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Introduction

It is well known that anti-neoplastic drugs interfere with the structure and functions of DNA directly or indirectly. However, they sometimes not only affect target cells but also normal cells. In light of this, comprehension of the anticancer functions still require investigation in order to reduce the side effects before their use in the direct treatment of patients. Thus, there is not only the need to evaluate the impairment caused by anticancer drugs on the whole organism but also to investigate the effects of genotoxic alterations at a cellular level (1). Currently, numerous compounds from natural plants have been shown to induce cell death via the induction of cell apoptosis. However, the interruption of cell DNA damage is also needed because these effects can lead to cell death. Some anticancer drugs such as cisplatin or etoposide have been shown to induce DNA damage and eventually cell death (2). Thus, focusing on the ability of these compounds to interfere with DNA and produce DNA damage will be helpful and critical to understand how these compounds induce cell death.

Casticin, one of the ingredients derived from Fructus viticis (3), has been shown to exhibit anticancer activity in prostate (4), breast (5), colon (6,7), lung (8,9), cervical (10), gastric (11) and ovarian cancer (12), glioma (13) and leukemia (14) Recently it was reported that forkhead box O3 (FOXO3a) is a critical mediator of the inhibitory effects of casticin on apoptosis in breast cancer cells (3). Furthermore, casticin significantly induced cell apoptosis through the activation of the apoptosis signal-regulating kinase 1-c-Jun N-terminal kinase (ASK1-JNK)-Bim signaling cascade and the accumulation of reactive oxygen species (ROS) in colon cancer cells (15). However, there is no available information to show that casticin induces cell apoptosis in melanoma cancer cells. Furthermore, there is no report showing that casticin induces DNA damage and affects DNA repair-associated protein expression levels in melanoma cells.

After melanoma becomes metastatic melanoma, it is characterized by a high mortality rate (16) due to a universal resistance to standard chemotherapy (17). Hence, the motality rate from unresectable melanoma continues to rise (18). Presently, the ineffectiveness of the treatments available, encourage additional studies to identify novel therapeutic molecules, delivery systems, and/or combination therapies for the treatment of melanoma (19). Casticin may be a potential antitumor agent with both antitumor and anti-proliferative activities. However, the effects of casticin on DNA damage and repair with associated protein expression are not widely known. Thus, the objective of this study was to investigate DNA damage and repair of melanoma B16F10 cells and our results confirmed that casticin induced DNA damage and affected DNA repair systems in vitro.

Materials and methods

Chemicals and reagents

Casticin, dimethyl sulfoxide (DMSO), propidium iodide (PI), Trypsin-EDTA, penicillin-streptomycin, anti-MGMT (cat no. M3068), anti-PARP (cat no. P248), anti-p-ATMSer1981 (cat no. SBA4300100) and anti-β-actin (cat no. A5316) were all purchased from Sigma Chemical Co. (St. Louis, MO, USA). Anti-DNA-PK (cat no. PC127) was purchased from Calbiochem (San Diego, CA, USA). Anti-p-H2A.X (cat no. GTX80694) and anti-BRCA1 (cat no. GTX70111), were purchased from GeneTex Inc. (Irvine, CA, USA). Anti-p-p53 (cat no. sc-7997) was obtained from Santa Cruz Biotechnology (Santa Cruz, CA, USA). Anti-MDC1 (cat no. 05-1572) was purchased from Millipore (Billerica, MA, USA). Dulbecco's modified Eagle's medium (DMEM) and fetal bovine serum (FBS) were purchased from Gibco®/Invitrogen Life Technologies (Carlsbad, CA, USA).

Cell culture

The murine melanoma cell line (B16F10) was purchased from the Food Industry Research and Development Institute (Hsinchu, Taiwan). Cells were grown in 75-cm2 flasks with DMEM supplemented with 10% FBS, 100 U/ml penicillin and 100 µg/ml streptomycin at 37°C in 5% CO2 humidified incubators (20).

Cellular morphology and viability examination

B16F10 cells were plated at a density of 1×105 cells/well into 12-well plates in DMEM. After the required confluency was reached, cells were exposed to 0, 20, 30 and 40 µM of casticin for 24 and 48 h in a 5% CO2 incubator at 37°C. Cells were examined and their images were captured by contrast phase microscopy at ×200 magnification. Subsequently, the cells were collected, washed and stained with PI (5 µg/ml) in phosphate-buffered saline (PBS) and were analyzed by flow cytometry (Becton-Dickinson, San Jose, CA, USA) for the total percentage of viable cells as previously described (21).

4′,6-Diamidino-2-phenylindole dihydrochloride (DAPI) staining for DNA condensation examination

B16F10 cells (1.5×105 cells/well) were plated onto a 6-well plate for 24 h and then were exposed to casticin (30 µM) for 0, 6, 24 and 48 h. After treatment, 4% formaldehyde in PBS was used to fix cells for 10 min and then DAPI staining followed. After staining, the cells were examined and their images were captured using a fluorescence microscope at ×200 magnification as previously described (21).

Western blotting for examination of protein expression

B16F10 cells (1×106 cells/dish) were plated onto a 10-cm dish and were incubated with 30 µM of casticin for 0, 6, 24 and 48 h. Cells were collected, suspended in sodium dodecyl sulfate (SDS) sample buffer, sonicated, boiled for 10 min as previously described (21) and were centrifuged at 12,000 rpm for 15 min. The supernatant was collected and the concentrations of the total protein were determined by Bio-Rad protein assay kit (Bio-Rad Laboratories, Hercules, CA, USA). The cells were electro-phoresed by 10% sodium dodecyl sulfate-polyacrylamide gel (SDS-PAGE) and were transferred to polyvinylidene fluoride (PVDF) membranes (Millipore, Bedford, MA, USA). Immune complexes were formed by incubation of proteins with primary antibodies (anti-MGMT, anti-BRCA1, anti-PARP, anti-p-p53, anti-MDC1, anti-DNA-PK, anti-p-ATM and anti-β-actin) at 4°C (overnight) followed by incubation with a secondary antibody. Immunoreactive protein bands were visualized with a chemiluminescent detection system and the protein expression levels were measured as described by the manufacturer (20,21).

Confocal laser microscopy for examination of protein translocation

B16F10 cells were plated at a density of 1.5×105 cells/well on a 6-well plate and were treated with 30 µM of casticin for 48 h. After treatment, cells were fixed in 4% formaldehyde in PBS for 15 min and 0.1% Triton X-100 in PBS was added to permeable cells. Subsequently, cells were washed with PBS and blocked with 1% BSA in PBS for 60 min and then they were stained with primary anti-p-p53 and anti-p-H2A.X (green fluorescence) overnight followed by staining with a secondary antibody (FITC-conjugated goat anti-mouse IgG). After being washed, cells were stained using PI (red fluorescence) for nuclei. All samples were mounted and photomicrographed by using a Leica TCS SP2 confocal spectral microscope (Leica Microsystems, Heidelberg, Mannheim, Germany) as previously described (21).

Statistical analysis

The comparisons between the casticintreated and the untreated groups were performed using the Student's t-test, to determine the statistical significance of the differences between these groups. P<0.05 was considered to be significant.

Results

Casticin induces cell morphology and decreases the total viability of the B16F10 cells

B16F10 cells were treated with various concentrations of casticin (0, 20, 30 and 40 µM) at 24 and 48 h. Cells were examined for morphological changes and images were captured using a phase contrast microscope at ×200 magnification (Fig. 1). The results indicated that casticin induced cell morphological changes in a dose-dependent manner. Cells were collected in order to measure the percentage of viable cells by flow cytometric assay (Fig. 2). The results indicated that the percentage of total cell viability was decreased significantly after treatment with casticin and that this effect was dose-dependent. The treatment of casticin at 48 h had a higher effect than that at 24 h.

Figure 1 Casticin induces cell morphological changes of viable mouse melanoma B16F10 cells. Cells (1×105 cells/well) were placed in 12-well plates and were treated with casticin (0, 20, 30 and 40 µM) for 48 h. Cells were examined and images were captured using phase contrast microscopy at ×200 magnification.

Figure 2 Casticin decreases the percentage of viable mouse melanoma B16F10 cells. Cells (1×105 cells/well) were placed in 12-well plates and were treated with casticin (0, 20, 30 and 40 µM) for 24 (A) and 48 h (B). Cells were harvested and stained with PI (5 µg/ml) and analyzed by flow cytometry as described in 'Materials and methods'. *P<0.05 indicates a significant difference between the casticin-treated and the untreated groups.

Casticin induces nuclear DNA condensation of B16F10 cells

In order to further confirm whether casticin induced cell death via nuclear DNA condensation in the B16F10 cells, we selected 30 µM of casticin for treatment with cells at 0, 6, 24 and 48 h, Subsequently the cells were stained with DAPI to examine the formation of DNA condensation (Fig. 3A and B). The results indicated that casticin induced nuclear DNA condensation in the B16F10 cells and this effect was time-dependent.

Figure 3 DAPI staining for nuclear DNA damage and condensation in the B16F10 cells. Cells (1.5×105 cells/well) were maintained in 6-well plates for 24 h and were treated with 30 µM of casticin for 0, 6, 24 and 48 h. Then the cells were stained with DAPI and were examined, and their images were captured using a fluorescence microscope at ×200 magnification as described in 'Materials and methods'. (A) Representative images of DAPI staining; (B) mean of fluorescence (% of control). *P<0.05 indicates a significant difference between the casticin-treated and the untreated groups.

Casticin affects DNA damage of the associated proteins in the B16F10 cells

Cells were treated with 30 µM of casticin for 0, 6, 24 and 48 h and then DNA damage of the associated proteins such as O6-methylguanine-DNA methyltransferase (MGMT), p-H2A.X, breast cancer 1 and early onset (BRCA1), poly(ADP-ribose) polymerase (PARP), phospho-p53 tumor suppressor protein (p-p53), mediator of DNA damage checkpoint 1 (MDC1), DNA-dependent protein kinase (DNA-PK) and phospho-ataxia telangiectasia mutated kinase (p-ATM) were examined by western blot analysis (Fig. 4). The results indicated that casticin decreased the protein levels of MGMT and BRCA1 (Fig. 4A) and MDC1 (Fig. 4B), but increased the levels of p-H2A.X and PARP (Fig. 4A), p-p53 and p-ATM (Fig. 4B) in the B16F10 cells. These effects are associated with DNA damage and repair that may lead to cell death.

Figure 4 Western blot analysis of protein levels of the proteins associated with DNA damage and repair in the B16F10 cells. Cells (1×106 cells/well) were placed in a 10-cm dish and then were incubated with 30 µM of cas-ticin for 0, 6, 24 and 48 h. The total amount of proteins was determined and the amounts of protein from each treatment were measured by SDS-PAGE and immunoblotting as described in 'Materials and methods'. (A) MGMT, p-H2A.X, BRCA1 and PARP. (B) p-p53, MDC1, DNA-PK and p-ATM.

Casticin affects the translocation of p-p53 and p-H2A.X in the B16F10 cells

To further confirm whether casticin affects DNA damage in associated protein translocation in the B16F10 cells, cells were treated with 30 µM of casticin and then they were examined by confocal microscopy (Fig. 5). The results revealed that casticin increased the p-p53 (Fig. 5A) and p-H2A.X (Fig. 5B) expression levels in the cytoplasm when compared to the control groups and these observations indicate that casticin induces DNA damage and repair and may also regulate p-p53 and p-H2A.X in the cytoplasm in the B16F10 cells.

Figure 5 Confocal laser system microscopy was used to examine the protein expression in the B16F10 cells. Cells (1.5×105 cells/well) were kept on a 6-well plate and were incubated with 30 µM of casticin for 48 h. Subsequently, they were fixed in 4% formaldehyde in PBS for 15 min followed by 0.1% Triton X-100 in PBS for 15 min. Immunostaining was then performed as described in 'Materials and methods'. (A) p-p53; (B) p-H2A.X. Both samples were examined and images were captured using a Leica TCS SP2 confocal spectral microscope.

Discussion

Based on the review of the literature, casticin was found to induce cell death (cytotoxic effects) via both induction of cell cycle arrest and apoptosis in many types of human cancer cells, but there is no available information showing that casticin induces DNA damage and repair and affects associated protein expression in human cancer cells. Therefore, in the present study, we investigated the cytotoxic effects of casticin and whether, through the induction of DNA damage, it affected DNA repair and associated protein expression levels in mouse melanoma B16F10 cells in vitro. After B16F10 cells were exposed to various concentrations of casticin we found that i) casticin induced cell morphological changes (Fig. 1) and decreased the total cell viability (percentage of viable cells) in a concentration- and time-dependent manner (Fig. 2); ii) a time-dependent increase in nuclear DNA condensation was observed in the B16F10 cells after exposure to casticin, which was assayed by DAPI staining (Fig. 3); iii) casticin decreased the proteins levels of MGMT and BRCA1 (Fig. 4A), and MDC1 (Fig. 4B), and increased the levels of p-H2A.X and PARP (Fig. 4A), p-p53 and p-ATM (Fig. 4B) in the B16F10 cells and these effects were time-dependent; iv) casticin induced DNA damage and repair and may also regulate p-p53 (Fig. 5A) and p-H2A.X (Fig. 5B) which are increased in the cytoplasm when compared to the control groups in the B16F10 cells.

We observed that casticin induced cell morphological changes and decreased the total percentage of viable B16F010 cells in a dose-dependent manner at 20–40 µM. Thus, we further examined whether casticin induced cell death and was associated with induction of DNA damage in the B16F10 cells. Based on the results from Fig. 3, it was revealed that casticin induced DNA damage and condensation in the B16F10 cells which were assayed by DAPI staining, respectively. It has been reported that specific and bulky DNA lesions which trigger cell apoptosis have been identified (22). It has been well documented that DAPI staining can reveal DNA fragmentation and nuclear DNA condensation and our results indicated that casticin induced nuclear DNA condensation in a time-dependent manner. Recently, a new type of anti-neoplastic therapy has emerged, whose aim is to manipulate DNA damage response (DDR) (23–25) as DDR inhibition has been proven as an effective treatment for cancer. It has been reported that oxidative DNA damage has been recognized to be an etiological factor in aging and in the development of systemic diseases including cancer in the human population (26,27). In cells, DNA repair enzymes monitor chromosomes and correct damaged nucleotides to prevent these adverse effects (28).

In the present study, our findings are the first to provide information regarding casticin-induced DNA damage and the affect on the DNA repair system in the B16F10 cells (Fig. 3). Western blotting (Fig. 4) indicated that casticin induced DNA damage and affected repair in associated protein expression levels such as MGMT and BRCA1 (Fig. 4A), MDC1 and DNA-PK (Fig. 4B) but increased the levels of p-H2A.X and PARP (Fig. 4A) and p-p53 and p-ATM (Fig. 4B) in the B16F10 cells. It has been reported that MGMT is a DNA repair enzyme which eliminates O6 methylguanines (29) and that the inhibition of MGMT may be strategic in increasing tumor susceptibility to chemotherapy (30). In breast and ovarian cancers, BRCA1 plays an important role in DNA repair in the maintainance of genomic stability (31) and in breast cancer, BRCA1 promoter methylation was found to be positively associated with increased mortality (32). MDC1 and BRCA1 represent important assets in the repair of double-strand breaks after DNA damage occurs (33,34). Furthermore, MDC1 may affect the radiosensitivity of tumor cells (35). It has been reported that DNA-PK is a serine/threonine protein kinase and is expressed in most mammalian cells (36). DNA-PK plays an important role in the main repair pathway of DNA double-strand breaks and cells deficient in DNA-PK exhibit hypersensitivity to radio/chemotherapy (37–39). It has also been reported that anticancer drugs such as 5-FU induce DNA double-strand breaks, and the presence of p-H2A.X which is a phosphorylated form of the histone H2A.X which has been shown to be a specific marker for the detection of these DNA breaks was observed (40). Based on this observation, we suggest that casticin induced double-strand breaks in the B16F10 cells.

Moreover, it has been reported that the ATM/p53 pathway is involved in the apoptosis of various cancer cells induced by chemotherapy drugs (41). Herein, results from the western blot analysis indicated that casticin increased the protein levels of proteins p-p53 and p-ATM (Fig. 4B) in the B16F10 cells which was also confirmed by confocal laser system microscopy examination (Fig. 5). Notably, it has also been reported that antioxidant N-acetylcysteine (NAC) pretreatment enhances ATM and p53 phosphorylation, p53 acetylation and H2A.X phosphorylation in ovarian cancer cells (42). In view of this, the possible signaling pathways involved in the casticin-induced DNA damage and nuclear condensation in the B16F10 cells are summarized in Fig. 6. Thus, further studies should be conducted to elucidate the exact molecular mechanism of casticin-induced DNA damage and how to affect DNA damage and repair-associated signaling pathways.

Figure 6 The proposed signaling pathways involved in casticin-induced DNA damage and the effects on DNA repair proteins in the B16F10 cells.

Acknowledgments

This study was supported by grant CMU103-ASIA-01 from China Medical University (Taichung, Taiwan) and by grant 103-08 and 103-41 from Cheng Hsin General Hospital (Taipei, Taiwan). Experiments and data analysis were performed in part through the use of the Medical Research Core Facilities Center, Office of Research and Development at China Medical University (Taichung, Taiwan).

References