Cell Tissue Res DOI 10.1007/s00441-014-1938-1
REGULAR ARTICLE
Chitosan as a potential osteogenic factor compared with dexamethasone in cultured macaque dental pulp stromal cells Lisa R. Amir & Dewi F. Suniarti & Sri Utami & Basril Abbas
Received: 5 September 2013 / Accepted: 3 June 2014 # Springer-Verlag Berlin Heidelberg 2014
Abstract Chitosan, a natural biopolymer derived from chitin, is considered a promising scaffold material for bone tissue engineering. The ability of chitosan to promote the osteogenic differentiation of dental pulp stromal/stem cells (DPSCs) is unknown. We have evaluated the potential of chitosan to induce the osteogenic differentiation of macaque DPSCs in comparison with that of dexamethasone. DPSCs were cultured in mineralizing medium supplemented with 5 or 10 μg/ml chitosan or with 1 or 10 nM dexamethasone. The metabolic activity of DPSCs was measured by MTT assay. Their osteogenic differentiation was determined by the number of transcripts of RUNX2, alkaline phosphatase (ALP), and COL1A1 by using real-time polymerase chain reaction, by alizarin red staining for mineral deposition, and by the ALP activity released into the medium for their ability to support biomineralizaton. Addition of chitosan to the mineralizing medium significantly increased DPSCs metabolism after 7 and 14 days of culture (P≤0.0001). Chitosan at 5 μg/ml also significantly enhanced RUNX2 and ALP mRNA but not COL1A1 mRNA; chitosan tended to increase the release of ALP hydrolytic enzyme activity into the medium during the first week. Dexamethasone upregulated the osteogenic markers tested. Mineral deposition was similar in the chitosan and dexamethasone groups and was not statistically different This study was financially supported by a Universitas Indonesia Research Excellence Grant (DRPM/RUUI Unggulan/2010/I/4150). L. R. Amir (*) : D. F. Suniarti : S. Utami Department of Oral Biology, Faculty of Dentistry, Universitas Indonesia, Salemba Raya No.4, Jakarta Pusat 10430, Indonesia e-mail:
[email protected] L. R. Amir e-mail:
[email protected] B. Abbas Center for Application of Isotope and Radiation Technology, National Atomic Energy Agency, Jakarta, Indonesia
from that of the mineralizing control group. Thus, the potential of chitosan to stimulate DPSCs proliferation and early osteogenic differentiation is comparable with that of dexamethasone, but mineralization remains unaffected by chitosan treatment. In addition to its role as a three-dimensional scaffold for osteogenic cells in vivo, chitosan might also stimulate DPSCs proliferation and early osteogenic differentiation in vitro. Keywords Chitosan . Dexamethasone . Dental pulp stromal cells . Osteogenic supplement . Macaque
Introduction Tissue engineering is currently being developed to treat large bone defects. The technique involves the seeding of progenitor cells onto a scaffold material that serves as a temporary extracellular matrix to facilitate attachment, proliferation, and differentiation of the cells to form the desired tissues (Howard et al. 2008). The ideal scaffold should be biocompatible, biodegradable at a similar rate to new tissue formation and should mimic the natural environment of the tissue in vivo (Yang et al. 2001; Sachlos and Czernuszka 2003). Chitosan is now being actively investigated as a promising natural biopolymer scaffold material (Lee et al. 2000; Seol et al. 2004; Cho et al. 2004). Chitosan is a partially deacetylated form of chitin, a polysaccharide that biomineralizes and forms the shell of crustaceans. It appears to be biocompatible and is degraded by enzymes into oligosaccharides that are rapidly resorbed. It forms an insoluble complex with connective tissue molecules such as collagen and glycosaminoglycans to form a porous interconnected three-dimensional (3D) structure, making this biomolecule attractive as a scaffolding material for tissue engineering purposes (Mi et al. 2006; Yamada et al. 2007). Chitosan is highly versatile and useful for a wide range of applications: as injectable particles in solution, as solid porous
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3D structures, or as flat 2D membranes (Lee et al. 2000; Park et al. 2000a, 2000b; Amaral et al. 2005; Jiang et al. 2006). The feasibility of utilizing this biopolymer alone or in combination with other biomaterials as a scaffold for the purpose of bone/cartilage tissue engineering has been demonstrated (Hoemann et al. 2005, 2007; Jiang et al. 2006). Integrating chitosan with other biomaterials is important in order to improve its biological and mechanical properties suitable for clinical application. Chitosan has been used as a carrier for growth factor delivery (Lee et al. 2002; Park et al. 2000a, 2000b). The sequential release of bone morphogenetic protein-2 (BMP-2) followed by the sustained delivery of insulin-like growth factor-1 in a chitosan gel/gelatin microsphere increases early osteogenic differentiation (Park et al. 2000b). Other groups have reported the incorporation of chitosan with calcium-phosphate-based ceramic materials in the development of composite scaffolds. Mineralized tissue formation has been observed in chitosan/tricalcium phosphate sponge seeded with fetal rat calvarial osteoblasts (Lee et al. 2000). The application of chitosan powder onto bone fractures in an animal model increases the regeneration process and stimulates osteogenesis in vivo (Khanal et al. 2000). Similar results have been observed in studies in vitro (Seol et al. 2004; Amaral et al. 2006; Jiang et al. 2006; Wang et al. 2008). The differentiation of an osteoblast cell line is identical or even increased when cells are cultured on chitosan-coated disks, chitosan sponges, or chitosan membrane (Seol et al. 2004; Amaral et al. 2006). The numerous reports of the chemical and biological properties of chitosan demonstrate the increasing interest of this biopolymer for its potential use in bone tissue regeneration. However, whether chitosan can be used as an osteogenic factor to stimulate the differentiation of dental pulp stromal/ stem cells (DPSCs) into an osteoblastic lineage is not currently clear. DPSCs consist of a stroma-cell-like population that displays multi-differentiation potential including that of bone-forming cells (Gronthos et al. 2000). DPSCs have great clinical potential as a source of mesenchymal stromal cells (MSCs) that can be relatively easily harvested. MSCs can be stimulated into the osteogenic lineage by using glucocorticoid, BMP, and other growth factors (Cheng et al. 1994; Lee et al. 2000; Gharibi and Hughes 2012). Typical osteogenic differentiation medium contains dexamethasone for transcriptional activation, ascorbic acid for collagen synthesis, and βglycerophosphate supplementation for mineralization (Geesin et al. 1991; Igarashi et al. 2004). Dexamethasone is a synthetic glucocorticoid that is frequently used to induce the differentiation of osteoblasts (GuzmánMorales et al. 2009). Whether chitosan can be used as an alternative for dexamethasone in osteogenic differentiation media for DPSCs is unknown. The aim of the study has been to compare the effect of chitosan with dexamethasone with respect to their ability to induce osteogenic differentiation in DPSCs.
Materials and methods Shrimp chitosan was obtained from the Center for Application of Isotope and Radiation Technology, Indonesia National Atomic Energy Agency. Briefly, chitosan was prepared by the alkaline deacetylation of chitin with 50 % sodium hydroxide at 90ºC for 8 h and precipitated with 30 % hydrochloric acid. The degree of deacetylation was approximately 80 % as measured by Fourier transform infrared spectroscopy. Chitosan was sterilized by 25 kGy of gamma irradiation. The procedure reduced the molecular weight of chitosan from an initial 75 kDa to 34.8 kDa following gamma irradiation as determined by measuring its intrinsic viscosity (Kasaai et al. 2000). Chitosan particles (10 mg) were dissolved in 10 ml 1 % v/v acetic acid (0.1 M) on a rotary mixer for 8 h and were kept as a stock solution (1 mg/ml). Final chitosan concentrations of 5 and 10 μg/ml were diluted in cell culture medium. Our pilot study indicated that 5 μg/ml was the minimum chitosan concentration that induced metabolic activity in DPSCs. Dexamethasone (Sigma, USA; 1 mg) was dissolved in 1 ml ethanol (2.5 mM) and further diluted 25× in sterile medium to give a stock solution (100 μM). The final concentrations of dexamethasone used in this study were 1 nM and 10 nM: the common doses of dexamethasone to induce in vitro osteogenic differentiation (Guzmán-Morales et al. 2009; Chadipiralla et al. 2010). The final ethanol concentration did not exceed 0.05 % v/v ethanol (Castañeda and Kinne 2000). The β-glycerol phosphate, ascorbate-2-phosphate, and alizarin red were purchased from Sigma-Aldrich. Dulbecco’s modified Eagle medium, fetal bovine serum, penicillinstreptomycin, fungizone, trizol, and collagenase I were purchased from Gibco (USA). We obtained real-time polymerase chain reaction (PCR) primers (Table 1) from Invitrogen, USA, dispase from Roche (Japan), CD70, CD90, and CD45 from BD Biosciences (USA), an alkaline phosphatase (ALP) kit from Bioassay systems (USA), a cDNA synthesis kit from ThermoFisher Scientific (Lithuania), and Sybr Green from Applied Biosystems (UK). DPSC isolation and identification DPSCs were collected from three Macaque nemestrima upper and lower first incisors (6 teeth) at the Primate Research Center, Bogor Agricultural University, Indonesia. This study was a survival study, whereby all animals remained in the facility following tooth extraction. The procedure received ethical clearance from the Animal Care and Use Committee (ACUC No.11-B005-IR). Dental pulp tissues were removed from the root canal by an extirpation needle and finely chopped with a sterile scalpel followed by enzymatic degradation with 3 mg/ml collagenase I and 4 mg/ml dispase at 37 °C under 5 % CO2 in an incubator for 1 h. The tissue was dissociated by serial pipetting every 15–20 min. Cells were
Cell Tissue Res Table 1 Primers used for realtime polymerase chain reaction
Gene
Primer
Sequence (5′ to 3′)
ALP (alkaline phosphatase)
Forward Reverse Forward Reverse Forward
gCTTCAAACCgAgATACAAgCA gCTCgAAgAgACCCAATAggTAgT ATgCTTCATTCgCCTCAC ACTgCTTgCAgCCTTAAAT TCCAACgAgATCgAgATCC
Reverse Forward Reverse
AAgCCgAATTCCTggTCT ATggggAAggTgAAggTCg TAAAAgCAgCCCTggTgACC
RUNX2 (Runt-related transcription factor-2) COL1A1 (collagen type 1A1) GAPDH (D-glyceraldehyde-3-phosphate dehydrogenase)
subsequently cultured at 37 °C, under 5 % CO2 in air, in an incubator until they reached about 90 % confluency. Subsequently, 107 cells were prepared for cell sorting by fluorescence-activated cell sorting (FACS) in an FACSAria (BD Biosciences, USA) by using CD71, CD90, and CD45 antibodies (BD Biosciences). CD71- and CD90-positive cells and CD45-negative cells indicated the presence of DPSCs. Passages 4–6 were used for the experiments. Cell culture DPSCs (1 × 104) were cultured in 24-well plates (Nunc, Roskilde, Denmark) in mineralizing and experimental media as described above. Mineralizing medium (MM) consisted of DMEM supplemented with 10 % FBS, penicillin 100 IU/ml, 100 μg/ml streptomycin, 1.25 μg/ml fungizone, 100 μg ascorbic acid, and 10 mM β-glycerophosphate, and cells grown in this medium served as the control group. The experimental groups consisted of cells grown in MM with the addition of chitosan (5, 10 μg/ml) or dexamethasone (1, 10 nM). DPSCs were incubated in MM, in chitosan-containing medium, or in dexamethasone-containing medium for 7 and 14 days, and the media were refreshed twice a week. Cell culture media were then collected for protein analysis, whereas the attached cells were kept in TRIzol for 24 h in a freezer at −80 °C for RNA isolation. MTT assay Cell metabolism was analyzed by using the MTT (3–[4,5dimethylthiazol-2yl]−2,5-diphenyl-2H-tetrazolium bromide) assay and was used as an indirect measure of the cell proliferation of the cells. DPSCs (1 × 103) were plated in 96well plates and cultured in the control or experimental medium as described above for 7 and 14 days. MTT assay was performed by adding 15 μl MTT solution (5 mg/ml; Sigma, USA) to each well, followed by incubation for 3 h at 37 °C under 5 % CO2. Next, isopropopanol was added, followed by incubation for 1 h on a shaker at room temperature to dissolve the formed formazan crystals. The optical densities (OD) of the samples were determined by using a microplate reader (Benchmark,
Biorad) at 655 nm. Data were corrected for blank values without cells. Samples were examined in quadruplicate, and the experiments were repeated twice. ALP activity assay ALP activity in DPSC culture media was detected by measuring the release of p-nitrophenol (pNP) from p-nitrophenyl phosphate (pNPP; QuantiChrom Alkaline Phosphatase Assay Kit; Bioassay Systems, USA). Total protein in the collected medium was measured by the Bradford assay (BioRad protein assay kit; Bio-Rad, USA), and the absorbance value was calculated at 595 nm by using a microplate reader (Benchmark, Bio-Rad). Total protein was standardized at 40 μg/ml and transferred to 96-well plates. The reaction mix consisted of assay buffer (pH 10.5), 5 mM magnesium acetate, and 10 mM pNPP liquid substrate was added to each of the sample wells. Spectrophotometric quantification of pNPP was performed in a microplate reader at 405 nm. ALP activity of the samples is presented in International units per liter (IU/l). Alizarin red staining At day 14, mineral deposition was examined by alizarin red staining. Cells in 24-well plates were washed with phosphatebuffered saline and fixed with cold ethanol for 30 min. The fixed cells were incubated in 2 % alizarin red in Milli-Q water (pH 4.2) for 20 min at room temperature and rinsed twice to remove non-specific staining. Images were captured by using a digital camera (Canon, Japan). Red staining representing mineralized nodules was quantified by using ImageJ 1.45r (National Institute of Health, Bethesda, Md., USA). Samples were measured quadruplicate, and the experiments were repeated twice. RNA isolation and real-time PCR Total RNA from cultured cells was isolated by using TRIzol reagent (Gibco, USA); 5 μg glycogen (Invitrogen, USA) was added to isopropanol to increase RNA yield. The RNA content was determined by measuring the absorbance in water at
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260 nm with a spectrophotometer (Ultrospec 4300 pro, Amersham Pharmacia Biotech, UK). cDNA synthesis was performed by PCR (iCycler Biorad) with 2 μg total RNA in a 20-μl final reaction volume consisting of 5 μM random hexamer primer, 5 μM Oligo (dT)18 primer, 5× reaction buffer, 20 U/μl RiboLock RNase Inhibitor, 1 nM dNTP Mix, 200 U/μl RevertAid M-MuLV reverse transcriptase (RT). The reverse transcription step was performed by incubating the RNA at 70ºC for 5 min, followed by addition of MMuLV RT mix and incubation at 37ºC for 1 h and then at 70ºC for another 10 min to deactivate the enzyme. Real-time PCR was performed on a StepOne Real-Time PCR System (Applied Biosystems, USA). The gene for the housekeeping enzyme glyceraldehyde-3-phosphate dehydrogenase (GAPDH) served as the endogenous reference to normalize RUNX2, ALP, and COL1A1 expression (Table 1). For amplification of the PCR product, 10 ng cDNA was added to the PCR Master Mix consisting of SYBR Green I Dye and 300 nM of each primer in a final volume of 15 μl. The cycle conditions were as follows: denaturation at 95 °C for 15 s followed by an annealing and extension step at 60ºC for 1 min for 45 cycles. Relative expression was calculated by using the comparative CT method. Relative expressions of the target genes were expressed as 2–(ΔΔCT) as described by Livak and Schmittgen (2001). RT-PCR products were analyzed by electophoresis in 2 % agarose gels and visualized by gel documentation (Quantity One Universal Hood; Bio-Rad, USA). Statistical analysis Data were calculated by using GraphPad Prism 6 for Mac OS X and are presented as means and standard deviation. The mean values were analyzed for their normality by using a Shapiro-Wilk normality test. All data passed the normality test and were tested for significant differences by using a one-way analysis of variance, except for the data concerning the relative expression of RUNX2 mRNA at day 7 for which we used the non-parametric Kruskal-Wallis test. Significance was accepted at P≤0.05.
Results Effect of chitosan and dexamethasone on DPSCs metabolism First, we analyzed chitosan metabolism as an indirect measure of DPSCs proliferation. The metabolism of DPSCs was markedly increased in the chitosan group compared with the control group (P≤0.05; Fig. 1a, b). Both chitosan concentrations enhanced MTT values to the same degree for 2 weeks of culture. Dexamethasone gave the same values as both
Fig. 1 Metabolism of dental pulp stromal cells (DPSCs) after incubation with chitosan (Dex dexamethasone, MM mineralizing medium). a Cell metabolism at day 7. b Cell metabolism at day 14. Note the higher cell metabolism in the chitosan group at days 7 and 14. Means±SD; *P≤0.05, **P≤0.01, ****P≤0.0001
chitosan concentrations at 1 n,M but significantly lower values at 10 nM in comparison with the 5 μg/ml chitosan group (Fig. 1a).
Cell Tissue Res Fig. 2 Alkaline phosphatase (ALP) activity of DPSCs after incubation with chitosan (Dex dexamethasone, MM mineralizing medium). a ALP activity at day 7. b ALP activity at day 14. c ALP mRNA relative expression at day 7. d ALP mRNA relative expression at day 14. Note the higher ALP mRNA expression in medium containing 5 μg/ml chitosan at day 7 (P≤ 0.05) and a tendency for higher ALP mRNA activity in both the chitosan and dexamethasone groups. Means±SD
Effect of chitosan and dexamethasone on osteogenic differentiation of DPSCs Osteogenic differentiation of DPSCs was analyzed by measuring the normalized expression of RUNX2, ALP, and COL1A1 mRNA and the activity of ALP in the culture medium (Figs. 2, 3). The activity of ALP in the medium had a tendency to increase in both the chitosan and dexamethasone groups compared with the mineralizing control group, although this was not statistically significant. ALP activity was found to be highest in the 5 μg/ml chitosan group, with a six-fold increase, and the least in the dexamethasone group with a three-fold increase compared with the mineralizing control group (P>0.05). The expression patterns of RUNX2, ALP, and COL1A1 mRNA are presented in Figs. 2c, d, 3, 4. ALP mRNA expression and that of the osteoblast transcription factor for RUNX2 mRNA were significantly upregulated in the 5 μg/ml chitosan
group (P≤0.05, P≤0.001, Figs. 2c, 3a). Similarly, 10 nM dexamethasone induced RUNX2 mRNA expression after 1 week of culture, and much higher expression was found after 2 weeks. The lower concentration of chitosan used in this study seemed to induce higher RUNX2 mRNA expression. Dexamethasone at 10 nM upregulated COL1A1 mRNA expression four-fold (P≤0.01, Fig. 3c, d). A two- and three-fold increase in COL1A1 mRNA expression was found in the chitosan group after 1 week of culture (P>0.05). Treatment with chitosan particles and dexamethasone combined neither increased ALP activity nor upregulated COL1A1 mRNA expression, and even downregulated RUNX2 mRNA expression in comparison with treatment with chitosan or dexamethasone alone (data not shown). A small number of mineralization nodules was detected in the 10 μg/ml chitosan group and in the dexamethasone group. Quantification of the staining intensity revealed comparable values in all groups (P>0.05, Fig. 5a, b).
Cell Tissue Res Fig. 3 Osteogenic markers expressed by DPSCs (Dex dexamethasone, MM mineralizing medium). a Runtrelated transcription factor-2 (RUNX2) mRNA relative expression at day 7. b RUNX2 mRNA relative expression at day 14. c Collagen type 1A1 (COL1A1) mRNA relative expression at day 7. d COL1A1 mRNA relative expression at day 14. Means±SD; *P≤0.05, **P≤0.01, ***P≤0.001
Discussion In the past decade, DPSCs have been studied as an alternative source of MSCs given the ease of harvesting the tissue. Despite numerous reports of the potential of chitosan as a
Fig. 4 Gel electrophoresis of products from the reverse transcription plus polymerase chain reaction (Chit chitosan, Dex dexamethasone, MM mineralizing medium, NTC no template control, GAPDH D-glyceraldehyde-3-phosphate dehydrogenase)
scaffold material for bone tissue engineering, the influences of chitosan on the metabolism and differentiation of DPSCs into the osteoblastic lineage in vitro remain unclear. We have tested the hypothesis that, in addition to acting as a scaffold material, chitosan can also serve as an osteogenic factor in vitro. The proliferation of macaque DPSCs has been shown to be enhanced by treatment with chitosan, in agreement with data presented by others using different cells and testing different forms of chitosan (Cai et al. 2008; Pang et al. 2005; Wang et al. 2008). Chitosan-induced proliferative responses in human periodontal ligament fibroblasts (hPDLF) reach a plateau at 100 μg/ml chitosan (Pang et al. 2005). Chitosan-coated titanium surfaces stimulate neonatal rat calvaria osteoblasts to proliferate compared with uncoated titanium surfaces (Cai et al. 2008). However, no significant changes in DNA content have been observed in bone-marrow-derived mesenchymal stromal cells (BMSCs) exposed to chitosan (GuzmánMorales et al. 2009). Differences in the properties and
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Fig. 5 Alizarin red staining on DPSCs cultured with mineralized medium (MM), supplemented with dexamethasone (Dex) or chitosan. a Alizarin red staining of culture plates. b Quantification of staining intensity
characteristics of chitosan or the sensitivity of BMSCs for chitosan might explain these different results. In support of this, one of the earliest reports of DPSCs showed more colony-forming cells at various plating densities and more cells that could be labeled with bromodeoxyuridine than were found for BMSCs, indicating that, in a population of DPSCs, more clonogenic mesenchymal stem cells exist than in a population of BMSCs (Gronthos et al. 2000). Our data suggesting that dexamethasone reduces the proliferation of DPSCs are in agreement with the report that dexamethasone induces cell-cycle arrest leading to the osteogenic differentiation of human BMSCs in vitro (Guzmán-Morales et al. 2009). The osteogenic differentiation of MSCs can be achieved by incubating MSCs in standard culture medium supplemented with dexamethasone and 1α,25-dihydroxyvitamin D3, in combination with β-glycerophosphate and ascorbic acid (Cheng et al. 1994; Lee et al. 2000; Gharibi and Hughes 2012). The differentiation of MSCs toward the osteoblastic lineage is initiated with the expression of osteoblast-specific RUNX2, the main osteoblast transcription factor, followed by the expression of ALP, COL1A1, and other osteoblast-specific differentiation genes that play roles in bone formation (Karsenty and Wagner 2002). Our data show that the lowest used concentration of chitosan stimulates RUNX2 transcription significantly in the first week and in the second week of culture, reaching far higher levels in the second week than in the first week. The stimulation of DPSCs metabolism and the upregulation of RUNX2 mRNA have been observed, followed by a transient increase in ALP mRNA expression in the cells
and a tendency for higher ALP activity in the culture medium. Notably, despite the stimulation of RUNX2 and ALP mRNA expression in DPSCs cultured with chitosan, COL1A1 mRNA expression and mineral deposition in DPSCs cultured with chitosan remain comparable with those of the dexamethasone group and the mineralizing control group. Collectively, these data suggest that chitosan is able to induce the proliferation and early osteogenic differentiation of DPSCs in a similar way to dexamethasone, but that the stimulation of COL1A1 mRNA expression by dexamethasone is not detectable in the chitosan group. A longer incubation period might be necessary in order to observe any stimulating effects of chitosan on extracellular matrix maturation and on the mineralization process. The data seem to contradict previous reports that demonstrate enhanced mineralization following treatment with chitosan (Ambre et al. 2013; Lim et al. 2009; Wang et al. 2011). However, these studies have shown that human bone-marrowderived mesenchymal stromal cells (hBMSCs) secrete a higher calcium content in the mineral deposit when hBMSCs are cultured with a chitosan-hydroxyapatite composite scaffold. The osteoconductive property of hydroxyapatite has been widely studied. Thus, the positive stimulation on mineralization might be related to the incorporation of chitosan with this osteoconductive material (Ambre et al. 2013; Lim et al. 2009; Wang et al. 2011). Our results are partially consistent with the study reported by GuzmànMorales and colleagues (2009) concerning the osteogenic effect of chitosan in hBMSCs culture compared with that of dexamethasone. The authors have shown that the addition of chitosan particles in the mineralizing medium has no effect on collagen deposition and interferes with the mineralized matrix deposition (Guzmán-Morales et al. 2009). Dexamethasone induces osteoblast differentiation through the activation of the osteoblast transcription factor (Cheng et al. 1994). The positive effects of dexamethasone on MSCs differentiation are believed to be mediated by FHL2 (four-and-a-half LIM domains protein 2), a member of the LIM (Lin11, Isl-1, Mec-3) protein superfamily. Short-RNAmediated FHL2 silencing reduces the expression of osteoblast marker genes and diminishes dexamethasone-induced upregulation of Runx2 and Col1a1, two major phenotypic osteoblast genes (Hamidouche et al. 2008). Another report has demonstrated that dexamethasone induces osteoblast differentiation through the reduction of inflammatory factors such as interleukin-1α and the increase of cell attachment (GuzmánMorales et al. 2009). Whether a similar mechanism occurs in the osteogenic differentiation of DPSCs stimulated by chitosan remains unknown. Further studies are needed in order to verify the underlying mechanism of the osteogenic differentiation of MSCs by chitosan. In conclusion, under the present in vitro conditions, this study shows the potential stimulating ability of chitosan in DPSCs proliferation and early osteogenic differentiation
Cell Tissue Res
comparable with that of dexamethasone, but no significant stimulation on mineral deposition. In addition to its role as a 3D scaffold for osteogenic cells in vivo, chitosan might also stimulate DPSCs proliferation and early osteogenic differentiation in vitro. Acknowledgments The authors thank Dr. Antonius Bronckers (Academic Center for Dentistry Amsterdam, The Netherlands) for critically reading the manuscript and facilitating the FACS analysis.
References Amaral IF, Lamghari M, Sousa SR, Sampaio P, Barbosa MA (2005) Rat bone marrow stromal cells osteogenic differentiation and fibronectin adsorption on chitosan membranes: the effect of the degree of acetylation. J Biomed Mater Res Part A 75A:387–397 Amaral IF, Sampaio P, Barbosa MA (2006) Three-dimensional culture of human osteoblastic cells in chitosan sponges: the effect of the degree of acetylation. J Biomed Mater Res 76:335–346 Ambre AH, Katti DR, Katti KS (2013) Nanoclays mediate stem cell differentiation and mineralized ECM formation on biopolymer scaffolds. J Biomed Mater Res Part A 101:2644–2660 Cai K, Hu Y, Jandt KD, Wang Y (2008) Surface modification of titanium thin film with chitosan via electrostatic self-assembly technique and its influence on osteoblast growth behavior. J Mater Sci Mater Med 19:499–506 Castañeda F, Kinne RK (2000) Cytotoxicity of millimolar concentrations of ethanol on HepG2 human tumor cell line compared to normal rat hepatocytes in vitro. J Cancer Res Clin Oncol 126:503–510 Chadipiralla K, Yochim JM, Bahuleyan B, Huang CY, Garcia-Godoy F, Murray PE, Stelnicki EJ (2010) Osteogenic differentiation of stem cells derived from human periodontal ligaments and pulp of human exfoliated deciduous teeth. Cell Tissue Res 340:323–333 Cheng SL, Yang JW, RIfas L, Zhang SF, Avioli LV (1994) Differentiation of human bone marrow osteogenic stromal cells in vitro: induction of the osteoblast phenotype by dexamethasone. Endocrinology 134:277–286 Cho JH, Kim SH, Park KD, Jung MC, Yang WI, Han SW, Noh JY, Lee JW (2004) Chondrogenic differentiation of human mesenchymal stem cells using a thermosensitive poly (N-isopropylacrylamide) and water-soluble chitosan copolymer. Biomaterials 25:5743–5751 Geesin JC, Hendricks LJ, Gordon JS, Berg RA (1991) Modulation of collagen synthesis by growth factors: the role of ascorbatestimulated lipid peroxidation. Arch Biochem Biophys 289:6–11 Gharibi B, Hughes FJ (2012) Effect of medium supplements on proliferation, differentiation potential and in vitro expansion of mesenchymal stem cells. Stem Cell Trans Med 1:771–782 Gronthos S, Mankani M, Brahim J, Robey PG, Shi S (2000) Postnatal human dental pulp stem cells (DPSCs) in vitro and in vivo. Proc Natl Acad Sci U S A 97:13625–13630 Guzmán-Morales J, El-Gabalawy H, Pham MH, Tran-Khanh N, McKee MD, Wu W, Centola M, Hoemann CD (2009) Effect of chitosan particles and dexamethasone on human bone marrow stromal cell osteogenesis and angiogenic factor secretion. Bone 45:617–626 Hamidouche Z, Hay E, Vaudin P, Charbord P, Schule R, Marie PJ, Fromigue O (2008) FHL2 mediates dexamethasone-induced mesenchymal cell differentiation into osteoblasts by activating Wnt/βcatenin signaling-dependent Runx2 expression. FASEB J 22:3813– 3822 Hoemann CD, Sun J, Legare A, McKee MD, Buschmann MD (2005) Tissue engineering of cartilage using an injectable and adhesive chitosan-based cell-delivery vehicle. Osteoarthritis Cartilage 13: 318–329
Hoemann CD, Sun J, McKee MD, Chevrier A, Rossomacha E, Rivard GE, Hurtig M, Buschmann MD (2007) Chitosan-glycerol phosphate/blood implants elicit hyaline cartilage repair integrated with porous subchondral bone in microdrilled rabbit defects. Osteoarthritis Cartilage 15:78–89 Howard D, Buttery LD, Shakesheff KM, Roberts SJ (2008) Tissue engineering; strategies, stem cells and scaffolds. J Anat 213:66–72 Igarashi M, Kamiya N, Hasegawa M, Kasuya T, Takahashi T, Takagi M (2004) Inductive effects of dexamethasone on the gene expression of Cbfa1, osterix and bone matrix proteins during differentiation of cultured primary rat osteoblasts. J Mol Histol 35:3–10 Jiang T, Abdel-Fattah WI, Laurencin CT (2006) In vitro evaluation of chitosan/poly (lactic acid-glycolic acid) sintered microsphere scaffolds for bone tissue engineering. Biomaterials 27:4894–4903 Karsenty G, Wagner EF (2002) Reaching a genetic and molecular understanding of skeletal development. Dev Cell 2:389–406 Kasaai MR, Arul J, Charlet G (2000) Intrinsic viscosity-molecular weight relationship for chitosan. J Polym Sci Part B Polym Phys 38:2591– 2598 Khanal DR, Choontanom P, Okamoto Y, Minami S, Rakshit SK, Chandra K (2000) Management of fracture with chitosan in dogs. Indian Vet J 77:1085–1089 Lee JY, Nam SH, Im SY, Park YJ, Lee YM, Seol YJ, Chung CP, Lee SJ (2002) Enhanced bone formation by controlled growth factor delivery from chitosan-based biomaterials. J Control Release 78:187–197 Lee YM, Park YJ, Lee SJ, Ku Y, Han SB, Choi SM, Klokkervold PR, Chung CP (2000) Tissue engineered bone formation using chitosan/ tricalcium phosphate sponges. J Periodontol 71:410–417 Lim TY, Wang W, Shi Z, Poh CK, Neoh KG (2009) Human bone marrow-derived mesenchymal stem cells and osteoblast differentiation on titanium with surface-grafted chitosan and immobilized bone morphogenetic protein-2. J Mater Sci Mater Med 20:1–10 Livak KJ, Schmittgen TD (2001) Analysis of relative gene expression data using real-time quantitative PCR and the 2(−Delta Delta C (T)) methods. Methods 25:402–408 Mi FL, Huang CT, Liang HF, Chen MC, Chiu YL, Chen CH, Sung HW (2006) Physicochemical, antimicrobial, and cytotoxic characteristics of a chitosan film cross-linked by naturally occuring cross-linking agent, aglycone geniopocidic acid. J Agric Food Chem 54:3290– 3296 Pang EK, Paik JW, Kim SK, Jung UW, Kim CS, Cho KS, Kim CK, Choi SH (2005) Effects of chitosan on human periodontal ligament fibroblasts in vitro and on bone formation in rat calvarial defects. J Periodontol 76:1526–1533 Park YJ, Lee YM, Park SN, Sheen SY, Chung CP, Lee SJ (2000a) Platelet derived growth factor releasing chitosan sponge for periodontal bone regeneration. Biomaterials 21:153–159 Park YJ, Lee YM, Lee JY, Seol YJ, Chung CP, Lee SJ (2000b) Controlled release of platelet-derived growth factor-BB from chondroitin sulfate-chitosan sponge for guided bone regeneration. J Control Release 67:385–394 Sachlos E, Czernuszka JT (2003) Making tissue engineering scaffold work. Review: application of solid freeform fabrication technology to the production of tissue engineering scaffold. Eur Cells Mater 5: 29–39 Seol YJ, Lee JY, Park YJ, Lee YM, Young-Ku RIC, Lee SJ, Han SB, Chung CP (2004) Chitosan sponges as tissue engineering scaffolds for bone formation. Biotechnol Lett 26:1037–1041 Wang G, Zheng L, Zhao H, Miao J, Sun C, Ren N, Wang J, Liu H, Tao X (2011) In vitro assessment of the differentiation potential of bone marrow-derived mesenchymal stem cells on genipin-chitosan conjugation scaffold with surface hydroxyapatite nanostructure for bone tissue engineering. Tissue Eng Part A 17:1341–1349 Wang J, Boer J de, Groot K de (2008) Proliferation and differentiation of MC3T3-E1 cells on calcium phosphate/chitosan coatings. J Dent Res 87:650–654
Cell Tissue Res Yamada S, Ganno T, Ohara N, Hayashi Y (2007) Chitosan monomer accelerates alkaline phosphatase activity on human osteoblasts cells under hypofunctional conditions. J Biomed Mater Res A 83:290–295
Yang S, Leong KF, Du Z, Chua CK (2001) The design of scaffolds for use in tissue engineering. Part I. Traditional factors. Tissue Eng 7:679– 689
