Views of the National Institutes of Health

ABSTRACT

EVALUATION OF BONE MORPHOGENETIC PROTEIN-2 RELEASE FROM KERATIN SCAFFOLDS IN VITRO AND IN VIVO

by Jingxuan Li

Recombinant human bone morphogenetic protein-2 (rhBMP-2) can be used clinically to promote bone healing as an alternative to bone grafting treatment. The rhBMP-2 can stimulate cellular differentiation of osteoprogenitor cells to promote bone healing. However, delivery of rhBMP-2 is a challenge since rhBMP-2 has a short half-life and has therefore been delivered from collagen sponges implanted at the injury site. While this has led to effective bone regeneration, ectopic bone growth associated with the rapid degradation of collagen and subsequent rhBMP-2 release are clinical problems. We are investigating keratins as alternative rhBMP-2 carriers. Keratins are structural intermediate proteins and can be extracted from human hair. Oxidatively extracted keratin (KOS) cannot achieve disulfide crosslinks whereas reductively extracted keratin (KTN) can form disulfide crosslinks. The rate of degradation of keratin can be tuned by mixing keratose and kerateine in varying ratios. The hypothesis guiding this thesis is that keratin can be formulated with varying ratios of KOS and KTN to modulate the rate of scaffold degradation and thereby control the releasing rate of rhBMP-2. The in vivo release kinetics of rhBMP-2 was assessed by a critically-sized rat femur defect model. The biodistribution of rhBMP-2 after implantation in the critically-sized femur model was assessed in the vital organs.

EVALUATION OF BONE MORPHOGENETIC PROTEIN -2 RELEASE FROM KERATIN SCAFFOLDS IN VITRO AND IN VIVO

A Thesis

Submitted to the Faculty of Miami University in partial fulfillment of the requirements for the degree Master of Engineering Department of Chemical, Paper and Biomedical Engineering

Jingxuan Li

MIAMI UNIVERSITY

Oxford, Ohio

2016

Advisor ______Justin M. Saul

Reader ______Lei Kerr

Reader ______Michael Robinson

TABLE OF CONTENTS

CHAPTER I-Introduction to Bone Regeneration and Keratin Biomaterials ...... 1 I.1. The Need for Effective Clinical Treatment for Bone Fracture ...... 2 I.2. Bone Tissue and Fracture Repair ...... 2 I.3. Bone Grafting: Allograft and Autograft ...... 4 I.4. BMP-2 and Osteoinduction ...... 5 I.5. A Current Clinical rhBMP-2 Carrier: Collagen ...... 6 I.6. Keratin Biomaterial as rhBMP-2 Carriers ...... 6 I.7. Rationale for Use of Fluorescent Quantification of rhBMP-2 Release ...... 9 I.8. Fluorescent Dyes for Quantification of rhBMP-2 Release ...... 10 I.9. Rationale, Hypothesis and Approach of Thesis Work ...... 10

CHAPTER II – Release Kinetics and Biodistribution of rhBMP-2 following Implantation of Keratin Scaffolds in a Critically-sized Rat Femur Defect Model ...... 17 II.1. Introduction ...... 18 II.2. Material and Methods ...... 19 II.2.1. Fluorescent Labeling of rhBMP-2 ...... 19 II.2.2. Keratin Hydrogel Fabrication ...... 21 II.2.3. In vitro rhBMP-2 Release Study ...... 21 II.2.4. Rat Femur Defect Model for Release of Fluorescently-tagged rhBMP-2 In Vivo ...... 22 II.2.5. Bruker System Imaging and Photoshop Quantification of Images ...... 24 II.2.6. Assessment of Fluorescently-tagged rhBMP-2 in Vivo Biodistribution ...... 25 II.3. Results/ Discussion ...... 25 II.3.1. In vitro rhBMP-2 release from keratin hydrogels and collagen scaffolds ...... 25 II.3.2. Rat Femur Defect Model for Retention of rhBMP-2 at Implant Site ...... 29 II.3.3. Biodistribution of rhBMP-2 in Vivo ...... 32

CHAPTER III – Discussion of Results and Future Work ...... 40 III.1. Discussion of Thesis Results ...... 41 III.2. Future Directions ...... 48

iii LIST OF TABLES

CHAPTER II – Release Kinetics and Biodistribution of rhBMP-2 following Implantation of Keratin Scaffolds in a Critically-sized Rat Femur Defect Model

Table 1. Experimental keratin formulation groups. Table 2. Biomaterial formulations for each time points for rats femur defect model for assessing retention of fluorescently-labeled rhBMP-2 in vivo

Table 3. 15% (weight/volume) 100%KOS Standard Curve with serial rhBMP-2 concentration.

Table 4. Statistical analysis of keratin degradation for samples with rhBMP-2 in vitro.

Table 5. Statistical analysis of keratin degradation for samples without rhBMP-2 in vitro

Table 6. Number of fluorescent signal positive organs over the total number of each kind of organ of rats with rhBMP-2 loaded collagen, 100%KOS, 50:50 KOS:KTN and 100%KTN scaffold implantation at Day 1 after implantation.

Table 7. Number of fluorescent signal positive organs over the total number of each kind of organ of rats with rhBMP-2 loaded collagen, 100%KOS, 50:50 KOS:KTN and 100%KTN scaffold implantation at Day 3 after implantation.

Table 8. Number of fluorescent signal positive organs over the total number of each kind of organ of rats with rhBMP-2 loaded collagen, 100%KOS, 50:50 KOS:KTN and 100%KTN scaffold implantation at Day 7 after implantation.

iv LIST OF FIGURES

CHAPTER II – Release Kinetics and Biodistribution of rhBMP-2 following Implantation of Keratin Scaffolds in a Critically-sized Rat Femur Defect Model

Figure 1. Initial standard curve of absorbance with AF488 tagged rhBMP-2 concentration

Figure 2. rhBMP-2 percentage releasing profiles of 100:0 KOS:KTN, 70:30 KOS:KTN, 50:50 KOS:KTN, 30:70 KOS:KTN, 0:100 KOS:KTN and collagen biomaterials

Figure 3. Standard curve of absorbance with diluted BSA concentration for Lowry protein assay (n=3).

Figure 4. Lowry protein assay cumulative protein mass profiles of 100:0 KOS:KTN, 70:30 KOS:KTN, 30:70 KOS:KTN, 0:100 KOS:KTN and collagen biomaterial with or without rhBMP-2

Figure 5. Three kinds of images from Bruker Imaging System for each rats: Fluorescent image (A), optical image (B) and X-ray (C) for experimental rats.

Figure 6. Fluorescent images for 15% (weight/volume) rhBMP-2 loaded KOS scaffold standard curve are shown with serial rhBMP-2 concentration for quantification

Figure 7. Standard curve of serial concentration of rhBMP-2 that loaded into KOS scaffolds (15% weight/volume)

Figure 8. Representative fluorescent images of the average pixel intensity of rhBMP-2 loaded collagen, 100%KOS, 50:50 KOS:KTN and 100%KTN biomaterial at 1, 3 and 7 days post-implant

Figure 9. Percentage rhBMP-2 retention from collagen, 100%KOS, 50:50 KOS:KTN and 100%KTN scaffold at 1, 3, or 7 days post-implant.

Figure 10. Biodistribution fluorescent image for spleen of each rat, which implanted with rhBMP-2 loaded collagen, 100%KOS, 50:50KOS:KTN and 100%KTN at 1, 3, or 7 days post-impalnt

v Figure 11. Biodistribution fluorescent image for liver of each rat, which implanted with rhBMP-2 loaded collagen, 100%KOS, 50:50KOS:KTN and 100%KTN at 1, 3, or 7 days post-implant.

Figure 12. Biodistribution fluorescent image for left lung (A) and right lung (B) of each rat, which implanted with rhBMP-2 loaded collagen, 100%KOS, 50:50KOS:KTN and 100%KTN at 1, 3, or 7 days post-implant.

Figure 13. Biodistribution fluorescent image for left kidney (A) and right kidney (B) of each rat, which implanted with rhBMP-2 loaded collagen, 100%KOS, 50:50KOS:KTN and 100%KTN at 1, 3, or 7 days post-implant.

Figure 14. Biodistribution fluorescent image for heart of each rat, which implanted with rhBMP-2 loaded collagen, 100%KOS, 50:50KOS:KTN and 100%KTN at 1, 3, or 7 days post-implant.

Figure 15. Biodistribution fluorescent image for brain of each rat, which implanted with rhBMP-2 loaded collagen, 100%KOS, 50:50KOS:KTN and 100%KTN at 1, 3, or 7 days post-implant.

Figure 16. Quantification of rhBMP-2 concentration in spleen for collagen, 100%KOS, 50:50 KOS:KTN and 100%KTN implanted rats at 1, 3, or 7 days post-implant.

Figure 17. Quantification of rhBMP-2 concentration in liver for collagen, 100%KOS, 50:50 KOS:KTN and 100%KTN implanted rats at 1, 3, or 7 days post-implant.

Figure 18. Quantification of rhBMP-2 concentration in lungs for collagen, 100%KOS, 50:50 KOS:KTN and 100%KTN implanted rats at 1, 3, or 7 days post-implant.

Figure 19. Quantification of rhBMP-2 concentration in kidneys for collagen, 100%KOS, 50:50 KOS:KTN and 100%KTN implanted rats at 1, 3, or 7 days post-implant.

vi ABBREVIATIONS

KOS: Keratose

KTN: Kerateine rhBMP-2: Recombinant Human Bone Morphogenetic Protein-2 rhBMP-7: Recombinant Human Bone Morphogenetic Protein-7

PBS: Phosphate Buffered Saline

TGF-β: Transforming growth factor-β

FGF-2: Fibroblast growth factor-2

PDGF: Platelet-derived growth factor

VEGF: Vascular endothelial growth factor

MCSF: Macrophage colony stimulating factor

TNF-α: Tumor necrosis factor-α

BSA: Bovine Serum Albumin

IACUC: Institutional Animal Care and Use Committee

vii

ACKNOWLEDGEMENTS

This thesis work on the in vitro and in vivo study of rhBMP-2 loaded to keratin biomaterial to finish my master degree and summarizes the study of previous academic years. First of all, I’d like to thank my advisor, Dr. Justin M. Saul, who gave me an opportunity to join the amazing bone regeneration program and also be the great guidance on my research work.

Thanks to my committee member Dr. Michael Robinson and Dr. Lei Kerr for providing me help and guidance on my thesis work and inspiring me on my thesis writing.

Thanks to my lab members, Judy Strassburger-Bohnert, Salma Haque and BianBian Xiong. You have been the “savior” to me during the past two years. Thanks Judy, who taught and helped me on lab techniques and for being a great partner on rat imaging. Thanks Salma, who helped on my research. Thanks to BianBian, who is the best assistance on everything.

Thanks to Katie LaSance (University of Cincinnati), who helped me obtain the in vivo ratfluorescent images critical to this thesis. We worked on the imaging together at University of Cincinnati Vontz Imaging Center many times and you also let me learn a lot on traditional American culture.

At the last, I would like to thanks to my parents give my support on oversea study. Thanks to my friends in Oxford. You gave warmlike family and we did have fun together in this Oxbox.

This thesis work was supported by the National Institutes of Health (JMS; R01AR061391) and the content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

viii

CHAPTER I – Introduction to Bone Regeneration and Keratin Biomaterials

Jingxuan Li

1 I.1 The Need for Effective Clinical Treatment for Bone Fracture New strategies for the repair critically-sized segmental bone defects is an urgent need for clinicians since bone injuries can have a highly deleterious impact on everyday life for patients and society. Approximately 500,000 patients undergo surgical procedures for bone fracture every year. In the adult human body, bone tissue has the ability to regenerate spontaneously, unless the defect exceeds a certain limit in size [1]. In bone, a defect that will not heal spontaneously without intervention is termed as “critically-sized defect”. In the classical definition, a critically-sized defect is the smallest size tissue defect that will not completely heal over the natural lifetime of an animal or human beings. Among the approximately 6.5 million fractures suffered in United States each year, approximately 15% have delayed or non-union [2]. The delayed union is the bone fracture does not heal in the expected length of time, but which is still progressing toward healing while a non-union is a fracture that fails to heal [3]. Thus, an effective treatment for bone healing in normal period time is urgent. To-date, for most of these difficult cases there is no therapy that is clinically satisfying. An increasing number of grafting procedures combined with less-than-desirable outcomes for current autograft treatments in orthopedic surgery have led to a quest for alternative methods to reconstruct critically-sized bone defects. Before discussing strategies that promote bone regeneration in critically-sized defects, it is important to first understand the native bone tissue healing process.

I.2 Bone Tissue and Fracture Repairing Bone is a highly dynamic tissue with complex structure and functions that are regulated by the interactions between cells, extracellular matrices, biomechanical forces, and gene and/or protein regulatory factors. For the bone structure, the outer-most layer of the bone is the periosteum, which contains osteoprogenitor and osteoblast cells (among others), which are recruited to the defect site during the bone repair process [4]. Between the periosteum and the bone marrow is bone tissue that includes cancellous and cortical bone. Cancellous bone provides structural support and organization while bone marrow is interspersed inside. Cortical bone surrounds the cancellous bone to give mechanical strength to bone [4]. Within the cortical bone are osteons that contain osteocytes, which are mature bone cells, and the Haversian canal in which blood vessels and nerves reside. The osteogenic progenitor is driven from mesenchymal cells and can later develop or differentiate into osteoblast cells. The osteoblast is the dominant player in bone regeneration and contributes to the maintenance of extracellular matrix for bone structural strength [5].

2 Fracture repair is a complicated process that involves a coordinated cascade of biological events. Typically, bone healing is partitioned into three stages: (1) the initial inflammatory response, (2) the reparative phase, and (3) the remodeling phase. From the cellular level, the main players are inflammatory cells, vascular cells, osteoprogenitor cells and osteoblasts [4]. As to the molecular level, fracture healing operates by pro-inflammatory cytokines and growth factors, pro-osteogenic factors and angiogenic factors [6]. These factors and cells establish a morphogenetic gradient to stimulate bone growth. During the inflammatory phase, the damaged soft tissue will be repaired and the soft callus and later hard callus will give a bridge at fracture site. Then the hard callus will eventually be remodeled and reorganized to the bone geometry and function of the damaged tissue [7]. The first stage is inflammation that mainly involves the secretion of a series of cytokines and bone growth factors including transforming growth factor-β (TGF- β), platelet-derived growth factor (PDGF), fibroblast growth factor-2 (FGF-2), vascular endothelial growth factor (VEGF), macrophage colony stimulating factor (MCSF), interleukins-1 and -6 (IL-1 and IL-6), bone morphogenetic proteins (BMPs), and tumor necrosis factor-α (TNF- α) [8][9]. These secretions will lead to a positive feedback and the osteoprogenitor cells or mesenchymal stem cells, which were recruited by the inflammatory cascade in periosteum and/or bone marrow, will be the starting point for bone regeneration in the repair phase [10][11]. In the second stage (repair), the growth factors TGF-β2 and TGF-β3, PDGF, fibroblast growth factor-1 (FGF-1), and insulin-like growth factor (IGF), among others, are secreted [8][9]. The key molecules in promoting cell proliferation and chondrogenesis are the BMP family (in particular, BMP-2, -4, -5 and -6). These growth factors have a role in initiating, promoting, and maintaining chondrogenesis and osteogenesis [12]. The bone growth factor has able to stimulate chondrocyte differentiation and fibroblasts proliferation in order to provide a mechanical support in forming the semi-rigid soft callus. During the second stage the soft callus is systematically remodeled followed by a period of osteogenesis that occurs do to high levels of osteoblast activity leading to bone matrix formation and bridging with the new hard callus [11]. The last stage of healing involves the hard callus remodeling to the bone into original configuration. The remodeling cascade includes three stages (activation, resorption and formation) for immature bone remodeling. The remodeling phase occupies 70% of the total bone repair time [13]. When the osteoclast, a large multinucleate bone cell that absorbs bone tissue during growth and healing, and osteoblast at the woven bone intracortical envelope are activated by hormonal or physical stimuli,

3 the remodeling phase will be initiated. At this time, the osteoclasts resorb tissue, allowing the osteogenic cells to migrate or differentiate into osteoblasts [14]. The osteoblasts differentiate, become polarized, and adhere to the mineralized surface to form a border of the bone [12]. As the result, the functional unit, osteon, of compact bone will be formed [15]. However, in some instances, which may be due to large injury, genetic defect, or other causes, the normal healing does not occur, leading to non- union. In the case of non-union, alternative methods of repair are required.

I.3 Bone Grafting: Allograft and Autograft

Bone grafting is a surgical process to implant tissue or other tissue substitutes into the defect or the spaces around the broken bone in order to promote healing and/or stability to the fracture site. There are three properties required in bone graft processes: osteogenesis, osteoinduction and osteoconduction [16]. The new bone formation by cells from the host or the graft is defined as osteogenesis. Osteoinduction describes the induction of the mesenchymal stem cells from the graft or the host tissue to differentiate into chondroblasts or osteoblasts [16]. During the recruitment and differentiation of mesenchymal stem cells, the growth factors like BMP's play a role in. Osteoconduction indicates the three dimensional structure of the graft, which can facilitate the ingrowth of capillaries and mesenchymal stem cells to promote new bone formation on the graft [8][9]. Traditionally, there are two types of bone grafting used in the clinic: allograft and autograft. The allograft is the transfer bone from another human source (typically cadaver) into the patient’s bone defect. Before the implantation, allograft bone needs to be sterilized. The allograft bone material can proceed by several modifications including low dose irradiation, antibiotic washing and physical debridement in order to reduce the antigenicity of the bone graft [18]. Further sterilization may be achieved through gamma irradiation, electron beam irradiation or ethylene oxide treatment [19]. However, these sterilization methods will typically result a dose dependent decrease in the mechanical properties of the graft [20]. For clinical use, the implantation grafts are either frozen or freeze dried. Although the frozen bone has minimal effect on the structural properties, it does not greatly reduce the immunogenicity of the graft. Demineralized bone matrix (DBM) consists of most of the non-mineralized components of bone [21]. The DBM obtained from allograft bone undergoes acid extraction in order to remove the mineral components of bone. As the result, the remaining components give allograft bone its osteoinductive potential ability. These components still maintain the collagen trabecular structure with its osteoconductive properties [22]. The DMB can lead to rapid revasularization and the releasing of

4 local cytokines and growth factors that can recruit mesenchymal stem cells for bone formation. DMB is an alternative to autograft, but without donor-site morbidity. However, DBM does not have the same osteoinductive potential as autografts [21]. The autograft, which is histocompatible, is bone tissue taken from the patient’s own healthy bone, typically at the iliac crest of the hip. There are three types of autograft: cancellous, cortical or a combination of cortical and cancellous. Both autologous cancellous and cortical grafts can promote cells capable of bone formation, but cancellous autografts with their trabecular structure lined with osteoblasts and large surface area set a good condition for much more potent osteogenesis [22]. Cancellous autograft can be vascularized and attract the host stem cells to the graft site in low oxygen content quickly [23]. However, cancellous autograft lacks significant structural qualities. Cortical autografts can provide structural support and supply osteoblasts. However, the cortical autograft is used less frequently due to donor site morbidity. For the cortical autograft, the incorporation stimulates osteoclasts, which resorb the dense cortical structure before accelerated revascularization [23] and this can achieve to a 75% reduction in the strength of the graft [24]. Although the bone grafting can repair fractures, there are several risks that are limitations of this operation. These include reaction to medications, bleeding, infection, host immune response, risk of disease transmission and pain at the explant site [25].

I.4 BMP-2 and Osetoinduction

An alternative to autografts and allografts is the use of so-called synthetic bone grafts based the bone morphogenetic proteins (BMPs), which have been shown to induce bone formation. In the 1960s, Urist discovered BMPs, a milestone in strategies to promote bone regeneration. The initial application of this discovery was the use of devitalized, demineralized bone matrix, which contained bioactive proteins that could promote regeneration [26]. Since this discovery, many other growth factors have been found to play a significant role in bone healing, as noted above in the natural healing process. Since many investigators have shown that BMPs are able to induce mesenchymal stem cells to differentiate to osteogenic cells, which can produce bone, much research has focused on members of bone morphogenetic protein (BMP) family. Two commercially available BMP-based products have been approved by the Food and Drug Administration (FDA) since 2002: BMP-2 and BMP-7. Each of these is approved for specific orthopedic indications [27]. Recombinant human protein (rhBMP-2) is currently available for orthopaedic usage in

5 the United States [28]. The rhBMP-2 can stimulate cellular differentiation of osteoprogenitor cells toward an osteoblast phenotype to promote bone healing [1]. However, BMP-2 has a short biological half-life in vivo (6.7 min in non-human primates and 16 min in rats), because BMP-2 is broken down by enzymes, which will lead to rapid clearance and non-localized actions [29][30]. For this reason, a carrier system to achieve implantation and controlled release in vivo is needed.

I.5 A Current Clinical rhBMP-2 Carrier: Collagen

The INFUSE® Bone Graft consists of rhBMP-2 adsorbed to an absorbable collagen sponge (ACS), and is a product sold by Medtronic®. This product serves as an off-the-shelf synthetic bone graft alternative to autografts and allografts. Conceptually, the rhBMP-2 is delivered to the defect site by incorporation into a delivery system (ACS) that gradually releases the rhBMP-2 to promote localized bone formation [31][32]. Indeed, the Infuse product has been a considerable commercial success, with Medtronic generating $300 million in annual sales from the BMP products in 2014 [33]. The delivery vehicle should not only maintain BMP-2 concentration within the defect site for a sufficient period of time for osteoprogenitor cells migrating to the target-healing site and differentiate into osteoblasts, but it also needs to be a biological and biomechanical compatible framework to enhance bone formation (i.e., an osteoconductive “scaffold” material). The current FDA-approved carrier for BMP-2 delivery is bovine collagen [34].

However, collagen sponges exhibit burst release of the rhBMP-2 following implantation and collagen degrades in vivo within several days. Although approved for lumbar spinal disk fusion and tibial fractures, rhBMP-2 loaded collagen carriers have given rise to safety concerns when used for off- label applications such as cervical spinal fusion [35] such as ectopic bone growth [1]. Collagen is resorbed within several days due to the activity of collagenase enzymes. Thus, although useful, collagen may not be the ideal carrier to promote bone healing [34]. Due to these problems associated with collagen sponges as BMP-2 carriers, several alternative carrier systems are being widely investigated. Ideally, the carrier would have a degradation rate that could be modified to match the rate of bone formation/regeneration [35].

I.6 Keratin Biomaterials as BMP-2 Carriers

It has been shown that BMP-2 requires being loaded to a biomaterial matrix to attain maximal

6 efficacy. In two previous in vivo studies, our team has evaluated keratin as the carrier for rhBMP-2 to promote bone regeneration. Keratins are a family of proteins that can be isolated from various sources including human hair (the source material used in this thesis) [36]. Extraction of keratin and further processing of the material allows it to be fabricated it into a variety of biomaterial forms that may be useful for bone regeneration or other systems including hydrogels, films, scaffolds, or fibers [36]. Collagen can be eroded by collagenase enzymes, whereas the human body dose not has keratinase enzymes that would break down keratin in an analogous fashion. This suggests that keratin biomaterials can be expected to exhibit longer-term erosion than collagen, which has been verified in previous studies [37][38]. Moreover, keratin has a high percentage of cysteine residues in its primary protein structure. Depending on the extraction technique, resulting disulfide crosslinking can also contribute to a slow erosion rate [39]. The keratin extracted from human hair can be classified into three types: alpha-, beta- and gamma-keratin. The alpha-keratin, which resides in the hair fiber cortex, has alpha-helical secondary structure with low sulfur content and 60-80 kDa molar mass. The beta-keratin, which is difficult to be extracted is primarily protective and form the majority of the cuticle. However, the beta- keratin dose not form especially useful reconstituted structures and has not been used in this thesis work. The gamma-keratin is globular with high sulfur content and lower molar mass, which around 15kDa. The sulfur content is higher in gamma keratins than in alpha keratins [40].

In order to be used, keratin proteins must be extracted from the hair. Two general extraction methods are typically used: oxidative extraction or reductive extraction. The keratins are removed from the cortex by use of chemicals in order to break the disulfide bonds, which are prevalent in keratinized tissues. Then the alpha and gamma keratin convert to the non-cross-linked forms by oxidation or reduction, with which cysteine is converted to either cycteic acid or cysteine, respectively. The solution contains extracted protein and denaturing solvent can be purified by filtration and dialysis [41]. If using oxidative extraction carries out the extraction, the cysteic acid is found on the sulfur atoms of the cysteine residues and the resulting form of keratin is referred to as keratose (KOS). If the reductive extraction is used, the cysteine residues have thiol groups capable of forming disulfide crosslinks, and this form of keratin is referred to as kerateine (KTN). Stated differently, keratose is the form of keratin that results from the oxidative extraction. This method results in sulfonic acid groups on the cysteine residues, which prevents disulfide crosslinking [41]. Kerateine is the form of keratin obtained by reductive extraction and leaves the thiol groups available to form disulfide crosslinks. Thus, the KTN is capable of forming disulfide crosslinks between cysteine residues [41].

7 The chemical state of the cysteine residues in KOS and KTN then affects the presence of crosslinking in hydrogels or scaffolds (freeze-dried hydrogels) prepared from the extracted proteins. Biomaterials (hydrogels or scaffolds) formed from KOS have chain entanglements, but lack covalent crosslinks, leading to rapid erosion [42]. Conversely, biomaterials formed from KTN have both chain entanglements and covalent disulfide crosslinks, leading to slower rates of erosion. In other words, keratiene can form disulfide cross-links but keratose does not present a covalently cross-linked network structure [36]. The fully disulfide crosslinking gives it is more stability and disulfide bonds can also be re-crosslinked through oxidative coupling of cysteine groups. Because of the polarization of the backbone of keratose caused by the electron withdrawing properties of sulfonic acid residues on the cysteine residues it cannot form disulfide crosslinks. The keratose is oxidatively derived keratin and the oxidative reaction. Recently, our group has shown the ability to modulate the rate of keratin biomaterial degradation by two different methods that regulate the levels of disulfide crosslinking within keratin materials [37]. In one method, rhBMP-2 interacted with keratins modified with alkyl groups at cysteine residues to achieve different degrees of disulfide crosslinking. The chemical modification of keratin proteins was achieved by alkylation with iodoacetamide. The modification process to KTN changed the binding affinity to different growth factors based on their intrinsic physical or chemical structure [37]. However, in a recent study by our group the crosslinking density of a hydrogel material can be regulated simply by mixing different ratios of KOS and KTN [43], which is a simpler process compared to the alkylation method noted in the previous paragraph. Our team assessed both the material properties and biological response to keratin formulations of KOS-KTN mixture hydrogels. The results showed that hydrogels composed of the two forms of keratin could be used to modulate their degradation profiles. Such control over degradation is important in tissue engineering approaches to control the growth factor release, provide for drug delivery, and ultimately allow for tissue regeneration. The alkylated keratins or keratin hydrogel mixtures have been used for the delivery of insulin-like growth factor 1 (IGF-1) and ciprofloxacin. These previous results suggested that keratin hydrogels may serve as a carrier system for treatment of bone injuries through rhBMP-2 delivery [37][39]. In this thesis work, will build on the use of these tunable-degradation rate keratin materials to investigate effects on in vitro rhBMP-2 release, retention of rhBMP-2 in a critically-sized rat femur defect model, and to assess the biodistribution of rhBMP-2 following release from keratin or collagen biomaterials in the same femur defect model. In order to conduct these studies, we have elected to

8 fluorescently label rhBMP-2 in order to follow its release both in vitro and in vivo.

I.7 Rational for Use of Fluorescent Quantification of rhBMP-2 Release

In this study, we have used fluorescently labeled rhBMP-2. With such labeled rhBMP-2, it is possible to visualize and quantitatively track rhBMP-2 release over time. There are several methods available to quantify rhBMP-2 release: radiolabeling, ELISA (enzyme-linked immunosorbent assay) and fluorescence. We describe each of these and provide our rationale for the use of fluorescence labeling below.

125I (iodine-125) is commonly used for labeling proteins, usually at tyrosine residues. Some radio nuclei are synthesized in particle accelerators and have short half-lives and lower the detection time [44]. However, the radioactivity is problematic for the specific surgical technique since the radiation protective casing is indispensable in these studies and the short half-life would pose challenges if long time frames could be to be monitored. In this thesis study, the 7 days is the longest times point for monitoring. Initially it was thought that rhBMP-2 release may occur over as long as 16 weeks, which would be two half-lives of 125I. So, even without any release, only ¼ of the signal would be present at the end of the study. In the result the weak signal detection could not present quantification accurately. While our results indicate that release occurred on a much shorter timescale, this was only determined later (see Chapters II and III for further discussion).

ELISA is a test that uses antibodies and color change to identify a substance. One major disadvantage with ELISA is that it requires washing and incubations between additions of reagents. That can substantially increase the hands-on time required to run this type of assay [45]. In addition, the detection of BMP-2 for solid in vivo samples could prove challenging as homogenization and extraction could be required. Further, these processes could degrade some of the rhBMP-2 and make its quantification impossible.

We, therefore, believed that a trackable quantification (through fluorescence) of rhBMP-2 would be more straightforward and informative for this study. Fluorescent molecules, also called fluorophores or simply fluors, respond distinctly to light. The use of fluorescent molecules in biological research is beneficial and reliable in many applications, and their application is continually increasing due to their low cost, versatility, sensitivity and suitability for quantitation. Among their myriad of uses, fluorescence dyes are employed for the detection of protein, tracking activation and quantification in

9 conformational changes, and monitoring biological processes in vivo. The fluorescent labeling is the chemical process of covalently attaching a fluorophore to target molecule, usually are protein or nucleic acid (rhBMP-2 in the case of this thesis work). This is generally achieved by using a reactive derivative of the fluorophore that selectively binds to the functional group contained in the target molecule [46]. Although fluorescent labeling may have some effects on rhBMP-2 interaction with the keratin (or collagen control) materials, it provides a rapid and reliable method for of rhBMP-2 release from keratin hydrogel formulations both in vitro and in vivo. For in vitro studies, rhBMP-2 release was be determined by fluorescence spectroscopy while rhBMP-2 was quantified in vivo with a Bruker Image System at the University of Cincinnati.

I.8 Fluorescent Dyes for Quantification of rhBMP-2 Release

For the fluorescent theory, when an electron of a fluorescent particle absorbs a photon of excitation light, the energy level of the electron will be raised to a higher energy excited state, which is an energy absorbing process. During this short excitation period, part of the energy will dissipate through molecular collisions or be transferred to a proximal molecule, and as a consequence the remaining energy will be emitted as a photon to relax the electron back to the ground state where is the lower energy level. The fluorescence signal will be coupled by the energy loss, which is from a singlet- excited state to the ground state [47]. Because the emitted photon usually carries less energy and therefore has a longer wavelength than the excitation photon, the emitted fluorescence can be distinguished from the excitation light [48]. The specific fluorophore has its particular wavelength, which corresponds to the maximum and minimum excitation and emission signal intensity. Alexa Fluor® dyes is a widely used dye due to high fluorescent yield. Due to the wide range of Alexa Fluor® dyes there are many options for multiplex detection. This series of superior fluorophores, spans from near-UV, visible and near-IR spectrum. We elected to use Alexa Fluor® 488 dye for in vitro release studies due to the stability and fluorescence intensity of this fluorophore. For in vivo studies we used the DyLight 800 dye due to its near IR emissions and also minimized the muscle tissue or fur autofluorescence interference since Dylight800 have large wavelength.

I.9 Rational, Hypothesis and Approach of Thesis Work Our team has developed a technique to exploit the differences in keratose and kerateine in order to tune the rate of hydrogel degradation simply by modification of the amounts of keratose (more rapidly degrading) and kerateine (more slowing degrading) included in formulations for keratin materials (e.g.,

10 hydrogels or scaffold materials). We have previously observed that tuning the rates of degradation is a method that can be used to control the rate of release of various therapeutic agents including antibiotics and growth factors [37][39]. Thus, by modifying the composition of keratin to modulate disulfide crosslinking there is the possibility to control the rate rhBMP-2 release. The goal of this thesis is to make use of these previous findings to assess the effects of tunable keratin degradation on rhBMP-2 delivery both in vitro and in vivo. With the fluorescently labeled rhBMP-2, it is possible to visualize and quantitatively track rhBMP-2 release over time. The hypothesis guiding this thesis is that keratin can be formulated with varying ratios of KOS and KTN to modulate the rate of scaffold degradation and thereby control the rate of rhBMP-2 delivery. The complexity of the proposed animal model precludes use of every cross-linked formulation. Thus, several formulations were selected to span the range of possible effects from no covalent cross-linking (KOS) to full cross-linking (KTN). We specifically tested the hypothesis with two aims in this thesis.

Aim 1. Assess release rate of fluorescently-tagged rhBMP-2 from keratin biomaterials with different disulfide crosslinking density in vitro.

This aim was assessed by generating keratin hydrogels of varying disulfide crosslink densities (achieved by mixing KOS and KTN in varying ratios) and loaded with fluorescently-tagged (AF488) rhBMP-2. Release was determined by elution of the fluorescently-tagged rhBMP-2 into phosphate- buffered saline (PBS) placed on top of the gels and subsequently assayed with a fluorescence plate reader. The effects of keratin cross-linking modification on rhBMP-2 release was examined by two techniques fluorescence spectroscopy and was correlated to keratin protein degradation by assaying keratin by a Lowry protein assay.

Aim 2. Assess the in vivo release kinetics and biodistribution of rhBMP-2 following implantation of keratin scaffold in a critically-sized segmental rat femur defect model.

This aim is to investigate release kinetics of fluorescently-tagged (DyLight 800) rhBMP-2 loaded into keratin scaffolds of varying disulfide crosslink densities within a rodent model of a critically-sized defect. The rhBMP-2 loaded keratin scaffold was implanted into a critically-sized femur defect in living

11 rats and assessed at specified time points on a Bruker imaging system. The biodistribution of rhBMP-2 was assessed in major organs (brain, liver, kidney, heart, spleen, and lungs) by explanting these tissues at the terminal time points of the study and imaging on the Bruker Imaging System for detection of the presence of fluorescently-tagged rhBMP-2.

The objective of this thesis related to the keratin biomaterial characterization of rhBMP-2 delivery, which can provide a new strategy for controllable rhBMP-2 release in critically-sized bone defects. Thus, the in vivo study of keratin biomaterial characterization of rhBMP-2 may provide a foundation for future preclinical testing. Assessing rhBMP-2 biodistribution in vivo is the important guideline for given the large dose of rhBMP-2 in preclinical/clinical use.

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CHAPTER II – Release Kinetics and Biodistribution of rhBMP-2 following Implantation of Keratin Scaffolds in a Critically-sized Rat Femur Defect Model

Jingxuan Li

17 II.1. Introduction

Keratins are structural intermediate filament proteins and can be extracted from human hair. There are three broad classifications of keratin: alpha, beta and gamma keratins [1]. Beta keratins may be most well known for forming the harder structures of the hair cuticle and are not discussed further in this thesis. Gamma keratins are lower molecular weight proteins that have higher cysteine content [1], but these keratins were not explored in this thesis in order to focus on the role of the more structurally important alpha keratins. Alpha keratins are characterized by a high molecular weight and alpha helical structure. Two techniques for extraction of keratin from human hair are widely used: oxidative and reductive extraction [2]. Kerateine (KTN) is the form of keratin obtained by reductive extraction while keratose (KOS) is the form obtained by oxidative extraction. Kerateine has thiol groups capable of forming disulfide crosslinks, but keratose has sulfonic acid groups on the cysteine residues following extraction, which prevents formation of a covalently cross-linked network structure [3]. Based on this structure, hydrogels composed of KOS degrade more rapidly because it only has chain entanglements whereas hydrogels composed of KTN contain both chain entanglements and S-S crosslinks [1]. In a previous studies, we found that KOS and KTN had very different rates of degradation in vitro and also different rates of release for several molecules including vascular endothelial growth factor (VEGF), insulin-like growth factor 1 (IGF-1) and basic fibroblast growth factor (bFGF) with KOS leading to more rapid delivery than KTN [4]. Subsequently, our team has developed a technique to utilize the differences in keratose and kerateine in order to tune the rate of hydrogel degradation simply by modification of the amounts of keratose (more rapidly degrading) and kerateine (more slowing degrading) included in the formulations of keratin materials such as hydrogels or scaffold [5]. We have also observed, via an alkylation approach to tuning disulfide crosslinking, that modification of disulfide crosslinking can be used to tune the rate of release of several therapeutic agents including antibiotic ciprofloxacin and IGF-1 [6]. Based on these previous results, we were interested in exploring whether modulating the KOS-KTN ratio in keratin biomaterials as a means to modulate disulfide cross-linking could also be used to control the rate rhBMP-2 release. In this chapter, we describe a set of new studies in which we assessed the effects of tunable keratin degradation on rhBMP-2 delivery both in vitro and in a critically-sized rat femur defect model in vivo. To conduct rhBMP-2 release studies and allow quantification of its release (in vitro) or retention (in vivo) or presence in organs (in vivo biodistribution), the rhBMP-2 was fluorescently labeled. Although fluorescent labeling may have some effects on rhBMP-2 interaction with the keratin (or

18 collagen control) materials, it provides a rapid and reliable method for measurement of rhBMP-2 release from keratin hydrogel formulations both in vitro and in vivo. For in vitro studies, rhBMP-2 release was determined by fluorescence spectroscopy after modification of rhBMP-2 with AlexaFluor488. The rhBMP-2 kinetics was quantified in vivo with a Bruker Image System at the University of Cincinnati after the rhBMP-2 was labeled with DyLight 800. Again, as noted in Chapter 1, the hypothesis guiding this thesis is that keratin can be formulated with varying ratios of KOS and KTN to modulate the rate of scaffold degradation and thereby control the rate of rhBMP-2 delivery. The studies described below use both in vitro and in vivo experiments allow us to assess this hypothesis. More specifically, these studies investigated release kinetics of fluorescently tagged rhBMP-2 loaded into keratin hydrogels (in vitro) or scaffolds (in vivo) of varying disulfide crosslink densities. An initial in vitro study was followed by a more substantial in vivo study in a model of non-union, which is referred to here as a critically-sized rat femur defect model. At specified time points, the defect site in animals receiving various treatments containing fluorescently labeled rhBMP-2 was imaged for fluorescence on a Bruker imaging system and compared to standards for known amounts of fluorescently-labeled rhBMP-2. Further, the biodistribution of rhBMP-2 was assessed in other organs by explanting these and again imaging on the Bruker Imaging System. Completion of these studies provides information regarding the effects of varying degradation profiles of keratin hydrogels (based on KOS: KTN ratios) on the rate of rhBMP-2 release. The study also indicates whether released rhBMP-2 is detectable in any of the major organs of the body, which could indicate potential effects at sites distant from the original implant site (femur).

II.2. Materials and Method II.2.1. Fluorescent Labeling of rhBMP-2 The rhBMP-2 (Medtronic) was hydrated with deionized water (DI water) according to the manufacturer’s recommendations with the exception that the amount of water added was used to give a 3 mg/mL final rhBMP-2 concentration. The resulting solution of rhBMP-2 was dialyzed exhaustively in a 3000 Da MWCO dialysis tube against DI water to remove Tris salts containing amine groups. For in vitro studies, Alexa Fluor 488-carboxyl-NHS (AF488, Invitrogen, Eugene, OR) was used to label rhBMP-2, as a filter set was available in our laboratory’s Bio-Tek Synergy platereader to measure this wavelength. In order to reduce the background interference from autofluorescence in vivo (e.g., from skin, fur, or other tissues), a large wavelength dye was selected for rhBMP-2 labeling. For rhBMP-2

19 labeling in rat femur model, DyLight800 (Thermo scientific, Rockford, IL), which is near-IR fluor was selected, as this fluorophore is photostable and gives high levels of fluorescent yield. A similar procedure was used to label rhBMP-2 with AF488 and DyLight 800 for in vitro and in vivo studies, respectively. Both methods used carboxyl-terminated fluorophores to covalently couple to the amine groups of rhBMP-2 by standard methods.

One vial of AF488 or DyLight800 dye was added to roughly 450µL 3mg/mL dialyzed rhBMP-2 solution. The reaction was allowed to proceed for one hour at room temperature. The fluorescent labeled rhBMP-2 was then dialyzed for 3 days with repeated dialysate changes (1-2 L each) against DI water at pH 5. For AF488 labeling, the AF488 labeled rhBMP-2 concentration and labeling efficiency was determined by measuring the absorbance at 280nm and 494nm correlated wavelength for protein and dye with a Synergy plate reader (Bio-Tek, Winooski VT). The protein concentration and labeling efficiency equations are shown in equation (1) and equation (2), respectively, as provided by the manufacturer and where we determined the extinction coefficient of the protein separately (εprotein).

!"# !"#$%"&'() !"#$%&'! !"! !"#$%"&'() !"#$%&'×!.!! ×!"#$%"&' !"#$%& �� = ...... (1) !!"#$%&'

!"! !"#$%"&'() !"#$%&'×!"#$%"&' !"#$%& �� = ...... (2) !",!!!×!"#$%&' !"#!$#%&'%("# (!)

Again, the εprotein represents the protein molar extinction coefficient. Labeling of rhBMP-2 with DyLight800 followed the same methods but different equations were used to determine labeling efficiency. Equation (3) is correction factor evaluation. The protein concentration and labeling efficiency equations shows in equation (4) and equation (5), respectively. In brief, both methods of quantification are similar.

!"# !"#$%"&'() �� = �� = ...... (3) !!! !"#$%"&'()

!"# !"#$%"&'() !"#$%&'! !!! !"#$%"&'() !"#$%&'×!" ×!"#$%"!" !"#$%& �� = ...... (4) !!"#$%&'

!!! !"#$%"&'() !"#$%&'×!"#$%"&' !"#$%& �� = ...... (5) !"#,!!!×!"#$%&' !"#!$#%&'%("# (!)

20 II.2.2. Keratin Hydrogel Fabrication Keratin hydrogels of varying disulfide crosslink densities were formulated as previously described [6]. The typical formulation in this thesis study is 15% keratin hydrogel/scaffold, which means 1mL solution with 150mg keratin powder. Briefly, for keratin hydrogel fabrication, KOS and KTN powders were mixed together in at the ratios indicated in Table 1. Then 1mL sterile water or 1 mL of water containing 100µg/mL fluorescently-tagged rhBMP-2 solution was added to the 150mg keratin powder for control or experimental group for each formulation in order to give gels of 15% (weight/volume). Five keratin formulations were assessed, as shown in Table 1 below.

Table 1. Experimental keratin formulation groups.

Group 1 Group 3 Group 3 Group 4 Group 5 KOS 100%(150mg) 70%(105mg) 50%(75mg) 30%(45mg) 0%(0mg) KTN 0%(0mg) 30%(45mg) 50%(75mg) 70%(105mg) 100%(150mg)

The hydrated keratins were then vortexed, mixed manually (by spatula) to further homogenize the formulation, vortexed again, mixed manually again, centrifuged, and then packed into a 1 mL syringe (no needed). 100µL of the keratin was then added (by injection) to each of three 1.5mL tubes. The actual weight of the gel added to each 1.5 mL tube was determined by weighing the tube before and after addition of the gel.

II.2.3. In vitro rhBMP-2 Release Study In order to better understand the effects of keratin hydrogel degradation rates on rhBMP-2 release, we conducted an in vitro release assay with keratin hydrogels containing rhBMP-2 labeled with AF488. Hydrogels were used in these studies because use of scaffolds (lyophilized hydrogels) led to absorption of the water and inaccurate measurements at early time points (data not shown). We do note that keratin scaffolds were used for in vivo experiments below but we are suggesting that the behaviors of gels and scaffolds (once wetted in vivo) would be similar. The keratin hydrogels were prepared as described above, incubated at 37 oC for 24 hours, and then the keratin hydrogels in 1.5 mL tubes were layered with 150µL phosphate buffered saline (PBS; Invitrogen, Carlsbad, CA). The rhBMP-2 release profile was determined by elution of the fluorescently- tagged rhBMP-2 into PBS placed on top of the gels. At specific times (1.5, 3, 6, 12, and 24 hours then daily for 1 week) PBS was removed from each keratin hydrogel and then 150µL fresh PBS was placed

21 into the 1.5mL tubes on top of the keratin hydrogels. The removed PBS was placed in 1.5mL tubes and stored at -80oC until quantification. At the time of analysis, samples were thawed to room temperature and 100 µL of sample was placed into a well of a 96-well plate. One reading was taken per sample and there were 3 replicates per experimental group (n = 3, experimental triplicate). Fluorescence of rhBMP- 2 in the PBS was assayed on a Synergy plate reader (Bio-Tek, Winooski VT) at excitation/emission wavelengths of 485/528nm in order to obtain rhBMP-2 concentration by comparison to the standard curve. To determine protein content in the eluted samples, a modified Lowry protein assay (Bio-Rad, Hercules, CA) was used. In brief, all collected samples were diluted at 1:10 ratio, which means 45µL of PBS and 5µL of each sample. The standard curve was made with serial dilution of 90 mg BSA with 45 mL PBS (2mg/mL). The standard curve BSA concentrations used were as follows: 2mg/mL, 1mg/mL, 0.5mg/mL, 0.25mg/mL, 0.125mg/mL, 0.0625mg/mL, 0.03125 mg/mL and blank (PBS). 5µL of each standard curve concentration solution or the 1:10 diluted samples were placed in the well of plate. Then 25µL of reagent A and 200µL reagent B were added to each well. The absorbance was then read at 750nm after 20 minutes of incubation at room temperature. Statistical analysis by student t-test was used to evaluate the differences between experimental groups and the control group as well as between each formulation. Data are given as mean ± standard deviation (STD). Probability values of p < 0.05 as determined by t-test were considered to indicate statistical difference.

II.2.4. Rats Femur Defect Model for Assessing Retention of Fluorescently-labeled rhBMP-2 In Vivo In order to investigate rhBMP-2 in vivo release kinetics and biodistribution, the fluorescently- tagged rhBMP-2 keratin scaffold implant to living rats model will be made. The Bruker fluorescent image was then analyzed and quantified to obtain the rhBMP-2 retention at the implant site. Since the collagen material is the clinical standard for rhBMP-2 carriers, collagen was used as a control group in this study. All animal experiments were approved by the Miami University Institutional Animal Care and Use Committee (IACUC). Three formulations of keratin scaffolds were selected for in vivo testing. 100%KOS, 50%: 50% KOS: KTN and 100%KTN were formulated with rhBMP-2 labeled with DyLight 800 (Thermo scientific, Rockford, IL), where rhBMP-2labeling in ratio 1:0.11 with Dylight 800 was achieved by the same methods used for AF488 (Invitrogen, Eugene, OR) described above. Collagen scaffolds (Infuse,

22 Medtronic, Mineapolis, MN) with 100µg/mL DyLight800-labeled rhBMP-2 served as control group because this is the current clinically used material. Keratin scaffolds were prepared as described in Table 2 for 15% (weight/volume) keratin scaffold and collagen with 100 µg/mL. In an initial standard curve to related fluorescence (by the Bruker system) to rhBMP-2 concentration, we used 100% KOS biomaterial with decreasing DyLight 800 labeled rhBMP-2 concentrations as shown in Table 3. Keratin scaffolds were prepared as described above for 15% (w/v) keratin hydrogels with the following exceptions. First, in instead of placing into a 1.5 mL tube, approximately 500 µL was placed into a Tygon tube of ~ 8 mm length. Second, after being allowed to gel at 37oC overnight, the keratin (in the tube) was placed on a Labconco Freezone freeze-drier for at least two days. After freeze-drying, scaffolds were stored under vacuum until the time of implantation. For control, the collagen scaffolds went through 15 minutes incubation under 37 oC.

Table 2. Biomaterial formulations for each time points for rats femur defect model for assessing retention of fluorescently- labeled rhBMP-2 in vivo

Time Point Biomaterial Formulations 1 Day (n=3) Collagen 100%KOS 50:50 KOS:KTN 100%KTN 3 Day (n=3) Collagen 100%KOS 50:50 KOS:KTN 100%KTN 7 Day (n=5) Collagen 100%KOS 50:50 KOS:KTN 100%KTN

Table 3. 15% (weight/volume) 100%KOS Standard Curve with serial rhBMP-2 concentration.

Biomaterial 100%KOS rhBMP-2 Concentration (µg/mL) 100 50 25 12.5 6.25 3.125 1.5625 0

A critically-sized rat femur defect model was used to assess in vivo retention of rhBMP-2 at the defect site [7][8]. Briefly, rats (males, Sprague Dawley, Charles river) of average weight 378g, were anesthetized, hair on the left leg was shaved and underlying skin was disinfected with povidone-iodine solution and alcohol (3 times). Then the left femur exposed by dissection of surrounding skin, fascia, and muscle. An internal fixator (Georgia Tech Research Institute, GTRI, Atlanta, GA) of two 316 stainless steel plates bridged by a Thermalux® polysulfone bridge, designed by the Guldberg Lab at Georgia Tech [8], was attached to the femur by drilling holes and screwing the plate to the femur with gold-plated stainless steel (grade 303) screws. An 8mm segment of the femur between the plates was removed by reciprocating saw. The defect site was flushed by saline to remove blood and tissue

23 fragments. The keratin scaffolds were then approximated to the length of the defect based on the removed bone length and carefully placed into the defect site. The incision site was closed in muscle and skin layers, and the skin was then stapled by using stainless steel stapler to close the wound. Animals were allowed to recover and housing separately until the time of imaging. An initial study was conducted in which varying concentrations (Table 3) of rhBMP-2 labeled with DyLight 800 were implanted. Immediately after surgery, animals were humanely euthanized and frozen at -20oC until transported to the University of Cincinnati Vontz Imaging Laboratory. The purpose of this experiment was to determine if the fluorescence signal was linearly proportional to the amount of rhBMP-2. In order to reduce surgery times and material, the assumption was made that the linearity of 50:50 KOS:KTN, 100%KTN and collagen are the same. Thus the 100%KOS standard curve could be representative. Later, a study was conducted in which 100 µg/mL of rhBMP-2 labeled with Dylight 800 was implanted for 1, 3, or 7 days. At these times, rats were humanely euthanized and frozen at -20oC until transport to the Vontz Core Imaging Laboratory at University of Cincinnati for imaging. Based on a power calculation prior to the study for expected levels of variability, an n of 4 was selected for the early time points (1 and 3 days) and n = 5 for each group at 7 days. All surgical procedures and experimental procedures were under a sterile environment in order to avoid infection and all animal protocols were approved by the Miami University Institutional Animal and Care and Use Committee (IACUC).

II.2.5. Bruker System Imaging and Photoshop Quantification of Images At the Vontz Imaging Center, hair was removed by depalitation cream in order to minimize fluorescence signal from the hair. Then skin and muscle were removed to expose the defect site. On the Bruker system, an X-Ray was collected initially to ensure proper orientation of the animal. Then, an optical image was collected followed by a fluorescent image. For the fluorescence image, the following parameters were used: Ex/Em 730/790 and exposure time of 60 seconds. The raw data images were converted to TIFFs with the same scaling for each image to ensure consistency between images. To assess retention of rhBMP-2 at the defect site, we used Adobe Photoshop (San Jose, CA). The amount of DyLight800-labeled rhBMP-2 from the keratin and collagen scaffold implant site was determined by image analysis methods in which the pixel intensity was scaled from 0 to 255. The region of rhBMP-2 was determined by presence of fluorescent signal. The region where fluorescence was observed was manually selected (magnetic lasso tool). The total (integrated) pixel intensity and

24 average pixel intensity (total intensity/number of pixels) were then determined.Flourescence images were also overlayed on the X-ray images to qualitatively visualize rhBMP-2.

II.2.6. Assessment of Fluorescently-tagged rhBMP-2 in Vivo Biodistribution The biodistribution of rhBMP-2 was assessed in major organs (brain, liver, spleen, kidney, heart, lungs) by explanting these at the terminal time points of the study and imaging on the Bruker Imaging System for detection of the presence of DL800-tagged rhBMP-2. Images were collected as described above, and total pixel intensity was determined as well as the average pixel intensity.

II.3. Results/Discussion II.3.1 In vitro rhBMP-2 release from keratin hydrogels and collagen scaffolds Briefly, AF488 was added to the dialyzed rhBMP-2 at a molecular ratio of 1.5 to 2 By quantifying and plotting absorbance with varying rhBMP-2 concentration, a standard curve was obtained as shown in Figure 1). The cumulative percentage rhBMP-2 release for each formulation was obtained from cumulative rhBMP-2 mass (µg) over the initial rhBMP-2 mass of 10µg (100 µL at 100 µg/mL). As shown in Figure 2, all rhBMP-2 release profiles were similar for the various keratin formulations and began to plateau after 3 days in a manner consistent with first order release. Also, the collagen sponge gave rhBMP-2 release with an initial burst release (see first 12-24 hours of collagen release) while the keratin hydrogels did not exhibit the same level of burst release as collagen. Surprisingly, the in vitro kinetics results show that the cumulative release of rhBMP-2 from KOS is the lowest, which is the opposite of our expectation and previous results with other growth factors (e.g., IGF-1, bFGF, and VEGF) where release was lowest with KTN. Despite difference in the in vitro rhBMP-2 releasing profiles for collagen and keratin biomaterials, all biomaterial has the functional ability to control BMP-2 releasing. From statistical analysis, rhBMP-2 releasing profile of each keratin formulation or collagen showed no significant difference within 168 hours with each other since the probability values of p < 0.05 in t-test were considered to indicate statistical difference. But at 12 hours time points, the cumulative rhBMP-2 percentage releasing showed significant difference with the other keratin formulations, which indicate rhBMP-2 initial burst from collagen biomaterial statistically.

25 60 rhBMP-2 concentration (ug/mL) = 0.0017*Fluorescence

(ug/ 50 R² = 0.99913 40 30

mL) 20 Concentrtion

10 0

rhBMP-2 0 5000 10000 15000 20000 25000 30000 -10 Fluorescence (RFU)

Figure 1. Standard curve relating absorbance with AF488 tagged rhBMP-2 concentration. N=3 and error bars represent standard deviation.

140 70:30 KOS:KTN 120 BMP-2 0:100 100 KOS:KTN BMP-2 80 50:50 KOS:KTN BMP-2 60 30:70 KOS:KTN 40 BMP-2 Collagen 20 Cumulative rhBMP-2 Percentage Release 0 100:0 0 20 40 60 80 100 120 140 160 180 KOS:KTN Time (hr) BMP-2

Figure 2. rhBMP-2 percentage releasing profiles of 100:0 KOS:KTN, 70:30 KOS:KTN, 50:50 KOS:KTN, 30:70 KOS:KTN, 0:100 KOS:KTN and collagen biomaterial. N = 3, error bars denote standard deviation. Statistically, there is no significant difference between rhBMP-2 releasing profiles of each formulation from 12-168 hours. And the cumulative rhBMP-2 percentage releasing showed significant difference with the other keratin formulations from 0-12 hours. p < 0.05 considered as significant difference in t-test.

26 The protein content eluted into each sample was determined by the modified Lowry protein assay (Bio-Rad, Hercules, CA). The standard curve was made with serial dilution of 90 mg BSA with 45 mL PBS (2mg/mL) as shown in Figure 3.

2.5 BSA concentration(mg/mL) = 5.2446A - 0.5085 2 R² = 0.99949 1.5

BSA mg/mL 0.5

0 0.1 0.15 0.2 0.25 0.3 0.35 0.4 0.45 0.5 -0.5 Absorbance

Figure 3. Standard curve of absorbance with diluted BSA concentration for Lowry protein assay (n=3), error bars denote standard deviation.

KOS only 30

70:30 KOS:KTN + BMP-2 25 KOS + BMP-2

50:50 KOS:KTN 20 only 30:70 KOS:KTN only 15 50:50 KOS:KTN + BMP-2 30:70 KOS:KTN + 10 BMP-2 KTN only 5 Cumulative Protein Mass (mg) KTN + BMP-2

0 Collagen + BMP-2 0 20 40 60 80 100 120 140 160 180 Collagen only Time (hr)

Figure 4. Lowry protein assay cumulative protein mass profiles of 100:0 KOS:KTN, 70:30 KOS:KTN, 30:70 KOS:KTN, 0:100 KOS:KTN and collagen biomaterial (15% weight/volume, n=3) with or without rhBMP-2 from 1.5hr to 7 days (168hr), error bars denote standard deviation.

27 Table 4. Statistical analysis of keratin degradation for samples with rhBMP-2 in vitro from 0-168 hours.

With rhBMP-2 70:30 KOS:KTN 50:50 KOS:KTN 30:70 KOS:KTN KTN Collagen KOS - - + + + 70:30 KOS:KTN - + + + 50:50 KOS:KTN - + + 30:70 KOS:KTN - + KTN +

Having significant difference (+) or no significant difference between each formulation of biomaterial, KOS, 70:30 KOS:KTN, 50:50 KOS:KTN, 30:70 KOS:KTN, KTN and collagen (n=3) by loading with rhBMP-2. p < 0.05, as determined by t- test.

Table 5. Statistical analysis of keratin degradation for samples without rhBMP-2 in vitro from 0-168hours.

No rhBMP-2 70:30 KOS:KTN 50:50 KOS:KTN 30:70 KOS:KTN KTN Collagen KOS - + + + + 70:30 KOS:KTN - - + + 50:50 KOS:KTN - + + 30:70 KOS:KTN + + KTN +

From statistical analysis, only KOS and 50:50 KOS:KTN exhibit significant difference between loaded with rhBMP-2 and loaded without rhBMP-2. From Table 4 and Table 5, the cumulative protein mass of collagen has significant difference with all keratin biomaterial formulations for either with rhBMP-2 or without rhBMP-2. The KOS protein degradation rate showed significant differences with lower KOS content formulations including 50:50 KOS:KTN, 30:70 KOS:KTN, KTN and collagen. On the other hand, the KTN protein degradation rate has significant difference with lower KTN content formulations like KOS, 70:30 KOS:KTN, 50:50 KOS:KTN and collagen. From quantitative analysis of protein cumulative mass (Figure 4), the degradation rates of keratin biomaterials are faster than collagen. All biomaterials have rapid protein degradation on the first 10 hours and the protein degradation rates

28 were slow after that. The cumulative protein mass curves for all biomaterials have not reach the plateau, which is the steady state at 168 hours.

II.3.2 Rat Femur Defect Model for Retention of rhBMP-2 at Implant Site From Bruker Imaging System, three images were collected for each single rat (Figure 5): the X- Ray (Figure 5C) was collected to ensure proper orientation of the animal, the optical image (Figure 5B) was collected, followed by a fluorescent image (Figure 5A).

Figure 5. Three kinds of images from Bruker Imaging System for each rat: Fluorescent image (A), optical image (B) and X- ray (C) for experimental rats. The images were selected from one rat and shown to demonstrate information collected for every animal used in these studies.

Figure 6. To determine if fluorescence was proportional to concentration so that in later time course studies, fluorescence would be known to be dependent on concentration, keratose scaffolds containing serially decreasing amounts of DyLight-

29 800-labeled rhBMP-2 were implanted and the defect site was then imaged as indicated in the Materials and Methods section above. The pixel intensity represents the concentration of rhBMP-2. Fluorescent images for 15% (weight/volume) rhBMP-2 loaded KOS scaffold standard curve are shown with serial rhBMP-2 concentration for quantification (n=3). The images are representative of the average image pixel intensity among the three samples for each rhBMP-2 concentration.

For the standard curve images (Figure 6), the fluorescence intensity was the highest is the “hottest” (highest signal intensity) at the highest rhBMP-2 concentration (100 µg/mL) and the integrated (total) pixel intensity decreases with descending rhBMP-2 concentration. By quantifying and plotting 15% KOS scaffold with varying concentration of rhBMP-2 labeled with DyLight800, a standard curved was obtained, as shown in Figure 7.

6.00E+06 Integrated pixel intensity = 39931*rhBMP-2 concentration(ug/mL) 5.00E+06 R² = 0.99298

4.00E+06

3.00E+06

2.00E+06 Integrated Pixel Intensity Integrated Pixel 1.00E+06

0.00E+00 0 20 40 60 80 100 120

rhBMP-2 Concentration (ug/mL)

Figure 7. Standard curve of serial concentration of rhBMP-2 that loaded on 100%KOS (15% weight/volume) with integrated pixel intensity of fluorescent images, N=3, data points represent the mean ± standard deviation.

After establishing that the fluorescence signal intensity from the images was linear with respect to rhBMP-2 concentration (Figures 6 and 7), we then assessed retention of rhBMP-2 labeled with DyLight800 at the implant site for 4 different formulations (collagen, 100:0 KOS:KTN, 50:50 KOS:KTN, and 0:100 KOS:KTN). Images were collected in the same manner as for the standard curve, but at later time points. As shown in Figure 8, there is clearly fluorescent signal at the femur defect site. However, the signal intensity even at 24 hours is much lower than immediately after surgery (compare to Figure 6). The results seem to show increased retention of rhBMP-2 with KTN and keratin mixtures in comparison to collagen. Moreover, the signal of fluorescent is decaying over tome for each formula.

30 To investigate this quantitatively, we again assessed the integrated pixel intensity for the formulations at each of the 3 time points. As shown in Figure 9, in agreement with the quantitative images, the formulations containing KTN showed increased levels of rhBMP-2 retention. From the rhBMP-2 percentage retention, 1-5% of rhBMP-2 remained at the defect site for the various formulations (collagen, keratose, kerateine, or the 50:50 mixture of keratose:kerateine), which means 95-99% of the rhBMP-2 was released, approximately. The rhBMP-2 percentage retention of collagen and KOS was not statistically significant at 1, 3 and 7 days，since p values were greater than 0.05 by t- test. In Figure 8, there are weak and small areas of fluorescent signal at day 1 for collagen and KOS, and no signal at day 3 and day 7. The rhBMP-2 percentage retention of collagen was significantly different than both 50:50 KOS:KTN and KTN at 1 and 3 days since p value is less than 0.05 by t-test. And also the rhBMP-2 percentage retention of KOS has significant difference with both 50:50 KOS:KTN and KTN at 1 and 3 days since p value is less than 0.05 by t-test. At day 3, the BMP-2 percentage retention in 50:50 KOS:KTN and KTN shows significant difference by t-test. At day 7, there is no significant difference between each biomaterial for rhBMP-2 percentage retention by t-test.

Figure 8. Fluorescent images nearest the average value (of N=4) of the pixel intensity of rhBMP-2 loaded collagen, 100%KOS, 50:50 KOS:KTN and 100%KTN biomaterial at 1, 3 or 7 days post-implantation. Increasing retention of rhBMP- 2 is observed for formulations containing KTN comparison to collagen or KOS materials.

Figure 9. Percentage rhBMP-2 retention from collagen, 100%KOS, 50:50 KOS:KTN and 100%KTN scaffold at 1 (N=4), 3 (N=4) and 7 (N=5) days. Error bars indicate standard deviation. . * and # denote values that differ significantly from collagen and 100%KOS respectively at Day 1, + and – donate values that differ significantly from collagen and 100%KOS respectively at Day 3, p < 0.05, as determined by t-test.

II.3.3. Biodistribution of rhBMP-2 in Vivo

Due to the large amount of rhBMP-2 release from the defect (that is, the small amount of retention), we decided to investigate whether any of the organs showed DyLight800 signal that might indicate distribution of the released rhBMP-2 (or its degradation products) to these sites. Table 6, 7 and 8 list out the number of organs positive for fluorescence signal over total number of each kind of organ of rats with rhBMP-2 loaded collagen, 100%KOS, 50:50 KOS:KTN and 100%KTN scaffold implantation at 1, 3 or 7 days post-implant. Brain, heart, spleen, liver, left lung, right lung, left kidney and right kidney were assessed.

32 Table 6. Number of fluorescent signal positive organs over the total number of each kind of organ of rats with rhBMP-2 loaded collagen, 100%KOS, 50:50 KOS:KTN and 100%KTN scaffold implantation at Day 1 after implantation.

Day 1 Brain Heart Spleen Liver Left Lung Right Lung Left Kidney Right Kidney Collagen 0/4 0/4 2/4 0/4 0/4 0/4 1/4 2/4 100% KOS 0/4 0/4 2/4 2/4 1/4 1/4 0/4 0/4 50:50 KOS:KTN 0/4 0/4 1/4 1/4 0/4 0/4 0/4 0/4 100% KTN 0/4 0/4 1/4 0/4 1/4 1/4 1/4 1/4 N = 4.

Day 3 Brain Heart Spleen Liver Left Lung Right Lung Left Kidney Right Kidney Collagen 0/4 0/4 2/4 0/4 0/4 0/4 1/4 0/4 100% KOS 0/4 0/4 1/4 1/4 0/4 0/4 0/4 0/4 50:50 KOS:KTN 0/4 0/4 2/4 0/4 0/4 0/4 0/4 0/4 100% KTN 0/4 0/4 0/4 1/4 0/4 0/4 0/4 0/4 N = 4

Day 7 Brain Heart Spleen Liver Left Lung Right Lung Left Kidney Right Kidney Collagen 0/5 0/5 3/5 2/5 0/5 1/5 2/5 2/5 100% KOS 0/5 0/5 1/5 1/5 0/5 1/5 1/5 0/5 50:50 KOS:KTN 0/5 0/5 2/5 2/5 0/5 0/5 1/5 1/5 100% KTN 0/5 0/5 0/5 0/5 0/5 0/5 0/5 0/5 N = 5

As reflected in Tables 6 – 8 and shown in Figures 10,11,12 and 13. there are weak fluorescent signals on the spleen, liver, lung and kidney. No fluorescent signals were observed on the brains or hearts of any animals (Figure 14 and 15). Among the organs, the collgen and KOS used as the rhBMP-2 carriers showed more fluorescent signal.

Figure 10. Biodistribution fluorescent image for spleen of each rat, which implanted with rhBMP-2 loaded collagen, 100%KOS, 50:50KOS:KTN and 100%KTN at day 1 (N=4), 3 (N=4) and 7 (N=5) time points.

From Figure 10, the spleen is where the highest fluorescent signal was observed. The rhBMP-2 loaded collagen implanted rats spleen showed more fluorescent signal and also the signal is increasing over time in the spleen and that more of the collagen spleens showed signal (3/5) than any of the other formulations, especially at day 7.

Figure 11. Biodistribution fluorescent image for liver of each rat, which implanted with rhBMP-2 loaded collagen, 100%KOS, 50:50KOS:KTN and 100%KTN at day 1 (N=4), 3 (N=4) and 7 (N=5) time points

From Figure 11, the livers showed weak and small areas of fluorescent signal. From Figure 12 and 13, weak fluorescent signals on day 1 were observed for both lung and kidney. The lungs (left and right) have only fluorescent signal on 100%KOS and 100%KTN at day 1. But more fluorescent signals

34 on kidney for collagen material at day 1 and day 7. Signals were weak or not observed for the other biomaterial formulations. No fluorescent signals on brain heart (Figure 14 and 15).

Figure 14. Biodistribution fluorescent image for heart of each rat, which implanted with rhBMP-2 loaded collagen, 100%KOS, 50:50KOS:KTN and 100%KTN at day 1 (N=4), 3 (N=4) and 7 (N=5) time points.

Figure 15. Biodistribution fluorescent image for brain of each rat, which implanted with rhBMP-2 loaded collagen, 100%KOS, 50:50KOS:KTN and 100%KTN at day 1 (N=4), 3 (N=4) and 7 (N=5) time points.

9 8 7 6 5 Day 1 4 3 Day 3 2 Day 7 1 0 Collagen 100%KOS 50:50KOS:KTN 100%KTN rhBMP-2 Concentration (ug/mL) Formulations

Figure 16. Quantification of rhBMP-2 concentration in spleen for collagen, 100%KOS, 50:50 KOS:KTN and 100%KTN implanted rats at day 1 (N=4), 3 (N=4) or 7 (N=5) time points. Data points represent average for organs positive for fluorescence signal. Error bars indicate standard deviation.

36 5 4.5 4 3.5 3 2.5 Day 1 2 Day 3 1.5 Day 7 1 0.5 0 rhBMP-2 Concentration (ug/mL) Collagen 100%KOS 50:50KOS:KTN 100%KTN Formulations

Figure 17. Quantification of rhBMP-2 concentration in liver for collagen, 100%KOS, 50:50 KOS:KTN and 100%KTN implanted rats at day 1 (N=4), 3 (N=4) or 7 (N=5) time points. Data points represent average for organs positive for fluorescence signal. Error bars indicate standard deviation.

20 18 16 14 12 10 Day 1 8 Day 3 6 Day 7 4 2 0 rhBMP-2 Concentration (ug/mL) Collagen 100%KOS 50:50KOS:KTN 100%KTN Formulations

Figure 18. Quantification of rhBMP-2 concentration in lungs for collagen, 100%KOS, 50:50 KOS:KTN and 100%KTN implanted rats at day 1 (N=4), 3 (N=4) or 7 (N=5) time points. Data points represent average for organs positive for fluorescence signal. Error bars indicate standard deviation.

37 18 16 14 12 10 Day 1 8 Day 3 6 Day 7 4 2 rhBMP-2 Concentration (ug/mL) 0 Collagen 100%KOS 50:50KOS:KTN 100%KTN Formulations

Figure 19. Quantification of rhBMP-2 concentration in kidneys for collagen, 100%KOS, 50:50 KOS:KTN and 100%KTN implanted rats at day 1 (N=4), 3 (N=4) or 7 (N=5) time points. Data points represent average for organs positive for fluorescence signal. Error bars indicate standard deviation.

Figure 16, 17, 18 and 19 shows the average rhBMP-2 concentration for the positive organs, which are spleen, liver, lungs and kidneys. The non-positive organs were excluded in the quantification analysis. Since the brain and heart have no fluorescent signal, no quantification analysis done for these two organs. The bar chart gives the rhBMP-2 concentration is decreasing or increasing with time for each single biomaterial. Most of organs express high fluorescent signal on day 1 time point. Among the weak fluorescent organs, the collagen and KOS implantation rats showed higher rhBMP-2 concentration.

In the following chapter we will provide further analysis and discussion of these results and their contribution to the literature. We will also describe potential future work to build on these studies.

38 References

[1] Rouse, J.G.; Van Dyke, M.E. A Review of Keratin-Based Biomaterials for Biomedical Applications. Materials 2010, 3, 999-1014.

[2] de Guzman, R. C. et al. Mechanical and biological properties of keratose biomaterials. Biomaterials 32, 8205–17 (2011).

[3] de Guzman, R. C. et al. Bone regeration with BMP-2 delivered from keratose scaffolds Biomaterials 34, 1644–1656 (2013).

[4] Tomblyn, Seth, et al. "Keratin hydrogel carrier system for simultaneous delivery of exogenous growth factors and muscle progenitor cells." Journal of Biomedical Materials Research Part B Applied Biomaterials (2015).

[5] Ham, Trevor R, et al. "Tunable keratin hydrogels for controlled erosion and growth factor delivery." Biomacromolecules (2015).

[6] Han, Sangheon, et al. "Alkylation of Human Hair Keratin for Tunable Hydrogel Erosion and Drug Delivery in Tissue Engineering Applications.." Acta Biomaterialia 23(2015):201-213.

[7] Guzman, Roche C, De, et al. "Bone regeneration with BMP-2 delivered from keratose scaffolds.." Biomaterials 34.6(2013):1644–1656.Spicer P.P, Kretlow J.D. Evaluation of bone regeneration using the rat critical size calvarial defect. Nature Protocols 7, 1918–1929 (2012)

[8] Oest, Megan E., et al. "Quantitative assessment of scaffold and growth factor-mediated repair of critically sized bone defects.." Journal of Orthopaedic Research 25.25(2007):941-50.

CHAPTER III – Discussion of Results and Future Work

Jingxuan Li

40 III.1. Discussion of Thesis Results

Keratins are natural polymers that can be extracted from various sources including human hair. The materials are characterized by high levels of cysteine residues [1]. Keratin is typically extracted by oxidative techniques or reductive methods, which lead to extracted forms of keratin known as keratose (KOS) and kerateine (KTN), respectively. Because these materials are biodegradable with non-toxic degradation products, biomaterials made from keratin have been used for numerous in vivo applications including nerve regeneration [2], skin/wound healing [3], and others. In many of these applications, the keratin biomaterials have been used alone, without any drug delivery/controlled release aspects. While keratin has been used alone (without any drug delivery) for bone applications [4][5], our team has used keratin to delivery rhBMP-2 in femur defect and mandibular models of bone injury. As to the bone regeneration, the growth factor needs a carrier for controlled release. If growth factor carriers that release growth factor too slowly are used, cells may not attain enough growth factor to reach the threshold for cellular cues such as chemotaxis or differentiation. In addition, the initial large burst release of growth factor by carriers may result in suboptimal bone injury healing [6]. In an effort to achieve more sustained release of rhBMP-2, our group has conducted two recent in vivo studies related to the use of keratin biomaterials to promote bone regeneration. More specifically, KTN has been used in a mandibular defect model [5] whereas KOS has been used in a rat femur defect model [7]. Further, we have also recently demonstrated that KOS and KTN lead to very different rates of release of several different growth factors commonly used in tissue engineering applications [8]. The rationale for use of keratin hydrogels in bone regeneration is based on the fact that it is a resorbable material that degrades more slowly in vivo than collagen, but with natural (protein-based) degradation products. With a slower rate of resorption in vivo, this should provide a “scaffolding” upon which osteprogenitor and osteoblast migration can be supported for the deposition of bone matrix at fracture sites, thereby leading to improved healing and safety profiles. Based on these previous in vivo findings, our group recently sought to use alkylation chemistry to tune the rates of keratin hydrogel carrier degradation. This approach also demonstrated the ability to tune the release of ciprofloxacin antibiotic and rhIGF-1 [4]. However, this approach is somewhat challenging and was not as readily tunable as desired. Because KOS and KTN have different physiochemical properties in terms of their ability to form disulfide crosslinks, we reasoned that materials composed of KOS and KTN in varying ratios could be used to tune the rate of degradation.

41 We recently established the ability to tune keratin hydrogel carriers based on the levels of disulfide crosslinking, which also led to tunable delivery of rhIGF-1 [9]. The goal of this thesis study is to have a better quantitative understanding of how the delivery of rhBMP-2 can be modified with by manipulating the formulation and the resultant rate of degradation of keratin biomaterials. We hypothesized the modulation of disulfide crosslinking levels in keratin materials (controlled by the ratios of KOS and KTN) could also be used to control rhBMP-2 release in a manner similar to rhIGF-1 release previously identified by our group. In our initial study, we evaluated the degradation of keratin hydrogels (to confirm previous results) and rhBMP-2 release in vitro. To determine protein content, a modified Lowry protein assay (Bio-Rad, Hercules, CA) was used. From Chapter 2 Figure 4, Table 4 and Table 5, the collagen protein degradation rate is significantly different with keratin biomaterial from 0 to 168 hours and also has lower degradation rate than others by observation. By conducting t-test between each formulation from 0 to 168 hours, the KOS protein degradation rate showed significant differences with lower KOS content formulations including 30:70 KOS:KTN, KTN and collagen. On the other hand, the KTN protein degradation rate has significant difference with lower KTN content formulations like KOS, 70:30 KOS:KTN, 50:50 KOS:KTN and collagen. Thus, we can conclude that KOS:KTN ratios can be used to tune the rate of keratin biomaterial degradation rate since the KOS has no disulfide crosslinking and KTN has full disulfide cross linking [10]. In the rhBMP-2 in vitro release study, keratin hydrogels were loaded with AF488-labeled rhBMP-2 and release was measured over time at 37°C. This in vitro study shown in Figure 1 clearly shows that rhBMP-2 can be released over a period of time from keratin hydrogels. From Figure 1, we found that the keratin hydrogels exhibited lower burst release than collagen sponges. In addition, each keratin formulation showed first order release. However, it was surprising that unlike IGF-1, bFGF, VEGF, and ciprofloxacin, the rate of rhBMP-2 delivery was not affected by the ratios of KOS and KTN. There was no obvious pattern related to the rhBMP-2 release based on the keratin formulation [6]. There are several mechanisms for rhBMP-2 release from hydrogels. These include specific material properties for varying diffusivities, covalent bonding with cleavable chains, and/or non-covaent interactions interactions that allow for adsorbtion to the material carriers [6]. In the previous study from our team, experiments showed that rhBMP-2 has high equilibrium binding affinity to keratin and also indicates the disulfide bond would not affect the affinity of rhBMP-2 to keratin biomaterial. These previous experiments also suggested that the rhBMP-2 would not dissociate as easily from keratin

42 compared to collagen [6][11][12]. Since the binding affinity between rhBMP-2 and collagen is relatively weak, this explains the initial large burst that was observed (Figure 1). Since the rhBMP-2 molecule is dimerzied with intersubunit angles, which could create two cavities. Cavity I has the positive charges that could interact with keratin biomaterial. Furthermore, the positive charge in cavity I could correlate with stronger electrostatic interaction [6]. At the beginning, we predict the KOS degradation rate was larger than KTN, the rhBMP-2 release from KOS should be faster and larger than from KTN. Five formulations of keratin, KOS, 70:30 KOS:KTN, 50:50 KOS:KTN, 30:70 KOS:KTN and KTN, were assessed. However the experimental results did not followed the way we predict. In the range of disulfide crosslinking content, KOS actually showed lower rates of rhBMP-2 release than KTN, but KOS and KTN did not have significant release in statistical speaking. One possible explanation is that the relatively large rhBMP-2 (compared to, e.g., rhIGF-1 and ciprofloxacin) remains sequestered with the keratin hydrogel network even though the KOS is degrading more rapidly than the KTN in terms of protein release (Figure 11). When we prepare the biomaterial implantation, the keratin biomaterial was made to freeze-dried scaffold but collagen was a sponge. After implantation, the keratin scaffolds start to absorb water from the surround tissue to become hydrogel and collagen sponge start to degrade and release rhBMP-2 to the surrounding tissue. Also by the function of collagenase enzyme, the collagen exhibited an initial burst. Despite difference in the in vitro BMP-2 releasing profiles for collagen and keratin biomaterials, all biomaterial showed the ability to achieve sustained release of rhBMP-2.

Despite the unexpected in vitro results, it is well known that in vivo behaviors are often different than in vitro due to the complexities of the native physiological systems into which materials are placed. To assess rhBMP-2 “release” in vivo, we elected to assess retention of the rhBMP-2 at the site of a critically sized rat femur defect. This study and model are particularly relevant because this in vivo study is the first time in literature, to our knowledge, that assesses the release of rhBMP-2 from keratin formulations in a rat femur model. More generally, there are few papers from literature research that describe in vivo rhBMP-2 release from a material in a bone defect model. Freiss and Uludagpublished a report in the International Journal of Pharmaceutics in 1999, which studied the in vivo release kinetics of 125I-rhBMP-2 from collagen sponge by using a rat ectopic implant model. But the interest of Freiss and Uludag’s study is assessing the effect of local BMP levels and osteoinductive activity in vivo and the interaction of rhBMP-2 with collagen [13]. Their research gave the idea that the rhBMP-2 retention

43 within collagen sponge carrier was variable, ranging from 10-75% after 3 hours. They claimed the rhBMP-2 releasing kinetics was strongly dependent on the implanted biomaterial, which is the collagen carrier of the rhBMP-2 [13]. Unlike Friess model, our rat femur defect model not only can provide an rhBMP-2 retention from keratin and collagen biomaterials at the implanted area but also tried to track all rhBMP-2 disposition from vital organs.

The standard rat segmental model used was consistent with those previously reported [7][14]. In brief, an internal fixator of two 316 stainless steel plates bridged by a polysulfone plate to achieve highly stable mechanical fixation is implanted into the left femur of male Sprague-Dawley rats. Upon implantation, a critically sized 8-mm femoral segmental defect is created by sagittal saw. Keratin materials and a collagen control, each loaded with DyLight800-labeled rhBMP-2 were implanted at the defect site. Initially, a standard curve to relate fluorescence intensity to rhBMP-2 concentration was conducted with KOS as the carrier (Figure 5). The assumption was made that the Dylight800-labeled rhBMP-2 would show similar signal when bound to KTN, 50:50 KOS:KTN or collagen as it did with keratose. Since these rats used to generate the standard curve were euthanized immediately after implantation surgery, the rhBMP-2 could not have been released from the defect site and no biomaterial degradation had yet occurred. As shown in Figure 6, the fluorescence signal was indeed linear, indicating that the subsequent time-course studies should have linear response of fluorescence signal to rhBMP-2 concentration at the defect site.

In order to get to how the biomaterial carriers control the release of rhBMP-2, we quantified the rhBMP-2 retention at the implant site. We then conducted a study in which the materials (collagen, or 3 keratin formulations loaded with DyLight800 rhBMP-2) were implanted for 1, 3, or 7 days. Since this study is the first time that we have assessed the rhBMP-2 releasing profile from rat femur defect model, and we wanted to assess a range of disulfide crosslinking levels in the keratin formulations. Thus, no disulfide crosslinking (KOS), 50% disulfide crosslinking (50:50 KOS:KTN) and fully disulfide crosslinking (KTN) keratin formulations were prepared. In the future, more keratin biomaterial formulations need to be assessed.

Since no significant change was noted between the 50:50 KOS:KTN and the 0:100 KOS:KTN (100% KTN) formulations, it is logical to further explore higher keratose values in the formulation such as 10, 20, 30 or 40% keratose (10:90, 20:80, 30:70 and/or 40:60 KOS:KTN formulations, respectively).

44 Figure 8 shows the levels of rhBMP-2 (as indicated by DyLight800 signal) at the 3 different time points for each formulation. Unlike the in vitro study, the in vivo study did show dependence on the rate of material degradation, where it is noted that collagen and KOS degrade quickly while the formulations containing kerateine degrade more slowly. Clearly, the rhBMP-2 retention of KOS and collagen is significantly low compared to the 50:50 KOS:KTN and KTN formulations, which had more signal at days 1, 3, and 7. This result is consistent with that previously reported by de Guzman, 2011 in which the KOS was degraded and resorbed over time following a rectangular hyperbolic regression from. In their mouse model, 8% KOS was retained at the defect area at 8 week post-implantation. The lack of covalent disulfide bonds enable the hydrogel material degrades relatively fast [1]. We believe that the low signal from the collagen carrier is related to enzymatic-degradation and thus the inability to retain rhBMP-2. Keratinase enzymes have not been found in humans, which may be advantageous for long-term drug delivery since it’s under slow degradation in vivo. Also the disulfide bond in KTN could lead to stability with time [7]. It should be noted that all of the fluorescence signals at days 1, 3, and 7 for all formulations were very low. The fluorescent signals from collagen, KOS, 50:50KOS:KTN and KTN were much weaker compared to the standard curve fluorescent images after 1 day post-implantation, but fluorescent signals from 50:50KOS:KTN and KTN remained consistent over 1 week. This observation indicates that a significant proportion of rhBMP-2 was released from collagen and KOS, but also from the formulations containing KTN. The reason for the more sustained presence of rhBMP-2 in the KTN formulations could be due the disulfide bonds in these materials, which would lead to more stability in vivo than observed in vitro where collagen degraded more slowly. Thus, if it is desirable to achieve more sustained release (more retention at defect site), keratin (particularly kerateine) appears to be more suitable than collagen. Also, modification of disulfide cross-linking content in keratin formulations by mixing KOS and KTN appears to achieve some level of control over the retention of rhBMP-2, presumably due to the tunable biomaterial degradation. Further experiments may be needed to determine the fine-tuning more clearly, but this result seems to validate our initial hypothesis, at least in vivo (but see above for in vitro discussion). The result is consistent with our initial expectation that both 50:50 KOS:KTN and KTN have the covalent disulfide bond, leading to relatively slow degradation rate in vivo. The results also explain the effects observed clinically with collagen as the carrier for rhBMP-2, as its quick degradation will lead a

45 burst of rhBMP-2 release to the defect surrounding area. This burst release and lack of tunability suggest the ability to achieve more favorable healing with alternative carrier systems. The results of this study are also consistent with a previous study from our group in a critically-sized rat mandibular defect model in which the KTN scaffolds showed significantly less ectopic bone formation compared to the absorbable collagen sponge [7]. Again, our results indicate that the reason for this observation may be related to a lower initial burst release. Initially it was expected that the in vitro study and in vivo study would correspond since the disulfide crosslinking content should affect to the degradation rate of biomaterials and allow tuning of the releasing rate of rhBMP-2 in either case. For the in vitro study, there are 6 biomaterial formulations, which include KOS, 70:30 KOS:KTN, 50:50 KOS:KTN, 30:70 KOS:KTN, KTN and collagen. But for in vivo study, only KOS, 50:50 KOS:KTN, KTN and collagen were involved since in vivo study takes time for rat femur surgery and the most promising formulations needed to be tested. But in the future time, more keratin biomaterial formulation should be assessed, as discussed above. One key difference between the two studies that should be considered, however, it that the in vitro study used hydrogels while in vivo study used freeze-dried scaffolds (freeze dried hydrogels). There is a technical reason for this difference. In vitro study, freeze dried scaffolds initially absorb PBS, making collection of PBS difficult during this initial material swelling. For the in vivo study, the freeze- dried plugs were necessary in order to shape the material to the cylindrical shape of the bone defect in the femur, which is necessary to make surgical implantation easier. While this difference seems small, it represents a potential problem in directly relating the in vitro and in vivo release profiles. After noting that greater than 95% of the rhBMP-2 was no longer present at the bone defect site, we elected to investigate whether rhBMP-2 (indicated by DyLight800 signal) could be detected in any of the organs. The implication is that if evidence of rhBMP-2 signal in vital organs was found that this could lead to non-local (distal) ectopic bone formation. Alternatively, given the various roles of BMP-2 in bone formation, neural tube formation, and vascularization, other effects could occur if rhBMP-2 were to accumulate at distal sites. Although this thesis did not explore any effects of rhBMP-2 in distal organs, we did set out to identify whether detectable levels of rhBMP-2 could be found. As shown in Figure 10-15 and Table 6-8, rhBMP-2 (as indicated by DyLight800 fluorescence) was detectable in some (but not all) of the following organs: lung, kidney, liver, and particularly the spleen. The reason for this distribution is likely that after the rhBMP-2 loaded keratin or rhBMP-2 loaded collagen scaffold are implanted, the rhBMP-2 molecules diffuse away. Given the highly

46 vascularized nature of bone and also the surrounding tissue, it is likely that some of the rhBMP-2 molecules were transferred to blood vessels by hydrostatic and concentration gradient. From Figure 14, there is obviously no fluorescent signal in heart. Likewise, from Figure 15, there is obviously no fluorescent signal in the brain. In terms of the brain, the lack of detectable signal could be due to the blood-brain barrier. The blood–brain is defined as highly selective permeability barrier that can separate the circulating blood from the brain extracellular fluid in the central nervous system. Only water, some gases and lipid-soluble molecules could pass the blood–brain barrier by passive diffusion. Moreover, the glucose and amino acid could cross the blood-brain barrier by the selective transport mechanism, which is crucial to neural function [6]. Since the BMP-2 molecule contains 115 amino acids and has a molecular mass about 26000 Dalton, it is unlikely to be able to cross blood-brain barrier. In terms of the detection in the spleen and the liver, the explanation here may be related to the accumulation of rhBMP-2 in these organs for metabolism or possibly even to its uptake by the reticuloendothelial system (RES), in which these two organs play a major role. For example, the liver plays an important role in daily metabolism with numerous functions, including regulation of glycogen storage, decomposition of red blood cells, plasma protein synthesis, hormone production and detoxification [15]. The liver is also responsible for metabolic process, so the metabolized protein (and DyLight800 dye) could be metabolized in the liver and then excreted via the kidney. This could explain the fluorescent signal observed in the kidney. From statistical analysis (p<0.05), there is no significant difference between each implantation material and also no significant difference between each time point for brain, heart, liver, spleen, lung and kidney. In order to lower the bias in evaluation of the images, image analysis was repeated by a blinded observer who was unaware of what formulation was present for a given organ. The same result was obtained by the blinded observer. There are several limitations to this study, which should be considered and may provide directions for future research. One is the selection of fluorescence detection methods. Radioactive molecules such as 125-I provide much greater sensitivity. However, when this study was conducted, the duration of the effect was unknown. At the initial thought, the study will last 16 weeks since the 16 weeks is the normal time for bone healing, which includes inflammation phase and reparative phase. And rhBMP-2 plays an important role in these two phases. So the radiolabeling was not selected due to the concerns of using 125-I being (1) exposure to personnel during and after surgery and (2) the 59 days half-life of 125-I meaning that only ¼ of the original signal would be present if a 16-week study were

47 conducted. The 1 week study was conducted first. Upon recognition that most of the signal was depleted by this time, the earlier time points (1 and 3 days) were investigated rather than later time points. At this point, the fluorescence method had already been selected. Clearly, the fluorescent signal detection with low BMP-2 concentration is low, though it is detectable for the 50:50 KOS:KTN and KTN formulations. The detection limit was found to be approximately 2 µg/mL as indicated by the in vivo standard curve (Figure 7). While the signal was detectable in the bone, it led to limited ability to detect fluorescence in the organs. This is important because the detection limit was found to be 1.5625µg/mL. However, BMP-2 is known to have important physiological effects even at ng or pg quantities. Thus, although fluorescence was not detected in all organs, accumulation of smaller amounts of rhBMP-2 could lead to physiological effects. It should also be noted that we are unsure if the fluorescence signal was of intact rhBMP-2 or degradation products. However, this problem would not be overcome by the use of 125-I or other methods of quantification since protein degradation would still occur with these molecules.

III.2. Future Directions

This thesis work has provided results that keratin can be formulated with varying ratios of KOS and KTN to modulate the rate of scaffold degradation and thereby control the rate of rhBMP-2 delivery in a model of bone injury. Clearly, not all questions about keratin as an rhBMP-2 carrier for bone regeneration have been assessed and answered and there are several areas of interest for future studies. For the rhBMP-2 retention at the femur defect implantation site, the standard curve for KOS with concentration gradient of rhBMP-2 was made and then the linear relationships were assumed to also apply to formulations containing KTN or collagen. Instead of doing three times more rat femur surgery, the linear standard curve was made as an assumption. For future work, the linear relationships may need to be performed with each implantation material in order to obtain the slope value. As noted above, the detection limit of approximately 1.5625µg/mL in not ideal, though it allows quantification of rhBMP-2 signal in the femur defect. The weak fluorescent signal detection was a more significant limitation of the biodistribution study with rhBMP-2. To overcome the low concentration detection, a further analysis methods need to be found to assess rhBMP-2 distribution in the organs. As an alternative method, a more sensitive label would be appropriate. As noted above, 125I could be used. Besides, for more

48 accurate rhBMP-2 biodistribution detection, immunohistochemistry (IHC) can be a method for detection of proteins in tissue sections. The IHC staining is achieved with antibodies, which can recognize the target protein. Due to antibodies are highly specific, the antibody will bind only to the protein of interest in the tissue section [16]. The antibody-antigen interaction can be visualized using either chromogenic detection, or fluorescent detection. For chromogenic detection, an enzyme conjugated to the antibody cleaves a substrate to produce a colored precipitate at the location of the protein. And for fluorescent detection, a fluorophore is conjugated to the antibody and can be visualized by fluorescence microscopy [17]. So, in this case we could stain for the presence of rhBMP-2 or the DyLight800 fluorophore to assess co-localization with specific cells in the liver, spleen, or other tissues.

Another limitation of the fluorescence method was that the study was not longitudinal in nature; that is, separate animals had to be used for each time point. The Bruker System camera could not penetrate as the depth as the femur defect site, also the autofluorescent inference will occur due to the surrounding tissue and skin. This could lead to larger variability than if a probe could be used that does not require individual animals for each time point. In the future work, an alternative tagging method might be needed in order to keep animal alive until the end of the study. Since this thesis study is just to assess the rhBMP-2 release (retention) kinetics, not quantify the bone regeneration, a long-term study could be conducted that not only tracks the rhBMP-2 release but also measure the quantity of the bone formation. Thus, radiolabeling will not be the appropriate method since both 125-I (59 days) and 131-I (only 8 days) have short half-life.

According to the hypothesis guiding this thesis the modulation of the disulfide cross-link density provides a built-in system for tunable degradation rate of hydrogel and also the release of therapeutic agents. This seems to be the case from the in vivo study, though not the in vitro study. Thus for further analysis, more formulations should be tested for the in vivo study to get a better sense of where (i.e., what KOS:KTN ratio) the retention of rhBMP-2 occurs. In addition, a study in which retention and bone regeneration are done together could allow for better understanding of which formulations might be ideal for a specific individual or type of fracture.

According to the rhBMP-2 release result, if greater bone regeneration is found with collagen and KOS, it will imply that weak affinity between rhBMP-2 is more desirable. That is, this would suggest that it is the burst release that plays the greatest role. The problem with such an approach however, is

49 that the burst release of rhBMP-2 results rhBMP-2 not only retains at the defect area but also will be transport to the other organs for metabolism, as shown in our biodistribution study. In addition the formation of ectopic bone becomes problematic. Conversely, for a regeneration study, if the more bone regeneration were found with KTN or keratin mixture, it would indicate that a biomaterial that can retain rhBMP-2 for a longer time would be more beneficial. This thesis assessed quantification for the rhBMP- 2 retention at the defect area, but did not assess bone regeneration. This thesis work also indicates that other formulation aspects of keratin should be considered. In Chapter 1 we noted that gamma keratin is globular with high sulfur content and lower molar mass, which is around 15kDa. The high sulfur content is due to the high amount of cysteine content that we now recognized can play a role in the function of disulfide crosslinking. Including gamma keratin in the formulations allow greater tunability in the future. To further enhance the rhBMP-2 retention at the defect site, a coating that outside the cylindrical implant biomaterial could be considered in the future. The coating material could have lower degradation rate than the biomaterial implantation and also have low immune response to the native body, thus, the released rhBMP-2 could be held longer around the defect area rather than diffuse away. The implantation coating cold enhances the sustained release of rhBMP-2 for greater or possibly more rapid bone regeneration. If the coating strategy were successful, the dose of rhBMP-2 could be lower than the amount we used in this thesis work since we could reduce the amount of rhBMP-2 diffuse away from the defect area. Scott Guelcher group introduced biodegradable-segmented polyurethanes be the ideal materials to characterize the effect of mechanical properties on tissue development. Like keratin, polyurethane with tunable degradation rates, that can support cellular infiltration and generation of new tissue[18]. From Guelcher research, the polyurethane scaffolds showed a two-stage release profile of platelet-derived growth factor. First stage was characterized by a 75% burst release within the 24 h and second stage was slower release profile thereafter [19]. Since the slower degradation of polyurethanes at the second stage, the polyurethanes could be considered as the coating material for keratin biomaterial. The keratin biomaterial is the ideal scaffold for tissue regeneration since keratin can provide consistent growth factor releasing without initial burst. In summary, keratin biomaterials extracted from human hair have shown the possibility being modified by composition of disulfide cross-link to control rate of rhBMP-2 delivery in a model of bone injury. Although, this work does not provide information on bone formation, it does provide information

50 on the suitability of keratin materials to serve as an alternative to collagen to retain rhBMP-2 at an implant site with lower amounts of burst release.

51 References

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