive species that in turn can couple to DNA bases (Marnett, 2000). GSH, as the major intracellular antioxidant defense, is involved in reactions with ROS to form oxidized glutathione (GSSG; Franco et al., 2007). Consumption of GSH leads to a decrease in the GSH/GSSG ratio (Balendiran et al., 2004) and potentially to enhanced ROS generation. In this study, AaC induced a significant increase in intracellular ROS generation and decreased the GSH/GSSG ratio, suggesting that the cellular redox balance was disrupted and the level of intracellular antioxidants was reduced. These results demonstrated that AaC induced oxidative stress in HepG2 and A549 cells. ROS have been reported to cause specific chemical modifications of purine and pyrimidine bases, DNA strand breaks and DNA–protein cross-links (Janssen et al., 1993). The reaction of ROS with DNA yields a variety of oxidized base lesions (Evans et al., 2004). Levels of 8-OHdG increased



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Figure 6. The relationships among 8-OHdG levels, OTM and TM in HepG2 and A549 cells. (A) The relationships among 8-OHdG levels, OTM and TM in HepG2 cells. (B) The relationships among 8-OHdG levels, OTM and TM in A549 cells. The data represent the results of three independent experiments.

in both types of cells, indicating that AaC could induce oxidative DNA damage. Then, the relationships among the levels of ROS, 8-OHdG and GSH/GSSG were studied. As observed in Figure 5, 8-OHdG is positively related to ROS and negatively related to the GSH/GSSG ratio. These results further confirm that AaC-induced oxidative stress leads to oxidative DNA damage in HepG2 and A549 cells. 8-OHdG, an oxidized nucleoside of DNA, has been reported to be a reliable biomarker for oxidative DNA damage. This DNA lesion has been shown to be mutagenic, causing GC!TA transversions in both bacterial and mammalian cells (Moriya, 1993). Oxidative DNA damage has been implicated in the mechanisms of genotoxicity of several other xenobiotics such as acrylamide, hydroquinone and olaquindox (Jiang et al., 2007; Peng et al., 2013; Zou et al., 2009). To further confirm the relationship between DNA damage and AaC treatment, the relationships among 8-OHdG levels, TM and OTM were studied. As observed in Figure 6, the level of 8-OHdG was positively related to TM and OTM. These results suggest that the genotoxic effects of AaC are related to oxidative DNA damage induced by ROS. AaC induced a significant increase in the levels of ROS and 8-OHdG at lower concentrations in HepG2 cells than in A549 cells. AaC induced stronger oxidative stress effects

and more serious oxidative DNA damage in HepG2 cells. This may partially explain that AaC induced significant DNA migration at lower concentrations in HepG2 cells than in A549 cells, and also supports the theory that ROS-induced oxidative DNA damage contributes to the genotoxic effects of AaC.

Conclusions In conclusion, our results demonstrate that AaC exerts significant genotoxicity, cytotoxicity and oxidative stress in HepG2 and A549 cells. The genotoxicity induced by AaC likely resulted from ROS-induced oxidative DNA damage. To the best of our knowledge, this is the first report on role of oxidative stress in genotoxicity induced by AaC in HepG2 and A549 cells.

Declaration of interest The authors report no conflicts of interest. This study was supported by the National Science and Technology Support Program (2012BAK01B03) of China.

References Balendiran GK, Dabur R, Fraser D. (2004). The role of glutathione in cancer. Cell Biochem Funct 22:343–52.


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An aqueous pomegranate seed extract ameliorates oxidative stress of human hepatoma HepG2 cells.

Toxic Effects of the Major Components of Diesel Exhaust in Human Alveolar Basal Epithelial Cells (A549).

Oxidative stress and mitochondrial injury-mediated cytotoxicity induced by silver nanoparticles in human A549 and HepG2 cells.

M cell cycle arrest and apoptosis in A549 human lung cancer cells.

Capsaicin-induced genotoxic stress does not promote apoptosis in A549 human lung and DU145 prostate cancer cells.

Effect of ulinastatin on paraquat-induced-oxidative stress in human type II alveolar epithelial cells.

Edaravone attenuates paraquat-induced lung injury by inhibiting oxidative stress in human type II alveolar epithelial cells.

Azithromycin attenuates cigarette smoke extract-induced oxidative stress injury in human alveolar epithelial cells.

Andrographolide antagonizes cigarette smoke extract-induced inflammatory response and oxidative stress in human alveolar epithelial A549 cells through induction of microRNA-218.

Metabolism and cytotoxic effects of phosphatidylcholine hydroperoxide in human hepatoma HepG2 cells.

M cell cycle arrest and apoptosis in human hepatoma HepG2 cells.

Trichomonas vaginalis induces cytopathic effect on human lung alveolar basal carcinoma epithelial cell line A549.

Determining oxidative and non-oxidative genotoxic effects driven by estuarine sediment contaminants on a human hepatoma cell line.

Cytotoxicity and apoptotic effects of tea polyphenol-loaded chitosan nanoparticles on human hepatoma HepG2 cells.

Impaired oxidative phosphorylation regulates necroptosis in human lung epithelial cells.

Long term and large-scale cultivation of human hepatoma Hep G2 cells in hollow fiber bioreactor. Cultivation of human hepatoma Hep G2 in hollow fiber bioreactor.

Global secretome characterization of A549 human alveolar epithelial carcinoma cells during Mycoplasma pneumoniae infection.

OXIDATIVE STRESS IN HUMAN LEUKEMIA (HL-60), HUMAN LIVER CARCINOMA (HepG2), AND HUMAN (JURKAT-T) CELLS EXPOSED TO ARSENIC TRIOXIDE.

Comparative study of cytotoxic and genotoxic effects induced by herbicide S-metolachlor and its commercial formulation Twin Pack Gold® in human hepatoma (HepG2) cells.

Cytotoxic effects of procyanidins from Castanea mollissima Bl. shell on human hepatoma G2 cells in vitro.

Effects Induced by Organic Acids in a Human Lung Alveolar Carcinoma Cell Line A549.

Glutathione content and growth in A549 human lung carcinoma cells.

Modulation of the DNA repair system and ATR-p53 mediated apoptosis is relevant for tributyltin-induced genotoxic effects in human hepatoma G2 cells.

Cytostatic and genotoxic effect of temephos in human lymphocytes and HepG2 cells.