|Home | About | Journals | Submit | Contact Us | Français|
Arsenic is a trace element that occurs naturally in the earth’s crust. It has been found to be a major contaminant in groundwater supply in several countries of the world. Whether ingested or inhaled, arsenic induces both systemic (skin disorders, cardiovascular diseases, anemia, peripheral neuropathy, liver and kidney damage) and carcinogenic (skin, lung, bladder and liver neoplasms) effects. However, its molecular mechanisms of toxicity are not completely understood. In this research, we used HepG2 cells as a model to study the cytotoxicity and oxidative stress associated with exposure to arsenic trioxide. We hypothesized that oxidative stress plays a role in arsenic trioxide induced cytotoxicity. To test this hypothesis, we performed both MTT [3-(4, 5-dimethylthiazol-2-yl)-2, 5-diphenyltetrazolium bromide] assay and trypan blue exclusion test for cell viability and the thiobarbituric acid test for lipid peroxidation. Data obtained from the MTT assay indicated that arsenic trioxide significantly reduced the viability of HepG2 cells, showing a LD50 value of about 23 μg/mL upon 24 h of exposure, indicating a dose-dependent response. Similar trend was obtained with the trypan blue exclusion test. Data generated from the thiobarbituric acid test showed a significant increase (p ≤ 0.05) in MDA levels in arsenic trioxide-treated HepG2 cells compared to control cells. Arsenic trioxide treatment significantly increased cellular content of reactive oxygen species (ROS), as evidenced by the increase in lipid peroxidation by-products. Taken together, these results indicate that arsenic trioxide is cytotoxic to HepG2 cells. This cytotoxicity is mediated by oxidative stress, a biomarker of cellular injury.
Arsenic is a naturally occurring element in the earth’s crust. It is estimated that several million people worldwide suffer the effects of chronic arsenic exposure resulting from environmental release. The majority of worldwide arsenic exposure stems from groundwater sources that contaminate the drinking water. Epidemiological findings show that exposure to arsenic results in vascular disorders, peripheral neuropathy, and various neoplastic diseases, such as, skin, lung, bladder, liver and kidney cancers (1, 2). It has been reported that oxidative stress is an important mechanism involved in heavy metal toxicity (3). It is through this process that oxidative damage occurs. Reactive oxygen species generated in response to arsenic exposure leads to an accumulation of intracellular hydrogen peroxide by activating flavoprotein dependent superoxide producing enzymes such as NADT+(NADPH) oxidase (4, 5). Arsenic induced oxidative stress has been shown to cause DNA damage through the production of superoxide and hydrogen peroxide radicals, specifically reactive oxygen species (6, 7). The present study was designed to use human liver carcinoma (HepG2) cells as a test model to determine whether oxidative stress plays a key role in arsenic trioxide-induced cytotoxicity.
Arsenic trioxide (As2O3), CASRN 1327-53-3, MW 197.84, with an active ingredient of 100% (w/v) arsenic in 10% nitric acid was purchased from Fisher Scientific in Houston, Texas. Growth medium DMEM-F12 containing 2.5 mM L-glutamine, 15 mM HEPES, 0.5 mM sodium pyruvate, and 1200 mg/L sodium bicarbonate was purchased from American Type Culture Collection (Manassas, VA). Ninety six- well plates were purchased from Costar (Cambridge, MA). Fetal bovine serum (FBS), antibiotics (penicillin G and streptomycin), phosphate buffered saline (PBS), G418 and MTT assay kit were obtained from Sigma Chemical Company (St. Louis, MO).
In the laboratory, cells were stored in the liquid nitrogen until use. They were thawed by gentle agitation of their containers (vials) for 2 minutes in a bath at 37°C. After thawing, the content of each vial of HepG2 cells were transferred to a 25 cm2 tissue culture flask, diluted with up to 10 mL of DMEM-F12 and supplemented with 10% (v/v) fetal bovine serum (FBS), 0.4mg/ml G418, 1% (w/v) penicillin/streptomycin. The 25 cm2 culture flasks (2 × 106 viable cells) were observed under the microscope, followed by incubation in a humidified 5 % CO2 incubator at 37° C. Three times a week, they were diluted to maintain under the same conditions at a density of 5 × 105/mL and harvested in the exponential phase of growth. The cell viability was assessed by the trypan blue exclusion test (Life Technologies) and manually counted using a hemocytometer.
Human liver carcinoma (HepG2) cells were maintained in DMEM-F12 and supplemented with 10% (v/v) fetal bovine serum (FBS), 0.4mg/ml G418, 1% (w/v) penicillin/streptomycin, and incubated at 37°C in humidified 5% CO2 incubator. To 180 μL aliquots in six replicates of the cell suspension (5× 105/mL) were seeded to 96 well polystyrene tissue culture plates, 20 μL aliquots of ATO solutions (1.25, 2.5, 5, 10, 20, and 40μg/mL) were added to each well using distilled water as solvent. Cells incubated in culture medium alone served as a control for cell viability (untreated wells). All chemical exposures were carried in 96 well tissue culture plates for the purpose of chemical dilutions. Cells were placed in the humidified 5% CO2 incubator for 24 hrs at 37°C. After incubation, 20 μL aliquots of MTT solution (5 mg/mL in PBS) were added to each well and re-incubated for 4 hours at 37° C followed by low centrifugation at 800 rpm for 5 minutes. Then, the 200 μL of supernatant culture medium were carefully aspirated and 200 μL aliquots of dimethylsulfoxide (DMSO) were added to each well to dissolve the formazan crystals, following by incubation for 10 minutes to dissolve air bubbles. The culture plate was placed on a Biotex Model micro-plate reader and the absorbance was measured at 550 nm. The amount of color produced is directly proportional to the number of viable cell. All assays were performed in six replicates for each concentration. Cell viability rate was calculated as the percentage of MTT absorption as follows: % survival = (mean experimental absorbance/mean control absorbance×100).
Data were presented as means ± SDs. Statistical analysis was done using one way analysis of variance (ANOVA) for multiple samples and Student’s t-test for comparing paired sample sets. P-values less than 0.05 were considered statistically significant. The percentages of cell viability and MDA levels were presented graphically in the form of histograms, using Microsoft Excel computer program.
Data presented in Figure 2 clearly demonstrate that arsenic trioxide has a significant cytotoxic effect on HepG2 cells. A LD50 value of 23.2 ± 6.03 μg/mL was computed upon 24 h of exposure. At low doses of exposure, arsenic trioxide produces a slight increase in cell viability. On the other hand, it produces a non-linear gradual decrease in cell viability at higher doses.
The standard curve generated from lipid peroxidation assay is presented in Figure 3, and the effect of arsenic trioxide on lipid peroxidation is presented in Figure 4. Results of this assay showed an elevated production of MDA in HepG2 cells with increasing doses of arsenic trioxide. Data presented in this figure demonstrated that arsenic trioxide treatment resulted in a significant increase in MDA level, an indicator of lipid peroxidation. These data indicate that arsenic trioxide induces lipid peroxidation (a biomarker of cellular injury) as result of oxidative stress.
We examined the cytotoxic effect of arsenic trioxide on HepG2 cells. Findings from our study showed that arsenic trioxide (ATO) is cytotoxic to this cell line upon 24 hours of exposure. A previous study has shown that when using 0.5 to 1 μM/L of ATO, apoptosis was induced in the monocytic cells line NB4 (8). Cytotoxicity studies of two multiple myeloma (MM) derived cell lines, RPMI 8226 and U266, found that 1.0 μM/L ATO inhibited cell proliferation resulting in a weak degree of apoptosis induction, and 2.0 μM/L ATO- induced cell apoptosis. These results showed that ATO exerts apoptosis inducing and growth-inhibiting effects on MM derived cells (9). Recently, we reported that ATO is cytotoxic to human liver carcinoma (HepG2) cells, showing a LD50 of 8.55 ± 0.58 μg/ml after 48 hrs of exposure (10, 11). Findings from other studies suggest that low doses of arsenic are effective in acute promyelocytic leukemia treatment, and show considerable promise in preclinical models of other tumor types (12).
To elucidate the possible involvement of reactive oxygen species production in arsenic-induced toxicity, we performed the lipid peroxidation assay. Our results from this assay show that arsenic trioxide significantly increases (p<0.05) MDA levels in human liver carcinoma (HepG2) cells. These results indicate that oxidative stress may play a critical role in arsenic induced toxicity. Oxidative stress is an imbalance between the production and disposal of reactive oxygen. Arsenic is known for generation of reactive oxygen species (ROS). Studies have shown that exposure to arsenic generates superoxide, hydrogen peroxide, and hydroxyl radicals in keratinocyte cells in vitro (13). The process of lipid peroxidation is initiated by the attack of a free radical produced by a heavy metal such as arsenic, on unsaturated lipids and the resulting chain reaction is terminated by the production of lipid breakdown products, lipid, alcohols, aldehydes and malondialdehyde (14).
Results from this study indicate at the cellular level that arsenic trioxide significantly reduced the viability of human liver carcinoma (HepG2) cells at higher level of exposure (10–40μg/mL) in a dose-dependent manner. However, a stimulatory effect was observed at lower level of exposure (1.2–5μg/mL), as characterized by the slight increase in cell viability in arsenic trioxide-treated cells compared to control cells. At the molecular level, arsenic trioxide treatment resulted in a significant increase in lipid peroxidation. Findings from this study indicate that arsenic trioxide is cytotoxic to HepG2 cells, and this cytotoxicity may be mediated through the generation of reactive oxygen species, and subsequent induction of lipid peroxidation.
This research was financially supported by the National Institutes of Health (Grant # 1G12RR13459), through the RCMI-Center for Environmental Health at Jackson State University.