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Platelet-rich Fibrin Increases Proliferation and Differentiation of Human Dental Pulp Cells
Basic Research—Biology
Platelet-rich Fibrin Increases Proliferation and Differentiation of Human Dental Pulp Cells Fu-Mei Huang, DDS, MMS, PhD,*† Shun-Fa Yang, PhD,‡ Jiing-Huei Zhao, DDS,† and YuChao Chang, DDS, MMS, PhD*† Abstract Introduction: Platelet-rich fibrin (PRF) by Choukroun’s technique is derived from an autogenous preparation of concentrated platelets without any manipulation. When delicately pressed between 2 gauzes, the PRF clot becomes a strong membrane with high potential in clinical application. However, the effect of PRF on dental pulp cells (DPCs) remains to be elucidated. This study was to determine the biological effects of PRF on DPCs. Methods: PRF samples were obtained from 6 healthy volunteers. Human DPCs were derived from healthy individuals undergoing extraction for third molars. Cell proliferation resulting from PRF was evaluated by colorimetric assay. Western blot was used to evaluate the expression of osteoprotegerin (OPG). Alkaline phosphatase (ALP) activity was examined by substrate assay. Results: PRF did not interfere with cell viability of DPCs (P > .05). DPCs were observed to attach at the edges of PRF by phase-contrast microscopy. PRF was found to increase DPC proliferation during 5-day incubation period (P < .05). PRF was found to increase OPG expression in a time-dependent manner (P < .05). ALP activity was also significantly upregulated by PRF (P < .05). Conclusions: PRF was demonstrated to stimulate cell proliferation and differentiation of DPCs by up-regulating OPG and ALP expression. These findings might serve as a basis for preclinical studies that address the role of PRF in reparative dentin formation. (J Endod 2010;36:1628–1632)
Key Words Alkaline phosphatase, dental pulp, osteoprotegerin, platelet-rich fibrin, proliferation
From the )Department of Dentistry, Chung Shan Medical University Hospital, †School of Dentistry, and ‡Institute of Medicine, Chung Shan Medical University, Taichung, Taiwan. Address requests for reprints to Prof. Yu-Chao Chang, School of Dentistry, Chung Shan Medical University, 110, Sec. 1, Chien-Kuo N. Rd., Taichung, Taiwan. E-mail address: cyc@ csmu.edu.tw. 0099-2399/$0 - see front matter Crown Copyright ª 2010 Published by Elsevier Inc. on behalf of the American Association of Endodontists. All rights reserved. doi:10.1016/j.joen.2010.07.004
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T
ooth injury can induce reactionary or reparative dentin formation. During reparative dentin formation, the differentiation of dental pulp cells into odontoblasts depends on the regulation of multiple interacting signaling pathways (1, 2). Signaling molecules playing important roles in the self-repair process can also be liberated in pathologic conditions. To help promote endodontic tissue regeneration and healing, the local application of growth factors and host modulating agents is being used to maximize the body’s healing potential (3). Transforming growth factor (TGF)–b has been implicated in the differentiation of odontoblast-like cells and in pulp tissue repair in vivo (4). A novel platelet-derived growth factor (PDGF) isoform, PDGF-C, was observed and might be involved in the pathogenesis of rat experimental periapical lesions (5). Recently, platelet-rich plasma (6) has shown clinical success in enhancing endodontic regeneration for periapical inflammatory lesion, although the long-term predictability remains questionable, and its anticipated benefits are moderate. Platelet-rich fibrin (PRF) protocol, a simple and free technique developed in France by Dohan et al (7), is a second-generation platelet concentrate that allows one to obtain fibrin membranes enriched with platelets and growth factors after starting from an anticoagulant-free blood harvest (8, 9). PRF looks like a fibrin network. PRF contains platelets, growth factors, and cytokines that might enhance the healing potential of both soft and hard tissues (10, 11). Many growth factors such as PDGF and TGF-b are demonstrated to release from PRF (8, 9). Recently, studies have demonstrated that the PRF membrane has a very significant slow sustained release of many key growth factors for at least 1 week (12) and up to 28 days (13), which means that PRF could release growth factors with its own biological scaffold for wound healing process. This leads to the idea of using PRF membrane as a capping agent for reparative dentin formation or as a biomaterial for pulp regeneration. Osteoprotegerin (OPG), a naturally occurring inhibitor of osteoclast differentiation, binds to receptor activator of nuclear factor-kB ligand (RANKL) and blocks RANKL from interacting with RANK (14). The intensity of OPG immunoreactivity in odontoblastic layer has shown highly expressed in healthy and inflamed peripheral pulp specimens (15). Alkaline phosphatase (ALP) is a membrane-bound glycoprotein, which is one of the osteogenic differentiation markers considered to indicate the differentiation of cells to odontoblasts and mineralization process (16). OPG and ALP expression were generally regarded as markers of odontoblastic differentiation. Dental pulp cells (DPCs) are considered as cells primarily concerned with providing physical barriers and structural components in dental pulp. Cultured primary DPCs can be induced to differentiate into odontoblast-like cells and generate dentin-like mineral structure in culture dishes (17, 18). However, currently no research has addressed the role of PRF on human DPCs and its potential function for dentinogenesis. In this study, we studied the biological functions of PRF on DPCs through measuring cell proliferation, protein expression of OPG, and ALP activity.
Materials and Methods PRF Preparation The human blood samples in this study were obtained under the guidelines of the Ethics Committee of the Chung Shan Medical University Hospital. Blood from 6 healthy volunteers were treated according to the PRF protocol (10, 11) with a PC-02 table centrifuge and collection kits provided by Process (Nice, France). Briefly, blood
JOE — Volume 36, Number 10, October 2010
Basic Research—Biology samples were taken without an anticoagulant in 10-mL glass-coated plastic tubes (Vacutainer; Becton Dickinson, Franklin Lakes, NJ) and immediately centrifuged at 3000 rpm for 10 minutes. Three layers are naturally formed in the tube: a base of red blood cells at the bottom, acellular plasma on the surface, and finally a PRF clot in the middle between the other 2 parts. The fibrin clot was easily separated from the lower part of the centrifuged blood. The PRF clot was gently pressed into a membrane with sterile dry gauze for the following experiments.
Cell Cultures Human DPCs were cultured by using an explant technique as described previously (19, 20). Briefly, impacted third molars were obtained from healthy patients of the Department of Dentistry, Chung Shan Medical University Hospital with informed consent. Teeth were sectioned horizontally below the cementoenamel junction with a no. 330 high-speed bur with water spray. Pulp tissues were removed aseptically in lamina flow, rinsed with Hanks’ buffered saline solution, and placed in a 60-mm dish. Pulp tissues were minced with a blade into small fragments and grown in Dulbecco modified Eagle medium (DMEM) supplemented with 10% fetal calf serum (FCS) and antibiotics (100 U/mL penicillin, 100 mg/mL streptomycin, and 0.25 mg/mL of amphotericin B). Cultures were maintained at 37 C in humidified atmosphere of 5% CO2 and 95% air. Confluent cells were detached with 0.25% trypsin and 0.05% ethylenediaminetetraacetic acid (EDTA) for 5 minutes, and aliquots of separated cells were subcultured. Cell cultures between the third and eighth passages were used in this study. Cell Viability Each PRF membrane was covered with 5-mL suspension of cells at a concentration of 5 104 cells/mL in 35-mm culture dishes. After day 1, day 3, and day 5 culture periods, the medium was removed, and 0.5 mL of 0.25% trypsin in phosphate-buffered saline (PBS) was added to each culture dish to detach the cells. One milliliter of medium was added to 0.5 mL of this cell suspension. Then 0.5 mL of calcium-free and magnesium-free PBS containing 0.25% trypan blue (wt/vol) was added to 0.5 mL of the cell suspension to stain nonviable cells. Fifteen-microliter cell suspension was dropped into a hemocytometer chamber (Cambridge Instruments, Buffalo, NY), and cell numbers were counted under a phase-contrast microscope (21). Cell viability is represented as viable cells/total cells. Cell Proliferation A 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) colorimetric assay was developed to monitor mammalian cell survival and proliferation in vitro. DPCs were plated at an initial density of 2 104 cells/well in DMEM containing 10% FCS. After overnight attachment, cells were incubated in fresh medium containing PRF for 1, 3, and 5 days. In the end of each final 4 hours, 50 mL of MTT (Sigma, St Louis, MO) was added to each well. The insoluble formazan was dissolved with 150 mL of dimethyl sulfoxide to each well. Optical density (OD) was determined by eluting the dye with dimethyl sulfoxide, and the spectrophotometric absorbance was measured at 550 nm by using a spectrophotometer (Hitachi, Tokyo, Japan). The percentage of the dehydrogenase activity at each time point, compared with that of the control, was calculated from the absorbance values. Western Blot Nearly confluent monolayers of DPCs were washed with serumfree DMEM for 24 hours and immediately thereafter exposed to PRF for additional 24 hours. Cell lyates were collected as described previously (22). Briefly, cells were solubilized with sodium dodecylsulfate JOE — Volume 36, Number 10, October 2010
(SDS)–solubilization buffer (5 mmol/L EDTA, 1 mmol/L MgCl2 , 50 mmol/L Tris-HCl , pH 7.5 and 0.5% Triton X-100, 2 mmol/L phenylmethysulfonyl fluoride, and 1 mmol/L N-ethylmaleimide) for 30 minutes on ice. Then, cell lysates were centrifuged at 12,000g at 4 C, and the protein concentrations were determined with Bradford reagent by using bovine serum albumin (BSA) as standards. Equivalent amounts of total protein per sample of cell extracts were run on a 10% SDS–polyacrylamide gel electrophoresis and immediately transferred to nitrocellulose membranes. The membranes were blocked with PBS containing 3% BSA for 2 hours, rinsed, and then incubated with primary anti-OPG (Abcam, Cambridge, MA) antibodies (1:1000) in PBS containing 0.05% Tween 20 for 2 hours. After 3 washes with Tween 20 for 10 minutes, the membranes were incubated for 1 hour with biotinylated secondary antibody diluted 1:1000 in the same buffer, washed again as described above, and treated with 1:1000 streptavidin-peroxidase solution for 30 minutes. After a series of washing steps, the reactions were developed by using diaminobenzidine (Zymed, South San Francisco, CA). The intensities of the obtained bands were determined by using a densitometer (AlphaImager 2000; Cell Biosciences, Santa Clara, CA). Each densitometric value was expressed as the mean standard deviation (SD).
Measurement of ALP Activity According to our recent study (23), the cell lyates were sonicated on ice bath, centrifuged at 1500g for 5 minutes, and measured the ALP activity in supernatant by ALP assay mixtures containing 0.1 mol/L 2amino-2-methyl-1-propanol, 1 mmol/L MgCl2, and 8 mmol/L p-nitrophenyl phosphate disodium. After incubation at 37 C for 30 minutes, the reaction was stopped with 0.1 N NaOH, and the absorbance was read at 405 nm. The OD of control cells was considered to be 100%. The relative ALP activity of U2OS cells was calculated by the formula (OD of experimental sample/OD of control cells) 100%. Each value was expressed as the mean SD. Statistical Analysis Three replicates of each time point were performed for each test. All assays were repeated 3 times to ensure reproducibility. Statistical analysis was by one-way analysis of variance (ANOVA). Tests showing differences in the treatments were analyzed by the Duncan test, and a P value of <.05 was considered statistically significant.
Results The cell viability of DPCs treated with or without PRF was shown by the same trypan blue dye uptake. No significant statistical differences were seen between the PRF stimulation and untreated control during 5-day culture period (P > .05). PRF exhibited no cytotoxic effects to DPCs. DPCs on the flat surface of the culture dishes demonstrated a spindle-shaped morphology. Moreover, DPCs were found to attach at the edge of PRF under observation by phase-contrast microscopy (Fig. 1a). Fig. 1b shows the mitogenic effects of PRF on DPCs. PRF was found to increase DPC proliferation in a time-dependent manner (P < .05). From MTT proliferation assay, the absorbance values increased about 1.5-, 3.2-, and 3.7-fold on days 1, 3, and 5, respectively, as compared with untreated control (P < .05). As shown in Fig. 2a, PRF was found to up-regulate OPG protein expression in DPCs during 5-day culture period (P < .05). The quantitative measurement by the AlphaImager 2000 is shown in Fig. 2b. The levels of the OPG increased about 2.8-, 2.1-, and 1.6-fold on days 1, 3, and 5, respectively. The effects of PRF on the ALP activity were determined by substrate assay. As shown in Fig. 3, the ALP activity was found to be significantly
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Basic Research—Biology
Figure 2. (A) Kinetics of OPG expression in DPCs exposed to PRF for 1, 3, and 5 days, respectively, by Western blot analysis. b-actin was performed to monitor equal protein loading. (B) Levels of OPG protein treated with PRF were measured by AlphaImager 2000. Relative level of OPG protein expression was normalized against b-actin signal, and control was set as 1.0. OD values represent the mean SD. )Significant difference from control values with P< .05.
Figure 1. (A) DPCs exhibited their spindle-shaped morphology when cultured with PRF membranes. DPCs were found to attach at the edge of PRF membrane. (B) Effects of PRF on cell proliferation of DPCs were measured by MTT colorimetric assay. Percentage of absorbance value of DPCs with PRF compared with that of each control was calculated. Each point and bar represent the mean SD. )Significant difference from control value at P < .05.
enhanced after PRF treatment (P < .05). The relative ALP activities were 2.0-, 3.6-, and 3.0-fold after exposure to PRF as compared with untreated control, respectively (P < .05).
Discussion PRF by Choukroun’s technique is produced in a totally natural manner, without using an anticoagulant during blood harvesting or bovine thrombin or calcium chloride for platelet activation and fibrin polymerization (9). The PRF clot forms a strong natural fibrin matrix and shows a complex architecture as a healing matrix. It is an autologous biomaterial and not an improved fibrin glue. Unlike the plateletrich plasma, PRF by Choukroun’s technique does not dissolve quickly after application; instead, the strong fibrin matrix is slowly remodeled in a similar way to a natural blood clot (12, 24). In this study, PRF exhibited no cytotoxic effect to DPCs. Similar results have shown that PRF demonstrated no cytotoxicity toward many cells including preadipocytes, dermal prekeratinocytes, osteoblasts, oral epithelial cells, periodontal ligament cells, and gingival fibroblasts (25–27). In addition, each DPC maintained its original morphology. DPCs were observed to attach at the edge of PRF 1630
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membrane by phase-contrast microscopy. Consistently, osteoblasts, periodontal ligament cells, and gingival fibroblasts were demonstrated to attach at the edge of PRF membrane (25). Moreover, osteoblasts were found to form a network at the surface and the periphery of the PRF membrane and organized the apposition of a bone matrix under scanning electron microscopy analysis (26). Thus, the biocompatibility of PRF is not cell type–specific. Taken together, PRF acts as a biomaterial to DPCs. In this study, PRF was found to increase DPC proliferation as a mitogen. Similar results were reported by He et al (13), Tsai et al (25), and Dohan Ehrenfest et al (26) that PRF stimulated cell proliferation of gingival fibroblast, periodontal ligament cells, and osteoblasts in vitro. The mechanism responsible for the cell proliferation by PRF might be explained as follows. Many growth factors such as PDGF and TGF-b are released from PRF (8, 9, 12, 13). These findings suggest that PRF possibly modulates DPC proliferation by PDGFrelated and TGF-b–related mechanisms. In the present study, the expression of OPG was stimulated by PRF. Two hypotheses could account for the mechanisms of the stimulation of OPG secretion in DPCs. The first is that OPG secretion is stimulated by increasing the number of OPG-secreted cells, namely stromal/osteoblastic cells. The second is that PRF can enhance the amount of OPG secreted from each cell. Our results imply that the effect of PRF on the proliferation of DPCs and the enhancement of OPG secretion might be attributed to the odontoblastic differentiation and mineralization process. Elevated levels of OPG are found in DPCs, raising the question of which factors, especially growth factors in PRF, result in the outcome
JOE — Volume 36, Number 10, October 2010
Basic Research—Biology References
Figure 3. Effects of PRF on ALP activity in DPCs. Percentages of ALP activity in the presence of PRF relative to that in the control are shown. Results were averaged from 3 independent experiments. )Significant differences from control values with P < .05.
observed in the present study. As already noted, the contribution of PRF to the proliferation of DPCs is the most important factor in the enhancement of OPG secretion. TGF-b, one of the main growth factors detected in PRF, has been demonstrated in pulp tissues after injury and is implicated in the differentiation of odontoblast-like cells and in pulp tissue repair (28, 29). Consistently, TGF-b was found to increase OPG expression in MC3T3-E1 cells (30) and MG 63 cells (31). Increase of ALP activity, essential for biomineralization, was considered to indicate the differentiation of cells to odontoblasts (32). In the present study, ALP activity was elevated by PRF in a timedependent manner. Similar results have reported that PRF could increase ALP activity in osteoblasts in vitro (13, 26). These results indicated that PRF might contribute to the differentiation of human DPCs. Choukroun’s PRF is derived from an autogenous preparation of concentrated platelets without any manipulation and is widely used in clinical dentistry as a vector for cell growth factors. The present study demonstrates that PRF can stimulate DPC proliferation. PRF was demonstrated to enhance OPG protein expression and ALP activity. Taken together, PRF can increase DPC proliferation and differentiation, suggesting potential applications of PRF as a biological molecule to promote the regeneration of lost or injured dental pulp tissues and stimulate reparative dentinogenesis. Future studies are required to elucidate the precise mechanism of action of PRF for dental pulp regeneration both in vitro and in vivo. In vitro experiments are very helpful to assay the biological effects of PRF, but they might be limited in their ability to simulate the clinical condition. It might be unrealistic to transfer in vitro findings to in vivo situations. The other factors that modulate the expression of odontoblastic differentiation markers such as dentin sialophosphoprotein, dentin matrix protein, and osteocalcin should be evaluated in further studies. It is also necessary to continue investigating the in vivo effects of PRF on DPCs for dentinogenesis.
Acknowledgments This study was supported by a research grant (CSMU-96RD-05) from Chung Shan Medical University by Prof. Chang. The authors disclose no conflicts of interest.
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1. Ruch JV, Lesot H, Begue CK. Odontoblast differentiation. Int J Dev Biol 1995;39: 51–68. 2. Smith AJ, Tobias RS, Murray PE. Transdentinal stimulation of reactionary dentinogenesis in ferrets by dentine matrix components. J Dent 2001;29:341–6. 3. Bashutski JD, Wang HL. Periodontal and endodontic regeneration. J Endod 2009; 35:321–8. 4. Tziafas D, Papadimitriou S. Role of exogenous TGF-beta in induction of reparative dentinogenesis in vivo. Eur J Oral Sci 1998;106:192–6. 5. Wang L, Zhang R, Peng B. Expression of a novel PDGF isoform, PDGF-C, in experimental periapical lesions. J Endod 2009;35:377–81. 6. Hiremath H, Gada N, Kini Y, Kulkarni S, Yakub SS, Metgud S. Single-step apical barrier placement in immature teeth using mineral trioxide aggregate and management of periapical inflammatory lesion using platelet-rich plasma and hydroxyapatite. J Endod 2008;34:1020–4. 7. Dohan DM, Choukroun J, Diss A, et al. Platelet-rich fibrin (PRF): a second generation platelet concentrate—part I: technological concept and evolution. Oral Surg Oral Med Oral Path Oral Radiol Endod 2006;101:E37–44. 8. Dohan DM, Choukroun J, Diss A, et al. Platelet-rich fibrin (PRF): a secondgeneration platelet concentrate—part II: platelet-related biologic features. Oral Surg Oral Med Oral Pathol Oral Radiol Endod 2006;101:E45–50. 9. Dohan DM, Choukroun J, Diss A, et al. Platelet-rich fibrin (PRF): a second-generation platelet concentrate—part III: leucocyte activation: a new feature for platelet concentrates? Oral Surg Oral Med Oral Pathol Oral Radiol Endod 2006;101:E51–5. 10. Choukroun J, Diss A, Simonpieri A, et al. Platelet-rich fibrin (PRF): a secondgeneration platelet concentrate—part IV: clinical effects on tissue healing. Oral Surg Oral Med Oral Pathol Oral Radiol Endod 2006;101:E56–60. 11. Choukroun J, Diss A, Simonpieri A, et al. Platelet-rich fibrin (PRF): a secondgeneration platelet concentrate—part V: histologic evaluations of PRF effects on bone allograft maturation in sinus lift. Oral Surg Oral Med Oral Pathol Oral Radiol Endod 2006;101:E299–303. 12. Dohan Ehrenfest DM, de Peppo GM, Doglioli P, Sammartino G. Slow release of growth factors and thrombospondin-1 in Choukroun’s platelet-rich fibrin (PRF): a gold standard to achieve for all surgical platelet concentrates technologies. Growth Factors 2009;27:63–9. 13. He L, Lin Y, Hu X, Zhang Y, Wu H. A comparative study of platelet-rich fibrin (PRF) and platelet-rich plasma (PRP) on the effect of proliferation and differentiation of rat osteoblasts in vitro. Oral Surg Oral Med Oral Pathol Oral Radiol Endod 2009; 108:707–13. 14. Lacey DL, Timms E, Tan HL, et al. Osteoprotegerin ligand is a cytokine thatregulates osteoclast differentiation and activation. Cell 1998;93:165–76. 15. Kuntz KA, Brown CE Jr, Legan JJ, Kafrawy AH. An immunohistochemical study of osteoprotegerin in the human dental pulp. J Endod 2001;27:666–9. 16. Yokose S, Kadokura H, Tajima Y, et al. Establishment and characterization of a culture system for enzymatically released rat dental pulp cells. Calcif Tissue Int 2000;66:139–44. 17. Tsukamoto Y, Fukutani S, Shin-Ike T, et al. Mineralized nodule formation by cultures of human dental pulp-derived fibroblasts. Arch Oral Biol 1992;37:1045–55. 18. About I, Bottero MJ, de Denato P, Camps J, Franquin JC, Mitsiadis TA. Human dentin production in vitro. Exp Cell Res 2000;258:33–41. 19. Chang YC, Tai KW, Huang FM, Huang MF. Cytotoxic and nongenotoxic effects of phenolic compounds in human pulp cell cultures. J Endod 2000;26:440–3. 20. Huang FM, Tsai CH, Chen YJ, Chou MY, Chang YC. Examination of the signal transduction pathways leading to upregulation of tissue type plasminogen activator by interleukin-1a in human pulp cells. J Endod 2006;32:30–3. 21. Tai KW, Chang YC. Cytotoxicity evaluation of perforation repair materials on human periodontal ligament cells in vitro. J Endod 2000;26:395–7. 22. Tsai CH, Yang SF, Lee SS, Chang YC. Augmented heme oxygenase-1 expression in areca quid chewing associated-oral submucous fibrosis. Oral Dis 2009;15:281–6. 23. Huang FM, Tsai CH, Yang SF, Chang YC. Effects of root canal sealers on alkaline phosphatase in human osteoblastic cells. J Endod 2010;36:1230–3. 24. Dohan Ehrenfest DM, Rasmusson L, Albrektsson T. Classification of platelet concentrates: from pure platelet-rich plasma (P-PRP) to leucocyte- and platelet-rich fibrin (L-PRF). Trends Biotech 2009;27:158–67. 25. Tsai CH, Shen SY, Zhao JH, Chang YC. Platelet-rich fibrin modulates cell proliferation of human periodontally related cells in vitro. J Dent Sci 2009;4:130–5. 26. Dohan Ehrenfest DM, Diss A, Odin G, Doglioli P, Hippolyte MP, Charrier JB. In vitro effects of Choukroun’s PRF (platelet-rich fibrin) on human gingival fibroblasts, dermal prekeratinocytes, preadipocytes, and maxillofacial osteoblasts in primary cultures. Oral Surg Oral Med Oral Pathol Oral Radiol Endod 2009;108:341–52. 27. Chang IC, Tsai CH, Chang YC. Platelet-rich fibrin modulates the expression of extracellular signal-regulated protein kinase and osteoprotegerin in human osteoblasts [published online ahead of print July 8, 2010]. J Biomed Mater Res Part A doi: 10.1002/jbm.a.32839.
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cells and inhibits the survival of murine osteoclast-like cells. Biochem Biophys Res Commun 1998;252:747–52. 31. Schwartz Z, Olivares-Navarrete R, Wieland M, Cochran DL, Boyan BD. Mechanisms regulating increased production of osteoprotegerin by osteoblasts cultured on microstructured titanium surfaces. Biomaterials 2009;30:3390–6. 32. Tsukamoto Y, Fukutani S, Shin-Ike T, et al. Mineralized nodule formation by cultures of human dental pulp-derived fibroblasts. Arch Oral Biol 1992;37: 1045–55.
JOE — Volume 36, Number 10, October 2010
Platelet-rich Fibrin Increases Proliferation and Differentiation of Human Dental Pulp Cells Fu-Mei Huang, DDS, MMS, PhD,*† Shun-Fa Yang, PhD,‡ Jiing-Huei Zhao, DDS,† and YuChao Chang, DDS, MMS, PhD*† Abstract Introduction: Platelet-rich fibrin (PRF) by Choukroun’s technique is derived from an autogenous preparation of concentrated platelets without any manipulation. When delicately pressed between 2 gauzes, the PRF clot becomes a strong membrane with high potential in clinical application. However, the effect of PRF on dental pulp cells (DPCs) remains to be elucidated. This study was to determine the biological effects of PRF on DPCs. Methods: PRF samples were obtained from 6 healthy volunteers. Human DPCs were derived from healthy individuals undergoing extraction for third molars. Cell proliferation resulting from PRF was evaluated by colorimetric assay. Western blot was used to evaluate the expression of osteoprotegerin (OPG). Alkaline phosphatase (ALP) activity was examined by substrate assay. Results: PRF did not interfere with cell viability of DPCs (P > .05). DPCs were observed to attach at the edges of PRF by phase-contrast microscopy. PRF was found to increase DPC proliferation during 5-day incubation period (P < .05). PRF was found to increase OPG expression in a time-dependent manner (P < .05). ALP activity was also significantly upregulated by PRF (P < .05). Conclusions: PRF was demonstrated to stimulate cell proliferation and differentiation of DPCs by up-regulating OPG and ALP expression. These findings might serve as a basis for preclinical studies that address the role of PRF in reparative dentin formation. (J Endod 2010;36:1628–1632)
Key Words Alkaline phosphatase, dental pulp, osteoprotegerin, platelet-rich fibrin, proliferation
From the )Department of Dentistry, Chung Shan Medical University Hospital, †School of Dentistry, and ‡Institute of Medicine, Chung Shan Medical University, Taichung, Taiwan. Address requests for reprints to Prof. Yu-Chao Chang, School of Dentistry, Chung Shan Medical University, 110, Sec. 1, Chien-Kuo N. Rd., Taichung, Taiwan. E-mail address: cyc@ csmu.edu.tw. 0099-2399/$0 - see front matter Crown Copyright ª 2010 Published by Elsevier Inc. on behalf of the American Association of Endodontists. All rights reserved. doi:10.1016/j.joen.2010.07.004
1628
Huang et al.
T
ooth injury can induce reactionary or reparative dentin formation. During reparative dentin formation, the differentiation of dental pulp cells into odontoblasts depends on the regulation of multiple interacting signaling pathways (1, 2). Signaling molecules playing important roles in the self-repair process can also be liberated in pathologic conditions. To help promote endodontic tissue regeneration and healing, the local application of growth factors and host modulating agents is being used to maximize the body’s healing potential (3). Transforming growth factor (TGF)–b has been implicated in the differentiation of odontoblast-like cells and in pulp tissue repair in vivo (4). A novel platelet-derived growth factor (PDGF) isoform, PDGF-C, was observed and might be involved in the pathogenesis of rat experimental periapical lesions (5). Recently, platelet-rich plasma (6) has shown clinical success in enhancing endodontic regeneration for periapical inflammatory lesion, although the long-term predictability remains questionable, and its anticipated benefits are moderate. Platelet-rich fibrin (PRF) protocol, a simple and free technique developed in France by Dohan et al (7), is a second-generation platelet concentrate that allows one to obtain fibrin membranes enriched with platelets and growth factors after starting from an anticoagulant-free blood harvest (8, 9). PRF looks like a fibrin network. PRF contains platelets, growth factors, and cytokines that might enhance the healing potential of both soft and hard tissues (10, 11). Many growth factors such as PDGF and TGF-b are demonstrated to release from PRF (8, 9). Recently, studies have demonstrated that the PRF membrane has a very significant slow sustained release of many key growth factors for at least 1 week (12) and up to 28 days (13), which means that PRF could release growth factors with its own biological scaffold for wound healing process. This leads to the idea of using PRF membrane as a capping agent for reparative dentin formation or as a biomaterial for pulp regeneration. Osteoprotegerin (OPG), a naturally occurring inhibitor of osteoclast differentiation, binds to receptor activator of nuclear factor-kB ligand (RANKL) and blocks RANKL from interacting with RANK (14). The intensity of OPG immunoreactivity in odontoblastic layer has shown highly expressed in healthy and inflamed peripheral pulp specimens (15). Alkaline phosphatase (ALP) is a membrane-bound glycoprotein, which is one of the osteogenic differentiation markers considered to indicate the differentiation of cells to odontoblasts and mineralization process (16). OPG and ALP expression were generally regarded as markers of odontoblastic differentiation. Dental pulp cells (DPCs) are considered as cells primarily concerned with providing physical barriers and structural components in dental pulp. Cultured primary DPCs can be induced to differentiate into odontoblast-like cells and generate dentin-like mineral structure in culture dishes (17, 18). However, currently no research has addressed the role of PRF on human DPCs and its potential function for dentinogenesis. In this study, we studied the biological functions of PRF on DPCs through measuring cell proliferation, protein expression of OPG, and ALP activity.
Materials and Methods PRF Preparation The human blood samples in this study were obtained under the guidelines of the Ethics Committee of the Chung Shan Medical University Hospital. Blood from 6 healthy volunteers were treated according to the PRF protocol (10, 11) with a PC-02 table centrifuge and collection kits provided by Process (Nice, France). Briefly, blood
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Basic Research—Biology samples were taken without an anticoagulant in 10-mL glass-coated plastic tubes (Vacutainer; Becton Dickinson, Franklin Lakes, NJ) and immediately centrifuged at 3000 rpm for 10 minutes. Three layers are naturally formed in the tube: a base of red blood cells at the bottom, acellular plasma on the surface, and finally a PRF clot in the middle between the other 2 parts. The fibrin clot was easily separated from the lower part of the centrifuged blood. The PRF clot was gently pressed into a membrane with sterile dry gauze for the following experiments.
Cell Cultures Human DPCs were cultured by using an explant technique as described previously (19, 20). Briefly, impacted third molars were obtained from healthy patients of the Department of Dentistry, Chung Shan Medical University Hospital with informed consent. Teeth were sectioned horizontally below the cementoenamel junction with a no. 330 high-speed bur with water spray. Pulp tissues were removed aseptically in lamina flow, rinsed with Hanks’ buffered saline solution, and placed in a 60-mm dish. Pulp tissues were minced with a blade into small fragments and grown in Dulbecco modified Eagle medium (DMEM) supplemented with 10% fetal calf serum (FCS) and antibiotics (100 U/mL penicillin, 100 mg/mL streptomycin, and 0.25 mg/mL of amphotericin B). Cultures were maintained at 37 C in humidified atmosphere of 5% CO2 and 95% air. Confluent cells were detached with 0.25% trypsin and 0.05% ethylenediaminetetraacetic acid (EDTA) for 5 minutes, and aliquots of separated cells were subcultured. Cell cultures between the third and eighth passages were used in this study. Cell Viability Each PRF membrane was covered with 5-mL suspension of cells at a concentration of 5 104 cells/mL in 35-mm culture dishes. After day 1, day 3, and day 5 culture periods, the medium was removed, and 0.5 mL of 0.25% trypsin in phosphate-buffered saline (PBS) was added to each culture dish to detach the cells. One milliliter of medium was added to 0.5 mL of this cell suspension. Then 0.5 mL of calcium-free and magnesium-free PBS containing 0.25% trypan blue (wt/vol) was added to 0.5 mL of the cell suspension to stain nonviable cells. Fifteen-microliter cell suspension was dropped into a hemocytometer chamber (Cambridge Instruments, Buffalo, NY), and cell numbers were counted under a phase-contrast microscope (21). Cell viability is represented as viable cells/total cells. Cell Proliferation A 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) colorimetric assay was developed to monitor mammalian cell survival and proliferation in vitro. DPCs were plated at an initial density of 2 104 cells/well in DMEM containing 10% FCS. After overnight attachment, cells were incubated in fresh medium containing PRF for 1, 3, and 5 days. In the end of each final 4 hours, 50 mL of MTT (Sigma, St Louis, MO) was added to each well. The insoluble formazan was dissolved with 150 mL of dimethyl sulfoxide to each well. Optical density (OD) was determined by eluting the dye with dimethyl sulfoxide, and the spectrophotometric absorbance was measured at 550 nm by using a spectrophotometer (Hitachi, Tokyo, Japan). The percentage of the dehydrogenase activity at each time point, compared with that of the control, was calculated from the absorbance values. Western Blot Nearly confluent monolayers of DPCs were washed with serumfree DMEM for 24 hours and immediately thereafter exposed to PRF for additional 24 hours. Cell lyates were collected as described previously (22). Briefly, cells were solubilized with sodium dodecylsulfate JOE — Volume 36, Number 10, October 2010
(SDS)–solubilization buffer (5 mmol/L EDTA, 1 mmol/L MgCl2 , 50 mmol/L Tris-HCl , pH 7.5 and 0.5% Triton X-100, 2 mmol/L phenylmethysulfonyl fluoride, and 1 mmol/L N-ethylmaleimide) for 30 minutes on ice. Then, cell lysates were centrifuged at 12,000g at 4 C, and the protein concentrations were determined with Bradford reagent by using bovine serum albumin (BSA) as standards. Equivalent amounts of total protein per sample of cell extracts were run on a 10% SDS–polyacrylamide gel electrophoresis and immediately transferred to nitrocellulose membranes. The membranes were blocked with PBS containing 3% BSA for 2 hours, rinsed, and then incubated with primary anti-OPG (Abcam, Cambridge, MA) antibodies (1:1000) in PBS containing 0.05% Tween 20 for 2 hours. After 3 washes with Tween 20 for 10 minutes, the membranes were incubated for 1 hour with biotinylated secondary antibody diluted 1:1000 in the same buffer, washed again as described above, and treated with 1:1000 streptavidin-peroxidase solution for 30 minutes. After a series of washing steps, the reactions were developed by using diaminobenzidine (Zymed, South San Francisco, CA). The intensities of the obtained bands were determined by using a densitometer (AlphaImager 2000; Cell Biosciences, Santa Clara, CA). Each densitometric value was expressed as the mean standard deviation (SD).
Measurement of ALP Activity According to our recent study (23), the cell lyates were sonicated on ice bath, centrifuged at 1500g for 5 minutes, and measured the ALP activity in supernatant by ALP assay mixtures containing 0.1 mol/L 2amino-2-methyl-1-propanol, 1 mmol/L MgCl2, and 8 mmol/L p-nitrophenyl phosphate disodium. After incubation at 37 C for 30 minutes, the reaction was stopped with 0.1 N NaOH, and the absorbance was read at 405 nm. The OD of control cells was considered to be 100%. The relative ALP activity of U2OS cells was calculated by the formula (OD of experimental sample/OD of control cells) 100%. Each value was expressed as the mean SD. Statistical Analysis Three replicates of each time point were performed for each test. All assays were repeated 3 times to ensure reproducibility. Statistical analysis was by one-way analysis of variance (ANOVA). Tests showing differences in the treatments were analyzed by the Duncan test, and a P value of <.05 was considered statistically significant.
Results The cell viability of DPCs treated with or without PRF was shown by the same trypan blue dye uptake. No significant statistical differences were seen between the PRF stimulation and untreated control during 5-day culture period (P > .05). PRF exhibited no cytotoxic effects to DPCs. DPCs on the flat surface of the culture dishes demonstrated a spindle-shaped morphology. Moreover, DPCs were found to attach at the edge of PRF under observation by phase-contrast microscopy (Fig. 1a). Fig. 1b shows the mitogenic effects of PRF on DPCs. PRF was found to increase DPC proliferation in a time-dependent manner (P < .05). From MTT proliferation assay, the absorbance values increased about 1.5-, 3.2-, and 3.7-fold on days 1, 3, and 5, respectively, as compared with untreated control (P < .05). As shown in Fig. 2a, PRF was found to up-regulate OPG protein expression in DPCs during 5-day culture period (P < .05). The quantitative measurement by the AlphaImager 2000 is shown in Fig. 2b. The levels of the OPG increased about 2.8-, 2.1-, and 1.6-fold on days 1, 3, and 5, respectively. The effects of PRF on the ALP activity were determined by substrate assay. As shown in Fig. 3, the ALP activity was found to be significantly
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Figure 2. (A) Kinetics of OPG expression in DPCs exposed to PRF for 1, 3, and 5 days, respectively, by Western blot analysis. b-actin was performed to monitor equal protein loading. (B) Levels of OPG protein treated with PRF were measured by AlphaImager 2000. Relative level of OPG protein expression was normalized against b-actin signal, and control was set as 1.0. OD values represent the mean SD. )Significant difference from control values with P< .05.
Figure 1. (A) DPCs exhibited their spindle-shaped morphology when cultured with PRF membranes. DPCs were found to attach at the edge of PRF membrane. (B) Effects of PRF on cell proliferation of DPCs were measured by MTT colorimetric assay. Percentage of absorbance value of DPCs with PRF compared with that of each control was calculated. Each point and bar represent the mean SD. )Significant difference from control value at P < .05.
enhanced after PRF treatment (P < .05). The relative ALP activities were 2.0-, 3.6-, and 3.0-fold after exposure to PRF as compared with untreated control, respectively (P < .05).
Discussion PRF by Choukroun’s technique is produced in a totally natural manner, without using an anticoagulant during blood harvesting or bovine thrombin or calcium chloride for platelet activation and fibrin polymerization (9). The PRF clot forms a strong natural fibrin matrix and shows a complex architecture as a healing matrix. It is an autologous biomaterial and not an improved fibrin glue. Unlike the plateletrich plasma, PRF by Choukroun’s technique does not dissolve quickly after application; instead, the strong fibrin matrix is slowly remodeled in a similar way to a natural blood clot (12, 24). In this study, PRF exhibited no cytotoxic effect to DPCs. Similar results have shown that PRF demonstrated no cytotoxicity toward many cells including preadipocytes, dermal prekeratinocytes, osteoblasts, oral epithelial cells, periodontal ligament cells, and gingival fibroblasts (25–27). In addition, each DPC maintained its original morphology. DPCs were observed to attach at the edge of PRF 1630
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membrane by phase-contrast microscopy. Consistently, osteoblasts, periodontal ligament cells, and gingival fibroblasts were demonstrated to attach at the edge of PRF membrane (25). Moreover, osteoblasts were found to form a network at the surface and the periphery of the PRF membrane and organized the apposition of a bone matrix under scanning electron microscopy analysis (26). Thus, the biocompatibility of PRF is not cell type–specific. Taken together, PRF acts as a biomaterial to DPCs. In this study, PRF was found to increase DPC proliferation as a mitogen. Similar results were reported by He et al (13), Tsai et al (25), and Dohan Ehrenfest et al (26) that PRF stimulated cell proliferation of gingival fibroblast, periodontal ligament cells, and osteoblasts in vitro. The mechanism responsible for the cell proliferation by PRF might be explained as follows. Many growth factors such as PDGF and TGF-b are released from PRF (8, 9, 12, 13). These findings suggest that PRF possibly modulates DPC proliferation by PDGFrelated and TGF-b–related mechanisms. In the present study, the expression of OPG was stimulated by PRF. Two hypotheses could account for the mechanisms of the stimulation of OPG secretion in DPCs. The first is that OPG secretion is stimulated by increasing the number of OPG-secreted cells, namely stromal/osteoblastic cells. The second is that PRF can enhance the amount of OPG secreted from each cell. Our results imply that the effect of PRF on the proliferation of DPCs and the enhancement of OPG secretion might be attributed to the odontoblastic differentiation and mineralization process. Elevated levels of OPG are found in DPCs, raising the question of which factors, especially growth factors in PRF, result in the outcome
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Basic Research—Biology References
Figure 3. Effects of PRF on ALP activity in DPCs. Percentages of ALP activity in the presence of PRF relative to that in the control are shown. Results were averaged from 3 independent experiments. )Significant differences from control values with P < .05.
observed in the present study. As already noted, the contribution of PRF to the proliferation of DPCs is the most important factor in the enhancement of OPG secretion. TGF-b, one of the main growth factors detected in PRF, has been demonstrated in pulp tissues after injury and is implicated in the differentiation of odontoblast-like cells and in pulp tissue repair (28, 29). Consistently, TGF-b was found to increase OPG expression in MC3T3-E1 cells (30) and MG 63 cells (31). Increase of ALP activity, essential for biomineralization, was considered to indicate the differentiation of cells to odontoblasts (32). In the present study, ALP activity was elevated by PRF in a timedependent manner. Similar results have reported that PRF could increase ALP activity in osteoblasts in vitro (13, 26). These results indicated that PRF might contribute to the differentiation of human DPCs. Choukroun’s PRF is derived from an autogenous preparation of concentrated platelets without any manipulation and is widely used in clinical dentistry as a vector for cell growth factors. The present study demonstrates that PRF can stimulate DPC proliferation. PRF was demonstrated to enhance OPG protein expression and ALP activity. Taken together, PRF can increase DPC proliferation and differentiation, suggesting potential applications of PRF as a biological molecule to promote the regeneration of lost or injured dental pulp tissues and stimulate reparative dentinogenesis. Future studies are required to elucidate the precise mechanism of action of PRF for dental pulp regeneration both in vitro and in vivo. In vitro experiments are very helpful to assay the biological effects of PRF, but they might be limited in their ability to simulate the clinical condition. It might be unrealistic to transfer in vitro findings to in vivo situations. The other factors that modulate the expression of odontoblastic differentiation markers such as dentin sialophosphoprotein, dentin matrix protein, and osteocalcin should be evaluated in further studies. It is also necessary to continue investigating the in vivo effects of PRF on DPCs for dentinogenesis.
Acknowledgments This study was supported by a research grant (CSMU-96RD-05) from Chung Shan Medical University by Prof. Chang. The authors disclose no conflicts of interest.
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