Epoxy Stearic Acid, an Oxidative Product Derived from Oleic Acid

Article pubs.acs.org/JAFC

Cite This: J. Agric. Food Chem. 2018, 66, 5237−5246

Epoxy Stearic Acid, an Oxidative Product Derived from Oleic Acid, Induces Cytotoxicity, Oxidative Stress, and Apoptosis in HepG2 Cells Ying Liu, Yajun Cheng, Jinwei Li, Yuanpeng Wang, and Yuanfa Liu* School of Food Science and Technology, Synergetic Innovation Center of Food Safety and Nutrition, Jiangnan University, 1800 Lihu Avenue, Wuxi, Jiangsu 214122, People’s Republic of China ABSTRACT: In the present study, effects of cis-9,10-epoxy stearic acid (ESA) generated by the thermal oxidation of oleic acid on HepG2 cells, including cytotoxicity, apoptosis, and oxidative stress, were investigated. Our results revealed that ESA decreased the cell viability and induced cell death. Cell cycle analysis with propidium iodide staining showed that ESA induced cell cycle arrest at the G0/G1 phase in HepG2 cells. Cell apoptosis analysis with annexin V and propidium iodide staining demonstrated that ESA induced HepG2 cell apoptotic events in a dose- and time-dependent manner; the apoptosis of cells after treated with 500 μM ESA for 12, 24, and 48 h was 32.16, 38.70, and 65.80%, respectively. Furthermore, ESA treatment to HepG2 cells resulted in an increase in reactive oxygen species and malondialdehyde (from 0.84 ± 0.02 to 8.90 ± 0.50 nmol/mg of protein) levels and a reduction in antioxidant enzyme activity, including superoxide dismutase (from 1.34 ± 0.27 to 0.10 ± 0.007 units/ mg of protein), catalase (from 100.04 ± 5.05 to 20.09 ± 3.00 units/mg of protein), and glutathione peroxidase (from 120.44 ± 7.62 to 35.84 ± 5.99 milliunits/mg of protein). These findings provide critical information on the effects of ESA on HepG2 cells, particularly cytotoxicity and oxidative stress, which is important for the evaluation of the biosafety of the oxidative product of oleic acid. KEYWORDS: cis-9,10-epoxy stearic acid, HepG2 cell, cytotoxicity, apoptosis, oxidative stress

■

INTRODUCTION Lipid oxidation is a main cause of quality deterioration in lipidand oil-containing foods. Lipid oxidation involves a wide variety of reactions, including degradation, hydrolysis, polymerization, etc., which not only gives rise to nutrient loss and off-flavor generation but also causes formation of potentially toxic compounds and, thus, decreases the product quality, sometimes even making foods unsuitable for consumption.1−5 Meanwhile, oxidized products, including triacylglycerol (TAG) polymers, TAG dimers, oxidized TAG monomers, diacylglycerols, and free fatty acids, are formed. Among them, epoxy groups linked to TAG molecules, one of oxidized TAG monomers, have the highest content and are easily absorbed by both animals and humans.6−8 In general, epoxy fatty acids are produced by the reaction of corresponding fatty acids and hydroperoxides9 and exert some adverse effects on health.10−12 cis-9,10-Epoxy stearic acid (ESA) was derived from thermal oxidation of oleic acid (OA) and has been widely found in many food matrices.13−15 Recently, our research group found ESA in frying oil and established an extraction method for ESA, and the content of ESA in frying oil samples reached up to 5900 mg/kg.16 In addition, on the basis of the electron spin resonance spectroscopy method, the formation process of ESA may be that OA loses hydrogen radicals to form alkyl radicals, which could react with oxygen to form hydroperoxides, and then the O−O bonds of hydroperoxides break to form alkoxyl radicals, which could abstract hydrogen from other OAs to form ESA (Figure 1). Most of the research relevant to epoxy fatty acids focused on epoxy OAs. Fukushima et al.10 found that 9,10-epoxy-12-octadecenoate could increase the risk of cardiovascular diseases. Hu et al.17 found that 9,10-epoxy-12octadecenoate induced pulmonary edema in rats. However, the © 2018 American Chemical Society

effect of ESA derived from OA on animals or cells remains largely unknown. The HepG2 cell line, which was selected because of its comparability to the normal hepatocytes in aspects of expression of specific enzymes and the enzyme activities,18−20 has been widely used as the human hepatoma model in the performance of the lipid metabolic process.21−23 The present study investigated the effect of ESA formed by thermal oxidation of OA on HepG2 human hepatoma cells by considering cell viability, cytotoxicity, apoptosis, intracellular ROS level, activities of antioxidant enzymes, and lipid peroxide level to explore cytotoxicity and oxidative stress of ESA, which is important for evaluating the biosafety of the oxidative product of OA.

■

MATERIALS AND METHODS

Chemicals. ESA (99% purity) was purchased from Toronto Research Chemicals (Toronto, Ontario, Canada). OA and dimethyl sulfoxide (DMSO) were purchased from J&K Chemical Technology (Shanghai, China). HepG2 cells were purchased from the Institute of Biochemistry and Cell Biology, Shanghai Institutes for Biological Sciences (SIBS), Chinese Academy of Sciences (CAS) (Shanghai, China). Minimum essential medium (MEM) was obtained from Shanghai BasalMedia Technologies Co., Ltd. (Shanghai, China). (3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) (MTT) was purchased from Sigma-Aldrich Co. (St. Louis, MO, U.S.A.). Trypsin, fetal bovine serum (FBS), and other cell culture materials were purchased from Gibco BRL, Life Technologies (Carlsbad, CA, U.S.A.). The reactive oxygen species (ROS), malondialdehyde (MDA), superoxide dismutase (SOD), catalase (CAT), and glutathione peroxidase (GSH-Px) assay kits were all purchased from Beyotime Biotechnology Co., Ltd. Received: April 14, 2018 Accepted: May 5, 2018 Published: May 6, 2018 5237

DOI: 10.1021/acs.jafc.8b01954 J. Agric. Food Chem. 2018, 66, 5237−5246

Article

Journal of Agricultural and Food Chemistry

Figure 1. Structure formulas of the formation of ESA derived from OA during thermal oxidation.

Figure 2. Effects of OA and ESA on the viability of HepG2 cells. Different lowercase letters in the same column indicate significant differences (p < 0.05) for the same sample. (Shanghai, China). The annexin V−fluorescein isothiocyanate (FITC) apoptosis detection kit and cell cycle analysis kit were also obtained from Beyotime Biotechnology Co., Ltd. (Shanghai, China). All chemicals and reagents were of analytical grade or higher. Cell Culture and Treatment. HepG2 human hepatocellular carcinoma cells were cultured in MEM containing 10% FBS, 100 units/mL penicillin, and 75 units/mL streptomycin (Gibco BRL, Life Technologies, Carlsbad, CA, U.S.A.). Cells were incubated at 37 °C with 5% CO2 in a humidified atmosphere. HepG2 cells were treated with OA and ESA at various concentrations (10, 20, 50, 100, 200, and 500 μM) for 12, 24, and 48 h. Cytotoxicity Measurements by the MTT Assay. Undifferentiated HepG2 cells were plated into a 96-well plate (5 × 104 cells/mL) and preincubated for 24 h to ascertain cell attachment at 37 °C. The viability of cells was determined by the MTT assay. HepG2 cells were treated with fatty acids at various concentrations for different times. Following treatment, 10 μL of MTT (5 mg/mL) reagent was added to the wells, and the cells were further incubated at 37 °C for 4 h. Then, the medium was replaced with 150 μL of DMSO and incubated for 15 min. The absorbance was measured at 490 nm using a microplate reader (Thermo, Waltham, MA, U.S.A.). The cell viability (%) was calculated using the following equation:

according to the instructions of the manufacturer. Briefly, HepG2 cells (1 × 106 cells/well) were seeded in 6-well plates and treated with fatty acids at various concentrations for different times. Cells were then collected, washed with annexin-binding buffer, and stained with annexin V−FITC and PI for 15 min at room temperature. Finally, cells were analyzed by flow cytometry (BD FACSCalibur, San Jose, CA, U.S.A.). Measurement of ROS. The level of ROS was determined by measuring changes in 20,70-dichlorofluorescein diacetate (DCFH-DA) fluorescence. After treatment with fatty acids, cells were incubated with DCFH-DA according to the instructions of the manufacturer. Subsequently, the formation of the fluorescent-oxidized derivative of dichlorofluorescein (DCF) was measured by flow cytometer (BD FACSCalibur, San Jose, CA, U.S.A.) at an emission wavelength of 525 nm and an excitation wavelength of 488 nm. Measurements of SOD, GSH-Px, CAT, and MDA. The assay for SOD, GSH-Px, CAT, and MDA was carried out using commercial assay kits. Briefly, the SOD activity was measured using its ability to inhibit the reduction of WST-8 according to the instructions of the manufacturer. SOD activity was monitored spectrophotometrically at 450 nm using a microplate reader (Thermo, Waltham, MA, U.S.A.). GSH-Px was detected by measuring the decreasing amount of NADPH and monitored at 340 nm according to the instructions of the kit. CAT was detected by measuring the absorbance of the red compound [N-(4antipyryl)-3-chloro-5-sulfonate-p-benzoquinonemonoimine] reacted by hydrogen peroxide and oxygen at 520 nm. The content of MDA was determined by measuring the absorbance of MDA−thiobarbituric acid (TBA) reacted by MDA and TBA at 532 nm. Statistical Analysis. Analytical determinations were performed in triplicate, and the results were expressed as the mean ± standard deviation of replicated measurements. Statistical comparisons were performed by one-way analysis of variance (ANOVA) combined with Duncan’s multiple-range test using the SPSS statistical package (version 19.0, SPSS, Inc., Chicago, IL, U.S.A.). p < 0.05 was considered significant.

cell viability (%) = A treated / Acontrol × 100% Cell Cycle Analysis. Cell cycle analysis was conducted using the cell cycle analysis kit (Beyotime, Shanghai, China), according to the instructions of the manufacturer. Briefly, HepG2 cells (1 × 106 cells/well) were seeded in 6-well plates and treated with fatty acids at various concentrations for different times. Then, the cells were harvested, washed with ice-cold phosphate-buffered saline (PBS) buffer, and fixed with 70% alcohol at 4 °C for 12 h. After that, DNA was stained with 10 μL of propidium iodide (PI, 1 mg/mL) and 10 μL of RNase A (10 mg/mL) for 30 min at room temperature. Cells were then subjected to flow cytometry (BD FACSCalibur, San Jose, CA, U.S.A.). The percentage of cells in G1, S, and G2 phases of the cell cycle was calculated using Cell Lab Quanta SC software (Beckman Coulter, Inc., Fullerton, CA, U.S.A.). Cell Apoptosis Analysis. Cell apoptosis was detected with an annexin V−FITC apoptosis detection kit (Beyotime, Shanghai, China)

■

RESULTS Effects of OA and ESA on HepG2 Cell Viability. Effects of OA and ESA on the viability of HepG2 cells are shown in Figure 2. The MTT assay demonstrated a gradual decrease in 5238


Article


Figure 3. Effects of (A) OA and (B) ESA on the cell cycle arrest of HepG2 cells.

S phase was observed in ESA-treated groups (Figure 3B). As shown in Figure 3, untreated cells had 51.39, 57.61, and 59.51% of cells in the G0/G1 phase and 41.90, 35.10, and 33.94% of cells in the S phase after 12, 24, and 48 h, respectively. However, 60.50, 66.94, and 67.02% of cells in the G0/G1 phase and 29.69, 30.62, and 30.98% of those in the S phase were detected when HepG2 cells were exposed to 500 μM OA for 12, 24, and 48 h, respectively. In particular, when the concentrations of ESA reached 500 μM, the fraction of cells in the G0/G1 phase increased dramatically (68.34, 78.11, and 82.83% compared to 51.39, 57.61, and 59.51% in untreated cells), which suggested that HepG2 cells underwent DNA fractionation, one of the biochemical events leading to apoptosis. Taken together, these results indicated that ESA could inhibit HepG2 cell proliferation significantly by blocking the G0/G1 to S phase transition in the cell cycle in a dose- and time-dependent manner. Effects of OA and ESA on Apoptosis in HepG2 Cells. In the present work, combined analysis of annexin V−FITC and PI based on flow cytometry was carried out to determinate the apoptotic rate of HepG2 cells induced by OA and ESA. In Figure 4, cells in Q2 are described as advanced apoptotic or necrotic, normal cells are seen in Q3, and cells in Q4 are classified as early apoptotic. As shown in Figure 4, the cell apoptosis of HepG2 increased slowly by treated with different concentrations of OA for 12 and 24 h. In terms of ESA, increasing concentrations of ESA for the 12 h treatment induced apoptotic events, especially early apoptosis, in a dose-dependent manner,

HepG2 cell viability with the increasing concentrations of fatty acids treated for 12−48 h. At the lower treatment dose (10−50 μM) of OA, the viability of cells was kept above 70%. When HepG2 cells were exposed to OA with a concentration of 100−500 μM, the cell viability slightly decreased and was maintained at a certain level ranging from 57 to 67%. In comparison to OA, ESA significantly inhibited cell viability, and cell viability decreased with the increasing concentration and time of ESA treatment. ESA treatment at 10−20 μM for 12, 24, and 48 h caused a 65−71, 51−56, and 50−52% loss in cell viability, respectively. While after treatment of 500 μM ESA for 12, 24, and 48 h, the number of alive cells decreased, with cell viability values of 30.01, 25.69, and 24.41%, respectively. The maximum inhibition (75.59%) was observed in cells treated with 500 μM ESA for 48 h. On the basis of these results, it was clear that ESA caused a dose- and time-dependent decrease in HepG2 cell viability. Effects of OA and ESA on Cell Cycle Arrest in HepG2 Cells. To investigate whether OA and ESA could induce cell cycle distribution in HepG2 cells, flow cytometric analysis of PI-stained nuclei cells was performed after treatment with OA and ESA at various concentrations (10−500 μM) for 12−48 h. As shown in Figure 3A, results revealed that HepG2 cells treated with OA presented a dose- and time-dependent increase in the cell population of the G0/G1 phase. Meanwhile, the proportion of cells in the S phase decreased. A similar concentration- and time-related increase in the fraction of cells in the G0/G1 phase and decrease in the fraction of cells in the 5239


Article

Figure 4. Flow cytometric analysis for apoptosis induction of HepG2 cells treated with OA and ESA.


5240


Article


a biomarker for the antioxidative status in the cell, was detected in HepG2 cells treated with OA and ESA. Results demonstrated that the SOD content decreased with the increasing concentration and time of OA and ESA treatments (Figure 6B). Treatment for 12−24 h with different concentrations of OA only slightly decreased the content of SOD in HepG2 cells. While SOD levels in the ESA treatment groups decreased rapidly (p < 0.05), the activity of SOD in cells that were exposed to ESA at 500 μM for 12, 24, and 48 h were reduced by 51.37, 69.95, and 84.64%, respectively, when compared to the control group. Furthermore, decreased activity of CAT in HepG2 cells treated with OA and ESA was also observed (Figure 6C). The activity of CAT in cells treated with different concentrations of OA for 12 and 24 h induced a slight decrease when compared to the control group from 100.04 and 92.49 units/mg of protein to 82.62 and 81.29 units/mg of protein, respectively. However, a significant decrease (p < 0.05) in CAT levels was shown after ESA treatments for 12 and 24 h in HepG2 cells; at 500 μM ESA for 12 and 24 h treatments, the activity of CAT in cells were reduced by 32.72 and 58.34%, respectively. Specially, in contrast to HepG2 cells treated for 12 and 24 h, intracellular CAT activity decreased dramatically (p < 0.05) after OA and ESA exposure for 48 h. GSH-Px could catalyze the reduction of lipid hydroperoxides to hydroxides using glutathione (GSH), in which oxidized GSH is produced by GSH and then reduced back to GSH with the glutathione reductase (GR) catalyst. As shown in Figure 6D, a reduction in the levels of GSH-Px was detected in HepG2 cells after treatment with OA and ESA compared to the control group. Specially, ESA treatment significantly reduced GSH-Px levels in a dose- and time-dependent manner. The presence of 500 μM ESA in the culture medium for 12 and 24 h induced a significant decrease in the activity of GSH-Px compared to the control group from 120.44 and 113.32 milliunits/mg of protein to 61.09 and 50.73 mU/mg of protein, respectively. Furthermore, after pretreatment of cells with different concentrations of OA and ESA for 48 h, levels of GSH-Px showed a trend to be far below those of cells treated with OA and ESA for 12 and 24 h. The increase in the MDA content and decrease in the SOD, CAT, and GSH-Px activities of cells demonstrated that exposure to ESA induced oxidative stress, and cells treated with ESA may lose the ability to maintain the balance between ROS and antioxidants.

while the percentage of late apoptosis and dead cells increased with the increased concentration of ESA for the 24 h treatment. At 500 μM ESA for 12 and 24 h treatments, the apoptosis was 32.16 and 38.70%, respectively. They were more than 8 and 6 times that in the control group, respectively. As the concentration of ESA increased, the apoptotic rate of the HepG2 cells elevated dramatically (p < 0.05). Specially, in comparison to the control group, HepG2 cells treated with 500 μM OA and ESA for 48 h increased the apoptosis rate from 15.51 to 56.10 and 65.80% apoptosis, respectively. Interestingly, after treatment of HepG2 with 200−500 μM OA for 48 h, the apoptosis rate was above 40%; one possible reason may be that cell death occurred for HepG2 cells after a long-time OA treatment and nutrient deficiency. These results suggested that ESA could induce dramatic apoptosis in HepG2 cells. Effect of OA and ESA on ROS Accumulation in HepG2 Cells. DCF fluorescence was used to measure the level of ROS that was induced by OA and ESA exposure. As shown in Figure 5, in comparison to the control group, the intracellular ROS level in HepG2 cells after different concentrations of OA treatment for 12 and 24 h showed a gradual and slight increase in a doseand time-dependent manner. A significant increase in ROS generation was observed over time ranging from 12 to 24 h in the cells treated with different concentrations of ESA (p < 0.05). After HepG2 cells were treated with 500 μM ESA for 12 and 24 h, cellular fluorescence intensity were 2.6- and 3.0-fold of the non-ESA-treated control HepG2 cells. However, in contrast to HepG2 cells treated for 12 and 24 h, although the intracellular ROS level increased after OA and ESA exposure for 48 h, it was maintained at a relatively low level after 48 h of exposure to all concentrations of OA and ESA. Leakage of the probe was not observed in cells in pre-tests. Therefore, the final fluorescence caused by extracellularly oxidized DCF could be excluded. The reason for this may be that long-time OA and ESA treatments and nutrient deficiency led to a high percentage of cell death; the level of intracellular ROS was therefore relatively low. Our results indicated that ESA treatment could induce abnormal accumulation of intracellular ROS in HepG2 cells. Effects of OA and ESA on the Activities of Antioxidant Enzymes and Lipid Peroxide Levels in HepG2 Cells. Because ESA might induce accumulation of intracellular ROS and lead to oxidative damage to HepG2 cells, MDA, a product of lipid peroxides induced by ROS, was also measured in HepG2 cells. As seen in Figure 6A, treatment for 12−48 h with different concentrations of OA could only slightly increase the content of MDA in HepG2 cells, indicating a relatively low level of lipid peroxidation in response to OA treatment in cells. The increased concentration of MDA in HepG2 cells treated with ESA was found to be dose- and time-dependent. Meanwhile, a statistically significant increase in MDA levels was observed after ESA treatments for 12−24 h in cells when compared to the control group. Interestingly, after cells were treated with OA for 48 h, levels of MDA showed a trend to be almost similar to those of control untreated cells, even after 48 h with 500 μM. Although the 48 h treatment of HepG2 with ESA evoked an increase in the cellular concentration of MDA, the levels of intracellular MDA were much lower than those treated for 12 and 24 h. This could be attributed to the lack of nutrients and long-time treatment, which was in line with the results of intracellular ROS levels. Cellular damage caused by ROS depends upon not only the intracellular ROS level but also the balance between ROS and endogenous antioxidant species. The intracellular SOD,

■

DISCUSSION Lipid oxidation products have been claimed to exert adverse effects on health in in vivo and in vitro studies,5,24−27 including rising risks of non-alcoholic fatty liver disease, atherosclerosis, and diabetes mellitus. Moreover, cytotoxicity and oxidative stress have been observed in subjects after the intake of oxidative products of lipid.28−31 However, the adverse effects of ESA produced from oxidized OA remain unknown. In the present study, the effects of ESA on cytotoxicity and oxidative stress of HepG2 cells were investigated. Our results showed that cell viability, as determined by the MTT assay, reduced remarkably in a dose- and time-dependent manner after HepG2 cells were exposed to ESA. The relatively little effect of OA on the HepG2 cell survival rate was also observed. This was consistent with the studies reported by Greene et al.32 and Cao et al.,33 in which epoxy fatty acids and TAG polymers (both derived from the lipid oxidation process) were used, respectively. In addition, cell cycle analysis were 5241


Article

Figure 5. continued


5242


Article


Figure 5. Effect of OA and ESA on the intracellular ROS level of HepG2 cells.

carried out to illustrate ESA-induced cell cycle arrest. Results showed that the percentage of cells in the G0/G1 phase increased and those in the S phase decreased in HepG2 cells exposed to ESA, indicating that the inhibition of ESA on HepG2 cells mainly occurred in the G0/G1 phase. The apoptotic effect of ESA on HepG2 cells was confirmed by the annexin V−FITC/PI assay. Results indicated that ESA treatment for 12 and 24 h mainly induced early and late apoptosis in HepG2 cells based on a dose-dependent manner, respectively. This finding was in agreement with other studies, where the ingestion of oxidized products, containing polar compounds, oxidized phospholipids, TAG polymers, and auto-oxidation products from cholesterol, inhibited cell proliferation and induced apoptosis compared to those in untreated cells.28,32−34 A further study found that the intracellular ROS levels in HepG2 cells were markedly elevated after ESA treatment, suggesting that ESA-induced cell damage may be related to excessive ROS production. A similar effect of 9,10-epoxy-12-octadecenoate on rats was observed by Ozawa et al.35 Ozawa and co-workers found that 9,10-epoxy-12-octadecenoate, which is biosynthesized by human neutrophil, led to stress response in the lung. These results strongly suggested that epoxy fatty acids could induce excessive generation of the intracellular ROS level. It has been reported that ROS could perform normal functions when the generation and elimination of ROS in normal cell systems is in equilibrium, which was maintained by the endogenous antioxidant system.33,36 However, excessive accumulation of ROS causes injury to cellular components, including nucleic acids, cellular proteins, and lipids, and activates cell apoptosis signaling pathways, leading to a state known as oxidative stress.37 Moreover, oxidative stress is related to the etiopathogenesis of several human chronic diseases, such as many cardiovascular diseases, aging, neurodegenerative diseases, diabetes, and cancer.38−40 An elevated intracellular MDA level suggested that lipid peroxidation significantly increased in HepG2 cells after ESA treatment with the increase of the ESA concentration and exposure time. MDA, which is the principal and most studied lipid oxidation product of polyunsaturated fatty acids, has been widely analyzed to assess the level of oxidative stress. It has been reported that an increase in the MDA value indicated an increased lipid peroxidation, which results in tissue injury and the failure of antioxidant defense mechanisms in preventing excess ROS formation.41,42 Therefore, our results implied damage to the antioxidant defense system in cells. A reduction of SOD activity, one of the major components of the antioxidant capacity to defense against ROS-mediated injury in tissue,43 was found in cells treated with ESA compared to the control group, indicating that generation of oxidative stress was accompanied by the reduced enzymatic antioxidant activity in HepG2 cells, which was in accordance with the results reported by Bhor and colleagues,44 in which intestinal oxidative stress occurred with a reduction of the enzymatic antioxidant activity in enterocytes. Moreover, a decrease in the CAT activity was seen in the HepG2 cells pretreated with ESA. Similar results has been reported in previous studies with the consumption of lipid oxidation products, in which the activities of SOD and CAT in the liver were both significantly lowered.29 CAT, an enzyme to catalyze the oxidation of various hydrogen donors, could decompose hydrogen peroxide to molecular oxygen and water. Reduced CAT activity in tissue causes oxygen intolerance and triggers some deleterious reactions, especially DNA oxidation and cell death.45 The GSH-Px activity, an essential antioxidant for the intracellular quenching of cell-damaging peroxide species, significantly 5243


Article


Figure 6. Effect of OA and ESA on the intracellular (A) MDA level and activity of antioxidant enzymes, including (B) SOD, (C) CAT, and (D) GSH-Px of HepG2 cells.

ORCID

decreased in cells treated with ESA compared to the control group. It is well-known that the cellular antioxidant enzyme system plays an important role in the defense against oxidative stress, and the activity of antioxidant enzymes could be a biomarker of the antioxidant response. Therefore, our results clearly demonstrated that ESA could reduce the antioxidative capacity and induce oxidative stress in HepG2 cells, which may aggravate the imbalance between oxidation and antioxidation in cells. In conclusion, this is the first time to investigate the effect of ESA on the cytotoxic and oxidative stress of human liver carcinoma cells. The current study clearly demonstrated that administration of ESA to HepG2 cells for 12−48 h could induce cytotoxicity, DNA damage, apoptosis, and oxidative stress. ROS may play an essential role in DNA damage and oxidative stress induced by ESA in vitro. Our results showed that ESA treatment induced apoptosis activity on HepG2 cells, including cell proliferation, apoptosis, possible genetic damage, and accumulation of the ROS level. In addition, ESA also enhanced the lipid peroxides and caused the decrease of enzyme activities of SOD, CAT, and GSH-Px, which were biomarkers of the cellular oxidative status. Further studies should be carried out to investigate the pathological mechanism of ESA derived from thermally oxidized OA on apoptosis and oxidative stress of HepG2 cells.

■

Yuanfa Liu: 0000-0002-8259-8426 Funding

This work was supported by the Natural Science Foundation of China (31671786), the Research Fund of the National 13th Five-Year Plan of China (2016YFD0401404), the Northern Jiangsu Province Science and Technology Projects (BN2016137), and the Fundamental Research Funds for the Central Universities (JUSRP51501). Notes

The authors declare no competing financial interest.

■

REFERENCES

(1) Homma, R.; Suzuki, K.; Cui, L.; Mcclements, D. J.; Decker, E. A. Impact of association colloids on lipid oxidation in triacylglycerols and fatty acid ethyl esters. J. Agric. Food Chem. 2015, 63, 10161−10169. (2) Shahidi, F.; Zhong, Y. Lipid oxidation and improving the oxidative stability. Chem. Soc. Rev. 2010, 39, 4067−4079. (3) Barden, L.; Decker, E. A. Lipid oxidation in low-moisture food: A review. Crit. Rev. Food Sci. Nutr. 2016, 56, 2467−2482. (4) Kanner, J. Dietary advanced lipid oxidation endproducts are risk factors to human health. Mol. Nutr. Food Res. 2007, 51, 1094−1101. (5) Mozaffarian, D. Trans fatty acids-Effects on systemic inflammation and endothelial function. Atheroscler. Suppl. 2006, 7, 29−32. (6) Erickson, M. D. Production and Composition of Frying Fats. In Deep Frying: Chemistry, Nutrition, and Practical Applications, 2nd ed.; Erickson, M. D., Ed.; AOCS Press: Urbana, IL, 2007; Chapter 1, pp 3−24, DOI: 10.1016/B978-1-893997-92-9.50007-4.

AUTHOR INFORMATION

Corresponding Author

*Telephone/Fax: 0510-85876799. E-mail: yfl[email protected]. 5244


Article

Journal of Agricultural and Food Chemistry (7) Zhang, Q.; Saleh, A. S. M.; Chen, J.; Shen, Q. Chemical alterations taken place during deep-fat frying based on certain reaction products: A review. Chem. Phys. Lipids 2012, 165, 662−681. (8) Kmiecik, D.; Kobus-Cisowska, J.; Korczak, J. The content of antinutritional components in frozen fried-potato products. LWTFood Sci. Technol. Nol. 2017, 85, 275−282. (9) Frankel, E. N. Hydroperoxide Formation. In Lipid Oxidation, 2nd ed.; Frankel, E. N., Ed.; Woodhead Publishing, Ltd.: Cambridge, U.K., 2012; Oily Press Lipid Library Series, Vol. 18, Chapter 2, pp 25−50, DOI: 10.1533/9780857097927.25. (10) Fukushima, A.; Hayakawa, M.; Sugiyama, S.; Ajioka, M.; Ito, T.; Satake, T.; Ozawa, T. Cardiovascular effects of leukotoxin (9,10-epoxy12-octadecenoate) and free fatty acids in dogs. Cardiovasc. Res. 1988, 22, 213−218. (11) Greene, J. F.; Newman, J. W.; Williamson, K. C.; Hammock, B. D. Toxicity of epoxy fatty acids and related compounds to cells expressing human soluble epoxide hydrolase. Chem. Res. Toxicol. 2000, 13, 217−226. (12) Wilson, R.; Fernie, C. E.; Scrimgeour, C. M.; Lyall, K.; Smyth, L.; Riemersma, R. A. Dietary epoxy fatty acids are absorbed in healthy women. Eur. J. Clin. Invest. 2002, 32, 79−83. (13) Marmesat, S.; Velasco, J.; Dobarganes, M. C. Quantitative determination of epoxy acids, keto acids and hydroxy acids formed in fats and oils at frying temperatures. J. Chromatogr. A 2008, 1211, 129− 134. (14) Mubiru, E.; Shrestha, K.; Papastergiadis, A.; De Meulenaer, B. Development and validation of a gas chromatography−flame ionization detection method for the determination of epoxy fatty acids in food matrices. J. Agric. Food Chem. 2014, 62, 2982−2988. (15) Velasco, J.; Marmesat, S.; Bordeaux, O.; Márquez-Ruiz, G.; Dobarganes, C. Formation and evolution of monoepoxy fatty acids in thermoxidized olive and sunflower oils and quantitation in used frying oils from restaurants and fried-food outlets. J. Agric. Food Chem. 2004, 52, 4438−4443. (16) Liu, Y.; Wang, Y.; Cao, P.; Liu, Y. Quantitative determination of epoxy stearic acids derived from oxidized frying oil based on solidphase extraction and gas chromatography. LWT-Food Sci. Technol. Nol. 2018, 92, 250−257. (17) Hu, J.-N.; Taki, F.; Sugiyama, S.; Asai, J.; Izawa, Y.; Satake, T.; Ozawa, T. Neutrophil-derived epoxide, 9,10-epoxy-12-octadecenoate, induces pulmonary edema. Lung 1988, 166, 327−337. (18) Liu, R.; Jin, Q.; Huang, J.; Liu, Y.; Wang, X.; Zhou, X.; Mao, W.; Wang, S. In vitro toxicity of aflatoxin b1 and its photodegradation products in hepg2 cells. J. Appl. Toxicol. 2012, 32, 276−281. (19) Iizaka, T.; Tsuji, M.; Oyamada, H.; Morio, Y.; Oguchi, K. Interaction between caspase-8 activation and endoplasmic reticulum stress in glycochenodeoxycholic acid-induced apoptotic hepg2 cells. Toxicology 2007, 241, 146−156. (20) Yeh, C. T.; Yen, G. C. Involvement of p38 MAPK and Nrf2 in phenolic acid-induced P-form phenol sulfotransferase expression in human hepatoma HepG2 cells. Carcinogenesis 2006, 27, 1008−1017. (21) Lin, Y.; Vermeer, M. A.; Bos, W.; van Buren, L.; Schuurbiers, E.; Miret-Catalan, S.; Trautwein, E. A. Molecular structures of citrus flavonoids determine their effects on lipid metabolism in HepG2 cells by primarily suppressing ApoB secretion. J. Agric. Food Chem. 2011, 59, 4496−4503. (22) Tu, Z.; Mosspierce, T.; Ford, P.; Jiang, A. Rosemary (Rosmarinus of f icinalis L.) extract regulates glucose and lipid metabolism by activating AMPK and PPAR pathways in HepG2 cells. J. Agric. Food Chem. 2013, 61, 2803−2810. (23) Wang, C.; Wang, L.; Li, W.; Yan, F.; Tian, M.; Wu, C.; Qi, L.; Wang, X.; Song, J.; Hou, X.; Chen, L. Irisin has no effect on lipolysis in 3T3-L1 adipocytes or fatty acid metabolism in HepG2 hepatocytes. Endocrine 2015, 49, 90−96. (24) Feldstein, A. E.; Lopez, R.; Tamimi, T. A.-R.; Yerian, L.; Chung, Y. M.; Berk, M.; Zhang, R.; McIntyre, T. M.; Hazen, S. L. Mass spectrometric profiling of oxidized lipid products in human nonalcoholic fatty liver disease and nonalcoholic steatohepatitis. J. Lipid Res. 2010, 51, 3046−3054.

(25) Leonarduzzi, G.; Gamba, P.; Gargiulo, S.; Biasi, F.; Poli, G. Inflammation-related gene expression by lipid oxidation-derived products in the progression of atherosclerosis. Free Radical Biol. Med. 2012, 52, 19−34. (26) Poli, G.; Sottero, B.; Gargiulo, S.; Leonarduzzi, G. Cholesterol oxidation products in the vascular remodeling due to atherosclerosis. Mol. Aspects Med. 2009, 30, 180−189. (27) Horie, K.; Miyata, T.; Maeda, K.; Miyata, S.; Sugiyama, S.; Sakai, H.; van Ypersole de Strihou, C.; Monnier, V. M.; Witztum, J. L.; Kurokawa, K. Immunohistochemical colocalization of glycoxidation products and lipid peroxidation products in diabetic renal glomerular lesions. Implication for glycoxidative stress in the pathogenesis of diabetic nephropathy. J. Clin. Invest. 1997, 100, 2995−3004. (28) Li, J.; Li, X.; Cai, W.; Liu, Y. Comparison of different polar compounds-induced cytotoxicity in human hepatocellular carcinoma HepG2 cells. Lipids Health Dis. 2016, 15, 30−35. (29) Dhibi, M.; Brahmi, F.; Mnari, A.; Houas, Z.; Chargui, I.; Bchir, L.; Gazzah, N.; Alsaif, M. A.; Hammami, M. The intake of high fat diet with different trans fatty acid levels differentially induces oxidative stress and non alcoholic fatty liver disease (NAFLD) in rats. Nutr. Metab. 2011, 8, 65−76. (30) Demozay, D.; Mas, J.-C.; Rocchi, S.; Van Obberghen, E. FALDH reverses the deleterious action of oxidative stress induced by lipid peroxidation product 4-hydroxynonenal on insulin signaling in 3T3-L1 adipocytes. Diabetes 2008, 57, 1216−1226. (31) Lobo, V.; Patil, A.; Phatak, A.; Chandra, N. Free radicals, antioxidants and functional foods: Impact on human health. Pharmacogn. Rev. 2010, 4, 118−126. (32) Greene, J. F.; Newman, J. W.; Williamson, K. C.; Hammock, B. D. Toxicity of epoxy fatty acids and related compounds to cells expressing human soluble epoxide hydrolase. Chem. Res. Toxicol. 2000, 13, 217−226. (33) Cao, W.; Wang, X.; Zhang, W.; Wang, X. Toxic effects of triacylglycerol polymer on macrophages in vitro. Eur. J. Lipid Sci. Technol. 2013, 115, 756−763. (34) Stemmer, U.; Dunai, Z. A.; Koller, D.; Pürstinger, G.; Zenzmaier, E.; Deigner, H. P.; Aflaki, E.; Kratky, D.; Hermetter, A. Toxicity of oxidized phospholipids in cultured macrophages. Lipids Health Dis. 2012, 11, 110. (35) Ozawa, T.; Sugiyama, S.; Hayakawa, M.; Satake, T.; Taki, F.; Iwata, M.; Taki, K. Existence of leukotoxin 9,10-epoxy-12octadecenoate in lung lavages from rats breathing pure oxygen and from patients with the adult respiratory distress syndrome. Am. Rev. Respir. Dis. 1988, 137, 535−540. (36) Zhang, X.; Wang, L.; Wang, R.; Luo, X.; Li, Y.; Chen, Z. Protective effects of rice dreg protein hydrolysates against hydrogen peroxide-induced oxidative stress in HepG-2 cells. Food Funct. 2016, 7, 1429−1437. (37) Nordberg, J.; Arnér, E. S. J. Reactive oxygen species, antioxidants, and the mammalian thioredoxin system 1. Free Radical Biol. Med. 2001, 31, 1287−1312. (38) Ceconi, C.; Boraso, A.; Cargnoni, A.; Ferrari, R. Oxidative stress in cardiovascular disease: Myth or fact? Arch. Biochem. Biophys. 2003, 420, 217−221. (39) Sohal, R.; Mockett, R.; Orr, W. Mechanisms of aging: An appraisal of the oxidative stress hypothesis. Free Radical Biol. Med. 2002, 33, 575−586. (40) Adly, A. A. M. Oxidative stress and disease: An updated review. Res. J. Immunol. 2010, 3, 129−145. (41) Del Rio, D.; Stewart, A. J.; Pellegrini, N. A review of recent studies on malondialdehyde as toxic molecule and biological marker of oxidative stress. Nutr., Metab. Cardiovasc. Dis. 2005, 15, 316−328. (42) Chow, C. K. Biological effects of oxidized fatty acids. In Fatty Acids in Foods and Their Health Implications, 3rd ed.; Chow, C. K., Ed.; CRC Press (Taylor and Francis Group): Boca Raton, FL, 2007; Chapter 36, pp 855−878, DOI: 10.1201/9781420006902.ch36. (43) Kangralkar, V. A.; Patil, S. D.; Bandivadekar, R. M. Oxidative stress and diabetes: A review. Int. J. Pharm. Appl. 2010, 1, 38−45. 5245


Article

Journal of Agricultural and Food Chemistry (44) Bhor, V. M.; Raghuram, N.; Sivakami, S. Oxidative damage and altered antioxidant enzyme activities in the small intestine of streptozotocin-induced diabetic rats. Int. J. Biochem. Cell Biol. 2004, 36, 89−97. (45) Halliwell, B.; Gutteridge, J. M. C. Free radicals and antioxidant protection: Mechanisms and significance in toxicology and disease. Hum. Toxicol. 1988, 7, 7−13.

5246


Epoxy Stearic Acid, an Oxidative Product Derived from Oleic Acid

Recommend Documents