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. Author manuscript; available in PMC: 2019 Jun 1.
Published in final edited form as: Clin Oral Implants Res. 2018 May 16;29(6):592–602. doi: 10.1111/clr.13257

Injectable, compression-resistant polymer/ceramic composite bone grafts promote lateral ridge augmentation without protective mesh in a canine model

Anne D Talley 1, Lauren A Boller 2, Kerem N Kalpakci 3, Daniel A Shimko 3, David L Cochran 4, Scott A Guelcher 1,2,5,*
PMCID: PMC6152823  NIHMSID: NIHMS960471  PMID: 30240051

Abstract

Objective

The objective of this study was to test the hypothesis that a compression-resistant bone graft augmented with recombinant human morphogenetic protein-2 (rhBMP-2) will promote lateral ridge augmentation without the use of protective mesh in a canine model.

Materials & Methods

Compression-resistant (CR) bone grafts were evaluated in a canine model of lateral ridge augmentation. Bilateral, right trapezoidal prism-shaped defects (13–14 mm long × 8–9 mm wide × 3–4 mm deep at the base) in 13 hounds (2 defects per hound) were treated with one of four groups: (1) absorbable collagen sponge + 400 µg rhBMP-2/ml (ACS, clinical control) protected by titanium mesh, (2) CR without rhBMP-2 (CR, negative control), (3) CR + 200 µg rhBMP-2 (CR-L), or (4) CR + 400 µg rhBMP-2 (CR-H). All animals were euthanized after 16 weeks. Ridge height and width and new bone formation were assessed by µCT, histology, and histomorphometry. The release kinetics of rhBMP-2 from CR bone grafts in vitro and in vivo in a femoral condyle defect model in rabbits were also evaluated.

Results

All four bone grafts promoted new bone formation (11– 31.6 volume%) in the lateral ridge defects. For CR grafts, ridge height and width increased in a dose-responsive manner with increasing rhBMP-2 concentration. Ridge height and width measured for CR-H without the use of protective mesh was comparable to that measured for ACS with a protective mesh.

Conclusions

At the same dose of rhBMP-2, an injectable, compression-resistant bone graft resulted in a comparable volume of new bone formation with the clinical control (ACS). These findings highlight the potential of compression-resistant bone grafts without the use of protective mesh for lateral ridge augmentation.

Keywords: Bone graft, bone morphogenetic protein, compression resistant, lateral ridge augmentation, canine model

Introduction

Mandibular bone defects due to craniomaxillofacial injuries can lead to significant morbidity if not properly repaired. Dental implants are often utilized to restore dentition following alveolar bone injury. However, adequate bone volume is necessary for successful implant dentistry (Rakhmatia et al. 2013). Current surgical techniques to increase bone volume at deficient defect sites prior to dental implant placement include the use of autografts, distraction osteogenesis, and guided bone regeneration. Autografts are considered the clinical gold standard, but have the disadvantage of causing donor site morbidity (generally the iliac crest (Kalk et al. 1996)). Furthermore, autografts are subject to resorption and at times fail to integrate, especially without site vascularization (Chim & Gosain 2009). As an alternative to autogenous bone, the local placement of bone grafts augmented with growth factors is a promising treatment option for enhancing the alveolar ridge in patients preceding dental implant surgery (Jung et al. 2008; Polimeni et al. 2010; Wikesjö et al. 2008). However, these common delivery systems rely upon the principle of guided bone regeneration and require space-maintaining protective polymeric or metal mesh to direct the growth of new bone (Liu & Kerns 2014). By acting as a barrier, the mesh excludes non-osteogenic tissue invasion from the defect site and maintains a suitable space for osteoblast infiltration and new bone formation. However, the mesh can lead to infection and wound failure (Herford et al. 2012; Ruskin et al. 2000). Additionally, the rigidity and stiffness of non-resorbable meshes can lead to an increased risk of mesh exposure (Buser et al. 1996; Ruskin et al. 2000). Furthermore, these meshes often require additional stabilization hardware and must be removed with a second surgical procedure (Liu & Kerns, 2014). Thus, there is a clinical need to develop new carriers for local delivery of growth factors in alveolar ridge augmentation which do not depend on mesh for space maintenance.

Bone morphogenetic proteins (BMPs) are a group of osteoinductive growth factors originally discovered by Urist (Urist 1965). BMPs delivered locally regulate the complex process of bone repair and restoration. Recombinant human bone morphogenetic protein-2 (rhBMP-2) is the most extensively investigated growth factor for bone regeneration and enhances new bone formation by stimulating recruitment of osteoprogenitor cells and osteoblast differentiation (Dimitriou et al. 2011). RhBMP-2 delivered via an absorbable collagen sponge (ACS) (INFUSE® bone graft, Medtronic) is FDA-approved for the treatment of posterior-lateral spinal fusions, fractures of the tibial mid-diaphysis, and extraction socket defects and sinus lift procedures.

Biodegradable lysine-derived poly(ester urethane) (PEUR) scaffolds have been extensively investigated for bone repair applications (Bonzani et al. 2007; Dumas et al. 2012). These materials breakdown by hydrolytic and cell-mediated oxidative degradation to non-toxic decomposition products (Hafeman et al. 2011; Zhang et al. 2002). PEUR scaffolds augmented with ceramic particles have shown improved mechanical properties and osteoconductivity (McEnery et al., 2016). In a recent study, PEUR/ceramic (CM) composite bone grafts exhibited compression-resistant mechanical properties and supported new bone formation in femoral condyle plug defects in sheep (Talley et al. 2016).

In a previous study, an injectable compression-resistant (CR) composite bone graft augmented with rhBMP-2 promoted new bone formation and maintained space in a canine mandibular saddle defect model without the use of protective mesh (Talley et al. 2016). However, the ability of the CR graft to promote bone healing comparable to that achieved with the use of mesh has not been investigated in a regulatory model of craniofacial bone regeneration. In this study, we hypothesized that the CR graft augmented with rhBMP-2 will maintain space and promote new bone formation in a dose-responsive manner without the use of protective mesh. We compared CR carriers augmented with varying concentrations (0, 200, or 400 µg /ml) of rhBMP-2 to the ACS carrier augmented with 400 µg/ml rhBMP-2 and protected by a titanium mesh in a canine model of lateral ridge augmentation. Ridge height and width and new bone formation were assessed by µCT, histology, and histomorphometry at 16 weeks post-grafting.

Materials and Methods

Materials

Lysine-triisocyanate prepolymer (LTI-PEG, 21.7% isocyanate (NCO)) was received from Medtronic Spinal & Biologics (Memphis, TN). Glycerol, stannous octoate, ε-caprolactone, and APTES were purchased from Sigma-Aldrich (St. Louis, MO). Glycolide and DL-lactide were purchased from Polysciences (Warrington, PA). Triethylene diamine and dipropylene glycol were purchased from Sigma Aldrich and mixed to obtain a 10% (w/w) solution of triethylene diamine in dry dipropylene glycol. MASTERGRAFT® Mini Granules (CM) received from Medtronic Spinal (Memphis, TN) were ground and sieved to 100–500 µm. Titanium mesh (ARTISAN™ Space Maintenance System) and rhBMP-2 were provided by Medtronic Spinal (Memphis, TN).

Synthesis of polyester triol

The polyester triol was synthesized as described previously (Guelcher et al. 2007; Guelcher et al. 2006). Glycerol was mixed with ε-caprolactone, glycolide, and DL-lactide monomers under argon at 140°C for 40 h to yield a viscous fluid that was washed with hexane (3×) and vacuum-dried at 80°C for 48 h. The backbone of the polyester consisted of 70% ε-caprolactone, 20% glycolide, and 10% DL-lactide, and the molecular weight was 450 g mol−1.

Fabrication of compression-resistant (CR) bone grafts

CM granules were ground and sieved to 100–500 µm to improve the handling properties of the injectable composite (Talley et al. 2016). The components were mixed using a two-step method. First, the polyester triol, CM particles (45 wt%), and triethylene diamine (1.1 pphp) were added to a 10-mL cup and mixed by hand for 30 s. The LTI-PEG and lyophilized rhBMP-2 were then added and mixed by hand for 60 s. The index (ratio of isocyanate:hydroxide equivalents × 100) was set at 115.

In vitro rhBMP-2 release

For in vitro release measurements, three replicate rhBMP-2-loaded porous scaffolds (~ 50 mg) were placed in a closed vial containing 1 mL of release media (α-MEM with 1% bovine serum albumin) in an incubator at 37°C. The composites contained 100 µg rhBMP-2/mL scaffold to mimic the in vivo study. Bovine serum albumin was included to minimize adsorption of rhBMP-2 to the sample vial (Li et al., 2010). Media was collected and replaced every day to minimize degradation of the rhBMP-2. The media samples were pooled as indicated on the release plot and concentration was measured for triplicate samples using a Human BMP-2 Quantikine ELISA kit (R&D systems).

rhBMP-2 radiolabeling

To determine in vivo release kinetics, a fraction of the rhBMP-2 was radiolabeled as described previously (Kempen et al. 2008). Briefly, 100 µl of a 1.43 mg/ml rhBMP-2 solution, 20 µl of a 0.1 M NaOH solution, and 2 mCi of Na125I were combined in a pre-coated iodination tube (Pierce, Rockford, IL). After incubation at room temperature for 15 min, the solution was dialyzed (10kDa molecular weight cutoff (MWCO) Slide-A-Lyzer®, Pierce) for 24 h with three times media change against an aqueous rhBMP-2 buffer containing 5mM glutamic acid, 2.5 wt% glycine, 0.5 wt% sucrose, and 0.01 wt% Tween 80 (pH 4.5). The dialyzed fraction was concentrated with a Vivaspin® centrifugal concentrator (10 kDa MWCO, Viva Products, Inc., Littleton, MA) and trichloroacetic acid precipitation indicated 99% precipitable counts. The labeled rhBMP-2 was mixed with unlabeled rhBMP-2 to obtain a hot:cold ratio of 1:8. The solution was aliquoted and lyophilized in appropriate doses.

Evaluation of in vivo rhBMP-2 release in a rabbit femoral plug defect model

In vivo release kinetics of radio-labeled rhBMP-2 was evaluated in rabbit femoral condyle plug defect model. Four healthy, skeletally mature, male New Zealand white rabbits between (~3.5 kg, 6 months of age) were used in this study. The animals were kept in separate cages and received food twice daily and had ad libitum access to water. All surgical and care procedures were carried out under aseptic conditions according to the approved IACUC protocol at Vanderbilt University. The study design is listed in Table 1. The individual components of the CR grafts were gamma-irradiated using a dose of 25 kGY. Prior to surgery, rabbits were pre-medicated subcutaneously with the analgesic buprenorphine (0.02mg/kg) and the antibiotic cefazolin (10mg/kg). General anesthesia was induced through intramuscular injection of ketamine hydrochloride/xylazine (35mg/kg+ 5mg/kg) and maintained with isoflurane (approximately 0.5–4%) in oxygen. Before incision, hair was shaved from the surgical sites and the skin was disinfected with 4% chlorohexidine. Bilateral critical-sized defects of 5 mm by 6–8 mm in depth were drilled in the lateral femoral condyles of each rabbit. The rabbits were randomly divided into two groups of two animals each. The groups were treated with CR (n=4) or ACS (n=4) grafts augmented with rhBMP-2. The ACS carrier was chosen as the clinical control due to its widespread use in bone repair and FDA approval. For the CR grafts, the reactive paste was injected into the defect site and allowed to cure for 10 min prior to soft tissue closure. For the ACS carrier, the lyophilized growth factor was reconstituted in sterile water and injected onto the collagen sponge. The sponge was allowed to adsorb the rhBMP-2 solution for at least 15 min prior to implantation. A dose of 100 µg/ml rhBMP-2 (based on defect volume) was investigated, considering a previous study reporting that a dose of 80 µg/ml rhBMP-2 with CR grafts was shown to enhance bone formation and bridging in rabbit calvarial defects (Dumas et al. 2012). The rabbits were given antibiotic cefazolin (10kg/mg) for 2 days post-surgery and the analgesic buprenorphine (.05mg/kg) as needed. In vivo retention of rhBMP-2 was measured on days 1, 2, 3, 7, 14, 21, and 28 using a scintillation probe collimated with a hollow tube wrapped in leaded tape (Model 44-3 scintillator, Ludlum Measurement Inc., Sweetwater, TX) and connected to a digital scaler (Model 2241-3 scaler, Ludlum Measurements Inc.). The rabbits were restrained with ketamine (20mg/kg) via intramuscular injection and radioactive counts were obtained over 1 min intervals (repeated twice at each time point). The 125I counts were normalized to values at initial time and corrected for radioactive decay. The rabbits were euthanized after eight weeks with an overdose of pentobarbital sodium (70mg/kg).

Table 1.

In vivo release of rhBMP-2 study design.

Treatment Group Particle
diameter
µm		 rhBMP-2
µg cm−3 defect
volume		 n
8 weeks
ACS N/A 100 4
CR 100 – 500 100 4
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Evaluation of CR bone grafts in a canine mandibular lateral ridge augmentation model

13 skeletally mature female hounds (~25 kg, not less than 11 months of age) were used in this study. The animals were kept in separate cages and received food twice daily and had ad libitum access to water. All surgical and care procedures were carried out under aseptic conditions according to the approved IACUC protocol at IBEX Preclinical Research. Four treatment groups were used in this study and are listed in Table 2. The canines were randomly assigned an animal ID number to randomize treatment groups per animal. The individual components of the CR grafts were gamma-irradiated using a dose of 25 kGY. Immediately prior to surgery, animals were given antibiotic ampicillin (5mg/kg). General anesthesia was induced by an intravenous injection of ketamine (10 mg/kg) and diazepam (0.3 mg/kg) followed by placement of an endotracheal tube. General anesthesia was maintained with isoflurane (approximately 0.5–4%) in oxygen. A fentanyl patch (50 µg/h) was applied to the skin to provide analgesia for up to 72 h. In the first surgery, the dogs underwent bilateral extraction of the four mandibular premolars and removal of a portion of the buccal plate to simulate dental loss in humans resulting from chronic disease (Figure 1A). Following the extraction procedure, antibiotic enrofloxacin (5mg/kg) was administered daily for 7 days via intramuscular injection. After at least a two-month healing period, the extraction sites were accessed and debrided. Right trapezoidal prism-shaped defects were created bilaterally in each mandible (2 defects per dog) measuring approximately 13–14 mm mesiodistally, 8–9 mm apico-coronally, and 3–4 mm bucco-lingually (Figure 1B). General anesthesia remained the same for the second procedure. For the ACS samples, lyophilized rhBMP-2 was reconstituted in sterile water, injected onto the collagen sponge, and allowed to sit for 15 min. The sponge was then placed in the defect and covered with the titanium mesh (ARTISAN™ Space Maintenance System, Medtronic) which was screwed into the mandible (Figure 1C). The individual components of the CR bone grafts were mixed with lyophilized rhBMP-2 at a Low (200 µg/mL) or a High (400 µg/mL) dose (n=6 per group) and injected into the defect site (Figure 1D). The CR grafts cured for approximately 10 min prior to soft tissue closure. Following the lateral ridge augmentation grafting procedure, antibiotic amoxicillin clavulinate (20mg/kg) was orally administered twice a day for 7 days. The dogs were euthanized with an intravenous injection of barbiturates (1mL/4.5kg body weight) after sixteen weeks. The mandibles were extracted and fixed in 10% formalin for two weeks prior to processing for histology.

Table 2.

Treatment groups evaluated in the canine lateral ridge defects

Treatment Group Particle diameter
µm		 rhBMP-2
µg cm−3 defect
volume		 n
16 weeks
ACS N/A 400 6
CR 100 – 500 0 6
CR-L 100 – 500 200 6
CR-H 100 – 500 400 6
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Figure 1.

Figure 1

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Surgical photographs. (A) Tooth extraction and removal of the buccal plate. (B) Second surgery to create the lateral ridge defect. (C) Placement of the collagen sponge and titanium mesh in the defect site. (D) Injection of CR grafts in the defect site.

Analysis of bone morphometry by Micro-Computed Tomography (µCT)

A µCT50 (SCANO Medical, Basserdorf Switzerland) was used to acquire scans of the extracted mandibles in formalin at 70 kVp energy, 200 µA source current, 1000 projections per rotation, 800 ms integration time, and an isotropic voxel size of 24.2 µm. Axial images were reconstituted using manufacturer-provided software. Attenuation values were converted to tissue mineral density through calibration with hydroxyapatite (HA) phantoms with densities of 0 to 780 mg HA cm−3 (calibrations checked weekly). Using the coronal boundary of the defect for alignment, the reconstructed image stack was re-oriented so that the apico-coronal direction was parallel to the z-axis. 3D reconstructions of the defect were generated, from which the total volume of new bone within the defect region was calculated to quantitatively assess ridge augmentation. The region of interest was contoured by first determining the midpoint of the defect, from which 50 slices on either side of the midpoint were contoured to create a 100-slice middle defect region. This region was then segmented from mastergraft and soft tissue using a threshold range of 300–400 mg HA/ccm, sigma 0.2, and support of 1. Bone volume fraction (BV/TV) was then calculated for this region using Scanco evaluation software.

Histology

After fixation in formalin, the explanted mandibles were dehydrated in a graded series of ethanol and embedded in poly(methyl methacrylate). Using an Exakt band saw, sections were cut from each block in the center of the defect (bucco-lingually) using the µCT images as reference. The sections were then ground and polished to <100 µm using an Exakt grinding system, and stained with Sanderson’s rapid bone stain. New bone stained red, residual CM black, and infiltrating cells blue.

Histomorphometry

For quantitative analysis, new bone and residual CM particles were measured from the histological sections. Ridge width was measured as a function of height at the mid-section of the defect. A right-angle trapezoid was drawn on the ridge using Metamorph software (Version 7.0.1, Waltham, MA) (Figure 5C). The top and bottom edges of the defect represent the 4 corners of the trapezoid. The trapezoid was divided into 4 right-angle trapezoidal areas of interest starting at the coronal base of the defect. Each area of interest measured 2 mm in height by the width of the defect. At each region, the ridge width was normalized to the width of the trapezoid base. Metamorph was utilized to color threshold the residual CM (black) and new bone (red) and quantify the area of each phase within the area of interest of each section. The diameter of the residual CM particles within the defect were measured to assess CM resorption.

Figure 5.

Figure 5

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Histological analysis of new bone formation and space maintenance. (A) Low- (scale bar represents 1 mm) and (B) high- (10×) magnification images of histological sections show new bone formation (red), infiltrating cells (blue), and residual CM (black) particles. New lamellar bone is indicated with yellow arrows, and osteoid is indicated with blue arrows. No residual polymer was observed. (C) Trapezoidal region used for histomorphometric analysis. (D) Normalized ridge width within the defect area as a function of the height above the baseline. (E) Histomorphometric analysis of new bone and residual CM within the defect area.

Statistical analysis

The µCT and histomorphometric parameters were compared between the four experimental groups using IBM SPSS Statistics Version 25 (IBM®, Armonk NY). A linear mixed model was used with rhBMP-2 dose and combination of doses within animal (due to the bilateral defect model) set as fixed effects and canine subject set as a random effect. Summary statistics including mean, standard deviation median values, and t-values were recorded for each defect. Statistical significance was considered for p < 0.05.

Results

rhBMP-2 release kinetics

To evaluate the in vitro release of rhBMP-2, lyophilized protein was incorporated into CR composites or ACS and the release kinetics measured by ELISA (Figure 2A). After one week, all of the growth factor was released from the ACS samples. In contrast, CR grafts released less than 50% release of the total amount of rhBMP-2 within the first week. By four weeks, approximately 75% of rhBMP-2 was released from the CR grafts.

Figure 2.

Figure 2

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Cumulative rhBMP-2 release kinetics from CR and ACS grafts. (A) In vitro release measured by ELISA. (B) In vivo release measured by 125I radiolabeling. (C) Fitting of in vitro and in vivo release kinetics data to the Weibull model.

In vivo release of rhBMP-2 was measured in femoral plug defects in rabbits (Figure 2B). The surgical procedures and subsequent healing period were uneventful. 8/8 samples obtained (n=4, per group) were used for analysis. This study provided data for the local release of rhBMP-2 from the defect site but did not investigate the systemic biodistribution of the growth factor. Similar to the in vitro results, the release from the ACS samples was complete after the first week. In the CR samples, there was a burst release of approximately 25% growth factor within the first 3 days and 50% within the first week. No growth factor was measured within the defect site after 4 weeks. The rhBMP-2 release was slightly faster for CR composites in vivo compared to in vitro conditions.

µCT analysis

Representative µCT images of lateral ridge samples after 16 weeks (Figure 3) show differences in healing between the treatment groups. More new bone is evident in all three planes (coronal, sagittal, and transverse) within the CR-H and ACS samples compared to the CR and CR-L treatment groups. Additionally, the width of the ridge was best maintained within the CR-H and ACS samples, and these grafts showed the most uniform healing within the defect area. Residual CM (bright white particles) is present in the CR-treated samples. The titanium mesh can be seen within the images of the ACS samples (bright white line) surrounding the defect space. After 16 weeks, BV/TV within the defects were comparable between the ACS (28.9%) and CR-H (31.6%) samples (Figure 4). Mean and median values for all groups are reported in Table 3. CR-H samples displayed greater BV/TV when compared with the CR group.

Figure 3.

Figure 3

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Representative µCT images of the defects within each treatment group after 16 weeks of healing showing the coronal, sagittal, and transverse views of the ridge. The residual CM shows up as bright white particles within the defect area for the CR treated groups. The titanium mesh surrounds the defect within the ACS group.

Figure 4.

Figure 4

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Morphometric parameter, bone volume/ total volume (BV/TV) within the defects of each treatment group after 16 weeks.

Table 3.

Volumetric measurements of µCT (n=6). BV/TV, bone volume/total volume.

BV/TV
CR Mean ± SD 0.17 ± 0.12
Median 0.11
t
CR-L Mean ± SD 0.24 ± 0.17
Median 0.16
t 0.773
CR-H Mean ± SD 0.27 ± 0.15
Median 0.28
t 1.17
ACS Mean ± SD 0.25 ± 0.09
Median 0.27
t 0.96
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Histology

The surgical procedures and subsequent healing process were mostly uneventful. In one of the canines, a technical failure due to graft displacement occurred and no new bone formation was observed. Thus, an additional canine was required to preserver the power of the study. 24/26 samples obtained were used for analysis. Figure 5A shows representative low-magnification images of all groups. In sections stained with Sanderson’s rapid bone stain, new bone appears red, cells are blue, and residual CM particles appear black. Similar to the µCT results, the histological sections reveal that the ACS and CR-H groups promoted the highest amount of new bone within the defect area. Less new bone was present in the CR or the CR-L groups. In the ACS-treated defects, the new bone did not bridge the entire gap between the defect wall and the titanium mesh, and the empty gap was filled with cells and fibrous tissue. There was minimal fibrous tissue within the CR-H group while CR-L treated defects contained some fibrous tissue in the defect adjacent to the new bone. All four groups showed no evidence of excessive inflammation. Residual CM particles (black) are evident in all of the CR treatment groups. The high-magnification images (Figure 5B) show both new woven bone and lamellar bone formation in all groups. In the CR groups, some of the CM particles are incorporated within the new bone structure and show appositional bone growth adjacent to the particles. Ridge width as a function of ridge height was measured for each image at 0, 2, 4, and 6 mm above the baseline of the defect (assuming a trapezoidal shape for each section) and plotted in Figure 5D. Mean and median values for all groups are reported in Table 4. In all groups, there was a trend of decreasing ridge width with increasing height above the defect baseline due to the anatomic contour of the ridge. The ridge width measured for the CR group at 6 mm was significantly lower than the CR-L (p=0.035), ACS (p <0.001), and CR-H (p = 0.0063) groups at 6 mm above the baseline.

Table 4.

Normalized ridge width measurements at 16 weeks (n=6). 0, baseline of the defect; 2,4,6, mm from the baseline.

0 2 4 6
CR Mean ± SD 1.00 ± 0.00 0.60 ± 0.39 0.51 ± 0.45 0.17 ± 0.24
Median 1.00 0.57 0.34 0.003
t
CR-L Mean ± SD 1.00 ± 0.00 0.76 ± 0.22 0.80 ± 0.40 0.49 ± 0.20*
Median 1.00 0.73 0.58 0.47
t 0.00 1.18 1.19 2.55
CR-H Mean ± SD 1.00 ± 0.00 0.95 ± 0.17 0.82 ± 0.09 0.58 ± 0.09*
Median 1.00 1.03 0.81 0.57
t 0.00 1.26 1.39 3.64
ACS Mean ± SD 1.00 ± 0.00 0.88 ± 0.10 0.82 ± 0.15* 0.65 ± 0.12*
Median 1.00 0.92 0.85 0.70
t 0.00 1.47 2.33 5.82
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*

Significantly different to CR group (p<0.05)

Histomorphometry

Area percent of new bone and residual CM within the defect area are displayed in Figure 5E. Mean and median values for all groups are reported in Table 5. The CR-H group exhibited the highest amount of new bone (49.6 %) within the defect area, although there were no significant differences between any of the groups. The area% of residual CM particles ranged from 5– 7 area%. Differences in area% CM particles at 16 weeks were not significant between any of the CR groups, but the CR-H samples had the lowest residual CM (5.0%). The diameters of measured residual CM particles ranged from 30–305 µm.

Table 5.

Histomorphometric measurements of %area new bone and %area residual ceramic particles at 16 weeks (n=6).

Bone CM
CR Mean ± SD 49.12 ± 12.81 5.73 ± 2.62
Median 53.38 5.01
t
CR-L Mean ± SD 46.57 ± 16.21 7.01 ± 2.52
Median 44.30 6.66
t −1.76 0.57
CR-H Mean ± SD 49.58 ± 18.42 4.98 ± 4.29
Median 47.43 4.67
t −0.63 1.72
ACS Mean ± SD 45.30 ± 8.90
Median 48.07
t −2.05
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Discussion

Prior to placement of dental implants, ridge augmentation is frequently performed to restore adequate bone volume (Petrungaro & Amar 2005). Guided bone regeneration utilizing rhBMP-2 carriers to promote new bone growth and support implant placement is a predictable and successful technique for ridge augmentation. However, currently available carriers must be delivered in conjunction with polymeric or metal space-maintaining devices. The most frequent complications from these devices include seroma and wound dehiscence leading to subsequent infection or mesh exposure (Jovanovic et al. 2007; Ruskin et al. 2000). Mesh exposure has been shown to have a detrimental effect on new bone formation (Lang et al. 1994; Machtei 2001) and resulted in 20% less new bone formation when compared with a completely submerged mesh (Souza et al., 2010). CR grafts augmented with rhBMP-2 are an advantageous alternative to guided bone regeneration in that they exhibit the compression-resistant mechanical properties of bone, support new bone formation, and maintain space without the use of non-degradable protective devices (Talley et al. 2016). We hypothesized that injectable and settable CR bone grafts augmented with rhBMP-2 would maintain space and support new bone formation comparable to the ACS carrier with titanium mesh (clinical control). CR grafts supported sustained release of rhBMP-2 for 30 days compared to 10 days for the ACS carrier, and CR augmented with 400 µg/ml rhBMP-2 maintained the height and the width of the canine alveolar ridge comparable to the ACS carrier with titanium mesh treatment at the same dose.

The CR composites tested in the lateral ridge defects contained 45 wt% CM particles (MasterGraft®, Medtronic Spine & Biologics). This FDA-approved ceramic was chosen based on its compression-resistant and osteoconductive properties observed when implanted in porcine mandibular continuity defects (Herford et al., 2012), sheep femoral plug defects (Talley et al. 2016), and posterolateral fusion in non-human primates (Akamaru et al. 2003). Additionally, block-type biphasic calcium phosphates supported local delivery of rhBMP-2 and new bone growth in a rabbit calvarial defect (Kim et al. 2012). In the present study, we evaluated the effects of rhBMP-2 dose on ridge width as a function of ridge height in a canine model of lateral ridge augmentation. Images of 2D µCT (Figure 3) and histological (Figure 5) sections showed enhanced new bone formation and space maintenance for the CR-H and ACS groups compared to the lower dose groups. Importantly, histomorphometric analysis revealed no significant differences in ridge width as a function of height above the baseline of the defect between the CR-H and ACS groups (Figure 5D–E). In a previous study, rhBMP-2 delivered from an ACS carrier combined with a ceramic osteoconductive bulking agent required a higher effect dose of rhBMP-2 (2.0 mg/mL) compared to delivery from the ACS carrier alone (1.5 mg/mL) in a mandibular continuity defect in rhesus macaques (Herford et al. 2012). However, in the present study, no significant differences in ridge width were observed between the CR-H and ACS groups at the same rhBMP-2 concentration (400 µg/mL). The more sustained release of rhBMP-2 from the CR-H carrier (30 days, Figure 2B) compared to the ACS carrier (10 days) is conjectured to enhance bone healing, as evidenced by previous studies reporting that sustained release of rhBMP-2 increased new bone formation compared to a burst release (Brown et al. 2011; Kolambkar et al. 2011).

While sustained release of rhBMP-2 from lysine-derived polyurethane grafts has been shown to promote bone healing in preclinical models of bone regeneration (Brown et al. 2011; Dumas et al. 2012; Li et al. 2009; Talley et al. 2016), the mechanism of rhBMP-2 release from these biomaterials, which undergo both hydrolytic and cell-mediated oxidative degradation (Hafeman et al. 2011; Talley et al. 2016), has not been previously investigated in vivo. Release of rhBMP-2 from CR composites followed a Fickian diffusion mechanism in vitro, which is consistent with previous studies (Li et al. 2009). However, under in vivo conditions, CR carriers released rhBMP-2 by a combined mechanism characterized by both Fickian diffusion and Case II transport (i.e., polymer erosion-controlled release), which is associated with b values in the range 0.75 < b < 1 (Papadopoulou et al. 2006) (Figure 2C). This difference between in vitro and in vivo in release kinetics is attributed to the accelerated in vivo resorption of the PEUR due to cell-mediated oxidative degradation, which has been previously described for lysine-derived polyurethanes (Hafeman et al. 2011; McEnery et al. 2016). The kinetics of rhBMP-2 release from the ACS carrier under in vivo conditions was comparable to that reported in previous studies (Seeherman & Wozney, 2005; Uludag et al. 2001) but faster than the CR carriers. Release of rhBMP-2 from the ACS carrier also followed a combined mechanism in vivo.

The rhBMP-2 dose did not affect the rate of resorption of the PEUR or CM component at 16 weeks. Images of µCT and histological sections revealed evidence of residual CM particles within the grafts, however, the diameters of residual CM particles decreased in size by 16 weeks. The PEUR was completely degraded at 16 weeks. CM particles in the CR-L and –H groups showed evidence of resorption and appositional bone growth, while those in the CR group were isolated within fibrous tissue (Figure 5B). These findings are consistent with a previous study investigating CR grafts in a canine saddle defect model which showed similar patterns of CM and PEUR resorption at 16 weeks (Talley et al. 2016). A limitation of this study was the single time point, since earlier time points may have provided information pertaining to the relative rates of CM and PEUR resorption and bone formation (Dumas et al. 2014). To our knowledge, CR grafts are the first injectable and settable carriers for rhBMP-2 that effectively maintain space and promote new bone formation in a canine lateral ridge augmentation model without protective mesh. In future studies, these compression-resistant carriers will be tested in a non-human primate model to determine whether CR grafts augmented with rhBMP-2 concentrations used to treat human patients (1.5 mg/ml) (Govender et al. 2002) promote lateral ridge augmentation.

Conclusions

This study was designed to test the hypothesis that injectable, settable, and compression-resistant bone grafts augmented with rhBMP-2 maintain space and promote bone regeneration comparable to the ACS carrier protected by titanium mesh. CR grafts supported sustained release of rhBMP-2 for up to 30 days, compared to 10 days for the ACS carrier. At concentrations of 400 µg/ml, local delivery of rhBMP-2 from CR grafts promoted lateral ridge augmentation comparable to the ACS carrier protected by titanium mesh. These observations highlight the feasibility of injectable, settable, and compression-resistant grafts as a new mesh-free approach to lateral ridge augmentation.

Acknowledgments

This material is based in part upon work supported by the National Science Foundation under Grant Number 0847711 (CAREER award to S.A.G.). Any opinions, findings, and conclusions or recommendations expressed in this material are those of the author(s) and do not necessarily reflect the views of the National Science Foundation. This work was supported by the Army, Navy, NIH, Air Force, VA and Health Affairs to support the AFIRM II effort, under Award No. W81XWH-14-2-0004. The U.S. Army Medical Research Acquisition Activity, 820 Chandler Street, Fort Detrick MD 21702-5014 is the awarding and administering acquisition office. Opinions, interpretations, conclusions and recommendations are those of the author and are not necessarily endorsed by the Department of Defense. Anne Talley acknowledges support from the Department of Education for a Graduate Assistance in Areas of National Need Fellowship under grant number P200A090323. Instrumentation for µCT analysis was purchased with funds from NIH grant S10RR027631.

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