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Original Article International Journal of Oral Research 2012:3;e1

Yuniardini Septorini Wimardani1 Dewi Fatma Suniarti2 Hans-Joachim Freisleben1 Septelia Inawati Wanandi3 Masa-Aki Ikeda4 Graduate Study Program in Biomedical Science, Faculty of Medicine, Universitas Indonesia, Indonesia 2 Department of Oral Biology, Faculty of Dentistry, Universitas Indonesia, Indonesia 3 Department of Biochemistry and Molecular Biology, Faculty of Medicine, Universitas Indonesia, Indonesia 4 Section of Molecular Embryology, Graduate School of Medical and Dental Sciences, Tokyo Medical and Dental University, Japan 1

Cytotoxic effects of chitosan against oral cancer cell lines is molecular-weightdependent and cell-type-specific Abstract

The elucidation of the anticancer mechanisms of many anticancer agents from natural sources with minimal toxicity to normal cells are still being performed. Chitosan is a polycation polysaccharide, which is an N-deacetylated derivative of chitin. It is naturally and abundantly present in crab and shrimp shells, and has been widely used as a multipurpose biomaterial. Antitumor activity is one of many attractive biological properties of chitosan. Report of its antitumor activities on oral squamous cell carcinoma (SCC) cells is scarce despite many in vitro and in vivo reports on other cancer types. Physical characteristics of chitosan have been reported to influence its antitumor activity, the effects of which vary depending on cell types. Therefore, this study examined whether cytotoxic effects and doses of chitosan are affected by its molecular weight (MW) in oral SCC and non-cancer cell line. Cytotoxic effects of two types of chitosan with different MWs (average 50 - 190 kDa and 310 - >375 kDa) were tested on three oral SCC (HSC-3, HSC-4 and Ca9-22) cell lines and a keratinocyte cell line (HaCaT) using MTT assay. However, chitosan had opposite effects on HaCaT cells at certain concentrations. Both Submitted July 19, 2011 types of chitosan induced proliferation of HaCaT cells at concentrations that Accepted December 23, 2011 showed cytotoxic effects on HSC-3 and Ca9-22 cells (200 – 300 mg/ml). In particular, HaCaT cells treated with the high MW chitosan exhibited 2-fold stronger proliferative activity compared to untreated cells (300 mg/ml, p < 0.01). In contrast, neither type of chitosan induced proliferation of oral SCC cell lines. HSC-3 and Ca9-22 cells were more sensitive to both types of chitosan, which inhibited cell proliferation in a dose-dependent manner, while Keyword: Chitosan, Oral squamous cell carcinoma, HSC-4 cells were resistant to the both types. The low MW chitosan exerted stronger cytotoxic effects on all cancer cells than the high MW one, indicatCell type specificity, Cytotoxicity, Cell ing that the cytotoxic activity of chitosan negatively correlates with its MW. proliferation, Keratinocytes Although cytotoxic properties of chitosan against oral SCC cells are varied among the cell lines tested, chitosan exerts selective toxicity to oral SCC cells and even the opposite effects on non-cancer keratinocytes. Corresponding author: Yuniardini Septorini Wimardhani, Graduate Study Program in Biomedical Science, Faculty of Medicine, Universitas Indonesia Jl. Salemba Raya No. 6 Jakarta 10430 Indonesia, email: [email protected]

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Introduction Squamous cell carcinoma (SCC) is the most prevalent type of cancer found in > 90 % of all oral cancer cases. Oral SCC contributes to worldwide health burden since it causes high morbidity and mortality with geographical diversity in incidence rates. In India, oral SCC represents a major health problem responsible for the most prevalent cancer in male and the third most common cancer in females (Sharma et al., 2010). Although oral carcinogenesis is a complex multistep process, several long-established predisposing factors of oral SCC, including tobacco smoking, alcohol drinking and betel quid chewing have been known. However, variation of prevalence rates may also correlate with genetic background and way of life of people in a certain geographical areas (Copper et al., 1995). The overall incidence and mortality of oral SCC are still a dilemma with only 50 % survival rates, despite several advances in diagnostic and therapeutic approaches (Carvalho et al., 2004). The type of treatment and prognosis for patients with oral SCC are influenced by the stage of disease at diagnosis. Chemotherapy for treatment of oral cancer is still practiced with the application of chemotherapeutic agents such as cisplatin, 5-fluorouracil (5-FU) and docetaxel which have challenged clinicians through serious adverse reactions such as myelo-, immune-, and gastrointestinal toxicity as well as body weight loss (Andreadisa et al., 2003; Shibuya et al., 2004). Research aiming to discover anticancer agents from natural sources with minimal toxicity to normal cells is still being performed. To develop better cancer treatments with fewer side effects, the main purpose of this study is to examine the effects of natural, less toxic, anticancer agents on oral SCC cell lines. Chitosan is polycation polysaccharide, which is N-deacetylated derivative of chitin. It is naturally and abundantly present in crab and shrimp shells and has randomly distributed (1-4)-linked D-glucosamine and N-acetyl-D-glucosamine composition (Dutta et al., 2004) and the number of removed N-acetyl unit would determine its degree of deacetylation (DD). It has been widely used as a multipurpose biomaterial because it is biocompatible, biodegradable, not toxic and adsorptive (Singla and Chawla, 2001; Suh and Matthew, 2000). Formerly, chitosan has been recognized for its plasma cholesterol lowering effects. Possible mechanisms

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of its effects are reducing the level of cholesterol absorption as well as complicating the absorption of bile acids (Gallaher et al., 2000; Sugano et al., 1980). Recently, literature has shown attractive biological chitosan activities, which include immuno-enhancing effects (Zaharoff et al., 2007), antimicrobial activities (Martinez et al., 2010), facilitating wound healing (Howling et al., 2001; Cai et al., 2010) and antitumor activities (Maeda and Kimura, 2004; Takimoto et al., 2004; Qi et al., 2005a). Correlation between molecular characteristics of chitosan and its antimicrobial activity has been reported (No et al., 2002; Seyfarth et al., 2008; Zhang et al., 2010). Reports of antitumor activities of chitosan on oral SCC cells are scarce despite many in vitro and in vivo reports on other cancer types (Maeda and Kimura, 2004; Qi et al., 2005a; Qi et al., 2006; Liu et al., 2010). Although the mechanism(s) of how chitosan interacts with cancer cells is still unclear, several possible mechanisms have been suggested by previous studies: extracellularly binding of bigger-molecule chitosan to cell membrane, or endocytosis or internalization of chitosan nanoparticle (Huang et al., 2004). These interactions might be initiated by ionic interaction between positively charge chitosan molecules with negatively charge tumor cell membrane (Yang et al., 2009), which may activate signalling pathways leading to apoptosis or authophage. Alternatively, such chitosan interaction with tumor cell membrane may disrupt cellular functions needed to maintain cell viability, resulting in necrosis. Results of in vitro studies showed that antitumor effects of chitosan depending on its molecular weight (MW), particle vary size, dosage, and incubation time. In vivo anticancer activity of chitosan on tumour weight and volume reduction was clearly different depending on its administration route with no effects on the animal body weight reduction and was significantly different from other conventional anticancer drugs (Qi et al., 2006; Liu et al., 2010). Cytotoxic effects of conventional anticancer agents such as cisplatin surpassed chitosan against cancer cells (Qi et al., 2006). However, effects of chitosan on normal cells might be different from those of cisplatin (Luanpitpong et al., 2011), since chitosan, but not cisplatin, considerably induced proliferation of fibroblast and keratinocyte cell lines in vitro (Howling et al., 2001), which are the types of cells surrounding oral SCC. Proliferative activity of chitosan to fibroblasts

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and keratinocytes was positively correlated with its DD (Howling et al., 2001) but negatively correlated with its MW (Wiegand et al., 2010). However, in cancer cells, such as A549, zeta potential seemed to determine the type of cellular uptake, but not cytotoxicity, of chitosan molecule and chitosan nanoparticle. It has been shown that changes in MW had bigger effects than DD in altering the zeta potential of chitosan molecules and chitosan nanoparticle (Huang et al., 2004). Therefore, physical characteristics of chitosan play significant roles in determining its effects on a given cell, which may contribute to the variations of effects on different types of cancer cells in previous studies (Pathak et al., 2007; Liu et al., 2009). Therefore, this study examined whether cytotoxic effects of chitosan are affected by its MW, and are different between oral SCC and noncancer keratinocyte cell lines.

Materials and Methods Characterisation of chitosan Low molecular weight chitosan (LMWC) and high molecular weight chitosan (HMWC) were purchased from Sigma-Aldrich (St. Louis, MO). The average molecular weight of LMWC (Sigma Cat No. 44,886-9) was 50 - 190 kDa and that of HMWC (Sigma-Aldrich Cat No. 41,941-9) was 310 - >375 kDa. A 1 % (w/v) stock solution of each type of chitosan was made using 1 % acetic acid. Particles of each chitosan solution were measured for size distribution and zeta potential in a Beckman Coulter Particle Size Analyzer provided by Nanosociety Indonesia. Cell culture HSC-3, HSC-4, Ca9-22 and HaCaT cell lines were maintained in Dulbecco’s Modified Eagle’s Medium (DMEM, GIBCO Cat.No.11965-092, Life Technologies, California, USA) supplemented with 10% fetal bovine serum (FBS; Caisson Laboratories, Inc., Utah, USA) with 100 IU/ml penicillin; 100mg/ ml streptomycin (Caisson Laboratories Inc, Utah, USA) and 25 µg/ml fungizone (Biobasic Inc, Ontario, Canada) at 37 oC in a humidified atmosphere containing 5 % CO2. All cell lines used in this study were provided

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by the Section of Molecular Embryology, Graduate School of Medical and Dental Sciences, Tokyo Medical and Dental University. HSC-3, HSC-4 and Ca9-22 were oral squamous cell carcinoma established at the Second and First Departments of Oral and Maxillofacial Surgery, Faculty of Dentistry, Tokyo Medical and Dental University. HSC-3 and HSC-4 (assigned as JCRB0623 and JCRB0624) were established from tongue carcinoma, while Ca9-22 (assigned as JCRB0625) was derived from gingival carcinoma. Each cell line carries p53 mutation with different mutated codon (Sakai and Tsuchida, 1992). HaCaT, a cell line of dermal keratinocyte origin that was immortalized but not tumorigenic (Boukamp et al., 1988) served as controls in order to analyze toxicity of chitosan on normal cells. Cell suspension containing 5.0 x 105 cells was inoculated in a 96-well flat-bottom plate (100 µl/well). Cells were stabilized for 1 h at 37 oC before exposure to different concentrations of chitosan. They were incubated for 24 h in experimental medium containing each type of chitosan at final concentrations of 150, 200, 250, 300, 1000 and 5000 µg/ml. Cell viability assay MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazoliumbromide) assay measures the activity of mitochondrial dehydrogenases. These enzymes cleave the tetrazolium ring to form formazan, which reaction can be used as an index of cell viability. Dead cells would not have this reaction since they lack of mitochondrial dehydrogenase. MTT (Sigma-Aldrich. Co, Cat. No. M2128, Saint Louis, USA) solution was freshly prepared by dissolving 5 mg/ml in 0.9 % NaCl and 50 µl were added to each well. The plate was incubated in a CO2 incubator for 3 h and the formazan crystals formed were dissolved by adding 100 µl of isopropanol to each well in order to reveal its purple color. The plate was then placed on an orbital shaker for 1 h at room temperature. The absorbance of the resulting purple color was measured spectrophotometrically at 490 nm using a microplate reader (BioRad, Benchmark, California, USA). Each experiment was performed in triplicate. The same cell lines without exposure of both chitosan were served as controls. The percentage of cell viability was calculated using the following formula: (mean

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experimental absorbance/mean control absorbance) x 100 %. If the resulting cell viability percentage is larger than 100 %, the results will be defined as proliferation instead of cytotoxicity. The cytotoxicity of each type of chitosan was stated as a value of IC50 (referred to the concentration that presented 50 % cytotoxiciy). IC50 values were obtained by analysis of the percentage of inhibition by each type of chitosan at six different concentrations and were calculated from the cytotoxicity curves (GraphPad Software, Inc, California, USA).

slightly different DD and MW, which have been stated by the manufacturer. Particle size of LMWC was lower than that of HMWC, whereas zeta potentials of both types of chitosan were 38.81 and 37.62 mV, respectively. The results showed that the MW of chitosan diluted in 1 % acetic acid positively correlated with their particle size and the particle charge of both types of chitosan were classified as stable, although the particles of LMWC had slightly more positive zeta potential than that of HMWC, in other words slightly more stable particle charge (Figure 1).

Statistical analysis

Effect of two types of chitosan on the cytotoxicity of oral SCC and normal cell lines

All experiments were performed in triplicates of three different experiments and statistical analysis was performed by Student t-test with GraphPad Prism 5 Software. Significance was determined at p < 0.05.

Results Characteristics of chitosan A 1 % (w/v) stock solution of each type of chitosan was prepared using 1 % acetic acid, which is commonly used for solving chitosan. The manufacturer of chitosan used in this study only specified the DD and MW. Particles of each chitosan solution were measured for size distribution and zeta potential. Zeta potential is defined as overall charges of chitosan particles in medium and is an indicator of its stability in a colloidal system. Characteristics of each chitosan type are presented in Table 1. The two types of chitosan used in this study had

Three oral SCC cell lines (HSC-3, HSC-4 and Ca-922) and a keratinocyte cell line (HaCaT) were treated with six different concentrations of two types of chitosan (LMWC and HMWC) and then cytotoxicity was measured by MTT assay. Based on three independent experiments, IC50 values were determined by using non-linear regression: Dose response-Inhibition (log (inhibitor) vs normalized response (variable slope)) in GraphPad Prism 5 software. As shown in Figure 2, concentrations of the two types of chitosan causing cytotoxic activity were dose-dependent but varied among the oral SCC cells tested. Overall, LMWC exerted stronger cytotoxic effects on all cancer cells than HMWC. The data showed that HSC-3 and Ca9-22 cells were more sensitive to both types of chitosan. The IC50 of HSC-3 and Ca9-22 cells for LMWC was 461.4 + 32.96 mg/ml and 779.1 + 132.1 mg/ml, respectively, whereas that for HMWC was 1012 + 554.8 mg/ml and 1031 + 192.2 mg/ml, respectively (Figure 3). The IC50 values of LMWC for HSC-3 and Ca9-22 cells were 5-8

Table 1. Characteristics of chitosan products prepared as 1 % (w/v) in 1 % CH3COOH Compound Degree of deacetylation (DD) LMWC HMWC

75 – 85 % > 75 %

Viscosity (cps) 20 - 200 800 - 2000

Average Molecular Weight Particle size Zeta Potential (kDa) (nm) (mV) 50 - 190 310 - > 375

2434.53 4332.95

38.81 37.62

Particles of each chitosan solution were measured for size distribution and zeta potential in a Beckman Coulter Particle Size Analyzer provided by Nanosociety Indonesia.


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A

B

C

D

Figure 1. Size distribution and zeta potential of LMWC (A, B) and HMWC (C, D) particles measured in a Beckman Coulter Particle Size Analyzer provided by Nanosociety Indonesia. Each type of chitosan was prepared in a 1% concentration in 1% CH3COOH.

A

B

Figure 2. The effect of exposure of low (A) and high (B) molecular weight chitosans against several oral cancer cells compared to HaCaT cells. The cells were treated with increasing concentrations of chitosans for 24 h incubation at 37 oC, 5 % CO2. Cell viability was measured with MTT assay. The results are means + SD from three different independent experiments. Bars, SD. *p < 0.05 versus the same treatment condition on HaCaT cells (t-test).

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LMWC

HMWC

Figure 3. IC50 of each type of chitosan for each cell line is presented as mean + standard deviation (SD) of at least triplicate of three independent experiments. *p < 0.05 and **p < 0.01 vs control (HaCaT cell) by t-test. Significant differences in IC50 of LMWC and HMWC against the same cell line were compared using t-test with a p < 0.05 and b p < 0.01. fold lower than for HaCaT cells, which were statistically significant (p < 0.05) (Figure 3). In contrast to HSC-3 and Ca9-22 cells, HSC-4 cells were resistant to both, with HMWC was significantly less cytotoxic to HSC-4 cells than LMWC (p < 0.01). While the IC50 value of LMWC for HSC-4 cells was only slightly lower, that of HMWC was significantly higher than those of HaCaT cells (p < 0.01), indicating that HSC-4 cells are resistant to HMWC. Importantly, both types of chitosan at certain concentrations (200 to 250 mg/ml in LMWC, 200 to 300 mg/ml in HMWC) induced proliferation of HaCaT cells, although they inhibited at higher concentrations (>1000 mg/ml). In contrast, both of them did not induce proliferation of oral SCC cell lines at the concentrations that induced proliferation of HaCaT cells. Furthermore, HMWC showed stronger proliferation activity than LMWC in HaCaT cells. In particular, the cells treated with HMWC at 300 mg/ml showed almost 2-fold proliferation activity compare to untreated cells (p < 0.01) (Figure 2).

Discussion Our study revealed that LMWC exerted stronger cytotoxic activity than HMWC in all oral SCC cell


lines tested, which is consistent with other studies reported that cytotoxicity effects of chitosan were negatively correlated with its MW (Maeda et al., 2004; Huang et al., 2004). Furthermore, while both types of chitosan inhibited the growth of HaCaT cells at higher concentrations (Figure 2), LMWC exhibited significantly higher cytotoxic effects than HMWC (Figure 3), indicating that the cytotoxicity of chitosan on HaCaT cells is also negatively correlated with its MW. Although mechanism(s) involved in this phenomenon still remains unclear, the particle size of chitosan has been shown to influence the levels of its accumulation on tumor cells. It has been hypothesized that higher proton transfer to the glucosamine unit of LMWC than that of HMWC may generate more binding sites to cell membrane, resulting in increasing cytotoxicity effect of LMWC (Ma et al., 2009). However, further time-course experiments analyzing the optimal action time of chitosan and expression of cell death-related molecules are needed to investigate the types of cell death following chitosan treatment and to understand more detailed anticancer mechanisms of chitosan. It has been shown that, besides MW, modifications of the chitosan structure into water-soluble forms or nanoparticles significantly affect cytotoxic properties of this biocompatible material (Maeda et al., 2004; Takimoto et al., 2004; Qi et al., 2005a; Qi et al., 2005b). In our study, concentrations of chitosan required for

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inducing cytotoxic effects in cancer cells were higher than those used in previous studies (Maeda et al., 2004; Takimoto et al., 2004), which might be explained by the fact that the types of chitosan used here were not water-soluble and not in the form of nanoparticles. Furthermore, several reports have shown that other physical properties of chitosan, such as zeta potential and DD, also influence chitosan biological properties. An aqueous dispersion of chitosan is stable, when its zeta potential is more positive than +30 mV or more negative than -30 mV. As the particles of LMWC had slightly more positive zeta potential than those of HMWC, (38.81 vs 37.62 mV), the formers have slightly more stable particle charges. This positive value may also influence the interaction of chitosan particles with the tumor cell membrane that was usually negatively charged. In addition, it was reported that DD influences the cytotoxic activity of chitosan. Chitosan with higher DD has higher charge density, thereby facilitating its interaction with cell membrane (Huang et al., 2004). However, in this study, there are only subtle differences in DD and zeta potential between two types of chitosan used. Therefore, it is unlikely that these properties of chitosan significantly influence our results. Nevertheless, in order to eliminate any interference of acetic acid used for dissolving chitosan on culture conditions, further studies are required to examine direct effects of chitosan to cells by using water-soluble LMWC with high DD. We show that chitosan induced proliferation of HaCaT cells at certain concentrations that cause cytotoxicity to all three oral SCC cell lines tested (Figure 2). While chitosan treatments were less effective to induce cytotoxicity to HSC-4 cells, both types of chitosan caused cytotoxicity at concentrations starting from 200 µg/ml in HSC-3 and Ca9-22 cells. In contrast, both of them induced proliferation of HaCaT cells at 200 - 300 µg/ml, and, in particular, the cells treated with HMWC exhibited 2-fold stronger proliferative activity than untreated cells at 300 µg/ml. These results suggest that chitosan exerts selective toxicity and even opposite effects on cancer and non-cancer cell lines at lower concentrations (200 - 300 µg/ml), although chitosan showed toxic effects at a higher concentration (5000 µg/ml). In agreement with this, chitosan has been reported to have proliferative activity to fibroblasts and keratinocytes (Howling et al., 2001).

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Although DD of both types of chitosan used in this study was almost in the same range (Table 1), it has been also reported that proliferative activity of chitosan on cells is positively correlated with the DD (Howling et al., 2001). The exact mechanism(s) explaining proliferative effects of chitosan on keratinocytes have not yet been fully understood. It is possible that, in cell culture experiments, chitosan interacts with growth factors present in culture medium, thereby potentiating their proliferative activity to cells and also protecting the factors from degradation. By contrast, at high concentrations, chitosan might absorb metal ions and sequester them from culture medium, resulting in the reduction of available ion levels in culture medium, which leads to cytotoxicity to cells (Howling et al., 2001; Kim, 2004). The mechanism(s) behind the opposite effects of chitosan on non-cancer cells and cancer cells might be related to the fact that the highly positively charges of amino groups in chitosan molecules may attract tumor cell membrane, which is negatively charged in a greater extent than normal cells (Zhang et al., 2010). Thus, chitosan might directly attack cancer cells through interaction with tumor cell membrane or extracellularly with a particular receptor or via endocytosis to induce cytotoxicity in vitro (Huang et al., 2004). Alternatively, chitosan might cause membrane disruption by electrostatic interaction leading to an increase of inflammatory cytokines IL-6 and IL-8, which have been shown to have mitogenic effects on normal fibroblasts and keratinocytes (Wiegand et al., 2010). Cancer cells might lose the ability to respond the membrane damaging effects of chitosan, such as the release of such inflammatory cytokines. This study also shows variation of IC50 in the three cell lines treated with chitosan. Ca9-22 and HSC-3 cell lines were sensitive to both types of chitosan, whereas the higher concentrations were needed to inhibit the growth of HSC-4 cells, at the concentrations of which were also toxic to HaCaT cells (Figure 3), indicating that HSC-4 cells were resistant to chitosan. LMWC exposure at 250 µg/ml resulted in a slight increase in the number of HSC-4 cells (Figure 2). The actual mechanism(s) underlying this phenomenon needs further analysis. However, it might be explained by the assumption that HSC-4 cells still retain normal


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properties in response to chitosan, and thus the particle of LMWC at this concentration would preferably interact with molecules which facilitate proliferation in HSC-4 cells, as observed in HaCaT cells. Several studies have also shown that HSC-4 cells are less sensitive than HSC-3 and Ca9-22 towards other anticancer agents (Okamura et al., 2008; Chu et al., 2009; Hoshikawa et al., 2010). A possible explanation for different sensitivities to chitosans between HSC-4 cells and other cell lines might be due to their different genetic backgrounds affecting their response to a given stimulus. In this regard, the involvement of p53 mutation types (Sakai and Tsuchida, 1992; Kamiya et al., 2005; Ichwan, et al., 2006) or alterations of other genes related to anticancer agent sensitivity (Kaneda et al., 2006; Liu et al., 2009) might affect the sensitivity to chitosan in these cell lines.

stability in vivo, chitosan has been considered as a safe biomaterial with oral toxicity over 16 g/day/kg body weight (LD50) (Hirano, 1996) and the half-life of a highly deacetylated form of chitosan has been shown to be up to 84 days (Ren et al., 2005). In vivo experiments testing antitumor activity of chitosan nanoparticles (CNP) on sarcoma-induced mice by various types of drug administrations have also found that no lethal toxicity found in the experimental animals treated by oral administration of CNP at 2.5 mg/kg (Qi et al., 2006). Nevertheless, further in vivo experiments are needed to determine tumor-specific effects and optimum doses of chitosan, which might provide deeper insights to evaluate long-term effects of chitosan on cancer cells in vivo.

A number of studies have investigated the development of composite compounds in combination of chitosan with other anticancer agents to minimize their nonbiocompatible properties (Liu et al., 2010, Obara et al., 2005). This study only analyzed cytotoxic effects of chitosan alone to oral SCC cell lines, since we investigated the possibility of its clinical applications as a sole anticancer material. However, concentrations of chitosan needed to cause cytotoxic activity on oral SCC cells was approximately 10-fold higher than those cisplatin (Liu et al., 2009), indicating that cisplatin still exerts superior toxicity on oral SCC cancer cells. Consistent with these observations, it has been reported that, at low concentrations, chitosan alone showed neither significant decrease in tumor volume and tumor weight nor increase in survival rates in vivo (Liu et al., 2010). However, when chitosan was used as encapsulation material for camptothecin, a considerable decrease of the above tumour parameters was revealed. Furthermore, chitosan hydrogel in a form of microspheres or nanoparticles has been used for cancer treatment (Ta et al., 2008) and tested for the delivery of cytotoxic drugs such as paclitaxel, camptothecin and doxorubicin in vitro and in vivo. These drug delivery systems in combination with chitosan would possibly reduce the dosage and side effects of anticancer agents, and thus improve the efficacy of oral cancer treatment.

Conclusion

Tumor environment in vivo is different from that of this study performed in vitro. Regarding toxicity and


This study shows that cytotoxic activity of chitosan is negatively correlated with its MW. Although it was not effective to all oral SCC cell lines tested, chitosan exerts selective toxicity to cancer cell lines and even opposite effects on non-cancer cells. Taken into account its cytotoxic effects, there is possibility that chitosan might serve as a promising material for a safer therapeutic option in the treatment of oral cancer.

Acknowledgements This work was supported by Laboratory-Based Collaboration Research Grant of Universitas Indonesia 2010 and partly by Risbin Iptekdok Grant Number 2011. The authors are grateful to all staffs at Oral Biology Laboratory Universitas Indonesia and Fellow students at Tokyo Medical and Dental University, Japan for their kind help and support throughout this study.

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