C2C12 myoblasts obtained from the American Type Culture Collection (ATCC, Manassas, VA, USA) were grown in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and 100 µg/ml penicillin/streptomycin (both from WelGENE Inc., Daegu, Korea) in a humidified 5% CO₂ atmosphere at 37°C. 7,8-DHF (purity, ≥98%; Sigma-Aldrich, St. Louis, MO, USA) was dissolved in dimethyl sulfoxide (DMSO; Sigma-Aldrich) and adjusted to final concentrations using complete DMEM prior to use. In order to investigate the effects of PI3K/Akt and MAPKs pathways on the induction of Nrf2, p-Nrf2 and HO-1 by 7,8-DHF, LY294002 (an inhibitor of Akt), PD98059 (an inhibitor of ERK), SB203580 (an inhibitor of p38) and SP600125 (an inhibitor of JNK) were obtained from Calbiochem (San Diego, CA, USA).

As a measure of the overall levels of cell viability, the C2C12 myoblasts were assessed by 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT; Sigma-Aldrich) assay. Briefly, the C2C12 myoblasts were seeded in 6-well plates at a density of 1×10⁵ cells/well. Following 24 h of incubation, the cells were treated with the indicated concentrations of 7,8-DHF (2.5, 5, 10, 15 or 20 µM) in the absence or presence of hydrogen peroxide (H₂O₂) and/or zinc protoporphyrin IX (ZnPP, a specific inhibitor of HO-1; Sigma-Aldrich) for the indicated periods of time (24 or 6 h). MTT working solution was then added to the culture plates followed by incubation at 37°C for 3 h. The culture supernatant was completely removed from the wells, and DMSO was added to dissolve the formazan crystals. The absorbance of each well was then measured at 540 nm using a microplate reader (Molecular Devices, Palo Alto, CA, USA). The protective effects of 7,8-DHF against growth inhibition were assessed as a percentage of cell viability, and the vehicle (0.05 mM DMSO)-treated cells were considered 100% viable.

The cell suspension was mixed with 0.5% low melting agarose (LMA) at 37°C, and the mixture was spread on a fully frosted microscopic slide precoated with 1% normal melting agarose (NMA). After the solidification of the agarose, the slide was covered with 0.5% LMA and then immersed in a lysis solution [2.5 M NaCl, 100 mM Na-ethylenediaminetetraacetic acid (EDTA), 10 mM Tris, 1% Triton X-100 and 10% DMSO, pH 10] for 1 h at 4°C. The slides were then placed in a gel electrophoresis apparatus containing 300 mM NaOH and 10 mM Na-EDTA (pH 13) for 40 min to allow for the unwinding of the DNA and the expression of alkali-labile damage. An electrical field was then used (300 mA, 25 V) for 20 min at 4°C to draw the negatively charged DNA toward the anode. Following electrophoresis, the slides were washed 3 times for 5 min at 4°C in neutralizing buffer (0.4 M Tris, pH 7.5), followed by staining with 20 µg/ml propidium iodide (PI; Sigma-Aldrich). The slides were examined under a fluorescence microscope (Carl Zeiss, Oberkochen, Germany).

Whole-cell protein extracts from the C2C12 myoblasts were prepared with cell lysis buffer (20 mM sucrose, 1 mM EDTA, 20 µM Tris-HCl, pH 7.2, 1 mM dithiothreitol, 10 mM KCl, 1.5 mM MgCl₂ and 5 µg/ml aprotinin) for 30 min. In a parallel experiment, nuclear proteins were prepared using nuclear extraction reagents (Pierce Biotechnology, Rockford, IL, USA) according to the manufacturer’s instructions. The protein extracts were quantified using the Bio-Rad kit (Pierce Biotechnology). For western blot analysis, equal amounts of protein extracts were separated by denaturing sodium dodecyl sulfate (SDS)-polyacrylamide gels and transferred electrophoretically onto nitrocellulose membranes (Schleicher & Schuell, Keene, NH, USA). The membranes were then incubated overnight at 4°C with primary antibodies, probed with enzyme-linked secondary antibodies [mouse IgG, HRP-linked whole antibody (NA931) and rabbit IgG, HRP-linked whole antibody (NA934), Amersham Corp., Arlington Heights, IL, USA)] for 1 h at room temperature, and detected using an enhanced chemiluminescence (ECL) detection system (all from Amersham Co.). The antibodies used were as follows: iNOS (1:500; SC-7271, mouse monoclonal; Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA), TNF-α (1:500; #3707S, rabbit polyclonal; Cell Signaling Technology, Inc., Danvers, MA, USA), IL-1β (1:500; SC-7884, rabbit polyclonal), NF-κB p65 (1:500; SC-109, rabbit polyclonal), IκBα (1:500; SC-371, rabbit polyclonal), Akt (1:500; SC-8312, rabbit polyclonal), p-Akt (1:500; SC-101629, rabbit polyclonal), ERK (1:1,000; SC-154, rabbit polyclonal; all from Santa Cruz Biotechnology, Inc.), p-ERK (1:500; #9106S, mouse monoclonal; Cell Signaling Technology, Inc.), p38 (1:1,000; SC-535, rabbit polyclonal; Santa Cruz Biotechnology, Inc.), p-p38 (1:500; #9211S, rabbit polyclonal), JNK (1:1,000; #9252S, rabbit polyclonal), p-JNK (1:500; #9255S, mouse monoclonal; all from Cell Signaling Technology, Inc.), Nrf2 (1:500; SC-13032, rabbit polyclonal; Santa Cruz Biotechnology, Inc.), p-Nrf2 (1:500; ab76026, rabbit monoclonal; Abcam, Inc., Cambridge, UK), HO-1 (1:500; SC-136960, mouse monoclonal), Lamin B (1:500; SC-6216, goat polyclonal) and β-actin (1:1,000; sc-1616, goat polyclonal; all from Santa Cruz Biotechnology, Inc.). Actin and poly(ADP ibose) polymerase (PARP) were used as the internal controls of the total cellular and nuclear proteins, respectively.

To quantitatively assess the cell apoptotic rate, a fluorescein-conjugated Annexin V (Annexin V-FITC) staining assay was performed according to the manufacturer’s instructions (BD Biosciences, San Jose, CA, USA). Briefly, the cells were stained with 5 µl Annexin V-FITC and 5 µl PI. Following incubation for 15 min at room temperature in the dark, the degree of apoptosis was quantified as a percentage of the Annexin V-positive and PI-negative cells by flow cytometry, as previously described (22).

The intracellular accumulation of ROS was determined using the fluorescent probe, 2′,7′-dichlorodihydrofluorescein diacetate (H2DCFDA; obtained from Molecular Probes, Eugene, OR, USA). In order to monitor the generation of ROS, the cells were treated with 5 mM N-acetyl-L-cysteine (NAC; Sigma-Aldrich) for 30 min and then treated with 7,8-DHF for 6 h. After the addition of 10 µM H2DCFDA for 20 min at room temperature in the dark, ROS production in the cells was monitored using a flow cytometer (BD Biosciences) with CellQuest Pro software, as previously described (23).

Nrf2 siRNA and control siRNA were purchased from Santa Cruz Biotechnology, Inc. The cells were transfected with the siRNAs according to the manufacturer’s instructions using Lipofectamine 2000 Transfection Reagent (Life Technologies, Carlsbad, CA, USA). For transfection, the cells were seeded in 6-well culture plates and incubated with the control siRNA or Nrf2 siRNA at 50 nM for 6 h in serum-free OPTI-MEM medium. Following incubation, the transfected cells were subjected to treatment as described in the figure legends and as previously described (24).

Data are expressed as the means ± standard deviation (SD). One-way analysis of variance (ANOVA) was used for comparisons in the experiments with multiple time points and concentrations. When ANOVA indicated statistical significance, Duncan’s multiple range test was used to determine which means were significantly different. A probability value of P<0.05 was used as the criterion for statistical significance.

We first determined the effects of 7,8-DHF on the viability of C2C12 myoblasts by MTT assay. The cells were treated with a range of 7,8-DHF concentrations, from 2.5 to 20 µM for 24 h (Fig. 1A). Treatment of the C2C12 myoblasts with up to 10 µM 7,8-DHF did not result in any cytotoxic effects, whereas cell viability decreased in a dose-dependant manner following treatment with 7,8-DHF at concentrations >10 µM (Fig. 1A). Therefore, the dose of 10 µM 7,8-DHF was selected as the optimal dose for examining the cytoprotective effects of 7,8-DHF against H₂O₂-induced cell damage. To examine the protective effects of 7,8-DHF against H₂O₂-induced cytotoxicity, we treated the C2C12 myoblasts with 10 µM 7,8-DHF 1 h prior to exposure to H₂O₂, and cell viability was then measured by MTT assay. Following exposure to 1 mM H₂O₂ alone, cell viability was reduced to approximately 60% at 6 h, whereas the H₂O₂-induced decrease in of cell viability was significantly attenuated by pre-treatment with 7,8-DHF (Fig. 1B). These results clearly indicate that the exposure of C2C12 myoblasts to 7,8-DHF confers a significant protective effect against oxidative stress.

As DNA strand breakage is considered one of the most frequent types of damage that can be induced by oxidative stress (25), we examined the effects of 7,8-DHF on H₂O₂-mediated damage to C2C12 cell DNA using single-cell gel electrophoresis (comet assay) and western blot analysis. Treatment with H₂O₂ alone induced significant DNA damage in the C2C12 myoblasts; however, this adverse effect was markedly reduced by pre-treatment with 7,8-DHF (Fig. 2A). In addition, our results revealed that the exposure of C2C12 myoblasts to H₂O₂ resulted in an upregulation in the levels of the phosphorylated histone variant H2A.X (p-γH2AX) at serine 139, a sensitive marker of DNA double-strand breaks (25); however, pre-treatment with 7,8-DHF resulted in a significant decrease in p-γH2AX expression (Fig. 2B). In order to investigate the protective effects of 7,8-DHF against H₂O₂-induced apoptosis, the frequency of apoptotic cells was detected by flow cytometry, and the results revealed that the treatment of the cells with 7,8-DHF prior to exposure to H₂O₂ protected the C2C12 myoblasts against apoptosis.

Using H2DCFDA assay, we then investigated whether 7,8-DHF affects intracellular ROS generation induced by exposure of the cells to H₂O₂. As expected, significantly increased levels of ROS were detected following the epxosure of the cells to H₂O₂ compared with the levels observed in the untreated cells; however, this increase in ROS levels was significantly inhibited by treatment with 7,8-DHF (Fig. 3A). Moreover, no fluorescence was detected in the cells treated with 7,8-DHF alone (data not shown), indicating that 7,8-DHF itself does not contribute to ROS generation. As a positive control, the ROS scavenger, NAC, was also used. Treatment with NAC attenuated the H₂O₂-induced ROS generation and decreased the apoptotic rate; it also reversed the decrease in cell viability induced by exposure to H₂O₂; these effects were similar to those of 7,8-DHF (Fig. 3B and C). The results indicate that the H₂O₂-induced induction of apoptosis and the reduction in cell viability are mediated by the generation of ROS, and that 7,8-DHF exerts a potent ROS-scavenging effect, protecting the C2C12 myoblasts against H₂O₂-damage.

It has been well documented that antioxidant enzymes play an important role against oxidative stress (1,2); thus, we hypothesized that the effects of 7,8-DHF may be mediated by the induction of antioxidant enzymes. Treatment of the C2C12 myoblasts with 7,8-DHF induced the protein expression of HO-1 in a time-dependent manner; however, the levels of the other antioxidant enzymes, NQO-1 and thioredoxin reductase 1 (TrxR1), were unaffected (Fig. 4A). Previous studies have demonstrated that, under normal conditions, Nrf-2 is inactive and bound in the cytosol by Keap1, and that the translocation of Nrf2 into the nucleus is essential for the transactivation of various target genes, such as HO-1. Moreover, the phosphorylation of Nrf2 at Ser40 by several kinases is also a critical process in its stabilization and nuclear translocation (6,7). Therefore, in this study, we examined the phosphorylation and subcellular localization of Nrf2 following treatment with 7,8-DHF in order to confirm the Nrf2-activating properties of 7,8-DHF. We observed that 7,8-DHF increased the expression levels of total and phosphorylated Nrf2 in a time-dependent manner (Fig. 4A). Furthermore, western blot analysis of the nuclear fraction revealed a significant augmentation of Nrf2 phosphorylation and nuclear accumulation following treatment with 7,8-DHF, in a time-dependent manner (Fig. 4B).

We developed a Nrf2 gene knockout model using siRNA transfection in order to demonstrate the importance of Nrf2 upregulation. The results of western blot analysis revealed that the silencing of Nrf2 using specific siRNA abolished the 7,8-DHF-induced increase in Nrf2 expression and HO-1 upregulation (Fig. 5A), which is evidence that the augmentation of HO-1 is mediated by Nrf2. To further confirm the involvement of Nrf2, the protective effects of 7,8-DHF against the H₂O₂-induced decrease in cell viability were determined in cells in which Nrf2 had been knocked down. As shown in Fig. 5B, siNrf2 transfection abrogated the cytoprotective effects of 7,8-DHF compared to the control siRNA-transfected cells, thus proving that 7,8-DHF diminishes the H₂O₂-induced decrease in cell viability through the activation of the Nrf2/HO-1 signaling pathway.

To further determine whether the 7,8-DHF-induced antioxidant and cytoprotective activities against oxidative stress in C2C12 cellsmyoblasts are mediated through the activation of the Nrf2/HO-1 pathway, the C2C12 myoblasts were pre-incubated with or without a specific inhibitor of HO-1, ZnPP, and the levels of ROS and cell viability were then assessed. ZnPP abrogated the protective effects of 7,8-DHF against the H₂O₂-induced production of ROS and the decrease in cell viability (Fig. 6). These results indicate that 7,8-DHF exerts its protective effects by inducing the cellular defense mechanism against oxidative stress through the Nrf2-related cytoprotective pathway, and that HO-1 plays a crucial role in this protection of C2C12 myoblasts.

A number of studies have noted that multiple phosphorylation cascades participate in the regulation of the translocation of Nrf2 and Nrf2-mediated HO-1 gene expression (26–28). Thus, to identify the upstream signaling events involved in the 7,8-DHF-mediated activation of Nrf2 and the induction of HO-1, the potential involvement of PI3K/Akt and mitogen-activated protein kinases (MAPKs) were explored. Although the total levels of Akt, a downstream target of PI3K, did not show a notable change, the Akt phosphorylation levels markedly increased following treatment with 7,8-DHF within 30 min (Fig. 7A). However, treatment with LY294002, a pharmacological inhibitor of PI3K, prevented the increase in the phosphorylation levels of Nrf2 and resulted in a blockade of Nrf2 and HO-1 induction which was by 7,8-DHF (Fig. 7B), suggesting that the 7,8-DHF-induced activation of the Nrf2/HO-1 pathway may be a process necessary to the PI3K cascade.

Subsequently, we investigated the effects of 7,8-DHF on the activation of MAPKs in C2C12 myoblasts. The increase in the phosphorylation levels of ERK and p38 MAPK were observed 30 min following treatment with 7,8-DHF and this increase was sustained for up to 2 h following treatment with 7,8-DHF. However, there were no notable changes observed in the phosphorylation levels of c-Jun N-terminal kinase (JNK) (Fig. 8A). When a selective inhibitor of ERK (PD98059) was utilized, the induction and phosphorylation of Nrf2 were blocked and, accordingly, HO-1 induction was diminished (Fig. 8B). By contrast, inhibitors of p38 MAPK (SB203580) and JNK (SP600125) did not reduce the 7,8-DHF-induced HO-1 and Nrf2 expression or Nrf2 phosphorylation. These data indicate that the 7,8-DHF-mediated activation of the Nrf2/HO-1 pathway involves the ERK pathway, but not the p38 MAPK and JNK pathways. Taken together, these observations support the hypothesis that 7,8-DHF activates the PI3K/Akt and ERK pathways, which subsequently induces Nrf2/HO-1 activation in C2C12 myoblasts.