The aim of periodontal therapy is to return the periodontal tissues to a healthy state, to regenerate lost attachment apparatus, and to achieve a gain in clinical attachment level .
Periodontal treatment, oral implant surgery, and maxillofacial reconstruction are highly dependent on successful bone regeneration. Bone regenerative techniques, including graft materials, some proteins (as growth factors), and barrier membranes, are often used to improve bone quality or bone quantity before or during these treatments [2,3].
The mechanisms and pathways that govern wound healing and tissue regeneration have been studied. The cellular and molecular events resulting after a traumatic injury are mostly shared by the different tissues of the body and include early and late inflammation phases, proliferation and migration of cells, angiogenesis, granulation tissue formation, and finally matrix formation and remodeling [4,5].
It is assumed that all the phases of tissue repair process are mediated and controlled by a wide range of growth factors (GFs) and cytokines that modulate cell function through direct physical interactions with the extracellular domain of transmembrane receptors. The latter transduce secondary signals, thereby controlling diverse aspects of subcellular biology. Although the role of all the GFs involved in tissue regeneration is only partially explained, the potential benefits of many of them have been shown. For example, platelet-derived growth factor is a powerful mitogen for connective tissue cells , transforming growth factor-β (TGF-β) not only stimulates osteoprogenitor cells to proliferate but also blocks in later stages of cell differentiation and mineralization , insulin-like growth factor-1 might promote the late-stage differentiation and the activity of osteoblasts, and vascular endothelial growth factor induces endothelial cell proliferation and migration, thus initiating the angiogenic response .
Some of these GFs are released by platelets that undergo active degranulation [9,10]. Factors released from the platelets include platelet-derived growth factor, TGF-β, platelet-derived epidermal growth factor, platelet-derived angiogenesis factor, insulin-like growth factor, and platelet factor 4 .
Platelet-rich plasma (PRP) gel obtained from autologous blood is used to deliver growth factors in high concentrations to the site of the bone defect or a region requiring augmentation . There are differences in the last step of PRP preparation, which includes the addition of an agent to start gelation and the activation of platelets resulting in the release of a cascade of growth factors from the platelet α granules. Some investigators suggested different agents, such as bovine thrombin or fibrin adhesive or sodium alginate [12–17].
Although promising results have been obtained with the use of PRP in clinical applications, the optimal calcium and thrombin concentrations for PRP use are still unknown. A study conducted by Lacoste et al.  shows that calcium and thrombin induce immediate GF release from PRP in a dose-dependent manner and suggests that PRP could stimulate blood vessel formation.
To date, there is no information available concerning the effects of sodium alginate on the release of growth factors from PRP.
The purpose of this study was to determine the effect of PRP gel preparation by sodium alginate and by various concentrations of calcium–thrombin on the quantity of TGF-βl released from PRP.
Materials and methods
This study was conducted on 20 adult male Wister rats weighing 180–250 g. Rats were housed under controlled temperature, humidity, and dark–light cycle at Ain Shams University animal house. A certified pellet diet and tap water were available ad libitum.
These rats were used for quantification of TGF-β1 in PRP gel preparations using an enzyme-linked immunosorbent assay (ELISA).
The animals were operated under general anesthesia, with a single intraperitoneal injection of sodium thiopental (40 mg/kg of body weight) (Thiopental-Sod., vial, 100 mg, Egyptian Int. Pharmaceutical Industries Co., Egypt), which was sufficient for the entire procedure in which 7–9 ml of blood was drawn from each rat into two 5 ml EDTA-coated test tubes. The blood was centrifuged for 10 min at 1800 rpm at room temperature [Universal 16 centrifuge, Andreas Hettich GmbH & Co.KG (Hettich), Tuttlingen, Germany]. The blood in each tube was separated into three components: red blood cells (RBC) at the bottom of the tube; a buffy coat, a very thin white layer in the middle of the tube; and platelet-poor plasma (PPP), at the top of the tube. The entire PPP and buffy coat components, including some RBCs at the very top of the RBC layer, were pipetted out from both tubes and placed into another sterile tube. This pipetted material was centrifuged again at 3600 rpm for 10 min, and the top yellow PPP was removed, except for 0.7–0.9 ml of plasma at the bottom of the tube, which was defined as PRP, and the platelets were counted using an automated hematology analyzer.
The release of TGF-β1 was examined as a function of time and mode of PRP preparation. PRP prepared from each rat was divided into four equal aliquots:
(1) Group A: 17 units of bovine thrombin and 17 μl of 10% calcium chloride were added to each 0.1 ml PRP of the first aliquot and allowed to gel.
(2) Group B: 5 units of bovine thrombin and 5 μl of 10% calcium chloride were added to each 0.1 ml PRP of the second aliquot and allowed to gel.
(3) Group C: A measure of 0.017 g of sodium alginate was added to each 0.1 ml PRP of the third aliquot and allowed to gel.
(4) Group D: Nothing was added to the fourth aliquot.
Fifty microliter from the supernatant of each aliquot was pipetted and transferred to separate labeled polypropylene tubes and stored at −70°C. The remaining aliquots were incubated at 37°C for 7 days. Again, 50 μl from the supernatant of each aliquot was pipetted and transferred to separate labeled polypropylene tubes after 1 day, 4 days, and 7 days of PRP preparation and stored at −70°C. TGF-β1 release was assessed at 0, 1, 4, and 7 days using Biosource ELISA kit (Biosource Europe S.A., Belgium), which uses a sandwich enzyme immunoassay technique. ELISA was performed according to the manufacturer's instructions.
Statistical analysis was carried out using the Statistical Analysis Systems (Release 6.03 ed.), SAS Institute Inc., Cary, North Carolina, USA; 1988. Factorial analysis [two-way analysis of variance (ANOVA) one-way repeated measurement] was run to test the effect of group, time, and their interaction on TGF-β1 concentration.
One-way ANOVA (procedure ANOVA of SAS) followed by Duncan's Multiple Range Test were used to test the effect of group within each time. The paired t-test (procedure means of SAS) was run to compare the effect of different intervals on TGF-β1 concentration within each group .
Platelet count study
Platelet counts confirmed that the platelet-rich plasma preparation technique used in this study produced samples of highly concentrated platelets in comparison with whole blood samples. The average platelet count in the whole blood samples was 514×103/μl. The average concentration of platelets in the PRP increased by almost four fold as it was 2028.4×103/μl.
Measurement of transforming growth factor-β1 levels in platelet-rich plasma preparations by enzyme-linked immunosorbent assay
With regard to the mean concentrations of TGF-β1 in all groups at different intervals, the mean concentrations of TGF-β1 for group A increased through the 7-day interval; it was 12 983.6±10 620.4 pg/ml at zero time, 14 408.3±6394 pg/ml after 1 day, 25 836.1±4457.7 pg/ml after 4 days, and 139 311.3±77 332 pg/ml after 7 days. Although in group B, it was 17 946.8±7021.2 pg/ml at zero time, then temporarily decreased to 12 499±2113.8 pg/ml after 1 day, then increased again to 16 980±6402.4 pg/ml after 4 days, and 197 853.1±41 588.1 after 7 days. In group C, it was 10 815.8±2046.7 pg/ml at zero time, then increased to 11 705.3±2549.6 pg/ml after 1 day, and this was the highest mean reached for group C, but decreased to 8061.5±1485.3 pg/ml after 4 days and 9132.6±4483.4 pg/ml after 7 days. Although in group D, it was 17 387±7774.9 pg/ml at zero time, but temporary decreased to 13 292.5±7019.1pg/ml after 1 day, then increased to 15 302.9±3583.1 pg/ml after 4 days, and 28 896.6±6307.7 pg/ml after 7 days.
Statistically, there was no significant difference between the four groups at zero time and after 1 day, whereas after 4 days, group A was better than group B followed by group D and then group C and there was a significant difference in the mean of TGF-β1 concentrations between all groups but not between groups B and D. After 7 days, group B was better than group A followed by group D and then group C and there was a significant difference in the mean of TGF-β1 concentrations between groups A and B on the one hand and groups C and D on the other; however, no significant differences between groups A and B or between groups C and D were observed (Fig. 1).
Table 1 compared the mean changes in the concentrations of TGF-β1 in all groups during different intervals and Table 2 shows their percentage changes. There was no significant difference between all groups in the mean change in the concentration of TGF-β1 from zero time to 1 day and from 1-day to 4-day intervals, although it was the highest in group A. There were significant differences in the 4–7-day interval between groups A and B on the one hand and groups C and D on the other. However, there was no significant difference between groups A and B or between groups C and D, with the highest mean change in group B with mean change in TGF-β1 concentration (180 873.1±39 788 pg/ml), which represented 100.537% change from the total change in TGF-β1 concentration (i.e. the change from zero time to 7 days). This was followed by group A whose mean of change in TGF-β1 concentration was (113 475.2±81 457.9) and then group D whose mean of change in TGF-β1 concentration was (13 593.7±7421.8).The lowest mean of change was in group C, with mean change in TGF-β1 concentration (1071.2±3348.5 pg/ml), which represented 63.64% change from the total change in TGF-β1 concentration. The mean change in the concentration of TGF-β1 from zero time to 7-day interval showed significant difference only between groups B and C, in which the mean change was the highest in the former (179 906.2±43 173.4 pd/ml) and the lowest in the latter (−1683±3383.9 pg/ml), (Figs 2–5).
Earlier studies showed that local application of growth factors alone or mixed with bone allograft is capable of increasing bone growth, accelerating soft tissue healing, and facilitating periodontal repair in animal and human studies . A major concern in the delivery of the growth factors to the site of bone healing has been their short half-lives . Some growth factors administered at surgery are degraded before being used in the osteogenic process, which occurs at least 2 weeks after trauma [22,23].
PRP has been introduced into the field of periodontal surgery for the purpose of periodontal tissue regeneration. In this process, PRP combines the advantage of an autologous fibrin clot that will aid in hemostasis and provide growth factors in high concentrations to the site of a tissue defect. PRP has the advantage that it can also act as a binding medium for bone grafts, making it easier to handle and place them into the graft site [1,12,13,24].
As a result of variation in procedures for centrifugation of the original blood samples (e.g. force, time), platelet density in PRP preparations was different; there were significant variations in growth factor concentrations between individuals, platelet and white blood cell content, and degree of platelet activation by the production process. In this, the centrifugation force and the time used in PRP preparation resulted in an increase in platelet concentrations by almost four fold in comparison with the original whole blood samples.
In addition in earlier studies, there are differences in the agents added to PRP to start gelation and the activation of platelets resulting in the release of growth factors from the platelet α granules [12–17]. It is believed that varying methods of PRP preparation, and consequently, concentrations of growth factor levels may explain the inconsistent clinical results. To the best of our knowledge, this study may be the first to evaluate the in-vivo effect of PRP activated by sodium alginate.
Determination of the effects of sodium alginate and various concentrations of calcium–thrombin on the release of TGF-βl from PRP was the aim in this study.
TGF-βl was selected in this study as it is generally considered as one of the growth factors mostly involved with general connective tissue repair and bone regeneration [25,26].
In this study, TGF-β1 levels in PRP preparations were measured by ELISA as a function of time (at 0, 1, 4, and 7 days) and mode of PRP preparation, which was either by adding 17 units of bovine thrombin and 17 μl of 10% calcium chloride to each 0.1 ml PRP or by adding 5 units of bovine thrombin and 5 μl of 10% calcium chloride to each 0.1 ml PRP or by adding 0.017 g of sodium alginate to each 0.1 ml PRP or without adding any activating agent. The first mode of PRP preparation referred to the concentrations reported by Marx et al. , who concluded that addition of PRP to bone grafts produced a quantifiably enhanced result in comparison with grafts alone. The second mode of PRP preparation referred to the concentrations reported by Gandhi et al. , who concluded that PRP normalized cellular proliferation and chondrogenesis and improved the mechanical strength of diabetic fracture healing. The third mode of PRP preparation referred to a study conducted by Okuda et al. , who concluded that treatment of intrabony periodontal defects with a combination of PRP and hydroxyapatite led to a more favorable clinical improvement than hydroxyapatite with saline.
Using these different modes in this study did not show a significant difference in the mean levels of TGF-β1 at zero time and after 1 day (Fig. 1). This might indicate that the release of TGF-β1 at these early intervals was because of the centrifugation process itself. Most TGF-β1 was released in the 4–7-day interval but it was significantly elevated when thrombin and calcium were used in either concentration in comparison with other modes of PRP preparation (Table 1, Fig. 2). This was in accordance with the studies performed by Puri , Davies et al. , and Lasne et al.  who concluded that platelet activation and degranulation by thrombin were mediated by the action of thrombin on its transmembrane receptor on the platelets, which led to an increase in intracellular calcium concentration. This came initially from intracellular stores and then later from an extracellular source, which was provided in this study in the form of calcium chloride.
Conversely, the findings of this study were not in accordance with two earlier studies; one was performed by Landesberg et al.  in which more than 81.4% of the TGF-β1 was released from PRP after 1 day of activation by thrombin and calcium and the other was performed by Lacoste et al.  in which calcium and thrombin induced immediate GF release from PRP (after 30 min) in a dose-dependent manner. This might be attributed to many factors, which include the limited sample size evaluated in their studies, difference between rat blood and human blood as they used the latter in their studies, different ELISA kits, and the different procedures used by Lacoste et al.  for centrifugation of the original blood samples (both centrifugations were hard spins).
The highest amount of TGF-β1 released from PRP was obtained at 4-day and 7-day intervals by using bovine thrombin and calcium chloride as evidenced by measurement of TGF-β1 levels by ELISA. The lowest amount of TGF-β1 released from PRP through all intervals was obtained by using sodium alginate.
The authors thank the members of the Medical Research Center, Ain Shams University, for their technical assistance in this study.
1. Hanna R, Trejo PM, Weltman RL. Treatment of intrabony defects with bovine-derived xenograft alone and in combination with platelet-rich plasma: a randomized clinical trial J Periodontol. 2004;75:1668–1677
2. Brunel G, Brocard D, Duffort JF, Jacquet E, Justumus P, Simonet T, et al. Bioabsorbable materials for guided bone regeneration prior to implant placement and 7-year follow-up: report of 14 cases J Periodontol. 2001;72:257–264
3. Zitzmann NU, Scharer P, Marinello CP. Long-term results of implants treated with guided bone regeneration: a 5-year prospective study Int J Oral Maxillofac Implants. 2001;16:355–366
4. Martin P. Wound healing--aiming for perfect skin regeneration Science. 1997;276:75–81
5. Polimeni G, Xiropaidis AV, Wikesjo UM. Biology and principles of periodontal wound healing/regeneration Periodontol 2000. 2006;41:30–47
6. Graves DT, Valentin Opran A, Delgado R, Valente AJ, Mundy G, Piche J. The potential role of platelet-derived growth factor as an autocrine or paracrine factor for human bone cells Connect Tissue Res. 1989;23:209–218
7. Maeda S, Hayashi M, Komiya S, Imamura T, Miyazono K. Endogenous TGF-beta signaling suppresses maturation of osteoblastic mesenchymal cells EMBO J. 2004;23:552–563
8. Ferrara N, Gerber HP, LeCouter J. The biology of VEGF and its receptors Nat Med. 2003;9:669–676
9. Khan SN, Bostrom MP, Lane JM. Bone growth factors Orthop Clin North Am. 2000;31:375–388
10. Maloney JP, Silliman CC, Ambruso DR, Wang J, Tuder RM, Voelkel NF. In vitro release of vascular endothelial growth factor during platelet aggregation Am J Physiol. 1998;275:H1054–H1061
11. Sanchez AR, Sheridan PJ, Kupp LI. Is platelet-rich plasma the perfect enhancement factor? a current review Int J Oral Maxillofac Implants. 2003;18:93–103
12. Marx RE, Carlson ER, Eichstaedt RM, Schimmele SR, Strauss JE, Georgeff KR. Platelet-rich plasma: growth factor enhancement for bone grafts Oral Surg Oral Med Oral Pathol Oral Radiol Endod. 1998;85:638–646
13. Anitua E. Plasma rich in growth factors: preliminary results of use in the preparation of future sites for implants Int J Oral Maxillofac Implants. 1999;14:529–535
14. Whitman DH, Berry RL, Green DM. Platelet gel: an autologous alternative to fibrin glue with applications in oral and maxillofacial surgery J Oral Maxillofac Surg. 1997;55:1294–1299
15. Landesberg R, Roy M, Glickman RS. Quantification of growth factor levels using a simplified method of platelet-rich plasma gel preparation J Oral Maxillofac Surg. 2000;58:297–300; discussion 300–301.
16. Sonnleitner D, Huemer P, Sullivan DY. A simplified technique for producing platelet-rich plasma and platelet concentrate for intraoral bone grafting techniques: a technical note Int J Oral Maxillofac Implants. 2000;15:879–882
17. Okuda K, Tai H, Tanabe K, Suzuki H, Sato T, Kawase T, et al. Platelet-rich plasma combined with a porous hydroxyapatite graft for the treatment of intrabony periodontal defects in humans: a comparative controlled clinical study J Periodontol. 2005;76:890–898
18. Lacoste E, Martineau I, Gagnon G. Platelet concentrates: effects of calcium and thrombin on endothelial cell proliferation and growth factor release J Periodontol. 2003;74:1498–1507
19. Statistical Analysis Systems (SAS). Statistical Analysis Systems (Release 6.03 ed.), SAS/STAT user's guide. 1988 Cary, NC SAS Institute, Inc.
20. Cochran DL, Wozney JM. Biological mediators for periodontal regeneration Periodontol 2000. 1999;19:40–58
21. Nimni ME. Polypeptide growth factors: targeted delivery systems Biomaterials. 1997;18:1201–1225
22. Amler MH. The time sequence of tissue regeneration in human extraction wounds Oral Surg Oral Med Oral Pathol. 1969;27:309–318
23. Koveker GB. Growth factors in clinical practice Int J Clin Pract. 2000;54:590–593
24. Marx RE. Platelet-rich plasma: evidence to support its use J Oral Maxillofac Surg. 2004;62:489–496
25. Roberts AB, Sporn MB. Physiological actions and clinical applications of transforming growth factor-beta (TGF-beta) Growth Factors. 1993;8:1–9
26. Miyazono K, Ten Dijke P, Ichijo H, Heldin CH. Receptors for transforming growth factor-beta Adv Immunol. 1994;55:181–220
27. Gandhi A, Doumas C, O'Connor JP, Parsons JR, Lin SS. The effects of local platelet rich plasma delivery on diabetic fracture healing Bone. 2006;38:540–546
28. Puri RN. Phospholipase A2: its role in ADP- and thrombin-induced platelet activation mechanisms Int J Biochem Cell Biol. 1998;30:1107–1122
29. Davies TA, Drotts DL, Weil GJ, Simons ER. Cytoplasmic Ca2+
is necessary for thrombin-induced platelet activation J Biol Chem. 1989;264:19600–19606
30. Lasne D, Donato J, Falet H, Rendu F. Different abilities of thrombin receptor activating peptide and thrombin to induce platelet calcium rise and full release reaction Thromb Haemost. 1995;74:1323–1328
31. Landesberg R, Burke A, Pinsky D, Katz R, Vo J, Eisig SB, et al. Activation of platelet-rich plasma using thrombin receptor agonist peptide J Oral Maxillofac Surg. 2005;63:529–535