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Oral cancer is one of the most common cancers in the world. Approximately 90% of oral cancers are subtyped to oral squamous cell carcinoma (OSCC). Despite advances in diagnostic techniques and improvement in treatment modalities, the prognosis remains poor. Therefore, an effective chemotherapy mechanism that enhances tumor sensitivity to chemotherapeutics is urgently needed. Methyl protodioscin (MP) is a furostanol bisglycoside with a wide range of beneficial effects, including anti-inflammatory and anti-cancer properties. The aim of the present study was to determine the antitumor activity of MP on OSCC and its underlying mechanisms. Our results show that treatment of OSCC cells with MP potently inhibited cell viability. Moreover, MP leading to cell cycle arrest at G2/M phase, which subsequently activates caspase-3, -8, -9 and PARP to induce cell apoptosis. Meanwhile, we also demonstrate that MP induces a robust autophagy in OSCC cells. The results indicate cathepsin S (CTSS) is involved in MP-induced apoptosis and autophagy by modulation of p38 MAPK and JNK1/2 pathways. These findings may provide rationale to combine MP with CTSS blockade for the effective treatment of OSCC.
Oral cancer, the most common head and neck cancer, is any cancerous tissue growth located in the oral cavity leading to more than 145,400 human deaths worldwide every year. Oral squamous cell carcinoma (OSCC) accounts for the vast majority of malignancies in the oral cavity1,2. Conventional treatment of OSCC includes surgery, radiotherapy, and chemotherapy3. Although the clinical outcome of OSCC patients has gradually improved in the last decades, the prognosis of patients with advanced-stage disease is still poor, reflecting limited advances in our understanding of pathogenesis of this disorder4. This unmet need highlights the necessity to develop novel therapeutic modalities for patients with advanced and resectable OSCC. Natural phytochemicals have received a great attention in drug discovery, which are becoming an emerging field for chemoprevention and chemotherapy in various diseases, including OSCC5,6,7. Methyl protodioscin (MP) is a furostanol bisglycoside isolated from the Dioscorea collettii var. hypoglauca (Dioscoreaceae), which is a traditional herbal medicine with anti-inflammatory and anti-tumor properties8,9. Previous studies have reported that MP induces G2/M cell cycle arrest and apoptosis leading to strong cytotoxicity across diverse cancer types9,10,11,12. However, cytotoxic effect of MP and its mechanism of action in OSCC cells are still unknown. In addition, accumulating studies suggest that protease activity is implicated in driving cancer progression via modulating both autophagy and apoptosis13,14,15,16. Two major routes of programmed cell death are closely associated with tumor resistance to anticancer drugs. Thus we tested whether proteases is involved in MP-induced cytotoxicity in OSCC cells. Here, we report that Cathepsin S (CTSS), a member of the cysteine cathepsin protease family, is involved in MP-induced cell death. CTSS is a lysosomal enzyme which is overexpressed in various cancer types and can promote lysosomal degradation of a variety of damaged or unwanted proteins13,17,18,19. There is increasing evidence to suggest CTSS plays a critical role in the regulation of autophagy and apoptosis16,20,21. In this study, we aim to determine the mechanisms by which MP regulates CTSS levels and subsequently leads to the apoptosis and autophagy in OSCC cells. Our studies clearly demonstrate that the combined use of MP with CTSS inhibitors results in significant synergy in increasing OSCC cell death, which might be a therapeutic approach to improve the prognosis of OSCC patients.
Methyl protodioscin (purity >98%) was purchased from Santa Cruz Biotechnology (Santa Cruz, CA). The compound was dissolved in dimethyl sulfoxide (DMSO) and stored at −20°C. Diluted in cell culture medium to the final concentration before use. The final concentration of DMSO for all treatments was consistently less than 0.1%. DAPI dye, propidium iodide (PI), RNase A, protease inhibitor cocktail, phosphatase inhibitor cocktail, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT), acridine orange (AO) and monodansylcadaverine (MDC) were purchased from Sigma-Aldrich (St Louis, MO). Antibody against Cdc2, Cyclin A, Cyclin B1, p21 Cip1, p27 Kip1, cleaved caspase-3, -8, and -9, cleaved poly (ADP-ribose) polymerase (PARP), LC3B, p62, Beclin1, Cathepsin S, p-AKT, AKT, p-p38, p38, p-ERK1/2, ERK1/2, p-JNK1/2, JNK1/2, and GAPDH were purchased from Cell Signaling Technology (Danvers, MA). Specific inhibitors for p38 MAPK inhibitor (SB203580) and JNK inhibitor (SP600125) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). The commercial Cathepsin S inhibitor Z-FL-COCHO was purchased from Calbiochem (San Diego).
The human oral squamous cell carcinoma (OSCC) cell lines (SAS and SCC9) were purchased from the American Type Culture Collection (ATCC) (Manassas, VA). SCC9 cells were cultured in Dulbecco’s modified Eagle’s medium-F12 supplemented with 10% fetal bovine serum (FBS), 1% NEAA, 1mM glutamine, 1% penicillin/streptomycin, 1.5g/L sodium bicarbonate, 25mM HEPES (pH 7.4), hydrocrostine (0.4mg/L), 1mM sodium pyruvate and 2mM glutamine (Sigma, St. Louis, Mo, USA). SAS cells were cultured in Dulbecco’s modified Eagle’s medium-F12 supplemented with 10% FBS, 1mM glutamine, 1% penicillin/streptomycin, 1.5g/L sodium bicarbonate, 25mM HEPES (pH 7.4) and 1mM sodium pyruvate. The cells culture was maintained at 37°C in a humidified atmosphere of 5% CO2.
MTT assay was used to evaluate the effect of MP on cell viability. Briefly, cells were seeded into 96-well plates at a density of 5000cells/well containing 100μl of culture medium. After overnight incubation to allow for attachment, the cells were incubated with indicated MP concentration. At every indicated time interval after MP treatment, 10μl of MTT (5mg/ml) was added to each well and incubated for further 4h at 37°C. The supernatant was then discarded, and 200μl of DMSO was added to each well to dissolve the formazan crystals. Optical density (OD) was evaluated by measuring the absorbance, with a test wavelength of 490nm and a reference wavelength of 630nm.
After being subjected to indicate treatment, cells were collected and fixed with 4% paraformaldehyde for 20min. Then, cells plated on the slides and stained with DAPI dye (50μg/ml) for 10min. After washing with phosphate-buffered saline (PBS), the morphological changes related to apoptosis were assessed by fluorescence microscopy (Lecia, Bensheim, Germany). Percentage of apoptotic cells was scored on at least 500 cells.
Cells were seeded in six-well dishes. After indicated treatments, cells fixed in 70% ethanol at −20°C for 16h. After washing with PBS, cells were incubated for 30min in the dark at room temperature with PI buffer (4mg/ml PI, 1% Triton X-100, 0.5mg/ml RNase A in PBS) and then filtered through a 40-mm nylon filter (Falcon, USA). The cell cycle distribution was analyzed by flow cytometry.
As previously described22. A Muse Annexin V & Dead Cell Assay Kit (Millipore) was used to quantify cell number in different stages of cell death. Briefly, 1×105 cells were resuspended in 100μl PBS (2% BSA). Add 100μl of Muse™ Annexin V & Dead Cell Reagent to each tube; the cell suspension was incubated for 20min at room temperature in the dark. Analyze by Muse Cell Analyzer flow cytometry and analysis data by the Muse® Cell Analyzer Assays (Millipore).
Cells were harvested and lysed in RIPA buffer containing protease inhibitor cocktail and phosphatase inhibitor cocktail. Protein concentration was determined by the BCA assay (Pierce). The equal amount of total protein was resolved by SDS-PAGE and transferred to PVDF membranes (Millipore, Bedford, MA). Membranes were blocked with 5% non-fat milk in TBST for 1h and then incubated at 37°C for 1h or at 4°C overnight with the indicated primary antibodies. Membranes were washed with TBST and incubated for 1h at room temperature with the appropriate secondary antibodies conjugated to horseradish peroxidase. Membranes were then washed and bound antibodies were visualized using a chemiluminescence (ECL) detection kit (Millipore).
Cells cultured in 6-well plates were either treated with indicated concentrations of MP or left untreated as control for 24h at 37°C. Thereafter, cells were stained with 1μg/ml AO or 50μM MDC in fresh medium and incubated for 30min at 37°C in the dark. After three times washing with PBS, cells were immediately visualized by a fluorescence microscope.
Cells were treated with or without MP for 24h in DMEM medium. Human protease antibody array (R&D Systems) was performed as per the manufacturer’s instruction. Briefly, the cell lysates (600μg) were mixed with array buffer and incubated with pre-blocked array membrane at 4°C for overnight. Membranes were then washed and incubated with primary antibody cocktail for 2h, followed by washing and incubated with secondary antibody for 30min. Membranes were then washed again and bound antibodies were visualized using ECL reagents and autoradiography.
The specific siRNA against CTSS and scramble control siRNA were purchased from Invitrogen. The target sequence of CTSS is 5′-CCACAACUUUGGUGAAGAA-3′. Gene silencing was performed using an RNAiMax reagent (Invitrogen) by following the manufacturer’s instructions. In brief, the cells were seeded onto 6-well culture plates and allowed to grow overnight to reach an appropriate confluence. The siRNA oligonucleotides and RNAiMAX reagent were separately diluted and then mixed. After 20min of incubation at room temperature, the cells were incubated with the transfection medium and incubated for overnight at 37°C in a humidified atmosphere of 5% CO2.
Full-length human CTSS plasmid (pCMV3-CTSS-His) was amplified by PCR using cDNA of human cells and cloned into the Kpnl and Xbal site of pCMV3-C-His vector (Sequencing primers: T7(TAATACGACTCACTATAGGG), BGH(TAGAAGGCACAGTCGAGG)). The constructed plasmid was transiently transfected into OSCC cells using FuGENE 6 (Roche Diagnostics, Basel, Switzerland). After 24h of transfection, the cells were used for the following experiments.
Values represent the means±standard deviation and the experiments were repeated at least three times. Statistical analyses were performed using the one-way analysis of variance (ANOVA) followed by Tukey’s post-hoc test was used when more than three groups were analyzed. Data comparisons were performed with Student’s t test (Sigma-Stat 2.0, Jandel Scientific, San Rafael, CA) when two groups were compared. In all cases, a p value<0.05 was considered to be statistically significant.
The chemical structure of Methyl protodioscin (MP) is shown in Fig. 1A. To examine the effect of MP on cell viability, SAS and SCC9 were treated with different concentrations of MP for 24, 48, and 72h. As shown in Fig. 1B,C, MP significantly inhibited cell viability in a dose- and time-dependent manner. These results indicate that MP can potently inhibit cell viability in different human OSCC cell lines.
To determine whether MP inhibits cell growth through the induction of apoptosis, DAPI staining was used to test cell apoptosis in OSCC cell lines. As shown in Fig. 2A, apoptotic cells with condensed chromatin were gradually increased in a dose-dependent fashion. The cell cycle distribution was also determined by PI staining and flow cytometry after exposing various doses of MP. As shown in Fig. 2B, MP induced a significant increase in accumulation of G2/M population of cells in a dose dependent manner. To further confirm that MP induces cell apoptosis, cells were stained with annexin V-FITC and PI, and subsequently analyzed by flow cytometry. The results revealed that MP induced a dose-dependent increase in the percentage of both early and late apoptotic cells (Fig. 2C). To understand the mechanisms by which MP induces G2/M phase arrest and apoptosis to inhibit cell growth, we next examined the effect of MP on the expression of key cell cycle regulators (Cdc2, Cyclin A, Cyclin B1, p21 Cip1, and p27 Kip1) and apoptosis-related proteins (cleaved caspase-3, -8, -9 and cleaved PARP) in SAS and SCC9 cells by Western blot. We observed variable alterations in the expression levels of cell cycle regulators (Fig. 2D). The expression levels of Cdc2, Cyclin A, and Cyclin B1 were significantly decreased in a concentration-dependent manner. In contrast, there was a dose-dependent increase in the expression level of negative regulators (p21 Cip1, and p27 Kip1) of cell cycle progression. Furthermore, MP also caused a dramatic dose-dependent increase in the protein level of cleaved caspase-3, -8, and 9 and cleaved PARP (Fig. 2E). These results indicate that MP is able to decrease the level of Cdc2, Cyclin A, and Cyclin B1 and to increase the level of p21Cip1 and p27Kip1 simultaneously resulting in cell cycle arrest at G2/M phase, which subsequently leads to apoptotic cell death through cleavage of caspase-3, -8, -9 and PARP.
Autophagy was proved to mediate cell death under specific circumstances, which can also be accompanied by apoptosis. Thus, we also examined whether or not MP induces autophagy in OSCC cells through biochemical and morphological analyses. As shown in Fig. 3A, we analyzed the expression levels of three autophagy-related proteins, namely LC3I/II, Beclin-1, and p62 by Western blot. The results showed that MP increased the protein levels of LC3I/II and beclin-1, while P62 was decreased in a dose-dependent manner, indicating marked induction of autophagy. To further confirm MP-induced autophagy, we examined the formation of autophagic vacuoles using the specific fluorescent dyes AO and MDC. MP treatment resulted in an increased formation of AVOs in SAS and SCC9 cells using fluorescence microscopy upon staining with the lysosomotropic agent AO (Fig. 3B). Untreated cells showed very little cytoplasmic red staining corresponding to lysosomes; however, MP-treated cells displayed many spots of red fluorescence, indicating the formation and accumulation of acidic vesicles. Consistent with AO staining, the green fluorescence intensities of MDC-labeled autophagic vacuoles were markedly increased in SAS and SCC9 cells after exposure to MP (Fig. 3C). Overall, these results clearly indicate that MP induces a robust autophagy in OSCC cells.
Accumulating evidence reveals that proteases have a potent role in modulating apoptosis and autophagy, which might be regulators of cancer progression and therapeutic response14,15,23. To assess the function of proteases, a protease array was used to screen for expressed levels of diverse proteases following stimulation of SCC9 cells with MP, compared with vehicle. As shown in Fig. 4A, the expressed level of CTSS was decreased under MP treatment. To confirm the effect of MP on CTSS expression, SAS and SCC9 cells were cultured in the presence of increasing concentrations of MP for 24h. The results showed that MP suppressed CTSS expression in a concentration-dependent manner (Fig. 4B). Furthermore, we also investigated whether overexpression or knockdown of CTSS in cells would mediate MP sensitivity. The results reveal that overexpression of CTSS could render cells more resistant to MP (Fig. 4C); while knockdown of CTSS could be sensitive to MP (Fig. 4D). In addition, the inhibition of CTSS activity by inhibitor was also significantly increase MP sensitivity (Fig. 4E). These findings suggest that CTSS is involved in MP-induced cell death.
We next investigated whether CTSS is involved in MP-induced apoptosis and autophagy in SAS and SCC9 cells. The involvement of apoptosis-related proteins (cleaved caspase-3 and cleaved PARP) and autophagy-related proteins (LC3 I/II and p62) were determined by Western blot analysis. As shown in Fig. 5A, overexpression of CTSS markedly attenuated MP-induced increase in the amount of cleaved caspase-3, cleaved PARP, and LC3 I/II but decrease in p62 expression. In contrast, knockdown of CTSS and inhibition of its enzymatic activity were significantly enhanced MP-induced apoptosis and autophagy as indicated by increased caspase-3 and PARP cleavage; LC3-II elevation and p62 degradation (Fig. 5B,C). We then evaluated the role of CTSS in MP-induced apoptosis by PI staining. As expected, overexpression of CTSS prevents MP-induced death, but CTSS depletion induces significant increase of the apoptosis cell population, as assessed by flow cytometry (Fig. 5D–F). Taken together, these results suggest that inhibition of CTSS is able to promote a synergistic cytotoxic effect with MP, leading to induction of both apoptotic and autophagic cell death.
Previously studies have reported that AKT and MAPKs signaling pathways are implicated in apoptosis and autophagy24,25,26,27. To further investigate the mechanism by which MP induces apoptosis and autophagy, we directly measured the phosphorylation of AKT and MAPKs in response to MP. The results revealed that stimulation of cells to MP induced an increase in phosphorylation of p38 MAPK and JNK in a dose- and time-dependent manner (Fig. 6A,B). To further clarify whether MP-induced apoptosis and autophagy depended on p38MAPK/JNK signaling pathways, cells were pre-treated with p38 MAPK inhibitor (SB203580) or JNK inhibitor (SP600125) for 1h followed by treatment with or without MP for 24h. The results showed that cleaved PARP and LC3-II were slightly decreased in MP co-treatment with SB203580 or SP600125 group compared with MP treatment alone (Fig. 6C,D). In addition, we further show that overexpression of CTSS blocked MP-induced activation of p38 MAPK and JNK (Fig. 6E). However, inhibition of CTSS activity resulted in significant enhancement of MP-induced activation of p38 MAPK and JNK (Fig. 6F). Collectively, these data indicated that the inhibition of CTSS led to sensitization of OSCC cells to MP cytotoxicity, which might be related to the activation of p38MAPK/JNK signaling pathways.
MP has been previously tested by NCI’s (National Cancer Institute, USA) anti-cancer drug screen on a panel of 60 human tumor cell lines. The results showed that MP had strong cytotoxic activity and might have a novel mechanism of anti-cancer action9. However, OSCC cells was not included in the NCI’s 60 cell lines, the bioactivity of MP on a human oral cancer cells requires further research. In this study, we investigated whether MP exert a strong cytotoxicity against human oral cancer cells. Here, our results indicate for the first time that MP can significantly repress the expression of CTSS leading to inducing autophagy and subsequent apoptosis in human OSCC cells. Furthermore, we found that the CTSS is involved in ROS-mediated p38MAPK/JNK signaling pathways in the regulation of autophagy and apoptosis in MP-treated cells.
Accumulating evidence suggests that MP, a furostanol saponin with four sugar units, has strong anti-tumor properties. In human chronic myelogenous leukemia cells, a study showed that MP inhibits cell proliferation via G2/M arrest and apoptosis, which are attributed to down-regulation of cyclin B1, Ca2+ homeostatic perturbation, mitochondrial dysfunction, and ROS generation12. Anticancer effects of MP with similar mechanisms of action have also been observed in human liver and lung cancer cells10,11. In addition to anti-cancer activities, MP has been recommended for the treatment of inflammation in the airway and intestine through modulation of immune responses8,28. Recently, inflammation has been considered as a key underlying mechanism for cancer development29. Notably, most anti-inflammatory compounds are potent anti-cancer agents. More importantly, several studies indicate MP is relatively safe without any side effects, suggesting that MP might serve as therapeutic approach for controlling oral cancer30,31.
In the present study, we demonstrated that inhibiting CTSS in OSCC cells can enhance MP-induced generation of ROS. Growing evidence in recent years suggests that ROS are involved in stimulation of autophagy during cancer initiation and progression32,33. Autophagy is self-digestion process that degrades intracellular long-lived proteins and damaged organelles in response to stress, which is implicated in both cell survival and death. Numerous reports have shown that extensive or inappropriate activation of autophagy can lead to apoptotic cell death34,35. Moreover, many anticancer therapies are able to increase ROS-induced autophagy leading to cell death36,37.
Autophagy is a dynamic multi-step process that involves fusion of the autophagosome with the lysosome to form the autolysosome, which have been shown to have an acidic pH and contain various proteases38. CTSS is a lysosomal protease that can promote degradation of proteins in the autolysosome39. Several studies have shown that CTSS plays a critical role in the regulation of autophagy and apoptosis16,40,41. In addition, CTSS has been shown to be highly expressed in a variety of cancers including brain42,43, lung44, gastric45,46, colorectal19,47, hepatocellular48, and prostate49,50 carcinomas, which might involve in clinical aggressiveness.
CTSS is therefore an attractive target for cancer therapy to prevent disease progression. Previous studies have demonstrated CTSS knockout mice causes reduced tumor vascularization and tumor growth18,51,52. These results indicate that the inhibition of CTSS activities may have clinical utility to abrogate tumor development. Over the last few years, inhibition of CTSS by a chemical inhibitor, RNA interference, or antibody has shown that targeting CTSS exhibits antitumor activity using in vitro and in vivo tumor models16,20,47,53,54,55,56. More recently, a study further demonstrated that a combination of the CTSS inhibitor and the EGFR tyrosine kinase inhibitor markedly promoted tumor apoptosis, indicating that CTSS is a potential novel therapeutic target for cancer treatment57.
In conclusion, our results reveal that MP can reduce CTSS expressed level to induce cell cycle arrest, autophagy, and apoptosis in human oral cell lines through mediated p38MAPK/JNK signaling pathways, suggesting that MP and CTSS are attractive candidates for tumor therapies. We believe that MP and CTSS may promise candidates for development of antitumor drugs targeting oral cancer.
How to cite this article: Hsieh, M.-J. et al. Inhibition of cathepsin S confers sensitivity to methyl protodioscin in oral cancer cells via activation of p38 MAPK/JNK signaling pathways. Sci. Rep. 7, 45039; doi: 10.1038/srep45039 (2017).
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This study was supported by grants from National Science Council, Taiwan (MOST 105-2314-B-371-003) (MOST 104-2314-B-371-008-MY2) and Changhua Christian Hospital (105-CCH-IRP-055) (105-CCH-IRP-094). The authors of the manuscript do not have a direct financial relation with the commercial identity mentioned in this paper.
The authors declare no competing financial interests.
Author Contributions M.J.H. and S.F.Y. conceived and designed the experiments; C.W.L., Y.S.L., Y.T.H. and M.K.C. performed the experiments; C.C.L. and M.J.H. analyzed the data; S.Y.C. and Y.C.C. contributed samples; J.C.C. wrote the paper.