Volume 33, Issue 8
Published by AEGIS Communications
Socket Grafting with Calcium Phosphosilicate Alloplast Putty: A Histomorphometric Evaluation
Lanka Mahesh, BDS, MBA; Maurice A. Salama, DMD; Gregori M. Kurtzman, DDS; and Frederic P.C. Joachim, DDS, MSc
BACKGROUND: Socket grafting with a bone graft substitute immediately after extraction is essential to preserve the ridge architecture for implant placement. Several bone graft substitutes have been tested for their ability to effectively regenerate osseous tissue in the sockets. Evidence suggests that socket bone typically regenerates during a period of 6 to 8 months or longer, depending on several factors including the original ridge dimensions, type of graft, and the overall systemic health of the individual. The purpose of this study is to histologically evaluate the bone regeneration potential of a novel synthetic calcium phosphosilicate putty (CPS) graft substitute.
METHODS: After extraction of the involved teeth, CPS putty graft was placed, and the sockets were covered with a collagen plug. Cores were taken from 20 patients for histological evaluation prior to implant placement. Ten cores were processed decalcified with hematoxylin and eosin (H&E) stain and the remaining 10 were processed undecalcified. Histomorphometric data obtained from both sets is presented.
RESULTS: Histomorphometric analysis revealed an average vital bone content of 49.5 (±20.7). A residual graft content of 4.3% (± 7.8) was observed following a healing time of 4.9 (± 0.8) months.
CONCLUSIONS: Clinical and histomorphometric data suggests that CPS putty is a good choice for socket bone regeneration in implant-related surgeries.
Periodontal disease, caries, or trauma may make it necessary to extract teeth.1 It is important to replace missing teeth, because ridge and site preservation at the time of extraction aid in long-term success and preservation of the osseous structure, irrespective of the procedure selected for tooth replacement.2
Several bone graft substitutes are currently available for replacing and regenerating the osseous structures. Autogenous bone has been the “gold standard” for its ability to supply the scaffold for osteoconduction, the growth factors for osteoinduction, and bone cells for osteogenesis.2
Bone graft substitutes can be classified based on the biological function they impart at the defect site: osteoinductive and/or osteoconductive. Osteoinduction is a biochemical process resulting in the recruitment and differentiation of the surrounding viable cells into bone-forming cells by the molecules contained within the graft, while osteoconduction is a physical phenomenon by which the matrix of the graft forms a scaffold on which cells in the recipient site are able to form new bone. It is a 3-dimensional (3-D) process of ingrowth of capillaries, perivascular tissue, and osteoprogenitor cells from the surrounding tissues into the graft. Most commercially available bone substitutes are osteoconductive, while some exhibit osteoinductive properties. The osteoinductive properties of a graft material vary, depending on donor tissue processing methods used. All xenografts and alloplasts are osteoconductive and provide an excellent scaffold for bone regeneration.
Biomaterial development has improved the characteristics and properties of potential synthetic bony substitutes.3 A new class of graft substitutes is bioactive—that is, they interact with the surrounding tissues both physically and chemically, unlike most osteoconductive graft substitutes, which are bioinert and provide only a physical scaffold for bone to grow through. These graft substitutes are helping to increase understanding of the graft-host interface, which has not been assessed completely to evaluate the bone fill potential.4
Bioactive glass-ceramics have demonstrated biocompatibility, resulting in direct contact with bone in the healed sites.5,6 The first report on a bioactive material appeared in 1971; it was a four-component oxide mixture consisting of 45% silicon, 24.5% sodium, 24.5% calcium, and 6% phosphorous.7 This product—a synthetic calcium phosphosilicate (CPS) putty—has evolved and is now being marketed as a pre-mixed, moldable material called NovaBone Dental Putty® (NovaBone Products, www.novabone.com) It consists of a bimodal bioactive phase with an additive and a glycerin binder. The CPS putty has been approved for bone regeneration in osseous defects throughout the body including spine, orthopedic, craniofacial, and dental defects.
This article’s purpose is to histologically evaluate CPS putty as a bone graft substitute when used in 44 human alveolar post-extraction sockets.
Materials and Methods
The study involved 42 patients—27 male, 15 female—between the ages of 25 and 79 (mean: 46). The study consisted of 49 alveolar sockets: 30 sockets were in the maxilla, with 16 in the anterior area (cuspid-to-cuspid) and 14 in the posterior area (premolar-molar); the remaining 19 sockets were in the mandible, with six in the anterior and 13 in the posterior area.
Usual case selection criterion excluded pregnant women and prospects with acute periodontal disease, the human immunodeficiency virus, and those with any systemic medical condition that could interfere with healing (eg, osteoporosis, steroid therapy, autoimmune diseases, etc).
The involved teeth were extracted atraumatically under local anesthesia to preserve as much socket architecture as possible. After extraction, the sockets were debrided, and an effort was made to completely remove all the inflammatory granulation tissue. CPS putty was injected into the sockets (Figure 1) and a spatula was used to gently adapt the material to the socket walls; care was taken not to compact the material too tightly. About 0.5 cc to 1.0 cc CPS putty was used in each socket. The putty consistency helped contain the graft substitute in the defect, and the unique cartridge delivery system enhanced the experience and minimized graft wastage. No membranes were used, but the socket was covered with Collaplug® (Zimmer Dental, www.zimmerdental.com) to help retain the material and the mucosa was sutured with resorbable sutures.
Standard post-operative instructions were given and the 27—among the 49 total patients—patients treated in India were given postoperative antibiotics (amoxicillin, 250 mg t.i.d.). All patients were placed on chlorhexidine oral rinse post-operatively. Pre- and immediate post-operative radiographs were taken. Patients were then recalled 2 to 3 weeks postoperatively to evaluate the clinical healing. Postoperative radiographs were taken between 4 and 6 months (average 4.9 months) to evaluate the bone regeneration prior to implant placement.
Prior to implant placement, a 2.7-mm inner diameter (3.5 mm outer diameter) trephine bur was used to obtain a bone core. All efforts were made to obtain the core from the center of the regenerated socket, where the implant was to be placed. The trephines with the biopsy cores were placed in 10% formalin for fixation and sent for histological study. All 20 cores were processed non-decalcified in two different locations. Of the 20 cores that were obtained, 10 cores were processed decalcified by the department of Oral Pathology, at Rajiv Gandhi University in Bangalore, India, and the remainder were processed non-decalcified by the hard tissue laboratory at the University of Minnesota in Minneapolis.
Decalcified histologies were performed by the Department of Oral Pathology at Rajiv Gandhi University. All samples were subjected to microwave decalcification with 5% nitric acid solution (95 ml deionized water with 5 ml nitric acid). The tissue specimen was immersed in the above solution and placed in the microwave and heated up to 800w for 20 seconds; this cycle was repeated three times with a 1-hour interval between each cycle. This was followed by routine automatic tissue processing, embedding, and sectioning, after which the sections were stained with hematoxylin and eosin (H&E).
Hard Tissue Histology
The Division of Pathology at the University of Minnesota performed non-decalcified histology and provided histomorphometric data on the remainder of the cases. Upon receipt, specimens were dehydrated with a graded series of alcohols for 9 days. Following dehydration, the specimens were infiltrated with a light-curing embedding resin (Technovit® 7200 VLC (Kulzer, www.kulzer-technik.de). This was then followed by 20 days of infiltration with constant shaking at normal atmospheric pressure, after which the specimens were embedded and polymerized by 450 nm light. Care was taken during specimen preparation to ensure that the temperature of the specimens never exceeded 40ºC. Specimens were then cut and ground; they were prepared in an apico-coronal direction parallel to the long axis and were cut to a thickness of 150 µm on a cutting/grinding system (XAKT Technologies, Inc, www.exaktusa.com). The cores were polished to a thickness of 45 µm to 65 µm with a series of polishing sandpaper disks from 800 to 2,400 grit, using a microgrinding system, which was then followed by a final polish with 0.3 µm alumina polishing paste. The slides were stained with Stevenel’s blue and Van Gieson’s picro fuchsin, and a cover slip was placed for histologic analysis using bright field and polarized microscopy.
Clinically and radiographically, all the sockets healed without any complications or adverse reactions. No signs of inflammation or infection were observed during the healing period. No particles of CPS putty were observed upon re-entry with the exception of one case (patient ID: 14/11-02) where a few particles were noted, and all the sockets appeared to be filled with a dense hard tissue. Clinically, there was no significant difference noted in the "tactile feel" when drilling into treated sites compared to adjacent non-treated sites, with bleeding in the graft site osteotomies showing clear evidence of vascular ingrowth of the grafted sites. Radiographs taken at 5 to 6 months (average 4.9) post grafting demonstrated dense bone fill in all study participants. The trabecular pattern within the grafted area presented with very similar trabecular patterns to the adjacent native bone, which was indicative of graft resorption and remodeling.
A 50-year-old woman in stable health presented with a failed root canal restoration and a tooth fracture associated with a bridge (lower right second premolar) (Figure 2). Following evaluation, the decision was made to extract the tooth and place an implant. The tooth was extracted atraumatically, at which time it was determined that the quality of bone was insufficient for immediate implant placement due to the loss of the buccal plate (Figure 3).
The socket was debrided and CPS putty was placed, filling the socket to the level of the lingual crest and shaping it on the buccal side to simulate the natural anatomy in the region (Figure 4). A 2-week follow-up visit revealed the tissue healing in the area and the contour of the ridge (Figure 5). The patient was recalled at 6 months post graft for evaluation. Clinically, the ridge looked healthy and well formed (Figure 6). Radiographically at 6 months, the site demonstrated good bone fill (Figure 7), which was confirmed clinically upon reentry with a full-thickness flap (Figure 8). The trabecular pattern in the regenerated area demonstrated similar characteristics to the trabecular pattern in the ungrafted area. It was noted that the defect completely regenerated, including the buccal wall, with no visual evidence of residual CPS putty graft particles.
Histologic images at 40x (Figure 10) and 100x (Figure 11) magnifications revealed dense interconnected vital bone with remnants of graft material. At 100x magnification, the osteoblasts in the lacunae could be clearly seen, which was indicative of healthy cancellous osseous tissue. An endosseous root form implant was placed into the site following core removal for histological analysis and site preparation. When drilled, the bone at the grafted site had a tactile feel of dense bone comparable to a D3 type bone typically found in the posterior mandible. The fixture was placed with a final insertion torque of 30 Ncm. A healing period of 4 months was allowed prior to restoration of the implant fixtures. After the healing period, a custom abutment was placed on the new fixture and both fixtures were restored with porcelain-fused-to-metal single fixed prosthetic units.
A 22-year-old woman presented with pain in the lower left first molar. Upon examination, the tooth was seen to be grossly decayed with no clinical crown (Figure 12). Radiographic examination demonstrated apical lesions on both root apices, furcation involvement, and decay, leaving the tooth a poor restorative prognosis (Figure 13). For best prognosis, it was decided that the tooth be extracted, the area be grafted with CPS putty, and that the area be re-entered after 4 months for implant placement. On the day of the surgery, the attached gingiva was detached using a Buser elevator (Hu-Friedy Mfg. Co. Inc., www.hu-friedy.com) under local anesthesia. The tooth was then extracted with luxators, and care was taken to preserve the buccal/lingual plates.
After extraction, the socket was debrided and irrigated with saline. CPS putty was placed into the socket using the unique cartridge delivery system (Figure 14). About 0.5 cc of the graft material was used to fill the socket, which was then covered with CollaPlug (Figure 15) and stabilized with 3-0 Cytoplast® sutures (Osteogenics Biomedical, www.osteogenics.com). Following a healing period of 4 months a radiograph was taken of the site to check graft organization (Figure 16). The site demonstrated organization of the graft and blend with the surrounding bone. A core was taken prior to implant placement and was processed by the Oral Pathology Department at Oxford Dental College, Bangalore, India. The histology slide (Figure 17) demonstrated good bone fill surrounded by vascular marrow tissue. Also noticeable were the osteocytes in mature bone. Some remnants of the bone graft substitute could also be seen. The site was enlarged with osteotomy burs to accommodate an endosseous threaded implant fixture, which was placed, and a cover screw inserted into the fixture. Following 3 months of healing to allow osteointegration of the fixture, a radiograph was taken to verify integration prior to the restorative phase of treatment (Figure 18).
Following non-decalcified histologic preparation, the cores were evaluated histomorphometrically. Cores were digitized at the same magnification using a Axiolab microscope (Carl Zeiss MicroImaging Inc, www.zeiss.com) and a digital camera (Coolpix® 4500, Nikon Americas Inc, www.nikonusa.com). Histomorphometric measurements were completed using a combination of programs (Adobe Photoshop Adobe® Photoshop® [Adobe Systems Inc, www.adobe.com] and NIH Image [National Institutes of Health, available to the public at http://rsb.info.nih.gov/nih-image]). Parameters evaluated within the study were total area of the core, percentage of new bone formation, and percentage of residual graft material. The remainder of the area was considered fibrous or marrow tissue. The primary section evaluated for each specimen was taken from the most central region of the obtained core. No comparison was made between the apical and coronal sections.
Histomorphometric evaluation of the non-decalcified cores revealed an average vital bone content of 49.5 (±20.7). In comparison, autogenous trabecular bone volumes, which can vary widely, have a range from under 20% to 40%.8 A residual graft content of 4.3% (± 7.8) was found for the CPS bone graft, following a healing time of 4.9 (± 0.8) months. The case of patient ID 14/11-02 was found to be densely filled with osteoid tissue, although the implant placed in the area was stable. The histomorphometric data from the 10 cases are presented in Table 1.
CPS putty is a third-generation bioactive graft material that not only provides a physical scaffold for the bone tissue to grow, but also interacts chemically with the surrounding tissue to impart an increased level of osteoblastic activity at the defect site. Upon implantation of the CPS putty, the smaller CPS particles release calcium and phosphorous ions into the area, with the binder material becoming absorbed over a period of 1 week, exposing the larger CPS particles to blood. Once the clot is organized, the dissolution of binder and the smaller CPS particles create a porous network for fluid flow through the clot, thus exposing the larger bioactive particles to blood. Within hours, the leached calcium and phosphate ions redeposit, resulting in formation of calcium phosphate nodules. These subsequently crystallize and a new surface apatite layer (hydroxycarbonite apatite) is produced. The graft substitute has been shown to bond to the connective tissue and also adjacent bone.9 This apatite layer assists in the stimulation of osteoprogenitor (undifferentiated) cells to produce transforming growth factor by the release of silicon from the surface.3,10-13 The surface reactions take place within a 2-day to 4-day time frame,14 with attachment of cells and the subsequent proliferation and differentiation of osteoblasts rapidly occurring on the surface of the bioactive material.15,16
Several publications have tried to explain the complex mechanisms and surface reactions of CPS particulate,17,18 which involves an ionic dissolution process that eventually stimulates genes that are involved in regulation of osteoblast differentiation and proliferation.14,19,20 This phenomenon has been termed osteostimulation. FDA recognizes the term and defines it as “an active stimulation of osteoblast proliferation and differentiation as evidenced during in vitro studies by increased levels of DNA synthesis and of the osteoblast markers osteocalcin and alkaline phosphatase.” In 2005, an FDA 510k was approved specifically for this claim and applies only to calcium phosphosilicate-derived products (FDA 510k #K052494 (http://www.accessdata.fda.gov/cdrh_docs/pdf5/K052494.pdf).
Osteostimulative materials support a higher level of osteoblast expression and activity than is seen with materials that are merely osteoconductive. According to a recent comparative histomorphometric study of two groups by Galindo-Moreno et al, no bone loss was observed radiographically or clinically in bone core biopsies taken 6 months after sinus grafting with either a bovine hydroxyapatite (HA) or CPS particles.21 Histologic analysis demonstrated that both grafts had high biocompatibility, but in the bovine HA-containing group, minimal xenogeneic graft absorption was noted. In contrast, the CPS group presented a high absorption rate, with occasional remaining particles embedded in new normal bone. Another recent publication by Kotsakis et al also evaluated the performance of CPS putty grafts in 12 extraction sockets. The average ridge width decreased by 1.1 mm (±0.44) and the ridge height decreased by 0.83 mm (±0.22) ) (P>0.05). Histomorphometric analysis revealed good bone regeneration with an average of 39.3% (±9.3) in the socket accompanied by rapid graft absorption (>90%).22
Historically, several bone graft substitutes have been used for socket grafting and ridge preservation including allografts and xenografts. Xenografts such as BioOss (Geistlich Biomaterials, www.bio-oss.com) have enjoyed a high level of popularity in implant-related bone graft surgeries.23,24 However, the material is slowly resorbable with high concentrations of residual bone volume 9 months post-grafting.25 This study uses a modified bio-col technique, which was first advocated by Sclar.26 Allografts have also been used to preserve sockets using a similar technique. Allografts demonstrate varying levels of resorbability dependent on the mineral content in the graft.27 Some studies have demonstrated presence of graft material in the socket surrounded by new bone and connective tissue as late as 6 months post-surgery.28
The results from this study show a very high level of bone regeneration as evidenced radiographically by a trabecular pattern that resembles closely the unrestored area and histologically with osteocytes in lacuna. A high degree of revascularization, essential for the support of new bone formation, was observed within the grafted area.
This study corroborates the findings from the earlier studies with CPS graft substitute. However, longer-duration studies are essential to demonstrate the absence of shrinkage associated with grafting in sites that will not receive implants or in large defects in which implants were placed. Additionally, no evidence was noted either clinically or histologically of any significant inflammatory reaction surrounding the graft material, suggesting good tissue compatibility.
Results suggest that CPS putty is a reliable choice for osseous regeneration in cases of socket grafting and implant-related surgeries. The putty consistency of the material and its unique delivery system also increases its clinical appeal.
NovaBone Products provided graft material for support of this project. Dr. Mahesh is a current consultant for NovaBone.
NovaBone Products (manufacturer of the bone graft material used in this study) provided partial research support for this project. The authors acknowledge Srinivas Katta, Director at NovaBone Products, for his assistance in writing the discussion section of this manuscript. The authors also gratefully acknowledge the assistance of Dr. Michael Rohrer, director, Hard Tissue Research Laboratory and Oral Pathology Laboratories, University of Minnesota School of Dentistry, for the preparation of the specimens, the histologic data used for histomorphometric analysis, and the assistance of Dr. Narayan Venkataraman at The Oxford Dental College, Oral Pathology Department, Bangalore, India, for the preparation of the non-decalcified specimens in India.
1. Klinge B, Hultin M, Berglundh T. Peri-implantitis. Dent Clin North Am. 2005;49(3):661-676.
2. Lekovic V, Kenney E, Weinlaender M, et al. A bone regeneration approach to alveolar ridge maintenance following tooth extraction: report of 10 cases. J Periodontol. 1997;68(6):563-570.
3. Anderegg CR, Alexander DC, Freidman M. A bioactive glass particulate in the treatment of molar furcation invasions. J Periodontol. 1999;70(4):384-387.
4. Hench LL, West JK. Biological applications of bioactive glasses. Life Chem Rep. 1996;13:187-241.
5. Bioceramics: material characteristics versus in vivo behavior. Ann N Y Acad Sci. 1988;523:1-298.
6. Gross U, Strunz V. The interface of various glasses and glass ceramics with a bony implantation bed. J Biomed Mater Res. 1985;19(3):251-271.
7. Hench LL, Splinter RJ, Allen WC, Greenlee TK. Bonding mechanisms at the interface of ceramic prosthetic materials. J Biomed Mater Res. 1971;5(6):117-141.
8. Watzek G, Ulm C, Haas R. Anatomic and physiologic fundamentals of sinus floor augmentation. In: Jensen O, ed. The Sinus Bone Graft. Chicago, IL: Quintessence Publishing; 1992:35-37.
9. Zhong JP, Latorre GP, Hench LL. The kinetics of bioactive ceramics: Part VII. Binding of collagen to hydroxyapatite and bioactive glass. In: Anderson OH, Yli-Urpo A, eds. Bioactive Ceramics: Theory and Clinical Application. Oxford, England: Butterworth-Heinemann; 1994.
10. Price N, Bendall S, Frondoza C, et al. Human osteoblast-like cells (MG63) proliferate on a bioactive glass surface. J Biomed Mater Res. 1997;37(3):394-400.
11. Wilson J, Low SB. Bioactive ceramics for periodontal treatment: comparative studies in the patus monkey. J Appl Biomater. 1992;3(2):123-129.
12. Cancian DC, Hochuli-Vieira E, Marcantonio RA, Garcia Júnior IR. Utilization of autogenous bone, bioactive glasses, and calcium phosphate cement in surgical mandibular bone defects in cebus apella monkeys. Int J Oral Maxillofac Implants. 2004;19(1):73-79.
13. Wilson J, Pigott GH, Schoen FJ, Hench LL. Toxicology and biocompatibility of bioglasses. J Biomed Mater Res. 1981;15(6):805-817.
14. Hench LL, Polak JM. A genetic basis for design of biomaterials for in situ tissue regeneration. Key Engineering Materials. 2008;377:151-166.
15. Xynos ID, Edgar AJ, Buttery LDK, et al. Gene-expression profiling of human osteoblasts following treatment with the ionic products of bioglass 45s5 dissolution. J Biomed Mater Res. 2001;55(2):151-157.
16. Orefice R, Hench L, Brennan A. Evaluation of the interactions between collagen and the surface of a bioactive glass during in vitro test. J Biomed Mater Res. 2009;90(1):114-120.
17. Hench LL. The challenge of orthopaedic materials. Current Orthopaedics. 2000;14(1):7-15.
18. Hench Ll, Polak JM. Third-generation biomedical materials. Science. 2002;295(5557):1014-1017.
19. Bombonato Prado KF, Bellesini LS, Junta CM, et al. Microarray based gene expression analysis of human osteoblasts in response to different biomaterials. J Biomed Mater Res. 2009;88(2):401-408.
20. Carinci F, Palmieri A, Martinelli M, et al. Genetic portrait of osteoblast-like cells cultured on perioglas. J Oral Implantol. 2007;33(6):327-333.
21. Galindo-Moreno P, Avila G, Fernandez-Barbero JE, et al. Clinical and histologic comparison of two different composite grafts for sinus augmentation: a pilot clinical trial. Clin Oral Implants Res. 2008;19(8):755-759.
22. Kotsakis G, Chrepa V, Katta S. Ridge Preservation with calcium phosphosilicate putty in 12 consecutive cases. Clin Oral Implants Res. 2011;22(9):1024.
23. Mardas N, D’Aiuto F, Mezzomo L, et al. Radiographic alveolar bone changes following ridge preservation with two different biomaterials. Clin Oral Implants Res. 2011;22(4):416-423.
24. Carmagnola D, Adriaens P, Berglundh T. Healing of human extraction sockets filled with Bio-Oss. Clin Oral Implants Res. 2003;14(2):137-143.
25. Artzi Z, Tal H, Dayan D. Porous bovine bone mineral in healing of human extraction sockets. Part 1: histomorphometric evaluations at 9 months. J Periodontol. 2000;71(6):1015-1023.
26. Sclar AG. Strategies for management of single-tooth extraction sites in aesthetic implant therapy. J Oral Maxillofac Surg. 2004;62(9 suppl 2):90-105.
27. Wood RA, Mealey BL. Histologic comparison of healing after tooth extraction with ridge preservation using mineralized versus demineralized freeze-dried bone allograft. J Periodontol. 2012;83(3):329-336.
28. Beck TM, Mealey BL. Histologic analysis of healing after tooth extraction with ridge preservation using mineralized human bone allograft. J Periodontol. 2010:18(12):1765-1772.
About the Authors
Lanka Mahesh BDS, MBA
New Delhi, India
Maurice A. Salama, DMD
Gregori M. Kurtzman, DDS
Silver Spring, Maryland
Frederic P.C. Joachim, DDS, MSc