Bone regeneration with BMP-2 delivered from ...

Biomaterials 34 (2013) 1644e1656

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Bone regeneration with BMP-2 delivered from keratose scaffolds Roche C. de Guzman a, Justin M. Saul b, Mary D. Ellenburg c, Michelle R. Merrill c, Heather B. Coan c, Thomas L. Smith a, Mark E. Van Dyke a, * a

Department of Orthopaedic Surgery, Wake Forest University Health Sciences, Winston-Salem, NC 27157, USA Department of Chemical and Paper Engineering, Miami University, Oxford, OH 45056, USA c Institute for Regenerative Medicine, Wake Forest University Health Sciences, Winston-Salem, NC 27157, USA b

a r t i c l e i n f o

a b s t r a c t

Article history: Received 22 October 2012 Accepted 1 November 2012 Available online 2 December 2012

InfuseÒ is used clinically to promote bone repair. Its efficacy is dependent on a crosslinked collagen carrier/scaffold system that has come under scrutiny due to an inability to control BMP-2 release, which may result in unwanted outcomes such as heterotopic ossification. In this study, keratose biomaterial was evaluated as a new carrier/scaffold. Keratose was mixed with BMP-2, fabricated into a scaffold, and implanted into a critical-size rat femoral defect. This construct showed bridging as early as 4 weeks and induced trabecular morphology characteristic of a remodeling hard fracture callus at 16 weeks. Compared to the normal cortical bone, the regenerated tissue had greater volume and mineral content but less density and ultimate shear stress values. Moreover, m-CT, biomechanics, FTIR-ATR spectroscopy, and polarized light microscopy data showed regeneration using keratose was similar to an Infuse control. However, unlike Infuse’s collagen carrier system, in vitro analysis showed that BMP-2 release correlated with keratose scaffold degradation. Surprisingly, treatment with keratose only led to deposition of more bone outgrowth than the untreated negative control at the 8-week time point. The application of keratose also demonstrated a notable reduction of adipose tissues within the gap. While not able to induce osteogenesis on its own, keratose may be the first biomaterial capable of suppressing adipose tissue formation, thereby indirectly enhancing bone regeneration. Ó 2012 Elsevier Ltd. All rights reserved.

Keywords: Keratin Keratose biomaterial BMP-2 delivery Critical-size defect Bone regeneration Tissue engineering

1. Introduction Segmental bone loss caused by high-energy trauma and disease represents an extreme medical scenario and is a major healthcare challenge [1]. Significant or critical-size gaps prevent the body from spontaneously-regenerating new mineralized bone tissues that completely bridges the defect. Thus, surgical and reparative interventions are needed. Treatment strategies for limb bone damage include limb shortening, bone transport distraction osteogenesis, and grafting in conjunction with metal plates, rods, or fixators to stabilize and accommodate the load-bearing function of the injured site. Graft implants that are in clinical use are recovered salvageable bone pieces, iliac crest and other vascularized and nonvascularized trabecular bone autografts, allografts and demineralized bone matrices, osteoconductive calcium compounds, and materials with osteogenic activity [2e4]. A potent osteoinductive factor, bone

* Corresponding author: School of Biomedical Engineering and Sciences, Virginia Polytechnic Institute and State University, Blacksburg, VA 23061, USA. Tel.: þ1 540 231 0048. E-mail address: [email protected] (M.E. Van Dyke). 0142-9612/$ e see front matter Ó 2012 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.biomaterials.2012.11.002

morphogenetic protein 2 (BMP-2), is currently being applied in patients at supraphysiological doses to stimulate production of bone growth through increased activation of the BMP signaling pathway [5]. The medical product InfuseÒ Bone Graft utilizes an absorbable formaldehyde-crosslinked type I collagen sponge carrier made from bovine Achilles tendon to locally-deliver BMP-2 [6]. It is U.S. Food and Drug Administration (FDA)-approved for anterior-approach lumbar spinal fusion caused by degenerative disc disease, sinus augmentation, localized alveolar ridge augmentation for defects associated with extraction sockets, and treatment of acute open tibial shaft fractures. In clinical studies of these applications, Infuse has been demonstrated to be a capable replacement to autografting procedures [7]. Off-label use of Infuse is also being performed in orthopaedic and dental applications [8], which includes repair and management of segmental long bone defects [9e11]. This is because of the urgent need for an easilyaccessible osteoregenerative material to deal with various bone defects. However, adverse side effects have been reported in both on- and off-label Infuse usage [12e16], most commonly in cervical spine fusions. Problems like chronic inflammation, graft rejection, and heterotopic ossification [17,18] may be linked to deficiencies of the collagen BMP-2 carrier. The chemically-crosslinked bovine

R.C. de Guzman et al. / Biomaterials 34 (2013) 1644e1656

collagen sponge can evoke a host antibody response [10] due to structural differences in human extracellular matrix (ECM) proteins compared to bovine. Hydration with BMP-2 solution collapses the bulk scaffold volume (Fig. 1), minimizing the ability of collagen to act as a provisional matrix for cells involved in regeneration. Consequently, Infuse is typically limited for use in conjunction with mechanically-stable materials such as bone allografts and calcium phosphate or calcium sulfate bioceramics in a “burrito-like” assembly [9,10], or in tightly-confined pockets to avoid graft translocation. Since the collagen doesn’t effectively bind to the BMP-2 at the molecular level, unintended compression can leach the absorbed BMP-2 into surrounding tissues, thereby effectively decreasing the growth factor load of the sponge and increasing the risk of ectopic bone formation. Hence, an alternative biomaterial vehicle that avoids these limitations may lower the incidence of detrimental outcomes, widen the range of clinical applications, and improve the performance of BMP-2. Keratin intermediate filament protein extracts obtained from human terminal hair strands have several properties suggested to be critical for the development of an ideal BMP-2 delivery system [19,20]. They are processed as oxidized and reduced forms termed keratose and kerateine, respectively. Implantation studies have shown their in vivo capacity as a biomaterial due to integration with host tissues, generation of minimal fibrous capsule response, tolerance to cellular and vascular infiltration, and aversion of foreign-body giant cell formation, chronic inflammation, and graft rejection [21]. Their human allogeneic nature, structural conservation across species, and absence of genetic elements make them non-antigenic and non-immunogenic [22]. Keratin biomaterials are readily-available, inexpensive to isolate, easy to handle, and can be gamma ray-sterilized. They have been employed as temporary support matrices for tissue engineering applications [23e25] because of their tissue compatibility and degradability [21] with potentially non-toxic peptide byproducts. Additionally, animals do not synthesize keratinase enzymes, so the in vivo degradation behavior of keratin biomaterials can be controlled predominately through manipulation of their material properties (e.g. molecular weight, effective crosslink density, oxidative state, etc.). Other investigators were able to show that a composite containing

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a keratin biomaterial was able to promote osseointegration in an ovine model [41]. Therefore, we hypothesized that a keratose scaffold would provide a stable depot for added BMP-2, allow for regenerative cell infiltration, produce a minimal inflammatory response, and facilitate bone regeneration across a critical-size gap. In this study, keratin in the form of a pure, freeze-dried keratose scaffold was utilized due to its relatively fast degradation rate (about 60% degradation in 1 week [21]) to allow for the deposition of fibrocartilage (soft) callus as in normal endochondral ossification fracture healing [26]. Other essential elements of a good BMP-2 carrier/scaffold system were also investigated, namely, binding interaction with BMP-2, maintenance of growth factor bioactivity, and effective dose delivery to induce regeneration of bone tissue in a non-spontaneously healing defect.

2. Materials and methods 2.1. Extraction of keratose and implant preparation The crude keratose biomaterial used in this study was obtained using our modified method [21], originally developed by Alexander and Earland [27]. Briefly, human hair clippings were oxidized via peracetic acid treatment followed by successive extraction of soluble keratin proteins in Tris base and deionized water. The pooled solution was neutralized and dialyzed in endotoxin-free water to remove traces of the oxidant, adjusted to pH 7.4 with NaOH, lyophilized into powder, and sterilized through exposure at 25-kGy to gamma irradiation. A 25% (w/v) homogenous keratose hydrogel was then prepared by dissolving the powder in sterile phosphate-buffered saline (PBS) and incubating overnight at 37 C. To make the keratose with BMP-2 implant (KB2), the 25% gel was thoroughly mixed with 1.25 mg/mL BMP-2 (in sterile water) component of the Infuse Bone Graft (Medtronic, Minneapolis, MN) using a positive-displacement pipette (Gilson, Middleton, WI), arriving at a final concentration of 20% keratose and 250 mg/mL BMP-2. The mixture was injected into the lumen of a NalgeneÒ 890 polytetrafluoroethylene tube (Nalge Nunc, Rochester, NY) (internal diameter ¼ 4.8 mm) at 200 mL each (length w 11 mm). Afterwards, samples were freeze-dried, removed from the cast, and re-sterilized by gamma irradiation. Each construct contained 50 mg of BMP-2. Control keratose scaffolds without the growth factor (KOS) were made similarly except for the use of water instead of the BMP-2 solution. In addition, a positive control implant was included using the Infuse system (INF). The sterile absorbable collagen sponge (bovine type I collagen) in the Infuse kit was first cut into a rectangular prism (4 4 12.5 mm3; V ¼ 200 mL) and 40 mL of BMP-2 at 1.25 mg/mL (50 mg) was then added through absorption. The properties of the implant groups are summarized in Table 1.

Fig. 1. A) A modular fixation device composed of 2 stainless steel plates and a polysulfone bridge held together by miniature gold-plated screws was used to stabilize the rat femur defect model. B) Representative crosslinked type I collagen sponge soaked with BMP-2 (INF) and lyophilized keratose gel with incorporated BMP-2 (KB2) are shown implanted into the 8-mm bone gap prior to wound closure.

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Table 1 Implants and animal groups.

2.5. Monitoring and evaluation of bone regeneration

Group

Biomaterial implant

BMP-2 mass

Scaffold volume

EMP (Empty) INF (Infuse)

None Bovine type I collagen sponge Human keratose lyophilized gel Human keratose lyophilized gel

0 50 mg

0 200 mL

0

200 mL

50 mg

200 mL

KOS (Keratose) KB2 (Keratose þ BMP-2)

2.2. In vitro scaffold degradation and BMP-2 release The sterile implants (INF, KOS, and KB2; n ¼ 3/group) were placed in a 1.5-mL flip-top tube, rehydrated with 250 mL of PBS, and incubated at 37 C. At multiple time points (0, 1.5, 3, 6, 12, and 24 h, and 2, 4, 7, 14, 21, and 28 d), 50 mL of the supernatant was collected and replaced by the same volume of fresh PBS. A final visual inspection was performed at 5 months and digital photographs of the residual scaffolds obtained. The collected samples were assayed for total protein content via DC Protein Assay (Bio-Rad, Hercules, CA). BMP-2 concentration was quantified using a BMP-2 ELISA kit (PeproTech, Rocky Hill, NJ). In both cases, dilution factors were noted and used for later adjustment of the computed final concentration. The BMP-2 release profile of KB2 was normalized to the KOS background ELISA signal. Cumulative percent release values were plotted versus time on a logarithmic scale.

2.3. In vivo BMP-2 functional assay To test the bioactivity of keratose-incorporated BMP-2, sterile 20% (w/v) keratose hydrogels were initially prepared as described earlier containing 0, 5, 25, and 50 mg of BMP-2 at a final volume of 100 mL and were loaded into 1-mL insulin syringes. All animal procedures were conducted with the approval of the Wake Forest University Health Sciences Animal Care and Use Committee (ACUC). Adult (6w old) male CD-1Ò mice (Charles River, Wilmington, MA) were placed under general anesthesia, hairs on the left leg were clipped, and the exposed skin regions were surface disinfected with povidone-iodine. Keratose gels were then injected (n ¼ 1 mouse/BMP-2 dose) into the hamstring muscles (posterior femoral muscles) of the left leg. Mice were euthanized at 14-d post-injection and micro-computed tomography (m-CT) scans were performed using the MicroCAT II system (Siemens Medical Solutions, Malvern, PA) with a 73-mm resolution. 3D images were reconstructed using AquariusNet (TeraRecon, Foster City, CA). Volumetric measurements of bone formation were performed using Amira software (Visage Imaging, San Diego, CA).

The progress of new bone formation was monitored using X-ray fluoroscopy (SIREMOBIL Compact L; Siemens Medical Solutions) at 1, 4, 8, 12, and 16-w postsurgery. To quantitatively evaluate bone tissue regeneration, animals (n ¼ 6 for EMP; n ¼ 8 for INF, KOS, and KB2) were subjected to cone-beam m-CT imaging (voxel size ¼ 0.0732 mm 0.0732 mm 0.0732 mm) at 8 and 16-w. DICOM image files obtained from COBRA (Exxim, Pleasanton, CA) were 3D-reconstructed using AquariusNet and Amira. The segmentation editor was used to select the region of interest (ROI) and the material statistics function to get bone volume readings (excluding internal radiolucent gaps). Raw 3D data files were imported into ImageJ (National Institutes of Health, Bethesda, MD) and mean gray values bound by the ROI were obtained (average of 10 measurements within the 8-mm defect region). A standard curve was generated from a phantom scan of known densities and employed to convert mean gray values to bone mineral density (BMD) data. The background level was subtracted from BMD values. Additionally, the tissue bone mineral content (BMC) was computed by multiplying BMD by the corresponding ROI volume that included the spaces bound by the structure. Contralateral long bones (right femurs) were also analyzed and served as uninjured normal (NOR) controls. 2.6. Bone harvest and biomechanical testing At the 16-w endpoint, rats were sacrificed by CO2 asphyxiation and left (injured treated) and right (uninjured normal) femurs were immediately dissected out. Internal fixator plates were carefully detached by removing the bone-penetrating screws and tissue samples were wrapped with gauze-soaked PBS. Femurs were briefly photographed and scanned by m-CT (used for BMD and BMC analyses described above). EMP and KOS groups did not exhibit segmental defect bridging to the point of stabilizing the injury and thus were excluded from biomechanical analysis. INF and KB2 specimens (n ¼ 6 each) with their contralateral NOR femurs were vertically potted (cylinder with a 25-mm length and 15-mm diameter) at their ends using Bondo (styrene-acrylic polymer) All-Purpose Putty (3M, St. Paul, MN), leaving a 10-mm exposed section of bone tissue in the femoral mid-diaphysis. Samples were subjected to torsional biomechanical testing using an 858 Mini BionixÒ II axial/torsion load frame (MTS, Eden Prairie, MN) with 647.02B Hydraulic Wedge Grip (MTS). A 2.5 kN/25 Nm axial/torsion load cell was used. Femurs were placed head down with epicondyles at the top grip and twisted at an angular frequency (u) of 2.5 /s (0.0436 rad/s), clockwise and counterclockwise directions for right and left femurs, respectively. Left femur torsional torques (T) were reversed in sign and torsional rotations (q) were reversed in order to properly compare with the right femoral values. Data were zeroed at the starting points and ultimate torsional torque (TU) and rotation (qU) measurements were identified at the instance of bone failure. Rotational stiffness (k) was obtained from the slope of the linear region of the T versus q plot. Shear value conversions were determined with the assumption that samples are hollow cylinders with R ¼ outer radius, r ¼ inner radius, L ¼ length, and J ¼ second moment of area ¼ p(R4er4)/2. Shear stresses (s) and strains (g) were derived using: s ¼ TR/J and g ¼ qR/L, respectively. Ultimate shear stresses (sU) and stains (gU) were reported, as well as the intrinsic shear modulus (G) properties, which was taken as the slope of the initial linear region of the seg curve.

2.4. Establishment of a critical-size bone defect model and biomaterial implantation 2.7. Infrared spectroscopy of powdered samples The implants were assessed for their capacity to induce bone regeneration in an 8-mm critical-size defect model based on Oest et al. [28]. Again, animal procedures were approved by the Wake Forest University Health Sciences ACUC. A modular internal fixation device composed of two stainless steel (SS) grade 316 plates and a ThermaluxÒ polysulfone bridge, originally designed by the Guldberg Lab at Georgia Tech, was modified to reduce its vertical profile using SolidWorks 3D CAD software (Dassault Systèmes, Waltham, MA). The fixator (Fig. 1A) was assembled using 4 gold-plated SS (grade 303) miniature screws and then sterilized by exposure to ethylene oxide. Male 8-month old (m ¼ 405 24 g) Fisher 344 (F344) inbred rats (Harlan, Indianapolis, IN) were acquired and acclimated for at least 24 h before surgery. Under isoflurane general anesthesia and oxygen, hair on the dorsal left leg area was shaved and the underlying skin was disinfected with povidone-iodine solution. A 4-cm skin incision was created and the muscles surrounding the left femur were separated and pulled away from the bone. The internal fixator was clamped on the anterior aspect of the femur diaphysis between the knee and hip joint. Four holes were drilled (2 on each end of the clamp) with a wire drill bit into the femoral cortices and the fixator was secured with screws. After removal of the clamp, an 8-mm bone segment was cut off using a micro-sagittal saw. The defect site was flushed with saline to remove blood and tissue fragments. Biomaterial implants (INF, KOS, and KB2; n ¼ 8 each) were then carefully placed into the bone gap (Fig. 1B). For the negative control group (EMP; n ¼ 6), the defect site was left empty. Muscle layers above and below the exposed femur (quadriceps femoris) were sutured together and skin flaps were stapled using a ProximateÒ SS stapler (Ethicon Endo-Surgery, Cincinnati, OH) to close the wound. Animals were allowed to recover, housed individually, and skin staples were removed after 7 d. Important physical measurements of excised femoral segments were determined to be: mass ¼ 94.2 12.9 mg and outer circumference (mean of proximal and distal ends) ¼ 10.9 0.8 mm.

Unpotted bone diaphysis segments (w8-mm in length) utilized earlier for biomechanical testing (left and right femurs of INF and KB2; n ¼ 5 each) were freezedried, finely-ground by a mortar and pestle, and analyzed using a Spectrum 400 FTIR (Fourier transform infrared) spectrometer with a Universal ATR (attenuated total reflectance) Sampling Accessory attachment (FTIR-ATR) (PerkinElmer, Waltham, MA). Powdered samples were placed on the diamond/ZnSe crystal surface and the pressure arm was tightened to force gauge ¼ 125e135 N. Transmittance readings were obtained at wavenumbers 4000-650 cm1 with 4 cm1 resolution. Data tuneup and peak table functions were processed via Spectrum software (PerkinElmer). Pure diaphysis bone and hydroxyapatite controls were also analyzed. 2.8. Histological evaluation of recovered femurs Recovered bones and their surrounding tissues (n ¼ 3/group; 1 sample each for INF and KB2 have been used for torsional testing) were fixed in 10 volume of 10% neutral-buffered formalin for 3 d, washed with water, and soaked in ImmunocalÒ (Decal, Tallman, NY) for 7 d (decalcifier solution changed daily). Complete decalcification was verified via X-ray fluoroscopy. Samples were then neutralized with CalArrestÔ (Decal) for 3 min, rinsed with water for 10 min, placed in 60% EtOH for storage, and paraffin-embedded. Longitudinal 5-mm sections from the middle aspect of the tissues were obtained using a microtome and stained for Masson’s trichrome. Bright field light microscope images were taken and assembled to re-create the region encompassing the entire 8-mm segmental defect in treated animals and the corresponding diaphysis in NOR femurs. Images were captured (5 random fields-of-view/sample) through circularlypolarized light microscopy employing a filter oriented parallel to the plane of aligned collagen in the outer cortex of the femoral diaphysis for maximum

R.C. de Guzman et al. / Biomaterials 34 (2013) 1644e1656 birefringence. The ratio of the area occupied by the signal was quantified using ImageJ and normalized to the NOR mean value. Tissue sections of recovered EMP and KOS samples were analyzed in ImageJ for adipose tissue content by selecting the area occupied by fat deposits divided by the total area of non-bony tissues within the 8-mm defect sites. Fibrous tissues composed of fibroblasts and collagen fibers were also identified and their area ratios were determined (fibrous tissue area/total area). Finally, the amounts of other tissues (including blood vessels) were measured using the equation: other tissue ratio ¼ 1(adipose tissue ratio þ fibrous tissue ratio). 2.9. Statistical analyses Scatter plot, bar graph, tabular, and individual data point values were reported as average 1 standard deviation. Logarithmic and linear fits obtained using Office Excel (Microsoft, Redmond, WA) for the in vitro BMP-2 release and scaffold degradation were represented by their respective equations and coefficients of determination (r2). Statistical analyses were conducted using Prism software (GraphPad, San Diego, CA) at 95% confidence intervals and probability of type I error (a) ¼ 0.05. Comparison of scatter plot curves, whole femoral volumes of BMP-2 treated rats, regenerated bone spectral peak transmittance, and non-bony tissue contents in unregenerated defect samples were performed using un-paired Student’s t-test. A paired t-test was employed for evaluation of recovered femur volume, BMD, and BMC measurements in treated versus the corresponding contralateral normal bone samples. Analysis of variance (ANOVA) and Tukey’s post-hoc tests were used in multiple-group comparison of bone volume in the 8-mm gap, biomechanical properties, and polarized light microscopy values.

3. Results

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into the medium at the 1-month time point approached that of the KB2 construct, at 75 14%. The in vitro keratose scaffold degradation exhibited a polymerlike dissolution behavior [29] in which the PBS solvent infiltrates and swells the bulk material, forms a gel, and facilitates the movement of disentangled polymer chains out of the intact scaffold. This erosion process was found to be not significantly different between KOS and KB2 (p ¼ 0.6928), indicating that the presence of BMP-2 in KB2 does not affect the construct’s bulk degradation properties. Both KOS (r2 ¼ 0.9517) and KB2 (r2 ¼ 0.9058) breakdown followed a logarithmic trend line while INF degraded in a zero-order linear fashion (r2 ¼ 0.9942). At 5 months, INF was completely degraded (Fig. 2 inset) but both KOS and KB2 showed a residual intact network. Extrapolation using regression analysis supports these observations and predicted that about 7% of the starting keratose scaffold mass is still present at the 5-month time point. Interestingly, the BMP-2 release curve from KB2 (Fig. 2) superimposes to the KB2 scaffold degradation profile (p ¼ 0.9568), which suggests direct interaction of BMP-2 with keratose proteins leading to BMP-2 release as the keratin scaffold undergoes dissolution. In contrast, BMP-2 did not exhibit binding to the collagen carrier and leached out independently from the scaffold. This was evidenced by the significant (p ¼ 0.0005) difference between INF scaffold degradation and INF-BMP-2 release behaviors.

3.1. Release profile of BMP-2 in relation to keratose bulk degradation

3.2. Muscle injection of keratose gels with BMP-2

BMP-2 within the keratose scaffold of the KB2 implant displayed a logarithmic (r2 ¼ 0.9198) release profile (Fig. 2). At the 24-h time point, 61 26% of BMP-2 was burst-released into the PBS medium, and by 28 days, the cumulative release was 80 16%. Type I collagen sponge in the INF implant provided a comparatively slower logarithmic (r2 ¼ 0.9654) BMP-2 delivery at the early time period (36 20% for 1 d). However, due to the steeper slope of the release ratio versus log (time) curve, the amount of BMP-2 released

Keratose hydrogels without (0 mg) and with 5 mg of BMP-2 did not induce any bone formation within the mouse hamstrings. The 25 mg dose, however, produced 3 mm3 of new bone tissue (Fig. 3) 14 days after local leg muscle injection. Doubling the amount of the osteoinductive factor from 25 to 50 mg generated a 25-fold increase in bone volume (76 mm3). Results demonstrated the dosedependent in vivo functional activity of BMP-2 delivered using the keratose biomaterial.

Fig. 2. Scaffold degradation versus BMP-2 release profiles. Keratose scaffolds (KOS and KB2) degrade faster than the crosslinked type I collagen control (INF) at early time points. However, after 5 months in vitro, residual keratose scaffolds in gel form were still intact while the collagen sponge was completely degraded (inset image). BMP-2 release from KB2 fits the logarithmic keratose degradation curves. In contrast, BMP-2 delivery from INF also behaved in a logarithmic trend but does not fit the collagen zero-order degradation curve.

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Fig. 3. 3D-reconstructed m-CT images display the dose-dependent effect of keratose-delivered BMP-2 for induction of ectopic bone in mouse leg muscles.

3.3. Bone regeneration in a critical-size femur diaphysis defect X-ray fluoroscopy in situ confirmed that excision of the 8-mm rat femoral diaphysis is indeed a critical-size model since minimal spontaneous bone regeneration and no bridging occurred when the gap was left untreated (empty; EMP) over a span of 16 weeks (Fig. 4 and Fig. 5A) in 6/6 animals. One week post-surgery, the defect sites in all groups remained clear of radiodense materials. However by 4 weeks, those with 50 mg of BMP-2 (INF and KB2) showed mineralized tissues connecting the proximal and distal bone stumps. The INF group was qualitatively observed to have more spread and diffuse mineralization as opposed to the more compact and tighter appearance produced by KB2 implants. Increase in radiopacity was discernible from 4 to 16 weeks. By the 16-week time point, images of INF and KB2 regenerated tissues appeared to have similar morphologies. Untreated (EMP) and keratose only-treated (KOS) defects displayed only small amounts of bone growth over time characterized by tapering of severed bone ends but no full gap bridging. Reconstruction of m-CT data clearly depicted the progress of bone regeneration in INF and KB2 groups (Fig. 5A) and allowed the volumetric analysis of mineralized bones. At the 8-week time point, the amounts of callus calcification in both INF and KB2 within the 8mm defect did not differ (p > 0.05) from that of the contralateral

un-injured normal (NOR) femur (Fig. 5B). By 16 weeks, the quantities of dense materials in INF and KB2 were significantly greater than the contralateral bone (150%; p < 0.05). The sizes of regenerated bones in INF and KB2 groups at 8 and 16 weeks were statistically similar (p > 0.05). Recovered whole femurs also showed that the BMP-2-treated bones (INF and KB2; Fig. 6) had 1.40 times more (p < 0.0001) mineralization in comparison to their respective NOR controls (Fig. 5C). The whole femur volume of INF was found to be not significantly different from that of KB2 (p ¼ 0.9105). The KOS group appeared to produce more bone within the 8mm gap than the EMP (Fig. 5B). However, comparative statistical analysis revealed that the means were statistically different (p < 0.0001) only at 8 weeks, but not (p ¼ 0.0649) at 16 weeks. The newly-regenerated bones recovered from INF and KB2treated animals appeared to have similar external texture to the normal femur (Fig. 6 and Fig. 7A). However, the process produced an irregularly-shaped bone with widening in the medial aspect, deviating from the oval cylindrical geometry of the contralateral femoral diaphysis. Additionally, the region directly below where the polysulfone bridge was located (anterior mid-diaphysis) generated a concave morphology with decreased mineralization but increased fibrous soft tissues. Anterior diaphysis areas in contact with the stainless steel plates became flattened over the 16-

Fig. 4. X-ray radiographs of critical-size femoral defects with or without treatment. Bone regeneration was evident in rats treated with INF and KB2 as early as 4 weeks after implantation. The bone growth in INF (red arrow) was more diffuse than in KB2 (blue arrow). Progressive increase in radiopacity was observed over time, indicative of increase in mineralization. By contrast, EMP and KOS groups did not produce bone gap bridging and repair. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

R.C. de Guzman et al. / Biomaterials 34 (2013) 1644e1656 Fig. 5. A) m-CT images of rat femurs, showing BMP-2 delivered in type I collagen (INF) and keratose (KB2) enabled bone gap bridging and regeneration as shown at the 8 and 16-week time points. B) Bone volumes within the 8-mm defect in INF and KB2 were found to be similar (p > 0.05) to the contralateral uninjured normal (NOR) femur at 8 w; but by 16 w, mineralized bone amounts have significantly increased (*p < 0.05). EMP and KOS groups have less (p < 0.001) bone than NOR at both time points. At 8 w, the value of KOS is greater than that of the EMP (yyyp < 0.0001). C) Quantification of m-CT images of recovered whole femur volumes demonstrated that animals with INF and KB2 generated larger bones (***p < 0.0001) compared to normal femurs. INF versus KB2 volumes are statistically similar (p ¼ 0.9105).

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Fig. 6. Recovered femurs from INF and KB2 implantation have external textures resembling the contralateral uninjured NOR bones but their geometrical shapes are bigger and have medial protrusions. Anterior mid-diaphysis regions have less mineralization but more fibrous tissues.

Fig. 7. A) Isosurface and orthoslice views of recovered INF and KB2-implanted femurs and their corresponding contralateral controls are represented above. Bone cortices of regenerated tissues are thinner and less dense than those of normal femoral diaphysis as seen in cross-section and longitudinal radiographs. B) Analysis showed significantly less (***p < 0.0001) volumetric bone mineral density (BMD) values in INF and KB2 compared to NOR bones. BMD of INF and KB2 are similar (p ¼ 0.9771) to each other. C) Bone mineral content (BMC) measurements of BMP-2-treated groups were determined to be greater (***p < 0.0001 for INF and **p ¼ 0.0025 for KB2) than their respective contralateral bones. Comparison of INF and KB2 displayed similar values (p ¼ 0.6161).


w time span and excess bone formation was observed surrounding the plates in both INF and KB2 animals. Orthoslice radiographs of INF and KB2-treated femurs showed formation of trabecular bone structures with less density and thinner cortices compared to the normal bone (Fig. 7A). Volumetric BMD results obtained from NOR femoral diaphysis of approximately 1.3 g/cm3 fell within published values [30]. BMD readings from regenerated bones of INF and KB2 were significant lower (p < 0.0001), at 49% of their normal controls (Fig. 7B). INF and KB2 values were found to be similar (p ¼ 0.9771). The femoral diaphysis BMC measurement, taken from the product of BMD and volume (including confined radiolucent spaces), of 115 mg is also within the expected result range [31]. It was determined that the amount of minerals in the NOR femur was less compared to the BMC in INF (p < 0.0001) and in KB2 (p ¼ 0.0025) (Fig. 7C). On average, INF and KB2-regenerated bones had 1.36 more minerals than NOR controls. The BMC fold difference was expected to be close to the bone volume change of 150% at the 16-week time point (Fig. 5B). INF and KB2 mineral contents were determined to be statistically similar (p ¼ 0.6161). 3.4. Recovered bone tissue biomechanics Torsional analysis of femurs harvested at 16 weeks post-surgery revealed that torques required to break the bones (TU) did not differ (p ¼ 0.2222) among the NOR, INF, and KB2 specimens (Table 2). However, it was observed that the ultimate angular rotations (qU) of the 3 groups were significantly different (p ¼ 0.0438). Specifically, INF was less (*p < 0.05) than the normal value. Stiffnesses (k) of the newly-regenerated INF and KB2 bones were found to be similar (p ¼ 0.2669) to the k value of contralateral femurs. Ultimate shear stresses (sU) were determined to be different (p ¼ 0.0049) among NOR, INF, and KB2 femurs. In particular, NOR > INF (*p < 0.05) and NOR > KB2 (*p < 0.05), but INF ¼ KB2 (p > 0.05), indicating that the normal femur mineral organization could withstand greater shear stress toward failure compared to the regenerated bone in the INF and KB2 groups. Ultimate shear strains (gU) and shear moduli (G) of the 3 femoral groups showed similarities (p ¼ 0.3960 for gU and p ¼ 0.2182 for G), which suggests that the regenerated tissues have the same flexibility as the uninjured normal control. This also demonstrates that they have similar shear strengths below the failure points. Initial axial forces were recorded during testing since the potted femurs were not perfectly parallel to the machine grips caused by irregular bone geometries. Generally, the axial forces gathered after the point of failure (end of the data collection) were detected to be close to the initial values, implying that only small un-avoidable axial components affected the torsional outcome measures. 3.5. Mid-infrared spectral profiles The FTIR-ATR spectral plots of INF and KB2 ground bones displayed similar and overlapping patterns (Fig. 8). Seven transmittance peaks at 2924e2923, 2854e2853, 1744e1743, 1643e1641, 1436e1414, 1015e1011, and 721e710 cm1 wavenumber ranges were observed (rectangular dots in Fig. 8) deviating from the

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contralateral normal bone spectrum, suggesting that these peaks distinguish the remodeling tissue from uninjured cortical bone. Student’s t-test comparison between INF and KB2 transmittance points showed no significant differences (p > 0.05), which signify resemblance in molecular and tissue composition of the 2 groups. The asymmetric (2924e2923 cm1) and symmetric (28542853 cm1) CH2 stretches, and C¼O stretch (1744e1743 cm1) peaks [32,33] found in INF and KB2 samples correspond to triglycerides and lipid hydrocarbons in bone marrow adipose tissues. Accordingly, smaller peaks were detected in normal bones, probably due to less bone marrow content. The rocking vibrations of CH2 in long-chain fatty acids caused the occurrence of the 721710 cm1 band. Peaks at 1436e1414 and 1015e1011 cm1 represent 3the respective carbonate (CO23 ) and phosphate (PO4 ) groups’ asymmetric stretching behaviors in carbonated hydroxyapatite [34] comprising the mineral phase of the bone (w65% of the bone mass) [32]. The remodeling bones were determined to have less carbonates but more phosphates relative to the normal diaphysis. The hydroxyapatite control spectra expectedly did not generate any carbonate bands. Type I collagen, which accounts for most of the organic bone phase, was also identified via the amide I (peptide C¼O stretching) at 1643e1641 cm1 and amide A at 3294e 3286 cm1 peaks. Amide band intensities indicated more collagen in the INF and KB2 samples than in the cortical bone. 3.6. Decalcified bone tissue histology Longitudinal sections of recovered decalcified tissues stained for Masson’s trichrome demonstrate that the newly-formed bones from INF and KB2 implants developed a cancelous macroscopic morphology unlike the compact structure of the normal femoral diaphysis (Fig. 9). Bone marrow was present within the network of trabecular bone spicules in both groups indicating that the new bone tissues integrated into the medullary cavity of adjacent uninjured bone. The periosteum lining the outer bone surface is the primary source of bone repair mesenchymal stem cells (MSCs) (other sources include bone marrow, muscles, and surrounding tissues). This connective tissue layer was found to be thicker due to the presence of more cells and secreted collagen ECM compared to the relatively less active periosteum of an uninjured femur. Cells in close proximity to the surface appeared to be active osteoblasts, characterized by their oval and less flattened shapes. Contrastingly, osteoprogenitors of the periosteum appear as flat and elongated cells. Cells farther away from the bone surface represent a mixture of osteoblasts and fibroblasts since their morphologies and staining properties are indistinguishable from each other. Multinucleated osteoclasts were also detected particularly in bone surfaceresorption cavities. Osteocytes were identified as individual cells embedded within the decalcified bone matrix. No blood vessels were found penetrating the INF and KB2 spongy bones. The bone layer immediately below the periosteum in normal diaphysis longitudinal sections can be seen to have parallel arrangement of collagens because of the outer circumferential lamellae organization. These aligned fibers were observed only in small areas of the outer INF and KB2 bone regenerates. The tissue

Table 2 Mechanical properties of recovered femurs. Femur

NOR INF KB2

Ultimate torsional TU ¼ torque (Nm)

qU ¼ rotation (rad)

0.513 0.133 0.379 0.125 0.443 0.123

0.317 0.0994 0.188 0.0667* 0.241 0.0477

k ¼ rotational stiffness (Nm/rad) 2.09 0.450 2.46 0.300 2.32 0.332

Ultimate shear

sU ¼ stress (MPa)

gU ¼ strain

52.9 13.4 30.8 11.0* 31.1 10.4*

0.0642 0.0204 0.0506 0.0179 0.0657 0.0110

G ¼ shear modulus (MPa) 871 250 706 94.0 686 170

1652 R.C. de Guzman et al. / Biomaterials 34 (2013) 1644e1656 Fig. 8. The FTIR-ATR spectra of INF and KB2-regenerated bones have similar profiles as illustrated in the split (left) and overlaid (right) graphical views. Transmittance peaks (-) show their characteristic identities compared to the spectral plots of contralateral normal femur (NOR), pure cortical diaphysis (bone), and hydroxyapatite (HA) mineral.


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Fig. 9. Left column images of Masson’s trichrome-stained decalcified longitudinal sections display the location of the 8-mm segmental defect created 16 w prior to tissue harvest in relation to the uninjured normal (NOR) control. The periosteum (p) on the outer layer of bone (b) in experimental groups (EMP, INF, KOS, and KB2) is active as evidenced by increased thickness caused by osteoblast proliferation. Tissues found in the gap of unregenerated EMP diaphysis are mostly composed of fat cells (f) and fibrous collagen (c) deposits. Keratose (k) remnants can still be observed in the KOS sample defect site. Bone marrow (m) tissues were detected in spaces between regenerated bone spicules in INF and KB2 femurs.

cancelous spicules display both woven (immature/primary) and lamellar (mature/secondary) structures, indicative of hard calluses in the process of being remodeled. The amounts of lamellar bone as detected by polarized light microscopy (Fig. 10) in INF and KB2 new tissues were not found to be statistically different (p > 0.05). However, they are still significantly less compared to the occurrence in normal compact bone (p < 0.05). Collagen fibers that exhibited an irregular and disorganized pattern in trichrome staining and produced no signal in polarized light filter correspond to deposited woven bone. The untreated femoral defect (EMP) and the keratose only (KOS) implantation groups showed periosteal layer activation via cell proliferation and partial bone growth at the tips of the uninjured bone stumps but were unable to fully bridge the defect. The EMP defect site was filled with primarily adipose tissues interspersed with fibroblasts and fibrous collagen deposits, blood vessels, and immune cells. Femoral muscles normally surround and contain these tissue structures. The patterns of cells and ECM did not

indicate cartilaginous callus formation. Significant reduction (**p ¼ 0.0023) of fat was observed within the defect region in animals treated with KOS scaffolds (Table 3). Quantities of the other tissues including blood vessels located in the bone gap were determined to be similar (p > 0.05) in KOS versus EMP groups. Residual keratose remnants filled with cells and blood vessels were also observed in KOS-implanted animal sections which displayed faint red color in Masson’s trichrome (Fig. 9). The availability of keratose materials in KOS but not in KB2 specimens imply less tissue activity and turnover when full-gap bone bridging is not achieved. 4. Discussion Peracetic acid oxidation of keratin proteins during the extraction process used to produce keratose biomaterials introduces sulfonic acids (eSO 3 ) in place of the highly-abundant cysteine and cystine thiol groups (eSH), which then confers net negative charges to the

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Fig. 10. Circularly-polarized light images of INF and KB2 spongy bone regenerates display the regions with oriented collagen fibers indicative of lamellar bone organization. Ratios of aligned collagen relative to the normal in INF and KB2 samples are not (p > 0.05) significantly different from each other, but both are less (p < 0.05) compared to the NOR value.

protein chains [21,27,35]. The recombinant human BMP-2 growth factor used in this study, conversely, is positively-charged at acidic and neutral pH levels due to its basic isoelectric point (pI) [6]. In the preparation of KB2 construct samples (Table 1), keratose in PBS at pH 7.4 was mixed with BMP-2 in water at pH 4.5 to get a mass ratio of 4 104 mg: 50 mg or 800: 1 (keratose : BMP-2) and a final pH value of slightly < 7.4. Consequently in this condition, it is expected that the keratose() scaffold in charge-excess will sequester all the BMP-2(þ) molecules via ionic/electrostatic interaction, serving as the basis for tightly-localized and controlled keratose delivery of BMP-2 into target sites. Our group has previously found that keratose may interact with the positively-charged drug ciprofloxacin for inhibition of bacterial growth [36]. Similarly, Tachibana et al. demonstrated that conversion of thiols to carboxylic acids (e COO) to create negative keratin backbone charges enabled attachment of proteins with a high (basic) pI such as BMP-2 [37]. In vitro kinetics results supported this proposed molecular attraction mechanism since the cumulative release of both keratose and BMP-2 proteins were correlated (Fig. 2). The possible keratoseBMP-2 association was shown here to maintain the biofunctionality of BMP-2 by inducing bone growth in mouse muscles (Fig. 3) and formation of hard fracture callus in the repair stages of segmental long bone defects (Figs. 4e9). BMP-2 exerts its effects by initially binding to specific BMP cell-surface receptors [5]. Thus, even the keratose-bound BMP-2 can still elicit downstream signaling functions, indicating the conservation of BMP-2 receptorinteraction regions. The loss of thiol groups in keratose prevents disulfide linkages (eSeSe) from assembling intra- and inter-chain crosslinks, thereby increasing the solubility of keratose in aqueous solvents [21]. Hence, polymer gelation and scaffold organization occur through physical chain entanglement of concentrated solutes and the bulk network subsequently destabilizes via polymer dissolution comprising solvent diffusion and chain disentanglement transport processes. Additionally, since keratoses are protein derivates and possess peptide bonds, they are prone to amide hydrolysis in water to yield smaller peptide byproduct fragments. The described mechanisms act together in driving the relatively fast keratose scaffold degradation rate seen in static protease-free PBS at 37 C

Table 3 Non-bony tissue in the unregenerated diaphysis gap. Defect site

Tissue content ratio (%) Adipose

Fibrous

Blood vessels and others

Total

EMP KOS

59.12 3.40 22.58 8.50**

23.37 8.71 49.88 17.24

17.51 5.33 27.54 9.27

100.00 10.76 100.00 21.34

(Fig. 2). By comparison, the zero-order (linear) in vitro degradation of crosslinked collagen sponge is likely governed solely by hydrolysis, a slow process especially at physiological pH. The actual resorption behavior of scaffolds in vivo at the bone defect site, however, is expected to be different due to the influence of surrounding structures, injury hematoma, colonizing cells, neovascularization, ECM deposition, tissue growth, micromotion, leg movement, unintended compression, and local temporal environment. Despite differences in the in vitro BMP-2 release and scaffold degradation profiles (Fig. 2), both BMP-2-loaded collagen and keratose carriers resulted in formation of new bones with similar gross tissue morphology, volume, density, mineral content, molecular composition, and strength properties at the end of the study (Figs. 5e10 and Table 2). Mineralization of the critical defect site was observed at the 4-week time point for INF and KB2 groups (Fig. 4), indicating that the first month of healing encompasses the endochondral ossification process up to the beginning of soft callus calcification. At this stage, it is expected that most of the temporary biomaterial matrix has been resorbed and replaced by the hostdeposited cartilaginous collagen that gets calcified and remodeled into woven bone [32,34,38,39], and the BMP-2 load has been exhausted. The only pronounced difference in response between INF and KB2 treatments were seen in the 4-week radiographs in which the keratose-BMP-2 implants produced a more cylindrical and compact bone mineralization following the shape of the scaffold, suggesting that the type of graft materials and BMP-2 delivery mechanism affects early bone healing (soft callus development and ossification) but not the woven bone remodeling phase. Additionally, bone remodeling proceeds independent of the initial structure of the hard callus since the 16-week morphologies of INF and KB2 regenerated tissues formed similarly. Thus, future work on improving the biomaterial-growth factor construct development for segmental bone defects should focus on the healing time frame leading to fibrocartilage synthesis and mineralization. Long bone fracture healing of gapless apposing bone segments is a well-defined biomechanical phenomenon that generally follows the endochondral ossification route involving the following sequence of events after the initial injury: hematoma (blood clot) buildup, acute inflammation, influx of MSCs, angiogenesis, cartilage/soft fibrocartilaginous callus formation, cartilage removal and calcification for synthesis of woven bone/hard callus, and tissue remodeling and lamellar bone maturation to produce the original bone shape and organization [38,39]. In the presence of a small (non-significant) gap, the process still proceeds similar to new bone deposition through the distraction osteogenesis technique, but is assumed to be slower. However in critical-size bone defect (and pathological fracture nonunion) situations, bone healing is impaired because of disruption in the actions of key regeneration


elements, namely, progenitor cells, growth factors, and appropriate milieu [40]. We found mostly fibrous and adipose tissues (and no cartilaginous structure) in the defect region of the empty unregenerated group (Fig. 9 and Table 3), which suggests interruption of the provisional soft callus development, particularly the chondrogenesis phase. The significant size of the gap may have prevented the full-length migration and proliferation of progenitor MSCs into the temporary hematoma fibrin mesh or may have permitted faster defect colonization by other competing cell populations (such as fibroblasts). MSCs may also have differentiated, under the specific inducive local microenvironment, into adipocytes instead of chondrocytes and osteoblasts. Implantation of exogenous graft materials into the gap site limits the amount of blood clot to fill-up the void volume, thereby changing the structural and biochemical matrix properties, and consequently the healing cascade response. In general, keratose scaffolds as implants offer beneficial attributes that on their face may provide advantages in segmental bone graft applications. They can be formulated as malleable constructs to fill-up the bulk injury gap without incorporation of viscosity thickeners and other support materials; they have a porous architecture and contain inherent cell-binding motifs [41], enabling cell and blood vessel infiltration into the material matrix [21] that are known to be important steps in bone repair [39]; they are not as susceptible to naturally-occurring ECM proteases released during wound healing and tissue repair, thereby allowing for greater control over the erosion-mediated release of incorporated payloads; and they demonstrate one of the lowest levels of inflammatory and foreign body responses among conventional biomaterials used in bone regeneration. Keratose on its own promoted more bone stump outgrowth at the 8-week evaluation point compared to the untreated defect (Fig. 5B). This effect may be linked to keratose’s ionic sequestration of injury-released positively-charged regenerative growth factors like certain BMPs, FGFs, IGFs, PDGFs, TGF-bs, and VEGFs [42], which would be expected to extend their bioavailability while not diminishing their bioactivity. A very interesting finding is that the presence of keratose appeared to profoundly decrease the adipose content in the repair site (Table 3), indicating an ability to downregulate adipogenesis and indirectly allow more efficient osteogenesis of bone-repair MSCs. MSCs are known to be chemotactically drawn to BMP-2 [43]. Hence, the keratose-BMP-2 construct (with sufficient BMP-2 load) may have been able to attract MSCs to span across the critical-size gap and induce their BMP-2-mediated differentiation into chondrocytes, and later osteoblasts [11], while perhaps inhibiting adipogenesis. Keratin biomaterials may then have a distinct advantage under conditions such as old age, diabetes, and osteoporosis where adipogenesis is a more strongly favored differentiation pathway for these progenitor cells. However, the data in this study do not provide a basis for a mechanistic postulate. Further investigation of MSCs signaling pathways during osteogenesis, chondrogenesis, and adipogenesis under the influence of a keratin-containing environment are warranted. 5. Conclusions The keratose biomaterial scaffold was shown to be a feasible BMP-2 carrier matrix for bone tissue engineering as it: 1) likely binds with BMP-2 through ionic interaction providing localized and controlled growth factor delivery during its bulk network degradation, 2) preserves the dose-dependent biological functionality of associated BMP-2, and 3) promotes the regeneration of critical-size segmental long bone defects with a small BMP-2 load. Implantation of medical-grade crosslinked collagen sponge (Infuse Bone Graft)

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and keratose with BMP-2 into the gap site led to formation of remodeling spongy bone hard fracture calluses with similar radiographical, histological, biomechanical, and spectroscopic (i.e. compositional) properties. In addition, keratose by itself (no BMP2) induces limited osteogenesis, possibly via sequestration and enrichment of host-secreted positively-charged pro-regenerative molecules like BMPs and influencing multipotent MSCs differentiation towards the osteogenic lineage by inhibition of adipogenesis. These findings indicate the potential use of a keratin-based platform for the development of scaffold delivery systems that expand the current applications of BMP-2 for bone repair and regeneration. Conflict of interest statement Dr. Van Dyke holds stock and is an officer of KeraNetics LLC, who provided partial funding for this research. Wake Forest University Health Sciences has a potential financial interest in KeraNetics through licensing agreements. Acknowledgments The authors would like to thank the following for their invaluable assistance: Luke Burnett and Richard St. Clair (scientific discussions), Maria Bahawdory, Jillian Richter, Bailey Fearing, and Lauren Pace (keratin preparation), Robert Guldberg Lab and Dennis Brown at Georgia Tech (fabrication of internal fixator plates), Erin Mitchell and ARP staff (animal care), Mandy Lockard (X-ray fluoroscopy), Debra Fuller, Sandra Kaminsky, and Josh Tan (m-CT scans), Peter Mente at North Carolina State (torsional testing), and Cathy Mathis and Bridgette Jones (histology). This work was supported by the US Army and KeraNetics, LLC. References [1] Keating JF, Simpson AH, Robinson CM. The management of fractures with bone loss. J Bone Jt Surg Brs 2005;87:142e50. [2] DeCoster TA, Gehlert RJ, Mikola EA, Pirela-Cruz MA. Management of posttraumatic segmental bone defects. J Am Acad Orthop Surg 2004;12:28e38. [3] Fuchs B, Ossendorf C, Leerapun T, Sim FH. Intercalary segmental reconstruction after bone tumor resection. Eur J Surg Oncol 2008;34:1271e6. [4] McKee MD. Management of segmental bony defects: the role of osteoconductive orthobiologics. J Am Acad Orthop Surg 2006;14:S163e7. [5] Lissenberg-Thunnissen SN, de Gorter DJ, Sier CF, Schipper IB. Use and efficacy of bone morphogenetic proteins in fracture healing. Int Orthop 2011;35: 1271e80. [6] Uludag H, Friess W, Williams D, Porter T, Timony G, D’Augusta D, et al. rhBMPcollagen sponges as osteoinductive devices: effects of in vitro sponge characteristics and protein pI on in vivo rhBMP pharmacokinetics. Ann N Y Acad Sci 1999;875:369e78. [7] Govender S, Csimma C, Genant HK, Valentin-Opran A, Amit Y, Arbel R, et al. Recombinant human bone morphogenetic protein-2 for treatment of open tibial fractures: a prospective, controlled, randomized study of four hundred and fifty patients. J Bone Jt Surg Am 2002;84-A:2123e34. [8] Guo J, Li C, Zhang Q, Wu G, Deacon SA, Chen J, et al. Secondary bone grafting for alveolar cleft in children with cleft lip or cleft lip and palate. Cochrane Database Syst Rev 2011:CD008050. [9] Schwartz ND, Hicks BM. Segmental bone defects treated using recombinant human bone morphogenetic protein. J Orthop 2006;3:e2. [10] Jones AL, Bucholz RW, Bosse MJ, Mirza SK, Lyon TR, Webb LX, et al. Recombinant human BMP-2 and allograft compared with autogenous bone graft for reconstruction of diaphyseal tibial fractures with cortical defects. A randomized, controlled trial. J Bone Jt Surg Am 2006;88:1431e41. [11] Ghodadra N, Singh K. Recombinant human bone morphogenetic protein-2 in the treatment of bone fractures. Biologics 2008;2:345e54. [12] Woo EJ. Adverse events reported after the use of recombinant human bone morphogenetic protein 2. J Oral Maxillofac Surg 2012;70:765e7. [13] Carreon LY, Glassman SD, Brock DC, Dimar JR, Puno RM, Campbell MJ. Adverse events in patients re-exposed to bone morphogenetic protein for spine surgery. Spine (Phila Pa 1976) 2008;33:391e3. [14] Wong DA, Kumar A, Jatana S, Ghiselli G, Wong K. Neurologic impairment from ectopic bone in the lumbar canal: a potential complication of off-label PLIF/TLIF use of bone morphogenetic protein-2 (BMP-2). Spine J 2008;8: 1011e8.

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[15] Balseiro S, Nottmeier EW. Vertebral osteolysis originating from subchondral cyst end plate defects in transforaminal lumbar interbody fusion using rhBMP-2. Report of two cases. Spine J 2010;10:e6e10. [16] McClellan JW, Mulconrey DS, Forbes RJ, Fullmer N. Vertebral bone resorption after transforaminal lumbar interbody fusion with bone morphogenetic protein (rhBMP-2). J Spinal Disord Tech 2006;19:483e6. [17] Joseph V, Rampersaud YR. Heterotopic bone formation with the use of rhBMP2 in posterior minimal access interbody fusion: a CT analysis. Spine (Phila Pa 1976) 2007;32:2885e90. [18] Boraiah S, Paul O, Hawkes D, Wickham M, Lorich DG. Complications of recombinant human BMP-2 for treating complex tibial plateau fractures: a preliminary report. Clin Orthop Relat Res 2009;467:3257e62. [19] Haidar ZS, Hamdy RC, Tabrizian M. Delivery of recombinant bone morphogenetic proteins for bone regeneration and repair. Part B: delivery systems for BMPs in orthopaedic and craniofacial tissue engineering. Biotechnol Lett 2009;31:1825e35. [20] Geiger M, Li RH, Friess W. Collagen sponges for bone regeneration with rhBMP-2. Adv Drug Deliv Rev 2003;55:1613e29. [21] de Guzman RC, Merrill MR, Richter JR, Hamzi RI, Greengauz-Roberts OK, Van Dyke ME. Mechanical and biological properties of keratose biomaterials. Biomaterials 2011;32:8205e17. [22] Kirfel J, Magin TM, Reichelt J. Keratins: a structural scaffold with emerging functions. Cell Mol Life Sci 2003;60:56e71. [23] Sierpinski P, Garrett J, Ma J, Apel P, Klorig D, Smith T, et al. The use of keratin biomaterials derived from human hair for the promotion of rapid regeneration of peripheral nerves. Biomaterials 2008;29:118e28. [24] Thilagar S, Jothi NA, Omar AR, Kamaruddin MY, Ganabadi S. Effect of keratingelatin and bFGF-gelatin composite film as a sandwich layer for full-thickness skin mesh graft in experimental dogs. J Biomed Mater Res B Appl Biomater 2009;88:12e6. [25] Dias GJ, Mahoney P, Swain M, Kelly RJ, Smith RA, Ali MA. Keratin-hydroxyapatite composites: biocompatibility, osseointegration, and physical properties in an ovine model. J Biomed Mater Res A 2010;95:1084e95. [26] Mann FA, Payne JT. Bone healing. Semin Vet Med Surg 1989;4:312e21. [27] Alexander P, Earland C. Structure of wool fibres; isolation of an alpha and beta-protein in wool. Nature 1950;166:396e7. [28] Oest ME, Dupont KM, Kong HJ, Mooney DJ, Guldberg RE. Quantitative assessment of scaffold and growth factor-mediated repair of critically sized bone defects. J Orthop Res 2007;25:941e50. [29] Miller-Chou BA, Koenig JL. A review of polymer dissolution. Prog Polym Sci 2003;28:1223e70.

[30] Ferretti JL, Gaffuri O, Capozza R, Cointry G, Bozzini C, Olivera M, et al. Dexamethasone effects on mechanical, geometric and densitometric properties of rat femur diaphyses as described by peripheral quantitative computerized tomography and bending tests. Bone 1995;16:119e24. [31] Suntornsaratoon P, Wongdee K, Krishnamra N, Charoenphandhu N. Femoral bone mineral density and bone mineral content in bromocriptine-treated pregnant and lactating rats. J Physiol Sci 2010;60:1e8. [32] Figueiredo MM, Gamelas JAF, Martins AG. Characterization of bone and bonebased graft materials using FTIR spectroscopy. In: Theophanides T, editor. Infrared spectroscopy e life and biomedical sciences. Rijeka, Croatia: InTech; 2012. p. 315e38. [33] Gazi E, Dwyer J, Lockyer NP, Gardner P, Shanks JH, Roulson J, et al. Biomolecular profiling of metastatic prostate cancer cells in bone marrow tissue using FTIR microspectroscopy: a pilot study. Anal Bioanal Chem 2007;387:1621e31. [34] Slosarczyk A, Paszkiewicz Z, Paluszkiewicz C. FTIR and XRD evaluation of carbonated hydroxyapatite powders synthesized by wet methods. J Mol Struct 2005;744:657e61. [35] Alexander P, Hudson RF, Fox M. The reaction of oxidizing agents with wool. 1. the division of cystine into two fractions of widely differing reactivities. Biochem J 1950;46:27e32. [36] Saul JM, Ellenburg MD, de Guzman RC, Van Dyke ME. Keratin hydrogels support the sustained release of bioactive ciprofloxacin. J Biomed Mater Res A 2011;98:544e53. [37] Tachibana A, Nishikawa Y, Nishino M, Kaneko S, Tanabe T, Yamauchi K. Modified keratin sponge: binding of bone morphogenetic protein-2 and osteoblast differentiation. J Biosci Bioeng 2006;102:425e9. [38] Ulstrup AK. Biomechanical concepts of fracture healing in weight-bearing long bones. Acta Orthop Belg 2008;74:291e302. [39] Bastian O, Pillay J, Alblas J, Leenen L, Koenderman L, Blokhuis T. Systemic inflammation and fracture healing. J Leukoc Biol 2011;89:669e73. [40] Kwong FN, Harris MB. Recent developments in the biology of fracture repair. J Am Acad Orthop Surg 2008;16:619e25. [41] Sando L, Kim M, Colgrave ML, Ramshaw JA, Werkmeister JA, Elvin CM. Photochemical crosslinking of soluble wool keratins produces a mechanically stable biomaterial that supports cell adhesion and proliferation. J Biomed Mater Res A 2010;95:901e11. [42] Devescovi V, Leonardi E, Ciapetti G, Cenni E. Growth factors in bone repair. Chir Organi Mov 2008;92:161e8. [43] Fiedler J, Roderer G, Gunther KP, Brenner RE. BMP-2, BMP-4, and PDGF-bb stimulate chemotactic migration of primary human mesenchymal progenitor cells. J Cell Biochem 2002;87:305e12.