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Molecular Analysis of Healing at a Bone-Implant Interface

¹ Department of Orthopaedic Surgery, University of California, San Francisco, CA 94110-1342, USA;
² Department of Biomedical Engineering, Jonsson Engineering Center, Rensselaer Polytechnic Institute, Troy, NY 12180-3590, USA;
³ Faculty of Dentistry, Université de Montréal, Canada; and
⁴ Department of Plastic and Reconstructive Surgery, Stanford University, 257 Campus Drive, Room GK207, Stanford, CA 94305, USA

Correspondence: ^* corresponding author, brunsj{at}rpi.edu

	ABSTRACT

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While bone healing occurs around implants, the extent to which this differs from healing at sites without implants remains unknown. We tested the hypothesis that an implant surface may affect the early stages of healing. In a new mouse model, we made cellular and molecular evaluations of healing at bone-implant interfaces vs. empty cortical defects. We assessed healing around Ti-6Al-4V, poly(L-lactide-co-D,L,-lactide), and 303 stainless steel implants with surface characteristics comparable with those of commercial implants. Our qualitative cellular and molecular evaluations showed that osteoblast differentiation and new bone deposition began sooner around the implants, suggesting that the implant surface and microenvironment around implants favored osteogenesis. The general stages of healing in this mouse model resembled those in larger animal models, and supported the use of this new model as a test bed for studying cellular and molecular responses to biomaterial and biomechanical conditions.

Key Words: bone • healing • implants • osteoblast

	INTRODUCTION

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Despite high success rates, implants still fail, due to wear debris (Wang et al., 2004), excessive micromotion (Soballe et al., 1992; Brunski, 1999), or excessive loading (Hoshaw et al., 1994; Isidor, 1997; Esposito et al., 2000). An understanding of the biological mechanisms of implant success and failure is fundamental to the development of preventive or remedial strategies for the treatment of loosened implants. Numerous studies (Brånemark et al., 1977; Lazzara et al., 1999; Schatzker, 2002; Berglundh et al., 2003; Watzak et al., 2005) have indicated that a typical initial interface will have some regions with direct bone-implant contact and other regions with bone-implant gaps. Regions of contact provide initial mechanical stability, despite surgical damage to bone and bone remodeling, which transiently increase porosity (Hoshaw et al., 1995; Huja et al., 1999). Gaps heal through blood clotting, matrix remodeling, angiogenesis, cell differentiation, woven bone formation, and turnover of woven bone (Berglundh et al., 2003; Davies and Hosseini, 2000). However, the source of skeletal progenitor cells, the rate-limiting steps that control their differentiation into bone, the signaling molecules that direct this process, and the role of biomechanical and biomaterial factors on cell fate decisions remain to be elucidated. We have developed a murine model for studying such topics at cellular and molecular levels. Here, we compared healing around implants with healing of empty implant sites, to assess whether the presence of an implant accelerated the initiation of bone healing.

	MATERIALS & METHODS

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Implants
Ti-6Al-4V implants ("Ti alloy", Whaledent, Inc., Akron, OH, USA), 1.0 mm diameter by 2.0 mm length, were lightly sand-blasted with 25 µm Al₂O₃ powder and ultrasonically cleaned for 5 min. Poly(L-lactide-co-D,L,-lactide) implants were made from "BioPin" implants (Imtec Corp., Ardmore, OK, USA). We modified pre-sterilized BioPin implants (resembling miniature thumbtacks with 1.0 mm shank diameter and 3.5 mm length) by cutting off the pointed tip, leaving a blunt tip and shank length 1.5 mm. Stainless steel implants ("303 SS", Insect Pins, Fine Science Tools, Foster City, CA, USA) were 303 medical-grade stainless steel, 0.25 mm diameter.

Preparation of Empty Holes and Placement of Implants
All procedures followed protocols approved by the Institutional Committee on Animal Research. Adult wild-type mice (males, 3–5 mos old) were anesthetized. For empty holes, Ti alloy, and BioPin implants, a 0.8-mm hole was drilled in the anterio-proximal tibia and enlarged via a 1.0-mm drill to minimize bone damage, as previously described (Colnot et al., 2005). Implants were press-fitted into slightly undersized holes, and wounds were closed. For 303 SS implants, each pin was transfixed percutaneously in the proximal tibia, leaving only a small segment across the leg. Following surgery, mice received subcutaneous injections of buprenorphine for analgesia and were allowed to ambulate freely. Mice were killed at days 3, 5, 7, 10, 14, 21, and 28 post-surgery (n = 2 d3, 2 d7, and 3 d10 for 303 SS implants; n= 3 d3, 2 d5, 2 d7, 2 d10, 1 d14, 2 d21, and 2 d28 for Ti alloy implants; and n = 3 d3, 4 d5, 5 d7, 5 d10, 2 d14, 4 d21, and 3 d28 for BioPin implants).

Tissue Processing, Histology, and in situ Hybridization
Tibiae were dissected and processed as described (Colnot et al., 2003). Ti alloy and 303 SS implants were gently removed after decalcification and prior to dehydration, while BioPin implants dissolved during processing. Tissue sections were prepared for histology with Safranin-O Fast green (SO/FG) and Trichrome (TC), and for histochemistry with tartrate-resistant acid phosphatase (TRAP) and alkaline phosphatase (AP). Adjacent sections were subjected to in situ hybridization analyses with collagen type I (Col1), collagen type (Col2)II, osteocalcin (Oc), osteopontin (Op), and runx2 (cbfa1) probes as previously described (Colnot et al., 2003).

Surface Characterization of Implants
Implant surface roughness was measured by optical interferometry (MicroXam^TM; ADE Phase-Shift, Tucson, AZ, USA) at a resolution of 0.05 nm (vertical) and 0.3 µm (horizontal), with S_a = arithmetic average value of vertical departures of the profile or surface from the mean line throughout the sampling length or area; S_t = maximum peak-to-valley height of the entire measurement trace; and S_q = root mean square of values of all points of the profile. Implant surfaces were analyzed by ESCA (electron spectroscopy for chemical analysis) at NESAC/BIO (Univ. of Washington, Surface Science Instruments S-probe spectrometer), with x-ray spot size ~ 800 µm and sampling depth ~ 50 Å.

	RESULTS

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Healing in Empty Implant Sites
We first determined how an implant site healed in the absence of any device, as a baseline for comparing healing around various implants. We examined the program of bone healing using histological stains to detect cartilage and bone. At all time-points (post-surgical d3-28), we never detected cartilage (data not shown) and found evidence of only new bone (Fig. 1, Trichrome panels). Thus, empty implant sites healed exclusively through intramembranous ossification.

View larger version (166K): [in this window] [in a new window]   Figure 1. Establishing the time-course and type of healing in the implant bed. A 1.0-mm drill hole was created in the proximal tibiae; samples were then collected at d5, d7, and d28, and stained with Trichrome (TC) and tartrate-resistant acid phosphatase (TRAP). (A–B) At d5, there is no evidence of new bone formation. The injury site is filled with densely packed fibroblasts, (B) while bone debris from the drilling process is engulfed by TRAP-positive osteoclasts. (C–D) By d7, new bone is first detected and undergoes rapid remodeling. (E–F) By d28, woven bone has been replaced by cortical bone fused with the old cortex. Scale bar = 100 microns. (See color version of this Fig. in the online APPENDIX.)

Histological analyses indicated that new bone formed in the empty implant site by d7. To pinpoint the actual onset of osteogenesis, rather than its histological manifestation, we conducted molecular analyses at earlier time-points. We examined expression of 3 genes: Collagen type I (Col I), Osteocalcin (Oc), and Osteopontin (Op). On d3, we identified cells in the injury site that expressed Col I (Fig. 2A, arrow), suggesting initiation of an osteogenic program. The lack of Oc and Op expression in adjacent sections indicated that cells had not yet differentiated into osteoblasts (Figs. 2B, 2C). The progressive increase in Col1, Oc, and Op expression from d5 to d7 indicated that cells initiated differentiation into osteoblasts during this window (Figs. 2D–2I). These analyses provided a molecular map illustrating initiation of osteoblast differentiation and bone formation in an empty site.

View larger version (132K): [in this window] [in a new window]   Figure 2. Molecular analyses of healing in the implant bed. Localization of mRNA for collagen type I (Col 1), Osteocalcin (Oc), and Osteopontin (Op) by in situ hybridization on sections adjacent to those shown in Fig. 1. (A–C) By d3, low expression levels for Col 1 are found in the marrow cavity (arrow), while Oc and Op are not expressed. (D–F) By d5, Col 1 is up-regulated in and around the defect within the marrow cavity. Oc mRNA is still not detected. Op-expressing cells are localized near the cortical defect (asterisk). (G–I) By d7, strong expression of all three markers is detected in the defect and in the marrow space underlying the defect. Scale bar = 200 microns. (See color version of this Fig. in the online APPENDIX.)

View larger version (72K): [in this window] [in a new window]   Figure 3. Healing at the bone-implant interface. Schematic representations of Ti-6Al-4V alloy and BioPin implants, and photomicrographs of these implants after placement (below). At post-surgical d5, longitudinal sections through (A) the Ti-6Al-4V implant and the (B,C) BioPin implant show evidence (by Trichrome staining) of new bone formation (arrowheads). (D–F) Bone formation around the implants was increased at d7. (G–I) Bone encasing the implants underwent remodeling until only a thin shell remained in the bone marrow cavity. A schematic drawing representing a direct interface and a minimal gap interface around an implant. Areas in the black box and the red box are illustrated on the tissue sections stained with Safranin-O Fast green (SO/FG) (J–M) and Trichrome (TC) (N–O). (J) On d3, the direct bone-implant interface was not populated by cells, whereas in (K), the gap interface was filled with a fibrous hematoma (bracket). (L) By d28, new bone was juxtaposed to the implant (arrowheads indicate stained nuclei of new osteoblasts), while in the gap (M), a new bone interface has been created by the mineralization of the fibrous matrix (bracket), and stained nuclei distinguish new osteoblasts from old osteocytes. Scale bar in A and corresponding magnifications = 0.5 mm; C and other high-magnification images = 50 microns. (See color version of this Fig. in the online APPENDIX.)

Osteoblast Differentiation is Accelerated in the Presence of an Implant
Since new bone was detected earlier by histological analyses around implants vs. empty defects, we further analyzed early stages of healing. With implants present, Trichrome staining revealed a faint amount of Aniline-blue-positive matrix at d5 (Fig. 4A, arrowhead). Alkaline phosphatase activity indicated the onset of mineralization (Fig. 4B, arrows), and TRAP activity indicated matrix remodeling (Fig. 4C). Thus, relative to an empty site, implant presence resulted in accelerated differentiation of peri-implant cells into osteoblasts, and acceleration in the remodeling of new bone matrix. We confirmed this using in situ hybridization, noting that cells around the implant up-regulated the osteoblast-related transcription factors Runx2 and Op at d3 (Figs. 4D, 4E). Lack of Collagen type II (Col II) expression around the implant indicated that intra-membranous ossification was the mechanism of new bone formation at implant sites (Fig. 4F).

View larger version (147K): [in this window] [in a new window]   Figure 4. The presence of an implant accelerated osteoblast differentiation. Cellular and molecular analyses at the Ti-6Al-4V (A–C) and 303 SS (D–F) implant surfaces. (A) At post-surgical d5, Trichrome staining indicates the initiation of endosteal bone formation (arrowhead). (B) AP staining corresponds to the areas of new mineralization (arrow), (C) which are undergoing extensive osteoclastic remodeling. (D–F) In situ hybridization on d3 indicates that cells surrounding the implant up-regulate Runx2 and Op; these cells are osteo- rather than chondroprogenitors, because of the absence of Col II expression. Scale bar = 200 microns. (See color version of this Fig. in the online APPENDIX.)

	DISCUSSION

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Cortical Bone Defect Healing and Interfacial Healing in the Mouse Model vs. Larger Animal Models
This study identified similarities and differences between cortical defect and interfacial healing. Early stages of healing in both were characterized by the formation of a hematoma, recruitment of matrix-resorbing cells, and deposition of woven bone in the empty defect or surrounding the implant. Woven bone was remodeled into lamellar bone, bridging the two cortical bone ends of the empty defect, or leaving behind a thin interfacial layer of bone at the implant surface. Our data on cortical defect healing complement previous findings of defect healing in rodent and larger animal models (Pritchard, 1964; Schenk and Hunziker, 1994; Chiba et al., 2001; Uusitalo et al., 2001; Campbell et al., 2003), wherein small islands of cartilage were sometimes also observed on periosteal surfaces. Interfacial healing in our mouse model shared some key features with interfacial healing around implants in other animal models, such as rats (Nanci et al., 1994; Masuda et al., 1997), rabbits (Schenk and Hunziker, 1994), dogs (Hoshaw et al., 1994), sheep (Plenk and Zitter, 1996), monkeys (Watzak et al., 2005), and humans (Lazzara et al., 1999)—with intramembranous bone healing in empty defects and around implants.

Differences in Timing of Bone Formation around Implants and in Empty Holes
Analysis of our cellular and molecular data indicates that: (1) cells surrounding implants initiated differentiation into osteoblasts sooner than when no implant was at the site; and (2) the timing of osteoblast differentiation and new bone matrix deposition was equivalent among the three implant biomaterials. One explanation for this is that an implant provides a surface onto which osteoblasts can adhere and deposit a matrix that mineralizes, and that this surface was similar among our implants. The range of roughness values for our implants was narrow (S_a, 0.185–0.709 µm) and at the lower end of values for "smooth" (S_a, 0 to 0.4 µm) and "minimally rough" (S_a, 0.5 to 1.0 µm) implants (e.g., S_a ~ 0.46 for machined Brånemark implants). Also, our implants were not as rough as "moderately rough" (S_a, 0.5 to 1.0 µm) or "rough" (S_a, > 2.0 µm) implants, e.g., S_a ~ 0.91 µm for "OsseoSpeed" implants; and S_a ~ 1.6 µm for SLA implants (Albrektsson and Wennerberg, 2004; Ellingsen et al., 2004; Sul et al., 2005). While a recent review (Shalabi et al., 2006) reported "a positive effect on the bone response...from R_a/S_a of ~ 0.5 µm up to ~8.5 µm", our narrow S_a range was at the lower end of that range. Moreover, our ESCA data showed comparable values of carbon on surfaces of all implants. (Commercial Ti implants can also have high carbon levels on their surfaces; Wieland et al., 2000; Massaro et al., 2002.) Ultimately, quantitative analyses would complement our qualitative spatial and temporal gene expressions. For example, our in situ data from mice are consistent with Ogawa and Nishimura’s (2006) RT-PCR data from tissues near Ti implants and at osteotomy sites in rat tibiae, which showed a 1.5- to two-fold up-regulation of collagen I, osteopontin, and osteocalcin expression (but not Runx2 and Bmp2) at 3 and 7 days after surgery.

Is Osseointegration Equivalent to Fracture Healing?
Concerning the oft-claimed analogy between fracture healing and interfacial healing (Brånemark et al., 1977; Brånemark, 1985; Pilliar and Simmons, 2002; Schatzker, 2002), both begin with a breach in an intact skeletal element, an immune response, neovascularization, and recruitment of skeletal progenitor cells. However, in a typical fracture, some skeletal progenitor cells differentiate into chondrocytes, while others differentiate into osteoblasts, followed by endochondral ossification. Around an implant, all skeletal progenitor cells differentiate directly into osteoblasts, followed by intramembranous ossification. Perhaps the key difference between fracture healing and interfacial healing is that the latter involves cellular and molecular responses that may be influenced by biomaterial surface texture, chemical composition, and implant biomechanics. In that context, advantages of the mouse model include its ability to allow for detailed molecular analyses, and the use of mouse mutants for the study of bone healing.

	ACKNOWLEDGMENTS

 
This study is supported by: an NIH/NIBIB grant (R01 EB 000504) to J.B., J.H., A.N., and J.C.; by an NIH/NIA grant (R01AG23218-01, Zena Werb); by an NIH/NIDCR grant (R03 DE16701), and by the Musculoskeletal Transplant Foundation (C.C.) and FA9550-04-1-0075 for J.H. We thank S. Hadwin (IMTEC Corp., Ardmore, OK, USA) for BioPins, Dr. G.W. Marshall (UCSF, San Francisco, CA, USA) for Ti alloy implants, L. Gamble (NESAC/BIO, University of Washington, Seattle, WA, USA) for ESCA, and Dr. A. Wennerberg (University of Göteborg, Sweden) for roughness data.

	FOOTNOTES

 
A supplemental appendix to this article is published electronically only at http://www.dentalresearch.org.

Received for publication October 11, 2006. Revision received April 21, 2007. Accepted for publication May 4, 2007.

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Journal of Dental Research, Vol. 86, No. 9, 862-867 (2007)
DOI: 10.1177/154405910708600911


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This Article Abstract Figures Only Full Text (PDF) Data Supplement Alert me when this article is cited Alert me if a correction is posted Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Add to Saved Citations Download to citation manager Request Permissions Request Reprints Add to My Marked Citations Citing Articles Citing Articles via Google Scholar Citing Articles via Scopus Google Scholar Articles by Colnot, C. Articles by Helms, J.A. Search for Related Content PubMed PubMed Citation Articles by Colnot, C. Articles by Helms, J.A. Pubmed/NCBI databases Compound via MeSH Substance via MeSH Hazardous Substances DB LACTIC ACID TITANIUM Social Bookmarking                         What's this?