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Silymarin, a standardized extract from milk thistle fruits has been found to exhibit anti-cancer effects against various cancers. Here, we explored the anti-cancer activity of silymarin and its molecular target in human oral cancer in vitro and in vivo. Silymarin dose-dependently inhibited the proliferation of HSC-4 oral cancer cells and promoted caspase-dependent apoptosis. A human apoptosis protein array kit showed that death receptor 5 may be involved in silymarin-induced apoptosis, which was also shown through western blotting, immunocytochemistry, and reverse transcription-polymerase chain reaction. Silymarin increased cleaved caspase-8 and truncated Bid, leading to accumulation of cytochrome c. In addition, silymarin activated death receptor 5/caspase-8 to induce apoptotic cell death in two other oral cancer cell lines (YD15 and Ca9.22). Silymarin also suppressed tumor growth and volume without any hepatic or renal toxicity in vivo. Taken together, these results provide in vitro and in vivo evidence supporting the anti-cancer effect of silymarin and death receptor 5, and caspase-8 may be essential players in silymarin-mediated apoptosis in oral cancer.

Keywords Silymarinoral cancerdeath receptor 5caspase-8apoptosis


Oral cancer ranks as the sixth most common neoplasm and is a significant cause of cancer morbidity and mortality worldwide.1 Only 40%–50% of patients with oral cancer have an overall survival rate of 5 years.2 Despite significant improvements in chemotherapy for oral cancer, the mortality and 5-year survival rates have not changed much, which can be mostly attributed to local recurrences and distant metastasis.3,4 Thus, the development of more effective and less toxic new drug candidates may enhance patients’ quality of life and survival.

There is scientific evidence from nutritional epidemiology indicating an inverse association between the consumption of natural products such as fruits and vegetables and the risk of developing certain types of cancer.5 Silymarin is isolated from milk thistle (Silybum marianum) and contains a mixture of flavonolignans consisting of silydianin, silibinin, and silychristin.6 Silymarin has anti-inflammatory, antioxidant, and pro-apoptotic activities, which protect against human hepatic and pancreatic diseases.7,8 It has also shown anti-cancer effects in various types of cancers including liver, breast, cervical, skin, lung, and prostate cancers.812 Recently, several studies have reported that silymarin exerted in vitro anti-proliferative and pro-apoptotic activities in human oral cancer cell lines (FaDu and MC-3).13,14 However, there are no reports of its in vitro and in vivo effects on oral cancer. Based on the in vitro therapeutic efficacy of silymarin against oral malignancies, we for the first time evaluated the efficacy and associated molecular targets of silymarin on oral cancer in both cell culture and animal models.

Materials and methods

Cell culture and chemical treatment

HSC-4 and Ca9.22 cells were provided from Hokkaido University (Hokkaido, Japan), and YD15 cells were kindly obtained from Yonsei University (Seoul, Korea). All cultures were grown in Dulbecco’s Modified Eagle Medium/Nutrient Mixture F-12 (DMEM/F12) or Roswell Park Memorial Institute (RPMI) supplemented with 10% fetal bovine serum (FBS) and antibiotics at 37°C in 5% CO2 incubator. All experiments were prepared in cells cultured at 50%–60% confluence. Silymarin (S0292; Sigma-Aldrich, St. Louis, MO, USA) was purchased commercially from the manufacturer. The complex is composed of silybin A, silybin B, isosilybin A, isosilybin B, silychristin, silydianin taxifolin, and apigenin 7-glucoside (approximately 45% silybin by weight).15 It was dissolved in dimethyl sulfoxide (DMSO), aliquoted, and stored at −20°C. The final concentration of DMSO did not exceed 0.1%. Z-VAD and Z-IETD-FMK were purchased from R&D Systems (Minneapolis, MN, USA) and Calbiochem (San Diego, CA, USA), respectively.

Trypan blue exclusion assay

Cells were seeded in six-well plates and incubated with various doses of silymarin for 24 or 48 h. Cells were stained with 0.4% trypan blue (Gibco, Paisley, UK), and then, viable cells were counted using a hemocytometer. All experiments were performed independently three times with triplicate samples.

Cell proliferation assay (3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium assay)

The effects of silymarin on cell viability were determined with a CellTiter 96 Aqueous Non-Radioactive Cell Proliferation Assay (Promega, Madison, WI, USA) according to the manufacturer’s instructions. Briefly, cells were seeded in 96-well plates and incubated with various doses of silymarin for 24 or 48 h. Combined 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium (MTS)/phenazine methosulfate (PMS) solution was added to each well of a 96-well plate and maintained for 2–4 h in a 37°C CO2 incubator. The absorbance was measured at 490 nm using a Chameleon microplate reader (Hidex, Turku, Finland).

4′,6-diamidine-2′-phenylindole dihydrochloride staining

To detect nuclear morphological changes in apoptotic cells, cells were stained with 4′,6-diamidine-2′-phenylindole dihydrochloride (DAPI) solution (Sigma-Aldrich). Briefly, cells were fixed in 100% methanol at room temperature (RT) for 10 min, deposited on slides, and stained with DAPI solution (2 μg/mL). The morphological changes of cells were observed under fluorescence microscopy with the appropriate excitation and emission filters (Leica DMi8, Wetzlar, Germany).

Live/dead assay

The cytotoxic effect of silymarin was evaluated with a Live/Dead Viability/Cytotoxicity assay (Life Technologies, Grand Island, NY, USA). The polyanionic dye Calcein-AM is retained within live cells, producing an intense green fluorescence through intracellular esterase activity. Ethidium homodimer-1 enters cells with damaged membranes and binds to nucleic acids, producing a bright red fluorescence in dead cells. Briefly, cells were stained with 2 μM Calcein-AM and 4 μM ethidium homodimer-1 and then incubated for 30 min at RT. Cells were analyzed under fluorescence microscopy.

Western blot analysis

Whole-cell lysates were extracted with radioimmunoprecipitation assay (RIPA) lysis buffer and the protein concentration in each sample was measured using a DC Protein Assay Kit (Bio-Rad Laboratories, Madison, WI, USA). After normalization, equal amounts of protein were separated by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and then transferred to immuno-blot polyvinylidene difluoride (PVDF) membranes. The membranes were blocked with 5% skim milk in tris-buffered saline with Tween20 (TBST) at RT for 2 h and then incubated with primary antibodies and corresponding horseradish peroxidase (HRP)-conjugated secondary antibodies. Antibodies against cleaved poly(ADP-ribose) polymerase (PARP), cleaved caspase-3, cleaved caspase-8, death receptor 5 (DR5), and Bid were purchased from Cell Signaling Technology, Inc. (Charlottesville, VA, USA). Actin antibody was obtained from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA, USA). The immunoreactive bands were visualized by ImageQuant™ LAS 500 (GE Healthcare Life Sciences, Piscataway, NJ, USA).

Human apoptosis antibody array

HSC-4 cells were treated with DMSO or silymarin (80 μg/mL) for 24 h, and we then analyzed the relative expression of 35 apoptosis-related proteins using the Human Apoptosis Antibody Array Kit (R&D Systems). Briefly, nitrocellulose membranes were blocked with array buffer for 1 h at RT. Protein lysates were then diluted, added, and incubated overnight. After washing with 1× wash buffer to remove unbound proteins, membranes were exposed to a cocktail of biotinylated detection antibodies for 1 h at RT. Membranes were then washed and incubated with streptavidin–HRP for 30 min at RT. Each capture spot corresponding to the amount of apoptotic protein bound was detected with enhanced chemiluminescence (ECL) western blotting luminol reagent (Santa Cruz Biotechnology, Inc). The locations of controls and capture antibodies are listed in Supplementary Figure 1.

Reverse transcription–polymerase chain reaction

Total RNA was extracted using the easy-BLUE Total RNA Extraction Kit (iNtRON, Daejeon, Korea), and then, 1 μg of total RNA was transcribed into complementary DNA (cDNA) using TOPscript RT DryMIX (Elpis Biotech, Daejeon, Korea). cDNA was subjected to polymerase chain reaction (PCR) using HiPi PCR PreMix (Elpis Biotech). DR5 and ß-actin transcripts were amplified by PCR using the following specific primers: DR5 sense 5′-ATG AGA TCG TGA GTA TCT TGC AGC-3′ and DR5 anti-sense 5′-TGA CCC ACT TTA TCA GCA TCG TGT-3′, ß-actin sense 5′-GTG GGG CGC CCC AGG CAC CA-3′ and ß-actin anti-sense 5′-CTC CTT AAT GTC ACG CAC GAT TTC-3′. DR5 amplification was performed in 30 cycles (1 min at 95°C, 1 min at 57.8°C, and 1 min 30 s at 72°C) and ß-actin amplification was performed in 25 cycles (1 min at 95°C, 1 min at 60°C, and 1 min 30 s at 72°C). The PCR products were separated by electrophoresis on a 1.2% agarose gel and visualized with ethidium bromide.

Immunofluorescence staining

HSC-4 cells were seeded on four-well plates and treated with silymarin. After 24 h, cells were fixed and permeabilized using Cytofix/Cytoperm solution for 1 h at 4°C. Cells were blocked with 1% bovine serum albumin (BSA) in phosphate-buffered saline (PBS) at 1 h at RT and incubated overnight at 4°C with antibodies against DR5 or cytochrome c, followed by incubation with fluorescein isothiocyanate (FITC)-conjugated secondary antibody for 1 h at RT. Cells were observed using fluorescence microscopy with the appropriate filters for DAPI or FITC dyes.

Tumor xenograft models

The 4-week-old non-obese diabetic–severe combined immunodeficiency (NOD-SCID) male mice were purchased from Harlan Laboratories, Inc. (Envigo, Gannat, France). All mice were caged in a facility with a 12 h light/dark cycle and allowed the Teklad diet (2018S) and water ad libitum in accordance with protocols approved by Institutional Animal Care and Use Committee (IACUC) at CHA University (IACUC approval number 160012). HSC-4 cells were subcutaneously injected into the flanks of the mice, and then, the mice were assigned randomly to two treatment groups (n = 4 for each group). About 10 days after inoculation, tumor-bearing mice received vehicle control (distilled water (DW)) or 200 mg/kg/day of silymarin (p.o.) five times per week for 5 weeks. Tumor volume and body weight were monitored once a week. Tumor volume was measured by the formula: Volume = π/6{(D + d)/2}.3 The mice were sacrificed 5 weeks after silymarin treatment for examination of tumor weight and organ weight.

Histopathological examination of organs

Mice organs (liver and kidney) were fixed in 10% neutral buffered formalin. Tissue sections were cut at a thickness of 4 μm and stained with hematoxylin and eosin (H&E). Histopathological changes were analyzed using a microscope equipped with a DFC550 digital camera (Leica).

Statistical analysis

A two-tailed Student’s t-test was used for comparing two experiments, and one-way ANOVAs were applied for multiple comparisons to determine the significance of differences between the control and treatment groups. For in vivo studies, statistical evaluation was calculated with the Mann–Whitney U test in SPSS due to the use of non-parametric data; values of p < 0.05 were considered significant (*).


Effects of silymarin on the survival and apoptosis of HSC-4 human oral cancer cells

To assess the anti-proliferative effects of silymarin on HSC-4 cells, cells were treated with various doses (40, 60, and 80 μg/mL) of silymarin for 24 h and cell viability was measured using trypan blue exclusion and MTS assays. Silymarin suppressed the viability of HSC-4 cells in a dose-dependent manner (Figure 1(a) and (b)). A qualitative examination of the live/dead assay was next performed to investigate the cytotoxic effect of silymarin, which showed a decrease in the number of green fluorescence–stained cells (live cells; Figure 1(c), upper panel). To evaluate whether the observed growth inhibition and cell death were related to apoptosis, DAPI staining was used to quantitate nuclear condensation and fragmentation, which are typical apoptotic features. The results revealed the condensed and fragmented nuclei in the silymarin-treated group, but no changes in the DMSO-treated group (Figure 1(c), lower panel). We also detected apoptosis-associated marker proteins (cleaved caspase-3 and cleaved PARP) to confirm the apoptotic effect of silymarin in HSC-4 cells. The cleaved forms of the two proteins were clearly observed by silymarin in a dose-dependent manner indicating that silymarin activated the apoptotic pathway (Figure 1(d)). Z-VAD, a general caspase inhibitor that irreversibly binds to the catalytic site of caspase proteases, was used to demonstrate the role of caspase activation in the induction of apoptosis. Z-VAD markedly blocked apoptosis caused by silymarin in HSC-4 cells (Figure 1(e)). These results provide evidence that the anti-proliferative and cytotoxic effects of silymarin are due to caspase-dependent apoptosis in human oral cancer cells.


Figure 1. Effects of silymarin on the survival and apoptosis of HSC-4 human oral cancer cells. HSC-4 cells were treated with DMSO or the designated doses of silymarin for 24 h. The effect of silymarin on cell viability was examined using (a) a trypan blue exclusion assay or (b) an MTS assay. Data are expressed as the mean ± SD of triplicate experiments and the significance compared with the DMSO-treated group is indicated (*p < 0.05). (c) Live (green) and dead (red) cells were quantified with live/dead assay kit as mentioned in the “Materials and methods” section (magnification, 200×; upper panel). Cells were stained with DAPI solution and observed by fluorescence microscopy (magnification, 400×; lower panel). Data are expressed as the mean ± SD of triplicate experiments, and significance compared with the DMSO-treated group is indicated (*p < 0.05). (d) We performed western blotting using antibodies against cleaved PARP and cleaved caspase-3. Actin was used as a loading control. (e) HSC-4 cells were pre-treated with 7.5 μM Z-VAD (a pan-caspase inhibitor) for 1 h prior to silymarin treatment. Protein levels of cleaved PARP and cleaved caspase-3 were analyzed by western blotting.

Involvement of DR5 in silymarin-induced apoptosis in human oral cancer cells

To explore the potential molecular targets through which silymarin induces apoptosis in HSC-4 cells, we screened 35 apoptosis-related proteins using a human apoptosis antibody array. As shown in Figure 2(a), the spot density of DR5 (TRAIL-R2) protein was stronger in the silymarin-treated group than in the DMSO-treated group. To validate the apoptosis antibody array data, we performed Western blotting and immunofluorescence staining. The results were consistent with the observations from the apoptosis antibody array (Figure 2(b) and (c)). We also found that silymarin greatly increased the level of DR5 messenger RNA (mRNA; Figure 2(d)), indicating that it was transcriptionally regulated by silymarin. These results suggest that DR5 may be involved in silymarin-induced apoptosis in human oral cancer cells.


Figure 2. Involvement of DR5 in silymarin-induced apoptosis in human oral cancer cells. (a) HSC-4 cells were treated with DMSO or 80 μg/mL of silymarin for 24 h, and then, human apoptosis protein array was performed. Data are representative of two independent experiments. (b) HSC-4 cells were treated with DMSO or the designated doses of silymarin for 24 h. Whole-cell lysates were analyzed by western blotting to detect the expression level of DR5 protein. (c) DR5 protein was detected using immunofluorescence staining (magnification, 400×). Relative fluorescence intensity is shown as the mean ± SD of triplicate experiments, and IgG antibody was used as a negative control. (d) DR5 mRNA was determined by RT-PCR. Data are expressed as the mean ± SD of triplicate experiments, and significance compared with the DMSO-treated group is indicated (*p < 0.05).

Effects of silymarin on caspase-8 and Bid in human oral cancer cells

Since caspase-8 activation is critical for the extrinsic apoptosis pathway via DR5 protein, we investigated the effect of silymarin on caspase-8 activation. As shown in Figure 3(a), silymarin increased the expression level of cleaved caspase-8, which gave rise to truncated Bid. PARP cleavage was dramatically inhibited in the presence of a caspase-8 inhibitor, Z-IETD-FMK indicating that apoptosis was associated with an increase in caspase-8 expression and activation (Figure 3(b)). Silymarin also increased the expression levels of cytochrome c protein, implying that it may induce the loss of mitochondrial membrane potential (Figure 3(c)). These results suggested that caspase-8-mediated cleavage of Bid protein is required for silymarin-induced apoptosis in human oral cancer cells.


Figure 3. Effects of silymarin on caspase-8 and Bid in human oral cancer cells. (a) Protein levels of cleaved caspase-8 and Bid were evaluated by western blotting. (b) Cells were pre-treated with 15 μM zIETD-FMK (a caspase-8 inhibitor) for 1 h prior to 24 h of silymarin treatment. Protein levels of cleaved caspase-8 and Bid were evaluated using western blotting. (c) Cytochrome c protein was detected using immunofluorescence staining (magnification 400×).

Growth-inhibitory and apoptotic effects of silymarin via DR5/caspase-8 signaling in YD15 and Ca9.22 human oral cancer cell lines

We used two other human oral cancer cell lines (YD15 and Ca9.22) to show that the growth-inhibitory and apoptotic effects of silymarin were not limited to one human oral cancer cell line. The results showed that silymarin significantly suppressed the viability of both cell lines (Figure 4(a)) and led to dramatic increases in the expression levels of DR5 and cleaved caspase-8 to induce PARP cleavage (Figure 4(b)). The results of imaging analysis by DAPI staining and live/dead assay showed that silymarin increased apoptosis in YD15 and Ca9.22 cells (Figure 4(c)). These results suggest that apoptosis may represent a general mechanism for the anti-cancer effect of silymarin in human oral cancer cell lines.


Figure 4. Growth-inhibitory and apoptotic effects of silymarin via DR5/caspase-8 signaling in YD15 and Ca9.22 human oral cancer cells. YD15 and Ca9.22 cells were treated with DMSO or 80 μg/mL of silymarin for 48 and 24 h, respectively. (a) The effect of silymarin on cell viability was examined with a trypan blue exclusion assay. (b) Protein levels of DR5, cleaved caspase-8, and cleaved PARP were analyzed by western blotting. (c) Cells were stained with DAPI solution (left panel) or Calcein-AM/ethidium homodimer-1 dye using a live/dead assay kit (right panel).

Anti-tumorigenic effect of silymarin in NOD-SCID mice without toxicity

Based on these cell culture results showing strong biological activities of silymarin in oral cancer cell lines, we next investigated its effect on the growth of HSC-4 cell xenograft in NOD-SCID mice. HSC-4 cells were subcutaneously injected into NOD-SCID mice, and the mice were orally administered vehicle control (DW) or silymarin (200 mg/kg/day) for 5 weeks. Throughout the study, we observed that silymarin significantly suppressed the tumor volume of HSC-4 cell xenografts in NOD-SCID mice at day 35 (the end of the study), and it showed a 59% (p = 0.043) reduction in tumor volume in comparison to the control group (Figure 5(a) and Table 1). Silymarin also showed a strong trend toward significance for a decrease in tumor weight (p = 0.081, Figure 5(b)). To test whether silymarin could be an effective strategy to reduce in vivo tumor growth without any toxicity, we measured body and organ weights (liver and kidney). The result showed that body and organ weights did not change with silymarin treatment (Figures 5(c) and (d)). In addition, there was no difference in histopathological findings between the control and silymarin treatment groups (Figure 5(e)). Collectively, these results suggest that silymarin has an anti-tumorigenic effect without hepatic or renal toxicity.


Figure 5. Anti-tumorigenic effect of silymarin in tumor xenograft animal models. HSC-4 cells were subcutaneously injected into the flanks of the mice, which were then treated with vehicle control (DW) or 200 mg/kg/day of silymarin (p.o.) five times per week for 5 weeks. (a) Tumor volume, (b) tumor weight, and (c) body weight were measured. (d) After silymarin treatment, the liver and kidney were surgically removed and their weights were measured. Data are presented as the mean ± SD, and statistical significance calculated with Mann–Whitney test is indicated (*p < 0.05, #p = 0.081). (e) Histopathological changes in the liver and kidney were visualized by H&E staining. Representative images are shown (magnification 200×).


Table Table 1. Inhibition rate of tumor volume in silymarin-treated tumor xenograft model.


Table 1. Inhibition rate of tumor volume in silymarin-treated tumor xenograft model.

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Data from several studies have demonstrated the possibility of silymarin as a chemotherapeutic drug candidate against human cancers.12,16 For example, silymarin suppressed cell cycle progression in prostate cancer cells, which was associated with induction of cyclin-dependent kinase inhibitors such as p21 and p2717 and enhancement of apoptotic signals through regulation of the Akt or p38/JNK mitogen-activated protein kinase (MAPK) signaling pathway.18,19 In this study, we explored the in vitro and in vivo anti-cancer effect of silymarin in human oral cancer cell lines and a tumor xenograft animal model. The results showed that silymarin induced growth inhibition and caspase-dependent apoptosis in human oral cancer cells (Figures 1 and 4) and suppressed tumor growth without any toxicity in vivo (Figure 5). Su et al.13 reported that silymarin significantly decreased cell viability and induced apoptotic cell death in FaDu human pharynx squamous cancer cell lines through dephosphorylation of Akt, with an increase in the expression of PTEN. Recently, our previous study also revealed that mitochondria-mediated apoptosis was induced by increasing the expression of the pro-apoptotic Bcl-2 family member Bim in the MC-3 mucoepidermoid carcinoma cell line, which is a type of oral cancer.14 Previous studies are in line with this study. It can thus be suggested that silymarin is a valuable anti-cancer drug candidate for the treatment of oral cancer.

DR5 (TRAIL-R2) is an essential mediator for the extrinsic apoptotic pathway. In response to the extrinsic apoptotic signal, DR5 recruits Fas-associated death domain (FADD), TNFR1-associated death domain (TRADD), and pro-caspase-8 to form death-inducing signaling complex (DISC), which can finally trigger apoptosis.20 This prompted us to develop the potential compounds targeting DR5 for cancer therapy. Indeed, agonistic monoclonal antibodies targeting DR5 have been developed for cancer therapy and are currently being tested in pre-clinical and clinical trials.21 Thus, exploring a potential compound targeting DR5 is an attractive chemotherapeutic strategy. Previously, several studies reported that natural compounds such as ginsenoside compound K and bigelovin induced DR5 to activate the extrinsic apoptosis pathway in colon cancer cell lines.22,23 Our research group also reported that the methanol extract of Smilax china L. or β-phenylethyl isothiocyanate induced the DR5/caspase-8/Bid axis by regulating the extracellular signal–regulated kinase (ERK) or p38 MAPK signaling pathway, leading to caspase-mediated apoptosis in oral or cervical cancer cells.2427 These previous findings fully support DR5 regulation by natural products during apoptotic cell death pathways in cancer. In this study, our human apoptosis antibody array data showing that silymarin upregulates DR5 protein are consistent with these findings. These implied that DR5 can be a potential molecular target for the apoptotic effect of silymarin in oral cancer. Silibinin, which is a major bioactive component of silymarin, is known as a naturally occurring STAT3-targeted pharmacological inhibitor that shows the potential for use as cancer therapy.28,29 Thus, we cannot exclude the possibility that DR5 could be either directly or indirectly affected by the effects of silymarin on phospho-STAT3, a signal transducer and activation of transcription factor 3 that regulates apoptosis in human oral mucoepidermoid carcinoma.

Caspase proteins are divided into initiators (caspase-2, 8, and 9) and executioners (caspase-3, 6, and 7), which can facilitate cleavage of substrates to induce apoptosis. Cleavage of caspase-8 induced by an extrinsic apoptotic signal could result in Bid truncation as well as executioner caspases.30 Several lines of evidence suggest that caspase-8 mutations have been reported in human cancers including gastric, hepatic, and colon cancers; this mutation might lead to block the death signal transduction and contribute to the pathogenesis of various cancers.3134 In addition, oral cancer cell lines lacking caspase-8 were resistant to chemotherapeutic agents when compared to lines expressing high levels of caspase-8 protein.35 Caspase-8 activation is indispensable for increasing anti-cancer efficacy against oral cancer. In this study, we revealed that silymarin triggered caspase-8 cleavage and Bid truncation, followed by the accumulation of cytochrome c in the cytosol (Figure 3). This phenomenon commonly occurred in two other oral cancer cell lines (Figure 4). Our findings suggest that targeting DR5/caspase-8 signaling using silymarin might be a good chemotherapeutic strategy for the treatment of oral cancer.

The aim of the present research was to examine the anti-cancer efficacy of silymarin and its molecular target in human oral cancer in vitro and in vivo. The results of this investigation provide supporting evidence that silymarin exerts anti-cancer activities through the induction of DR5/caspase-8 apoptotic signaling. This research will serve as a base for the development of an attractive drug candidate against oral cancer using silymarin or its derivatives.

Declaration of conflicting interests
The author(s) declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

The author(s) disclosed receipt of the following financial support for the research, authorship, and/or publication of this article: This research was supported by the Research Resettlement Fund for the new faculty of Seoul National University (2016) and the Basic Science Research Program through the National Research Foundation of Korea (NRF), funded by the Ministry of Science, ICT, and Future Planning (2017R1A2B2003491).

Sung-Dae Cho

Research ethics
Institutional Animal Care and Use Committee (IACUC) guidelines of CHA University (IACUC-160012).


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