Scaffold Design and Optimization for Integrative
Cartilage Repair
Margaret K. Boushell
Submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in the Graduate School of Arts and Science
COLUMBIA UNIVERSITY
2016
© 2015
Margaret K. Boushell
All Rights Reserved
ABSTRACT
Scaffold Design and Optimization for Integrative Cartilage Repair
Margaret K. Boushell
Osteoarthritis, a degenerative joint disease that affects nearly 30 million Americans, is characterized by lesions of articular cartilage that often lead to severe pain and loss of joint function. The current economic burden of osteoarthritis is estimated to be approximately $190 billion, and with the prevalence of arthritis expected to rise due to the aging population, the associated costs are forecasted to increase. Debilitating osteoarthritis is managed clinically by the surgical implantation of a cartilage graft or cartilage cells to replace the damaged tissue; however, current repair methods often result in poor long- term outcomes due to inadequate integration of the graft with host cartilage and bone. Thus, there is a significant clinical need for approaches that enable functional connection of grafting devices to the host tissue. To address this challenge, the strategy described in this thesis is a versatile, cup-shaped fibrous scaffold system designed to promote the simultaneous integration of the cartilage graft with both the host cartilage and subchondral bone. This thesis is guided by the hypotheses that 1) graft integration with native cartilage can be strengthened by inducing chondrocyte migration to the graft-cartilage junction through chemotactic factor release from the walls of the cup, and 2) graft integration with host bone and the formation of calcified cartilage can be facilitated by pre-incorporation of calcium phosphate nanoparticles in the base of the cup.
To test these hypotheses, a microfiber-based integration cup was designed with degradable, polymer-based walls that release insulin-like growth factor-1, which is well-established for inducing chondrocyte migration, and a base consisting of polymer with calcium deficient apatite nanoparticles. In the first aim of this thesis, the dose of insulin-like growth factor-1 in the cup walls was optimized to enhance the migration of cells from surrounding cartilage into the scaffold, and this design was tested in vitro to ensure that the scaffold supports chondrocyte viability, growth, and biosynthesis of a cartilage-like matrix. In the second aim of this thesis, the composition and dose of calcium phosphate in the base of the cup was optimized to support chondrocyte growth and the production of calcified cartilage-like tissue.
Subsequently, in the third aim, the independently developed walls and base were joined into a scaffold
that was tested in vitro and in vivo, using a simulated full thickness defect model, to examine its potential for clinical translation. Results from these studies demonstrate that the cup system can be implemented with autologous tissue and cell-based grafting strategies as well as with tissue engineered hydrogel grafts to promote integration with host tissue. Moreover, these investigations have yielded new insights into both chemical and structural parameters that direct chondrocyte migration and calcified cartilage formation.
In summary, this thesis describes the design and optimization of a novel, multi-functional device for improving integration of cartilage grafts with host tissues. The impact of the studies in this thesis extends beyond cartilage integration, as the interface scaffold design criteria elucidated here are readily applicable to the formation of interfaces between other grafts and host tissues.
TABLE OF CONTENTS List of Figures and Tables ...... vi List of Abbreviations ...... viii Acknowledgements ...... ix Dedication ...... xi Chapter 1: Introduction ...... 1 1.1 Specific Aims ...... 2 1.2 Background and Significance ...... 5 1.2.1 The Structure and Function of Healthy Articular Cartilage and the Osteochondral Interface ...... 5 1.2.2 Age- and Disease-Related Changes ...... 8 1.2.3 Challenges in Cartilage Repair ...... 9 1.2.4 Current Efforts in Cartilage-Cartilage Integration ...... 10 1.2.5 Current Efforts in Cartilage-Bone Integration ...... 12 1.2.6 Summary ...... 13 Chapter 2: Effect of IGF-1 Dose on Chondrocyte Migration ...... 15 2.1 Introduction ...... 16 2.1.1 Background and Significance ...... 16 2.1.2 Objectives ...... 17 2.2 Materials and Methods ...... 18 2.2.1 Scaffold Fabrication and Characterization ...... 18 2.2.2 Scaffold Contraction Analysis ...... 18 2.2.3 IGF-1 Release Characterization ...... 19 2.2.4 Cartilage Harvest and Culture ...... 19 2.2.5 Cell Imaging and Quantification ...... 20 2.2.6 Statistical Analysis ...... 20 2.3 Results ...... 20 2.3.1 Polymer Blend Optimization ...... 20 2.3.2 IGF-1 Scaffold Characterization ...... 20 2.3.3 Cell Migration ...... 21 2.4 Discussion ...... 21 2.5 Conclusions ...... 23 Chapter 3: Chondrocyte Response on IGF-1 Microfiber Scaffolds ...... 29 3.1 Introduction ...... 30 3.1.1 Background and Significance ...... 30 3.1.2 Objectives ...... 30 3.2 Materials and Methods ...... 31 3.2.1 Scaffold Fabrication ...... 31 3.2.2 Cells and Cell Culture ...... 31 i 3.2.3 Cell Imaging and Number ...... 32 3.2.4 Matrix Deposition ...... 32 3.2.5 Mineralization Potential ...... 33 3.2.6 Statistical Analysis ...... 34 3.3 Results ...... 34 3.3.1 Cell Distribution, Viability, Number, and Alkaline Phosphatase Activity ...... 34 3.3.2 Matrix Production ...... 34 3.4 Discussion ...... 34 3.5 Conclusions ...... 36 Chapter 4: The Effect of Ceramic Crystallinity on Deep Zone Chondorcyte Response and Calcified Cartilage Formation ...... 40 4.1 Introduction ...... 41 4.1.1 Background and Significance ...... 41 4.1.2 Objectives ...... 42 4.2 Materials and Methods ...... 42 4.2.1 Ceramic Characterization ...... 42 4.2.2 Cells and Cell Culture ...... 43 4.2.3 Scaffold Fabrication and Culture ...... 43 4.2.4 Cell Number and Distribution ...... 44 4.2.5 Matrix Deposition ...... 44 4.2.6 Mineralization ...... 45 4.2.7 Chondrocyte Hypertrophy ...... 46 4.2.8 Media Ion Analysis ...... 46 4.2.9 Statistical Analysis ...... 47 4.3 Results ...... 47 4.3.1 Ceramic Characterization ...... 47 4.3.2 Cell Distribution and Number ...... 47 4.3.3 Matrix Deposition ...... 48 4.3.4 Mineralization Potential and Hypertrophy ...... 48 4.3.5 Media Ion Concentrations ...... 49 4.4 Discussion ...... 49 4.5 Conclusions ...... 52 Chapter 5: The Effect of Ceramic Dose on Deep Zone Chondrocyte Response and Calcified Cartilage Formation in Agarose Scaffolds ...... 63 5.1 Introduction ...... 64 5.1.1 Background and Significance ...... 64 5.1.2 Objectives ...... 64 5.2 Materials and Methods ...... 65 5.2.1 Cells and Cell Culture ...... 65 5.2.2 Scaffold Fabrication, Characterization, and Culture ...... 65
ii 5.2.3 Cell Viability and Number ...... 66 5.2.4 Mineralization ...... 66 5.2.5 Matrix Deposition ...... 67 5.2.6 Statistical Analysis ...... 68 5.3 Results ...... 68 5.3.1 Scaffold Characterization ...... 68 5.3.2 Cell Viability and Number ...... 68 5.3.3 Matrix Deposition ...... 69 5.3.4 Mechanical Properties ...... 70 5.3.5 Mineralization ...... 70 5.4 Discussion ...... 70 5.5 Conclusions ...... 73 Chapter 6: Effect of Ceramic Dose in Microfiber Scaffolds on Deep Zone Chondrocyte Response and Calcified Cartilage Formation ...... 82 6.1 Introduction ...... 83 6.1.1 Background and Motivation ...... 83 6.1.2 Objectives ...... 84 6.2 Materials and Methods ...... 84 6.2.1 Scaffold Fabrication and Characterization ...... 84 6.2.2 Cells and Cell Culture ...... 85 6.2.3 Cell Viability and Number ...... 86 6.2.4 Matrix Deposition ...... 86 6.2.5 Mineralization ...... 87 6.2.6 Statistical Analysis ...... 88 6.3 Results ...... 88 6.3.1 Scaffold Characterization ...... 88 6.3.2 Cell Viability and Number ...... 89 6.3.3 Matrix Deposition ...... 89 6.3.4 Mineralization ...... 90 6.4 Discussion ...... 91 6.5 Conclusions ...... 93 Chapter 7: In Vitro Evaluation of Interface Formation in an Osteochondral Explant Model ...... 102 7.1 Introduction ...... 103 7.1.1 Background and Significance ...... 103 7.1.2 Objectives ...... 104 7.2 Materials and Methods ...... 104 7.2.1 Explant Harvest and Culture ...... 104 7.2.2 Cells and Cell Culture ...... 104 7.2.3 Scaffold Fabrication ...... 105
iii 7.2.4 Defect Repair and Culture ...... 106 7.2.5 Histology Analysis ...... 107 7.2.6 Integration Strength ...... 107 7.2.7 Statistical Analysis ...... 108 7.3 Results ...... 108 7.3.1 Full Thickness Defect Model Characterization ...... 108 7.3.2 Gross Morphology of Repaired Full Thickness Defects ...... 108 7.3.3 Histological Analysis ...... 109 7.3.4 Integration Strength ...... 109 7.4 Discussion ...... 110 7.5 Conclusions ...... 113 Chapter 8: In Vivo Evaluation of Interface Formation in an Osteochondral Explant Model ...... 123 8.1 Introduction ...... 124 8.1.1 Background and Significance ...... 124 8.1.2 Objectives ...... 124 8.2 Materials and Methods ...... 124 8.2.1 Explant Harvest and Culture ...... 124 8.2.2 Cells and Cell Culture ...... 125 8.2.3 Scaffold Fabrication ...... 125 8.2.4 Sample Preparation ...... 126 8.2.5 Subcutaneous Implantation and Culture ...... 127 8.2.6 Histological Analysis ...... 127 8.2.7 Statistical Analysis ...... 128 8.3 Results ...... 129 8.3.1 Animal Surgery ...... 129 8.3.2 Gross Morphology of Implants ...... 129 8.3.3 Autograft Control ...... 129 8.3.4 Autograft with Control Cup ...... 130 8.3.5 Autograft with IGF Cup ...... 130 8.3.6 Chondrocyte Implantation Control ...... 131 8.3.7 Chondrocyte Implantation with IGF Cup ...... 131 8.3.8 Hydrogel Graft Control ...... 132 8.3.9 Hydrogel Graft with IGF Cup ...... 132 8.3.10 Hydrogel Graft with IGF Cup and Agarose-CDA Composite Base ...... 133 8.4 Discussion ...... 133 8.5 Conclusions ...... 137 Chapter 9: Summary and Future Directions ...... 151 9.1 Summary ...... 152 9.1.1 Scaffold Design and Optimization for Cartilage-Cartilage Integration ...... 153
iv 9.1.2 Scaffold Design and Optimization for Cartilage-Bone Integration ...... 153 9.1.3 Evaluating the Clinical Potential of the Integrative Scaffold System ...... 154 9.2 Future Directions ...... 154 9.2.1 Human Cartilage Organ Model Testing ...... 154 9.2.2 Intra-Articular in Vivo Testing ...... 155 9.2.3 Stem Cell-Seeded Scaffold ...... 155 Reference List ...... 156
v LIST OF FIGURES AND TABLES
Figure 1.1 Schematic of the integrative scaffold. Figure 1.2 Thesis aims.
Figure 2.1 Polymer blend optimization. Figure 2.2 Scaffold fabrication and groups. Figure 2.3 Insulin like growth factor-1 release from microfiber scaffolds. Figure 2.4 Experimental setup. Figure 2.5 Migration from cartilage explants onto scaffolds.
Figure 3.1 Study design. Figure 3.2 Cell viability, number, and alkaline phosphatase activity. Figure 3.3 Matrix deposition.
Table 4.1 Primer sequences for gene expression. Figure 4.1 Ceramic characterization. Figure 4.2 Study design. Figure 4.3 Cell number and distribution. Figure 4.4 Collagen deposition. Figure 4.5 Day 14 collagen immunohistochemistry. Figure 4.6 Glycosaminoglycan deposition. Figure 4.7 Alkaline phosphatase activity and mineral distribution. Figure 4.8 Gene expression. Figure 4.9 Media ion concentrations.
Figure 5.1 Study design. Table 5.1 Acellular scaffold characterization for agarose constructs with varied CDA content. Figure 5.2 Cell number, viability, and distribution. Figure 5.3 Collagen production. Figure 5.4 Collagen immunohistochemistry. Figure 5.5 Glycosaminoglycan production. Figure 5.6 Mechanical properties. Figure 5.7 Mineralization.
Figure 6.1 Study design. Figure 6.2 Incorporation of calcium deficient apatite in microfiber scaffolds. Figure 6.3 Characterization of composite scaffolds composed of microfibers and calcium deficient apatite. Figure 6.4 Scaffold degradation over time. Figure 6.5 Cell viability and number. Figure 6.6 Collagen deposition. Figure 6.7 Glycosaminoglycan content. Figure 6.8 Mineralization.
Figure 7.1 Osteochondral tissue harvest and full thickness defect generation. Figure 7.2 Characterization of the full thickness defect model. Figure 7.3 Cup fabrication. Figure 7.4 Study design. Figure 7.5 Images of repaired explants after in vitro culture. Figure 7.6 Autograft-based repair. Figure 7.7 Hydrogel-based repair. Figure 7.8 Cell implantation-based repair. Figure 7.9 Integration strength.
vi Figure 8.1 Study design. Figure 8.2 Subcutaneous rat study surgical procedure. Figure 8.3 Gross morphology after in vivo implantation. Figure 8.4 Histology overview. Figure 8.5 Autograft control repair. Figure 8.6 Autograft with control cup repair. Figure 8.7 Autograft with IGF cup repair. Figure 8.8 Cell implantation control repair. Figure 8.9 Cell implantation with IGF cup repair. Figure 8.10 Hydrogel graft control repair. Figure 8.11 Hydrogel graft with IGF cup repair. Figure 8.12 Hydrogel graft with IGF cup and calcium phosphate agarose base repair. Figure 8.13 Mineralization.
Figure 9.1 Synovium derived stem cells on microfiber scaffolds with and without IGF.
vii LIST OF ABBREVIATIONS ALP: Alkaline phosphatase ANOVA: Analysis of variance BSA: Bovine serum albumin CDA: Calcium deficient apatite DCM: Dichloromethane DMEM: Dulbecco’s Modified Eagle’s Medium DMF: Dimethylformamide DMB: Dimethylmethylene blue DNA: Deoxyribose nucleic acid DW: Dry weight DZC: Deep zone chondrocyte EDXA: Energy dispersive X-ray analysis ELISA: Enzyme-linked immunosorbent assay FBS: Fetal bovine serum FTC: Full thickness chondrocyte FTIR: Fourier transform infrared spectroscopy GAG: Glycosaminoglycan HA: Hydroxyapatite H&E: Hematoxylin and eosin ICP: Inductively coupled plasma IGF-1: Insulin-like growth factor-1 ITS: Insulin transferrin, and selenous acid Ihh: Indian hedgehog IR: Infrared MMP: Metalloproteinase MSC: Mesenchymal stem cell NEAA: Non-essential amino acids PBS: Phosphate buffered saline P/S: Penicillin/Streptomycin PCL: Polycaprolactone PCR: Polymerase chain reaction PEGDA: Poly(ethylene glycol) diacrylate PLGA: Polylactide-co-glycolide pNP: P-Nitrophenol PTHrP: Parathyroid hormone-related protein RT-PCR: Reverse transcription – polymerase chain reaction SEM: Scanning electron microscopy T3: Thyroid hormone or triiodothyronine TCP: Tricalcium phosphate TGA : Thermogravimetric analysis TMOS: Tetramethyl orthosilicate WW: Wet weight XRD: X-ray diffraction
viii ACKNOWLEDGEMENTS
Throughout the years I have spent at Columbia University, I have had the privilege of working with an exceptional group of faculty, students, and staff. I would like to thank my advisor, Professor Helen
H. Lu, who has provided guidance for my research studies as well as my professional development. In addition, my thesis committee—including Professor Gerard A. Ateshian, Professor Clark T. Hung,
Professor Ernst B. Hunziker, and Professor Eric J. Strauss—has been a source of knowledge, wisdom, and inspiration. I am also grateful to the many collaborators who have been extremely generous with their time and continuously provided valuable insight to guide my research.
I would like to thank my colleagues in the Biomaterials and Interface Tissue Engineering
Laboratory who played an instrumental role in helping me perform the research outlined in this thesis. I am particularly grateful to my fellow graduate students and post-doctoral scholars in my lab, namely
Kristen Moffat, Nora Khanarian, Siddarth Subramony, Philip Chuang, Sagaw Prateepchinda, Xinzhi
Zhang, Nancy Lee, Danielle Bogdanowicz, Dovina Qu, Christopher Mosher, Yichen Zhao, Cevat Erisken,
Marissa Solomon, and Jennifer Robinson as well as the undergraduate and high school students that worked directly with me, including Wendy Sun, Mary Quien, Elizabeth Dente, Kevin Caparino, Andrew
Zhao, Gurbani Suri, Mishaelle Gomez, and Regan Dvoskin. Furthermore, I would like to acknowledge the contributions of several faculty members, staff members and graduate students from other laboratories and departments, including Dr. Alissa Park, Keith Yeager, Bin Zhou, Eric Yu, Michelle Cintron, Paulette
Louissant, and Shila Maghi. I would also like to gratefully acknowledge the contributions of the research faculty and staff at the Hospital for Special Surgery, including Dr. Stephen Doty, Orla O’Shea, and
Anthony Labissiere, the staff at the Columbia University animal facility, including Dr. Rivka Shoulson, Dr.
Samuel Baker, and Dr. Nicole Herndon as well as Professor Raquel Z. LeGeros from New York
University.
In addition to the people that have directly supported my research and professional development as a graduate student, I would like to thank the outstanding research advisors that inspired me and encouraged me to pursue a career in scientific research, starting in my childhood. I am forever thankful to my first scientific mentor, my Dad, for introducing me to science at a young age and cultivating a deep curiosity for scientific exploration within me. I am also thankful for Frank LaBanca, who mentored me in ix my first independent research project in high school and for Professor David Kaplan, who highlighted the merit of an engineering training and biomaterials research, leading me to pursue a degree in chemical engineering, complemented by biomedical engineering research. I would like to gratefully acknowledge
Professors Curtis Frank, Helen Blau, and Paula Hammond for giving me the opportunity to work in their labs as a visiting summer student during my undergraduate tenure and Drs. Karen Havenstrite, Penny
Gilbert, Anita Shukla, Michaela Regan, and Charu Vepari for their mentorship.
Research funding for the studies in this thesis has been provided by the National Institute of
Arthritis and Musculoskeletal and Skin Diseases, the National Institutes of Health (RO1-AR055280 and
T32 AR059038), New York Stem Cell Foundation (C029551), The Columbia Center for Technology,
Innovation and Community Engagement, and by Neil and Mindy Grossman. The collagen X antibody developed by Thomas F. Linsenmayer, was obtained from the Developmental Studies Hybridoma Banks developed under the auspices of the NICHD and maintained by the University of Iowa, Department of
Biology, Iowa City, IA 52242.
x DEDICATION
This thesis is dedicated to my husband who provides indefatigable support
and makes me smile every day.
xi CHAPTER 1: INTRODUCTION
1 1.1 Specific Aims
Osteoarthritis, a degenerative joint disease, is a leading cause of disability among Americans [1]. This condition is characterized by lesions of articular cartilage that often lead to severe pain and loss of joint function. Since cartilage has a limited capacity for self-repair [2], surgical intervention is often required for treatment. Current repair methods are associated with poor long-term outcomes due to unwanted fibrocartilage formation and inadequate repair tissue integration with both host cartilage and subchondral bone [3-6]. A variety of hydrogel-based cartilage grafts have been investigated for cartilage repair with promising results [7-12]; however, the integration of cartilage grafts with host cartilage remains a translational barrier. In addition to structural contiguity between neo and host cartilage, the presence of a structural barrier between the healing cartilage and bone is also critical to limit osseous invasion and maintain the integrity of the repaired cartilage [13].
To address the challenge of graft integration, the goal of this thesis is to develop and optimize a bioactive integration scaffold to enable the biological fixation of the cartilage graft with both the host cartilage and subchondral bone. For cartilage-cartilage integration, the ideal scaffold should promote cell migration into the region between the graft and native cartilage and encourage neo-cartilage formation.
For cartilage-bone integration, the scaffold should serve as a temporary barrier during healing while integrating the graft to the bone by promoting the formation of calcified cartilage tissue that is rich in collagen II and proteoglycans.
The research strategy described in this thesis centers on the design of a cup-shaped scaffolding polymer cup
system in which the walls of the cup are designed for growth factor cartilage cartilage bioactive cap cartilage-cartilage integration, and the base of the cartilage nanoparticles graft hydrogel cup is designed to promote cartilage-bone integration Figure 1.1 Schematic via osteochondral interface regeneration (Fig. 1.1). of the integrative scaffold with optimized phases for Poor graft integration with native cartilage results integration with the surrounding bone from reduced cell density at the wound edges caused bone and cartilage by cell death associated with surgical intervention. Stimulating cell migration to the graft-cartilage boundary with chemotactic factors has been reported to improve cartilage-cartilage integration [14]; thus,
2 the walls of the cup will be designed to promote cartilage integration by releasing insulin-like growth factor-1 (IGF-1), which will be optimized in Aim 1. For cartilage-bone integration, hydrogel scaffolds promote the formation of calcified cartilage-like matrix [11;15], fibrous ceramic scaffolds have been shown to enhance scaffold-bone integration [16], and the combination of hydrogel and fibrous scaffolds has been shown to result in organized calcified cartilage formation [17]. Therefore, in Aim 2, both hydrogel- and fiber-based ceramic composites will be investigated as components for the base of the cup. Ceramic properties and dosage will be optimized to promote calcified cartilage formation. In Aim 3, the walls will be joined to the base to form a cup, and the system will be evaluated in vitro and in vivo using a full thickness defect model. The working hypothesis of this thesis is that a scaffold with IGF-1-releasing walls and a mineral-containing base can enhance graft integration, thereby improving long-term success of cartilage repair. Accordingly, the three research aims and related hypotheses are:
Aim 1: Design, characterize, and optimize a polymer fiber-based scaffold for cartilage- cartilage integration Hypothesis: IGF-1 release can be optimized to produce a scaffold that homes cells and supports cell-mediated cartilage formation
Aim 2: Design, characterize, and optimize a polymer-ceramic composite scaffold for cartilage-bone integration Hypothesis: Ceramic composition and dose can be optimized to enhance chondrocyte viability, growth, and matrix production in agarose and microfiber scaffolds
Aim 3: Fabricate and evaluate the cup scaffold system in an explant model Hypothesis: The phases optimized in Aims 1 and 2 can be combined to support the integration of a cartilage graft with the surrounding tissue without impairing viability or matrix maintenance of native cartilage and bone
Specifically, Aim 1 will center on the fabrication and optimization of a bioactive polymeric scaffold to promote cartilage-cartilage integration. An IGF-1-releasing, degradable scaffold will be fabricated to bridge the gap between the cartilage graft and native tissue, supporting localized matrix elaboration over time. Insulin-like growth factor-1 promotes chondrocyte and stem cell migration [14;18] and will therefore be incorporated into the scaffold to attract cells. Electrospinning will be used to generate fibrous scaffolds because it allows tuning of material properties [19;20] and incorporation of growth factors [21-23]. The dose of incorporated IGF-1 will be optimized to maximize cellular infiltration (chapter 2), and the response of chondrocytes, which will likely be responsible for neo cartilage formation in the cup walls will be
3 determined (chapter 3). Overall, the anticipated benefit of the cup walls is that IGF-1 release will induce cell migration to the native tissue-graft junction, resulting in matrix deposition that integrates the host and graft cartilage.
Aim 2 will focus on the optimization of a scaffold that promotes cartilage-bone integration through the regeneration of calcified cartilage. Recent preliminary results from a rabbit in vivo study revealed that the combination of agarose-hydroxyapatite (HA) and poly(lactide-co-glycolide) (PLGA)-HA scaffolds resulted in more organized interface formation than a hydrogel or fibrous scaffold alone. This result suggests that a polymer-hydrogel-ceramic hybrid scaffold may be optimal for calcified cartilage regeneration [17].
Furthermore, it is hypothesized that matrix elaboration can be further enhanced with the incorporation of a bioactive ceramic. To this end, the ceramic phase of the scaffold will be optimized in terms of composition
(chapter 4) by evaluating deep zone chondrocyte (DZC) proliferation and matrix production over time in an agarose hydrogel scaffold. The dose of the bioactive ceramic will be optimized in both agarose
(chapter 5) and microfiber (chapter 6) scaffolds for the base of the cup. It is anticipated that a polymer- hydrogel-ceramic scaffold with an optimal bioactive ceramic dose will support the generation of a dense, mineralized matrix that is rich in both proteoglycans and collagen, similar to the calcified cartilage.
After the completion of Aims 1 and 2, the translational potential of the integrative scaffold system will be evaluated in Aim 3. Specifically, the cartilage-cartilage integration scaffold will be attached to the cartilage-bone integration scaffold to build a cup, and the optimized agarose-ceramic scaffold will be added to the base. A full thickness defect model will be developed, characterized, and used to test the efficacy of the scaffold system in terms of integration with both cartilage and bone in vitro (chapter 7) and in vivo (chapter 8). Matrix formation will be evaluated using histology, with a focus on cellularity as well as proteoglycan and collagen deposition at the graft-native tissue junctions.
In order to promote integrative cartilage repair, the studies outlined in this proposal consist of (1) complex scaffold design and optimization, as well as (2) development and implementation of a full thickness cartilage defect model. The proposed approach is unique because it aims to address both the integration of a cartilage graft with the native cartilage and with bone through osteochondral interface regeneration. It is anticipated that the optimized scaffold system will improve integration of cartilage grafts and long-term clinical success. Moreover, these studies have significance in fields beyond cartilage
4 repair, as they will likely establish key parameters for modulating cell-ceramic response, establish scaffold design criteria that can be applied at other graft-tissue junctions, and develop organ culture parameters that can be extended to new explant culture systems.
Aim 1 Cartilage-to-cartilage
cartilage integration cartilage cartilage graft . Optimize IGF-1 dose . Characterize cell response
polymer cup Aim 3
growth factor Scaffold testing cartilage cartilage in explant model bone cartilage graft Evaluate interface formation in an explant model: Aim 2 . in vitro Cartilage-to-bone . in vivo
integration cartilage cartilage cartilage graft Optimize ceramic: bone . composition . dose in agarose . dose in microfibers
bioactive CaP nanoparticles hydrogel bone Figure 1.2 Thesis aims
1.2 Background and Significance
1.2.1 The Structure and Function of Healthy Articular Cartilage and the Osteochondral Interface
Healthy cartilage tissue is composed of liquid and solid phases that together facilitate load bearing and joint articulation [24]. The extracellular matrix is actively maintained by resident chondrocytes and is rich in proteoglycans [25;26] as well as collagen II, with smaller amounts of collagen III, VI, IX, X, and XI present [27-30]. The charged proteoglycans attract water that swells the tissue, allowing it to support compressive loads, while the tensile properties of the collagen network contribute swelling resistance and shear strength [24;31-35]. In addition, interstitial fluid pressurization, combined with molecules released from the cartilage surface and synovium, lubricate the joint surfaces, facilitating painless joint motion [36].
5 Cartilage is organized into four structurally contiguous zones defined by variations in matrix composition, structural organization, and chondrocyte phenotype [37-39]. The surface zone, located at the articulating edge of the cartilage, accounts for approximately 10% of the height of the total tissue. This zone consists of thin, elliptical chondrocytes combined with progenitor cells that are surrounded by a matrix with high water content (~78%), low proteoglycan content relative to the other zones, and collagen fibrils (4-12 nm) that are oriented parallel to the surface [40-50]. The cells in this zone produce superficial zone protein that contributes to the lubrication of the joint [51;52]. Below the surface, the next 40-60% of the cartilage depth consists of the middle zone, within which spherical chondrocytes reside in a matrix rich in proteoglycans and randomly aligned collagen fibrils (9-60 nm) [44;46-49]. The deep zone, located below the middle zone, comprises approximately 30% of the cartilage depth and is marked by spherical chondrocytes oriented in stacks that are perpendicular to the articular surface. These cells, while sparsely distributed, maintain a matrix of relatively high glycosaminoglycan (GAG) content, relatively low water content (~68%), and radially oriented collagen fibrils (60-140 nm) [41;45;47-50]. Although the collagen fibrils generally increase in diameter from the surface to the deep zone, fine fibrils have been noted throughout the depth of the cartilage in human specimens [45-47].
Between the deep zone and the subchondral bone is the osteochondral interface which is comprised of hypertrophic chondrocytes embedded in a mineralized matrix rich in collagen II and proteoglycans. In humans, the calcified cartilage is reported to range from 20 to 243 µm in thickness, with this wide range of reported values accounted for by variability in age and total cartilage thickness [39;53;54]. In a study that histologically evaluated decalcified healthy human cartilage after exposure to a variety of enzymatic digestions, Fawns et al. first coined the term “tidemark” to reference the line that marks the limit of calcification [55]. Using scanning and transmission electron microscopy (SEM and TEM, respectively),
Bullough et al. directly imaged the calcification front of adult canine cartilage harvested from the tibia plateau after removing the organic matrix either chemically or via exposure to high temperatures.
Although the calcified cartilage is tightly interlocked with the underlying subchondral bone, a distinct border between the two tissues was observed that is actively regulated by living chondrocytes [56]. While the subchondral bone is highly vascularized, SEM imaging of mature human, lapine, and canine cartilage-
6 bone junctions revealed that most blood vessels terminate at the interface in healthy tissue and are separated from the calcified cartilage by a layer of bone [57].
The calcified cartilage matrix is composed primarily of proteoglycans, mineral, and collagen II, IX, and
X [27;58-60]. Highly aligned collagen II fibers run continuously from the deep zone into the calcified cartilage layer [61-63]. The alignment of the collagen bundles at the transition and the interdigitated nature of the calcified cartilage-bone junction together serve to anchor the non-calcified cartilage to the bone [62;64]. In a study by Redler et al., 30 human osteochondral specimens were examined using SEM, and the authors proposed that the undulating junction provides an ideal geometric configuration to resist the shearing action of articulation [64]. In addition, the network of branching collagen fibrils at the interface also contributes to the load bearing capabilities of the joint by diffusing and distributing load during force transmission from the cartilage to bone, reducing stress concentrations at the interface
[62;64].
In addition to anchoring the cartilage to the bone, the collagen fibers also serve as a template for mineralization [55;56]. In a study that characterized the mineral in human bone and calcified cartilage,
Zizak et al. noted a striking change in the mineral particle orientation from perpendicular to the interface in the calcified cartilage to parallel to the interface in the bone, reflecting the underlying alignment of the collagen fibers [65]. While the collagen fibers run continuously from the calcified cartilage to the deep zone, the mineral is localized to the calcified cartilage and bone regions, with an exponential increase in mineral content between non-calcified and calcified regions that persists with age [66]. This calcified collagen and proteoglycan network imparts strength to the calcified cartilage which has been characterized via a three-point bending test and nanoindentation. Using a three-point bending test, bovine calcified cartilage stiffness has been measured to be approximately 0.32 ± 0.25 GPa, which is intermediate between uncalcified cartilage and bone [67]. A modulus ranging from 10-30 GPa [68;69] has been reported for nanoindentation measurements of human femoral calcified cartilage. Interestingly, a direct positive correlation between mineralization and mechanical properties was demonstrated in adult human femoral calcified cartilage [69].
The calcified cartilage mineral is poorly crystalline carbonated hydroxyapatite that is similar to bone mineral in terms of both chemistry and size (2-4.2 nm) [59;61;65;70]. It is reported that calcified cartilage
7 has a higher calcium content than bone, although values reported for the calcium content of the interface range from 1-28 wt% [65;68]. The dense, mineralized matrix acts as a barrier that limits diffusion and prevents osseous invasion from the subchondral bone [13]. In the calcified cartilage of mature murine [71] and equine [72] specimens, the diffusion coefficient of small molecules (~400 Da) was 0.26 and 0.9
µm2/s, respectively. In the equine model, the diffusion coefficient in the calcified cartilage was five-fold lower than in the uncalcified region, highlighting the barrier effect of the calcified cartilage layer. However, using fluorescence loss induced photobleaching (FLIP), patches of nonmineralized regions were found that may serve as transport pathways for small molecules, accounting for 22% of the volume of calcified cartilage in the murine model. This finding suggests that the calcified cartilage may have a complex, multifaceted role in the transport of molecules between the bone and articular cartilage [71].
1.2.2 Age- and Disease-Related Changes
Structural and functional changes in cartilage with age are well established and have recently been reviewed in detail [73]; however, a brief summary is provided here. Hallmarks of the normal aging process in human cartilage include decreased thickness and cellularity [74] as well as an accumulation of advanced glycation end-products that alter collagen crosslinking, disrupting matrix integrity [75]. The earliest changes are observed in the superficial zone, with a 50% decrease in chondrocyte density reported between 20 and 90 years of age. As cell density declines, it is likely that intrinsic repair is further diminished, leading to an increased risk of fibrillation and osteoarthritis [74].
Changes are also observed in the calcified cartilage with age. The thickness of human calcified cartilage decreases with age [53] and becomes less permeable as it transitions to serve as a barrier to solute diffusion from the bone in adult years [76]. A steady decline in the number of blood vessels is observed with age in human femoral and humoral heads. This decline continues into the seventh decade of life, at which point the trend reverses to increased vascularization [77]. Parallel trends in remodeling have been reported, suggesting that blood vessel presence may be correlated to an active remodeling state [77].
Accumulation of these age-related changes can lead to osteoarthritis. This disease is a result of the disruption between the inherent homeostasis of anabolic and catabolic processes and leads to fibrillation of the articulating surface, osteophyte formation, and subchondral bone thickening [49;78-80]. This
8 process may be initiated gradually by wear that occurs during aging or rapidly due to trauma, such as a ligamental or meniscal injury [81]. While the specific factors that initiate joint degeneration are unknown,
Bullough noted that age-related degeneration first occurs in areas of unloaded cartilage. From this observation, he proposed that, as the joint remodels with age, areas of cartilage which were previously unloaded and had thus degenerated are exposed to new levels of load, leading to overtaxing. This process results in further degeneration that may lead to osteoarthritis [82;83]. Conversely, osteoarthritis may begin with changes in the subchondral bone, such as cracking, that result from overloading and occur beneath the intact hyaline cartilage, leading to subchondral bone collapse and eventual lesion formation in the articular cartilage [84].
Although catabolic processes overtake anabolic activities, the disease state is accompanied by an increase in overall chondrocyte activity [85]. The net loss of proteoglycans and the disruption of the collagen network have been attributed to increased matrix metalloproteinase (MMP) and aggrecanase activity as well as changes in chondrocyte phenotype, such as collagen III production [86;87]. Shifts in the underlying matrix architecture consequently result in lower compressive and tensile mechanical properties [32;88-90], ultimately compromising joint function.
1.2.3 Challenges in Cartilage Repair
Inherently, cartilage has a limited capacity for self-repair due to its avascular and aneural nature [2].
Consequently, spontaneous healing does not occur in partial thickness defects. In full thickness defects, spontaneous repair tissue can be generated, as bleeding from the bone carries both cells and growth factors into the cartilage lesion. In such cases, the regenerated tissue is fibrocartilaginous in nature, mechanically inferior to native tissue, and usually does not persist over time [2;91;92].
Surgical intervention is often implemented to encourage a healing response in partial thickness defects and improve healing in full thickness lesions. Clinical treatments that provide access to the osseous vascular supply include subchondral drilling and microfracture [93-95]. Larger defects are often treated with osteochondral transplantation, in which plugs from cadavers or non-load bearing regions of the patient’s own joint are transplanted into the damaged area [96;97]. Cell-based approaches [98] and tissue engineered constructs have also emerged as a promising alternative to existing treatment options.
Despite their promise, current grafts have focused primarily on cartilage and bone regeneration, and as a
9 result, two challenges are integration of cartilage grafts with 1) native articular cartilage and 2) subchondral bone. Cartilage-cartilage integration is critical for stable repair because, in the absence of continuity between host and neo cartilage, micromotion between the graft and adjacent tissue can fuel further cartilage degeneration [99]. Regeneration of the osteochondral interface is also necessary for stable repair because this calcified cartilage layer contributes to graft-bone integration, mechanical functionality, and long-term stability. The importance of a structural barrier separating the bone and healing cartilage was demonstrated by Hunziker et al. using a full-thickness cartilage defect model. It was observed that a structural barrier, in this case a Gore-Tex® membrane (0.2 µm pore diameter), placed between the cartilage and bone compartments was necessary to maintain the integrity of the newly formed cartilage, largely by limiting vascular ingrowth from the subchondral bed and preventing ectopic mineralization [13].
1.2.4 Current Efforts in Cartilage-Cartilage Integration
Cartilage-cartilage integration is necessary for incorporation of graft systems into the defect over time.
Surgical preparation for cartilage graft placement involves the removal of the damaged cartilage directly surrounding the lesion to ensure that the graft is well shouldered by healthy tissue. This process can result in chondrocyte apoptosis and necrosis [100], leaving a hypocellular region surrounding the defect
[101]. As a result, the integration of the new cartilage graft with surrounding native tissue is a significant hurdle [102;103]. Approaches to overcome this challenge have largely focused on increasing cellularity in the boundary region via chemotactic agents that draw viable cells into the gap between the graft and native tissue or digestion rinses that break down the dense matrix at the border of autografts/allografts to facilitate cell migration [104-107]. The use of digestion reagents, such as hyaluronidase [104-106], collagenase [104-106], and chondroitinase ABC [104;107;108], have been investigated. Lee et al. demonstrated the utility of digestive rinses for improving cartilage integration by showing that treatment of cartilage explants with chondroitinase ABC (1 U/mL for 5 minutes or 0.5 U/mL or 1 U/mL for 15 minutes) led to enhanced chondrocyte adhesion, as measured by a micropipette micromanipulation system [109].
Similarly, treatment with collagenase at 10 U/mL [106] and 30 U/mL [105] increased cellularity in the edges of bovine full thickness explants in vitro. Combined treatment with hyaluronidase (0.1%, 24 hours) followed by collagenase (10 U/mL or 30 U/mL, 24 hours) resulted in cellular concentrations in the wound
10 edge that exceeded that of healthy tissue [106]. By adjusting the doses and treating with the two enzymes simultaneously (300 U/mL collagenase with 3% hyaluronidase for 1 hour), improved cartilage-cartilage integration was demonstrated histologically between bovine explants that were cultured subcutaneously in an athymic rat for 35 days [106]. The safety of these treatments has been investigated by Quinn &
Hunziker with regard to changes in cell density, structure, and cell-mediated matrix deposition that result from choindroitinase ABC treatment (1 U/mL for 5 minutes). While chondroitinase ABC treatment did appear to further decrease cell densities within 100 µm of the defect surfaces, the decrease was small compared to the effect of the defect itself and was contained to the area immediately adjacent to the defect (within 100 µm), demonstrating that digestive treatments can be spatially controlled [108].
Given the low cell density naturally surrounding defects and the further reduction in cell number immediately following digestive rinses, methods that actively recruit cells into the edge of the graft are an attractive option to draw viable cells to the wound edge. Chemotactic agents that have been explored for this purpose include platelet derived growth factor (PDGF) [14;110], insulin-like growth factor I (IGF-1)
[14;18;110], basic fibroblast growth factor (bFGF) [14;110], vascular endothelial growth factor (VEGF)
[110], and a variety of factors from the bone morphogenic protein (BMP) family [110]. Boyden chamber assay results indicate that PDGF, IGF-1, and bFGF effectively stimulate migration of bovine articular chondrocytes at 25, 50, and 100 ng/mL but are not effective at lower doses, such as 5 ng/mL [14]. A combination of matrix digestion and chemotactic stimulation was evaluated by McGregor et al. using bovine metacarpophalangeal joint cartilage, in which the zone of death resulting from the OATS™ harvesting tool was approximately 173 μm. Several growth factors alone and in combination with a collagenase rinse led to an improvement in the repopulation of the zone of death. Combined treatment with collagenase (0.6%, 10 mins) and IGF-1 (25 ng/mL) resulted in the greatest reduction in the depth of the death zone (~50% of the untreated control) at four weeks [14].
More recently, material-based approaches have been investigated. For example, Wang et al. designed a “glue” by functionalizing chondroitin sulfate with methacrylate and aldehyde groups that form chemical bonds with both the native tissue and an acrylate-based scaffold [111]. This glue was tested in vitro via application to a cartilage explant and in vivo using a subcutaneous mouse model. The glue results in fixation of the poly(ethylene glycol) diacrylate (PEGDA) hydrogel construct to the cartilage,
11 exhibiting an integration strength that exceeds the bulk strength of the PEGDA hydrogel. While this system is a promising solution for integrating PEGDA-based hydrogels to the surrounding cartilage, it requires specific reactive chemical groups in the hydrogel in order to be an effective adhesive. Maher et al. developed a biodegradable, chondrocyte-laden nanofibrous hydrogel for the gap between the native and newly formed cartilage in order to improve the mechanical stability of the interface [112]. When the seeded scaffold system was tested in an in vitro cartilage gap model and supplemented with transforming growth factor β3 in the media, matrix elaboration was enhanced, with a resultant increase in maximum push-out strength. While this study highlights the importance of growth factors and cellularity in achieving cartilage-cartilage integration, the scaffold system discussed did not provide a mechanism to achieve local growth factor delivery or promote increased cell migration into the scaffold in vivo.
1.2.5 Current Efforts in Cartilage-Bone Integration
Tissue engineering approaches to achieve cartilage-bone integration have included single phase, multiphase, and gradient scaffold designs. Single phase scaffold designs have been evaluated in which cells are seeded onto bone scaffolds in order to generate calcified cartilage and cartilage-like tissue atop a bone-like material [113-117]. In addition, single phase hydrogel scaffolds which aim to regenerate the calcified cartilage layer and are designed to integrate with the underlying bone have been investigated
[11;15]. Distinct from single-phase scaffolds, bi-layered scaffolds have been developed that combine cartilage and bone scaffolds using fibrin [118-121], suturing [122], polymer polymerization [123], layered hydrogel gelation [124], and electrospinning [125]. While these studies aimed to regenerate both cartilage and bone, interface regeneration was often not the focus, and therefore minimal analysis was performed to evaluate this end-point.
Tri-phasic scaffolds that include a third distinct phase between the cartilage and bone have been designed to more closely mimic the native osteochondral architecture. For example, Kon et al. developed an acellular, tri-layered osteochondral scaffold that utilized a combination of calcium phosphate and extracellular matrix components to recapitulate the interface. The upper, intermediate, and lower layers consisted of 100% collagen I, 60% collagen I and 40% HA, and 30% collagen I and 70% HA, respectively
[126]. The layers were joined via freeze drying and tested in an equine osteochondral defect. After in vivo culture, distinct unmineralized and mineralized regions were present. More recently, this scaffold was
12 tested in a pilot clinical trial of 30 patients in which safety and potential clinical benefit were demonstrated after a two year follow-up [127]. A novel scaffold composed of agarose hydrogel and composite
PLGA/45S5 bioactive glass microspheres supported the region-specific co-culture of chondrocytes and osteoblasts. This design led to the production of three distinct yet continuous regions containing cartilage, calcified cartilage and bone-like matrices [128]. Marquass et al. introduced a tri-phasic scaffold consisting of a TCP layer to mimic bone and a collagen I hydrogel to mimic cartilage that were joined together by an intermediate activated plasma phase. When seeded with autologous mesenchymal stem cells (MSCs), the tri-phasic scaffold performed comparably to osteochondral autografts after one year in an ovine model
[129]. In a study by Cheng et al., MSC-collagen microspheres were pre-differentiated into chondrocytes and osteoblasts using a novel collagen microencapsulation technology [130]. The microspheres were aggregated to form chondrogenic and osteogenic layers in an osteochondral scaffold with a middle layer of undifferentiated MSCs. Formation of a calcified cartilage interface occurred only in the group with all three layers, while no interface was observed when undifferentiated MSCs were cultured on bi-layered scaffolds with either chondrogenic or osteogenic layers. Heymer et al. fabricated a tri-layered scaffold with a hydrophobic interface to separate the cartilage and bone portions of the scaffold, comprised of collagen I fibers and polylactic acid, respectively [131]. When stem cells were seeded on the polylactic acid portion above the interface and cultured for three weeks in vitro, cartilage-like tissue was formed.
Scaffold designs utilizing a compositional gradient have also been developed for osteochondral tissue repair [132-140]. While gradient scaffolds may offer a promising approach to achieve integration between cartilage- and bone-like phases, many of these scaffolds have yet to be tested in vitro or in vivo, and cellular studies that have been performed are preliminary, with a focus on achieving distinct cartilage and bone regions and little focus on the interface region.
1.2.6 Summary
Despite the promise of tissue engineering for osteochondral regeneration, achieving stable and integrative cartilage repair remains a significant clinical challenge. In addition, the native osteochondral interface, which is critical for cartilage maintenance over time, is not re-established using current tissue engineering approaches. As a result of these shortcomings, there is a significant clinical need for alternative repair solutions that encourage graft integration. In order to accomplish these goals, the
13 studies in this proposal consist of 1) complex scaffold design and optimization, as well as 2) development and implementation of a full thickness defect explant model. Scaffold phases for the cartilage-cartilage integration and cartilage-bone integration will be individually optimized. Subsequently, the two phases will be joined to form an integrative scaffold system to augment cartilage grafts and tested in an explant model.
14 CHAPTER 2: EFFECT OF IGF-1 DOSE ON CHONDROCYTE MIGRATION
15 2.1 Introduction
This thesis begins by focusing on the challenge of cartilage-cartilage integration. Hypocellularity at the graft-host interface contributes to the lack of integration between grafted and host cartilage [102;103].
The results of several published studies indicate that growth factors that induce cell homing improve integration [14;112] and repair [141]. Inspired by these findings, this study optimizes the release of insulin-like growth factor-1 from a microfiber scaffold that can be placed at the graft-host interface to promote cell migration from adjacent cartilage tissue.
2.1.1 Background and Significance
This study optimizes a microfiber scaffold, which releases insulin-like growth factor-1, that can be placed between a cartilage graft and the surrounding host tissue to promote cell migration to the healing cartilage-cartilage interface. Cartilage integration is hindered by the low cell density at the junction of the graft and native cartilage. In addition to the naturally low cellularity of cartilage [101], the trauma associated with surgical intervention can further reduce the cell density of the tissue at the wound edge via induction of chondrocyte apoptosis and necrosis [100], leaving a hypocellular region surrounding the defect [101]. To address this barrier to healing, it is envisioned that a thin, insulin-like growth factor-1- releasing scaffold, which lines the cartilage defect and encases the cartilage graft, will home cells directly to the interface of the graft and host tissue, thus facilitating cell-mediated integration.
To date, most cartilage-cartilage integration strategies have aimed to increase cellularity in the boundary region either by digestion rinses that break down the dense matrix at the border of autografts/allografts to facilitate cell migration [104-107] or chemotactic agents that draw viable cells into the gap between the graft and native tissue. Specifically, insulin-like growth factor-1 has been reported to increase chondrocyte migration in boyden chamber assays [14;18] and decrease the zone of cell death in cartilage explants when introduced via media supplementation at dose of 25 ng/mL [14].
The necessity of recruiting cells to the defect site has been demonstrated using a miniature pig model in which fibrin with incorporated chemotactic and mitogenic factors was used to repair a partial thickness defect [141]. This one-step repair approach demonstrated the promise of growth factor delivery from a scaffold to achieve cartilage repair. More recently, material-based integration approaches have been investigated. Maher et al. developed a biodegradable, chondrocyte-laden, nanofibrous hydrogel for the
16 gap between the native and grafted cartilage in order to improve the mechanical stability of the interface
[112]. When the seeded scaffold system was tested in an in vitro cartilage gap model and supplemented with transforming growth factor-β3 in the media, matrix elaboration was enhanced, with a resultant increase in maximum push-out strength. This study highlighted the importance of growth factors and cellularity to cartilage-cartilage integration; however, the reported scaffold system did not provide a mechanism to achieve local growth factor delivery or promote increased cell migration into the scaffold in vivo.
Building on these findings, this aim will design and optimize a microfiber scaffold that can locally deliver insulin-like growth factor-1 to the healing cartilage-cartilage interface. Since the scaffold is designed to encircle a cartilage graft, it is important that the mesh does not contract in physiologic conditions, as this would result in the withdrawal of the scaffold away from the host cartilage. To ensure structural stability, a combination of polycaprolactone (PCL), which does not contract in physiological conditions [142], and poly(lactide-co-glycolide) (PLGA), which has been previously used to culture deep zone chondrocytes, will be used [143]. These polymers are well-studied, biodegradable materials that support cartilage formation in vitro [138;144] and in vivo [144-146]. For the homing agent, several growth factors have been shown to enhance chondrocyte migration; however, insulin-like growth factor-1 is chosen for this application because, in addition to enhancing migration of chondrocytes and stem cells
[14;18], it promotes cartilaginous matrix deposition, [147;148] which is needed to connect the host and graft cartilage. Although there are promising findings reported in the literature for insulin-like growth factor-1 media supplementation in cartilage tissue engineering applications, the effects of releasing insulin-like growth factor-1 from microfibers on cell migration from cartilage has not yet been studied.
2.1.2 Objectives
The goal of this study is to fabricate microfibers that release insulin-like growth factor-1 and optimize the dose to maximize cell migration from cartilage explant tissue. It is anticipated that incorporation of insulin-like growth factor-1 incorporation will increase cell migration in a dose-dependent manner compared to an insulin-like growth factor-1-free control.
17 2.2 Materials and Methods
2.2.1 Scaffold Fabrication and Characterization
Unaligned microfiber scaffolds were fabricated by electrospinning [20;149]. For PLGA scaffold fabrication, a 54% (w/v) solution of PLGA (85:15, DL, High IV, Lakeshore Biomaterials) was mixed with
N,N-dimethylformamide (DMF, Sigma-Aldrich) and ethyl alcohol. For PLGA:PCL blended microfiber fabrication, PLGA and PCL (Sigma-Aldrich, Mw ≈ 70,000-90,000) were dissolved at varying ratios in a mixture of 60/40 dichloromethane (DCM, Sigma-Aldrich) and DMF. For fibers containing insulin-like growth factor-1 (IGF-1, Invitrogen), finely ground BSA (5% w/w, Sigma Aldrich) was added directly to a solution of 32% polymer (5:1 PLGA:PCL) in 60/40 DCM/DMF and vortexed continuously. After one hour,
IGF-1, suspended in distilled water at a concentration of 5 mg/mL, was added to the polymer melt, and the solution was vortexed for an additional hour. Each polymer solution was loaded into a 5 mL syringe with a stainless steel blunt tip needle (26.5 gauge for PLGA:PCL blends and 18 gauge for polymer-protein blends) that was 13 cm from the collecting target and electrospun at 8-10 kV using a custom electrospinning device. The polymer solution was deposited (1 mL/hour for PLGA and PLGA:PCL blend solutions, 0.8 mL/hour for polymer-protein solutions) onto a stationary collecting target using a syringe pump (Harvard Apparatus).
As-fabricated microfiber scaffolds were imaged with SEM (2 kV, Hitachi 4700, Hitachi Ltd.) to evaluate fiber morphology. Scaffolds were sputter-coated (Cressington 108auto) with gold-palladium to reduce charging effects. Microfiber diameter was quantified via image analysis of SEM micrographs using
ImageJ (National Institutes of Health, n=2).
2.2.2 Scaffold Contraction Analysis
To determine the contraction properties of blended polymer microfiber scaffolds, 10 mm scaffold disks were excised from meshes using a biopsy punch (Sklar) and sterilized via ultraviolet light exposure for 15 minutes on each side. Sterilized scaffold disks were submersed in DMEM at 37ºC and 5% CO2 and collected at 0, 2, 8, 24, and 72 hours. At each timepoint, the scaffold was imaged with a stereoscope
(Olympus SZ61) and the diameter was measured using ImageJ (National Institutes of Health, n=3). Fiber morphology was visualized using SEM (2kV,Hitachi S-4700, n=2).
18 2.2.3 IGF-1 Release Characterization
Scaffolds (thickness = 90-120µm) were cut into disks with a ten millimeter diameter using a biopsy punch (Sklar), sterilized via exposure to ultraviolet light for fifteen minutes on each side, and immersed in one milliliter of media (DMEM with 1% insulin, transferrin, selenous acid (ITS), 50 μg/mL proline, 0.1 μM dexamethasone, 0.9 mM sodium pyruvate, 50 μg/mL ascorbic acid) for three weeks at 37ºC and 5% CO2.
Media was collected and replaced every three days.
Insulin-like growth factor-1 release (n=4/group) was measured using an enzyme-linked immunosorbent assay (ELISA, R&D Systems) according to the manufacturer’s protocol. Briefly, samples were added directly to assay diluent in a prepared plate and incubated for two hours at 4°C prior to solution removal. Each well was washed four times before incubation for one hour with IGF-1 conjugate at 4°C. The conjugate was removed, the plate was washed four times, and the substrate solution was added to each well and allowed to react in the dark. The stop solution was added after 30 minutes and the absorbance was measured using a microplate reader (Tecan) at 450 nm and 570 nm, and the difference was used to calculate IGF-1 concentration.
2.2.4 Cartilage Harvest and Culture
Fresh immature bovine knee joints (Green Village Packing Co.) were soaked in soapy water followed by 70% ethanol for 20 minutes. The joint was opened in a sterile environment, and full thickness cartilage explants were extracted from the femoral groove and condyles using a sterile 6 mm biopsy punch (Sklar).
The top and bottom third of the cartilage tissue was removed using a scalpel blade, and the remaining middle zone cartilage was placed atop a microfiber scaffold for culture. To prevent motion of the cartilage, a Teflon® ring was placed on top of the fiber scaffold and around the cartilage ring (Fig. 2.4). Cartilage explants were cultured in 3 mL of media (DMEM with 1% insulin, transferrin, selenous acid (ITS),
50 μg/mL proline, 0.1 μM dexamethasone, 0.9 mM sodium pyruvate, 1% penicillin-streptomycin, 0.1% gentamicin sulfate, 0.1% antifungal (250 µg/mL amphotericin B), and 50 μg/mL ascorbic) acid in a twelve- well plate for two weeks at 37ºC and 5% CO2.
19 2.2.5 Cell Imaging and Quantification
Cell migration onto scaffolds was visualized using Calcein AM staining (n=6/group, Invitrogen) following the manufacturer’s suggested protocol. After washing in PBS, samples were imaged under confocal microscopy (Olympus Fluoview IX70) at excitation and emission wavelengths of 488 nm and 515 nm, respectively. Images were stitched together in PowerPoint (Microsoft), and the area of fluorescence was quantified for each scaffold using ImageJ (National Institutes of Health).
2.2.6 Statistical Analysis
Results are presented in the form of mean ± standard deviation, with n equal to the number of samples per group. One-way ANOVA was used to determine the effect of culture time on scaffold contraction, the effect of IGF-1 dose on fiber diameter, and the effect of IGF-1 dose on scaffold cell number. The Tukey-Kramer post-hoc test was used for all pair-wise comparisons, and significance was attained at p<0.05. Statistical analyses were performed with JMP IN (4.0.4, SAS Institute, Inc.).
2.3 Results
2.3.1 Polymer Blend Optimization
The contraction properties of the scaffold were characterized by measuring scaffold dimensions with
ImageJ after in vitro culture in media (Fig. 2.1). Before immersion, all scaffolds had a similar fiber diameter; however, after eight hours of in vitro culture, the scaffolds composed of PLGA:PCL at a ratio of
7:1, 9:1 and 100% PLGA were significantly smaller than the 5:1, 3:1, and 1:1 compositions. The 5:1, 3:1, and 1:1 compositions did not contract significantly during 72 hours of culture. Fiber morphology was visualized using SEM, and fiber curling was observed in contracted fiber groups after 72 hours (Fig. 2.1).
2.3.2 IGF-1 Scaffold Characterization
Scaffolds containing IGF-1 were visualized using SEM after fabrication, and ImageJ was used to analyze fiber diameter (Fig. 2.2). Fiber morphology was similar between the three meshes with varied IGF doses, with smooth, randomly aligned fibers observed for each group. All three scaffold groups exhibited fiber diameters of approximately one micron. IGF-1 release from the fibers was measured using an ELISA
(Fig. 2.3). Both scaffold groups had similar release profiles, with a burst release followed by slower,
20 sustained release thereafter. The burst release, which occurred within the first 24 hours, accounted for
78.5 ± 0.026% and 83.6 ± 0.002% of the IGF-1 which was released from the low and high dose scaffolds, respectively, over a three-week period. Twice as much IGF-1 was released from the high dose fibers compared to the low dose fibers after 22 days. Based on the amount released after 22 days, incorporation efficiency of active IGF-1 into the fibers is at least 8.6% ± 1.6% and 7.6% ± 1.4% for the low and high doses, respectively.
2.3.3 Cell Migration
Cell migration onto the scaffolds from cartilage explants after two weeks in culture was visualized using fluorescence microscopy (Fig. 2.5). Cells were observed on every scaffold in more than one location. Cell clusters were observed on scaffolds for all groups, most prominently in the control and high dose groups. For the control and low dose groups, the cells on the scaffold were randomly located in areas under the cartilage; however, the migrated cell population on the high dose scaffold mirrored the shape and size of the cartilage explant which had been cultured atop the scaffold, with dense areas of cells observed on every sample in this group. In order to compare the cell migration between groups, fluorescence was quantified using ImageJ for each scaffold, and the average area of fluorescence for each group was calculated (Fig. 2.5). No difference was found between the control and low dose group, but significantly higher fluorescence was measured for the high dose group with respect to both other groups.
2.4 Discussion
The goal of this thesis aim is to design a scaffold that can be placed between the host and graft cartilage to promote integration. Since cartilage-cartilage healing is hindered by low cellularity, the homing of cells to the wound edge through a scaffold that surrounds the cartilage graft may facilitate local, cell- mediated tissue generation that connects the graft to the surrounding host cartilage. This study specifically focuses on the fabrication and optimization of microfibers that release insulin-like growth factor-1, which is a known homing factor for chondrocytes and stem cells. In this study, microfibers with varying concentrations of insulin-like growth factor-1 are fabricated via electrospinning, and cell migration from cartilage explants cultured atop the scaffolds is evaluated using confocal image analysis. The
21 migration results demonstrate that, while cells migrate onto all scaffolds, insulin-like growth factor-1 incorporated at a dose of 100 ng/mg results in more cells on the scaffold after two weeks of culture than a lower dose (50 ng/mg) or scaffold without growth factor.
Microfibers with incorporated insulin-like growth factor-1 were reproducibly fabricated using the electrospinning process. The polymer base of the scaffold was first optimized to eliminate contraction behavior in physiologic conditions. Polycaprolactone, incorporated at 17% w/w (5:1 PLGA:PCL) confers structural stability. This formulation resulted in uniform fiber morphology that was stable over time in culture medium at physiologic conditions. Furthermore, the addition of polycaprolactone did not alter the fiber morphology or significantly change the fiber diameter compared to the poly(lactide-co-glycolide)- based scaffolds that were previously used for chondrocyte culture [150]. Addition of the growth factor also did not change the appearance or alignment of the fibers.
The scaffolds with high and low doses of incorporated insulin-like growth factor-1 released growth factor into the media when incubated in physiologic conditions. Both scaffolds exhibited a burst release in the first 24 hours followed by sustained release for at least three weeks. The burst release profile is consistent with other published reports of biological factor release from electrospun fibrous scaffolds
[22;151;152]. Consistent with the theoretical incorporation, the high dose scaffolds released twice as much insulin-like growth factor-1 compared to the low dose scaffolds over time, demonstrating that the incorporation method is scalable within this range of concentrations.
The growth factor-free and insulin-like growth factor-1 containing scaffolds supported cell migration and attachment when cultured under cartilage explants. While migrated cells were observed on all scaffold types, the high dose scaffolds had significantly staining for cells than the control and low dose groups after two weeks of culture. Furthermore, the area of the scaffold covered in cells in the high dose group resembled the shape of the cartilage explant (6 mm circle), which was not true of the other two scaffold groups. These findings may indicate that the high dose scaffolds induced more uniform migration from the surface of the cartilage explant than the other groups. Conversely, it is possible that the cells migrated from a focal area and then moved across the scaffold.
It is important to note that the insulin-like growth factor-1 delivered via the scaffold in this study is at a significantly lower dose than many published studies that use media supplementation. Based on the
22 measured growth factor that was released into a culture media volume of one milliliter (Fig. 2.3), the high dose scaffold resulted in an insulin-like growth factor-1 concentration of less than 2 ng/mL after the first media change. While there are inconsistencies between studies in the literature, the lowest dose at which insulin-like growth factor-1 has been reported to enhance migration is 10 ng/mL in a Boyden chamber assay performed with immature bovine chondrocytes [18]. The response measured at a lower dose in this study may suggest that the local availability of the insulin-like growth factor-1, released directly at the cut edge of the cartilage, increases efficacy of the growth factor for stimulating migration. However, in order to directly compare these doses, a media supplementation study would need to be conducted in parallel with a scaffold release study.
A limitation of this study is the inability to distinguish between cell proliferation and cell migration.
While migration studies can be conducted at shorter timescales (<24 hours) to reduce the confounding effect of cell proliferation, this study required a longer culture period because of the time required for the cells to move out of the tissue and onto the scaffolds. In a pilot experiment, earlier timepoints were investigated (4 and 7 days, data not shown); however, there were too few cells on the scaffolds to accurately quantify. In the next chapter, cell proliferation on the insulin-like growth factor-1 scaffolds will be tracked independently of a migration source, and these studies will help to elucidate the potential role of proliferation in the increased cell number reported here. Ultimately, the goal of this aim is to maximize cell number on the scaffold to optimize it for acellular in vivo implementation. Therefore, the scaffold with the most cells—the high dose insulin-like growth factor-1 scaffold—is ideal for the current application.
2.5 Conclusions
In summary, insulin-like growth factor-1 incorporation in polymer fiber scaffolds at a dose of 100 ng/mg increases cell number on scaffolds that are cultured adjacent to viable cartilage tissue over a two week period. Based on these findings, it is apparent that insulin-like growth factor-1 delivery from an electrospun scaffold can be used to home and support cells from cartilage tissue.
23 0.7 1:1 3:1 5:1 7:1 9:1 100% PLGA 5mm 1:1 3:1 5:1 7:1 9:1 100% PLGA 0.6 0 hour 0.5
2 0.4 * hours * * 0.3 * * * 8 * *
hours 0.2 Diameter(cm)
24 0.1 hours 0 72 0 20 40 60 80 hours Time (hours)
1:1 3:1 5:1 7:1 9:1 100% PLGA
0 hour
72 hours
5 µm
Figure 2.1 Polymer blend optimization. Scaffolds with PLGA:PCL ratio of 5:1 and lower are stable in physiologic conditions, with no significant change in dimensions after 72 hours (n=3, *p<0.05 for difference from original dimension). Scanning electron micrographs show changes in fiber morphology after soaking for 7:1, 9:1, and 100% PLGA compositions.
24 control 50 ng/mg 100 ng/mg
IGF-1 +
8-10 kV 0.8mL/hr PLGA:PCL 10 µm solution d (µm): 1.2 0.2 1.2 0.1 1.4 0.1
Figure 2.2 Scaffold fabrication and groups. Scaffolds were generated by electrospinning PLGA:PCL polymer with varying amounts of insulin-like growth factor-1 (IGF-1) incorporated. Scanning electron microscopy shows similar fiber morphology, alignment, and diameter (d, n=3) between groups.
25 30 24 Low Dose High Dose Low Dose High Dose 18 * 20
12 (ng) 1 -
IGF 10 6
Cummulative Released IGF (ng) 0 0 1 4 7 10 13 16 19 22 Total IGF-1 released after 22 days Days Figure 2.3 Insulin like growth factor-1 (IGF-1) release from microfiber scaffolds. IGF-1 was released from both low and high dose scaffolds over 22 days. High dose scaffolds released twice as much IGF-1 compared to low dose scaffolds after 22 days (n=4, *p<0.05 for difference between groups).
26 Custom Clamp Scaffold Migration Assay
microfiber scaffold cartilage Teflon® clamp o-ring
2 week culture
top view side view cross section
Figure 2.4 Experimental setup. Custom-designed Teflon® rings secured the microfiber scaffolds and ensured that cartilage explants remained on top of the scaffolds throughout the culture period.
27 microfiber scaffold Sample 1 Sample 2 Sample 3 Sample 4 Sample 5 cartilage explant
control
50 ng/mg
100 2.5 mm ng/mg
2 Control Figure 2.5 Migration from cartilage ) 2 Low Dose * explants onto scaffolds. Calcein AM High Dose stained cells that were attached to the scaffolds after two weeks of culture. After removing cartilage explants, the scaffolds 1 were imaged from the top view (top). Fluorescence was quantified using ImageJ (bottom left) and more fluorescence was measured for the high dose scaffolds (*p<0.05 for difference between groups). Areaof Fluorescence (mm 0 Week 2
28 CHAPTER 3: CHONDROCYTE RESPONSE ON IGF-1 MICROFIBER SCAFFOLDS
29 3.1 Introduction
In chapter 2, the dose of incorporated insulin-like growth factor-1 was optimized to promote cell migration from cartilage tissue into acellular, fiber-based scaffolds. It was determined that incorporating insulin-like growth factor-1 incorporation at a dose of 100 ng/mg increased cell number on scaffolds. In addition to promoting cell migration from cartilage tissue, the scaffold must also support cell viability and matrix production to bridge the gap between the two cartilage surfaces over time. This study will determine the response of chondrocytes on the polymer-fiber scaffolds because this cell population will likely be responsible for new cartilage formation in the cup walls after in vivo implantation.
3.1.1 Background and Significance
Healthy cartilage consists of dense extracellular matrix, rich in proteoglycans and collagen II [25;26], that is actively maintained by resident chondrocytes. Together, the charged proteoglycans and the collagen network support compressive loads endured by the joint and provide shear strength [24;31-35].
These significant structure-function relationships underscore the importance of rebuilding the cartilage matrix to restore functional properties; however, once damaged, cartilage possesses a limited intrinsic capacity for self-repair. Although surgical treatments are available, the resulting repair tissue is often fibrocartilaginous and does not persist over time. Moreover, for cases in which cartilage repair is achieved macroscopically, a gap in the matrix is observed microscopically at the interface between the cartilage graft and surrounding host cartilage [3]. This void in matrix may lead to micromotion between the graft and host tissue, contributing to graft failure and limiting long-term clinical outcomes [99].
To address this clinical limitation, the ideal integration scaffold will support matrix production by native chondrocytes to bridge the gap between healthy and healing cartilage. While the response of chondrocytes on electrospun scaffold systems has been investigated by many groups [153-157], the response of chondrocytes on fibers that release growth factors is relatively under characterized.
3.1.2 Objectives
The goal of this study is to investigate chondrocyte response on the optimized insulin-like growth factor-1 microfibers that will form the walls of the integration cup to determine if the cup walls support
30 chondrocyte attachment, viability and matrix production over time. It is anticipated that the scaffold will support cell attachment as well as matrix elaboration.
3.2 Materials and Methods
3.2.1 Scaffold Fabrication
Unaligned microfiber scaffolds (thickness = 90-120µm, Ø=10mm) were fabricated using the electrospinning process [20;149]. For the fabrication of PLGA:PCL blended microfibers, PLGA and PCL
(Sigma-Aldrich, Mw ≈ 70,000-90,000) were dissolved in a 5:1 ratio in a mixture of 60/40 dichloromethane
(DCM, Sigma-Aldrich) and dimethylformamide (DMF). For fibers containing IGF-1 (Invitrogen), finely ground bovine serum albumin (BSA , 5% w/w, Sigma Aldrich) was added directly the polymer melt and vortexed continuously. After one hour, IGF-1, suspended in distilled water at a concentration of 5 mg/mL, was added to the polymer melt, and the solution was vortexed for an additional hour. Each polymer solution was loaded into a 5 mL syringe with a stainless steel blunt tip needle (26.5 gauge for PLGA:PCL blends and18 gauge for polymer-protein blends) that was 13 cm from the collecting target and electrospun at 8-10 kV using a custom electrospinning device. The polymer solution was deposited (1 mL/hour for polymer solutions, 0.8 mL/hour for polymer-protein solutions) onto a stationary collecting target using a syringe pump (Harvard Apparatus).
3.2.2 Cells and Cell Culture
Primary articular chondrocytes were isolated from neonatal calf knees obtained from a local abattoir
(Green Village Packing Co.) according to published protocols [79]. Briefly, cartilage tissue was extracted from the femoral groove and condyles, minced, and incubated for 16 hours with 0.1% w/v collagenase
(Worthington) in Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 10% fetal bovine serum (FBS, Atlanta Biologicals), 2% antibiotics (10,000 U/mL penicillin, 10 mg/mL streptomycin), and
0.1% antifungal (250 µg/mL amphotericin B). The cell suspension was filtered to separate the cells from extracellular matrix debris before plating (30 μm, Spectrum). The isolated chondrocytes were maintained in high-density culture in fully-supplemented DMEM with 10% FBS, 1% non-essential amino acids, 1% antibiotics, and 0.1% antifungal for three days before seeding. All media supplements are purchased from
Cellgro-Mediatech unless otherwise specified.
31 To seed the scaffolds, 10 µL of concentrated cell suspension was pipetted onto each scaffold to
2 achieve a seeding density of 100,000 cells/cm and incubated for 25 minutes at 37°C and 5% CO2 before submersion in media. Cell-laden scaffolds were cultured at 37°C and 5% CO2 in 1.5 mL of media which was refreshed three times weekly. Cell culture media consisted of DMEM supplemented with 1% ITS+
Premix (BD Biosciences), 1% penicillin-streptomycin, 0.1% gentamicin sulfate, 0.1% antifungal
(250 µg/mL amphotericin B), and 40 µg/mL L-proline (Sigma).
3.2.3 Cell Imaging and Number
Prior to seeding the scaffolds, the cell membranes were stained using the Vybrant™ Cell-Labeling
Solutions kit (Molecular Probes) following the manufacturer’s protocol. The cells were trypsinized, rinsed with F/S medium, and counted. Cells were then spun down and resuspended at a density of 1x106 cells/mL. Dye (5 µL, Vybrant DiO solution) was added, and mixtures were incubated for 20 minutes in the dark under sterile conditions. Cells were collected via centrifugation, the dye was aspirated, and cells were resuspended in fresh F/S medium. This step was repeated three times for a total of four washes.
Cells were then recounted and seeded onto scaffolds. For imaging, medium was aspirated from wells and rinsed with sterile phosphate buffered saline (PBS). The samples were imaged under confocal microscopy (Olympus Fluoview, n=2) at 488 nm.
Cell proliferation (n=5) was determined using the Quanti-iT™ PicoGreen® dsDNA assay kit (Molecular
Probes, Eugene, OR) following sample digestion. The sample was rinsed with PBS and exposed to a freeze-thaw cycle in 500 μL of 0.1% Triton-X solution (Sigma) in order to lyse the cells. After desiccation for 12 hours in a CentriVap Concentrator (Labconco Co.), the samples were digested for 18 hours at 65°C with papain (8.3 activity units/mL) in 0.1 M sodium acetate (Sigma), 10 mM cysteine-HCl (Sigma), and 50 mM ethylenediaminetetraacetate (Sigma). For DNA content, a 25 μL aliquot of the sample was mixed with 175 μL of the PicoGreen® working solution, and fluorescence was measured with a microplate reader
(Tecan, Research Triangle Park, NC) at excitation and emission wavelengths of 485 and 535 nm, respectively. The conversion factor of 7.7 pg DNA/cell was used to determine cell number [158;159].
3.2.4 Matrix Deposition
Total collagen content (n=5) was quantified using a modified hydroxyproline assay [160] with bovine
32 collagen I solution (Biocolor, Carrickfergus, UK) as the standard. A 40 μL aliquot of sample digest was mixed with 10 μL 10 N sodium hydroxide and heated to 250°C for 25 minutes in order to hydrolyze the collagen. The hydrolyzate was then oxidized at room temperature for 25 minutes with 450 μL of buffered chloramine-T reagent prior to the addition of Ehrlich’s reagent (15% p-dimethylaminobenzaldehyde in 2:1 isopropanol/percholoric acid). Absorbance was measured at 555 nm with a microplate reader (Tecan).
Additionally, collagen distribution (n=2) was evaluated via histology. The samples were first fixed in neutral buffered formalin with 1% cetylpyridinium chloride (Sigma) for one day and stored in 0.01 M cacodylic acid at 4°C. Prior to processing, samples were soaked overnight in 5% polyvinyl alcohol (PVA,
Sigma-Aldrich), embedded in a frozen block of PVA, and sectioned at -21 °C using a cryostat (Model
OFT, Bright Instrument Company Microtome). Sections were soaked in distilled water for one hour to remove residual PVA, stained with picrosirius red staining for one hour, and exposed to 0.1 N hydrochloric acid for two minutes (n=2). For collagen I and II immunostaining, sections were incubated with primary antibody overnight. Cell nuclei were stained with 4',6-diamidino-2-phenylindole (Sigma). A FITC- conjugated secondary antibody (1:200 dilution, Abcam) was used and samples were imaged under confocal microscopy (Olympus Fluoview IX70) at excitation and emission wavelengths of 488 nm and
568 nm, respectively. Cover-slipped sections were imaged using a brightfield microscope (Zeiss, Axiovert
25).
Sample glycosaminogylcan content (GAG, n=5) was determined with a modified 1,9- dimethylmethylene blue (DMB) binding assay [161-163], with chondrotin-6-sulfate (Sigma) as the standard. The absorbance difference between 540 nm and 595 nm was used to improve the sensitivity in signal detection. In addition, histology was used to visualize GAG distribution. Sections were exposed to
3% acetic acid for three minutes, stained with alcian blue for 45 minutes, and rinsed twice with acid- alcohol (pH=1) for one minute (n=2) [159]. Cover-slipped sections were imaged using a brightfield microscope (Zeiss, Axiovert 25).
3.2.5 Mineralization Potential
Alkaline phosphatase (ALP) activity (n=5) was measured using a colorimetric assay based on the hydrolysis of p-nitrophenyl phosphate (pNP-PO4) to p-nitrophenol (pNP) [164]. The samples were lysed in
0.1% Triton™ X solution, exposed to a freeze-thaw cycle, and crushed with a mortar. A 25 μL aliquot was
33 added to pNP-PO4 solution (Sigma) and incubated for 10 minutes at 37°C. Absorbance was measured at
405 nm using a microplate reader (Tecan).
3.2.6 Statistical Analysis
Results are presented as mean ± standard deviation, with n equal to the number of samples per group. One-way ANOVA was used to determine the effects of IGF-1 dose on cell migration. The Tukey-
Kramer post-hoc test was used for all pair-wise comparisons, and significance was attained at p<0.05.
Statistical analyses were performed with JMP IN (4.0.4, SAS Institute, Inc.).
3.3 Results
3.3.1 Cell Distribution, Viability, Number, and Alkaline Phosphatase Activity
Full thickness chondrocytes were cultured on polymer scaffolds with and without IGF-1 for three weeks (Fig. 3.1). Cell distribution on the scaffolds was assessed via live-cell tracking (Fig. 3.2).
Chondrocytes were evenly distributed on the scaffolds at each timepoint. Quantitatively, cell number was similar on both scaffold types on day 1, with a significant increase in cell number measured for the control group between day 1 and day 14. Mineralization potential, assessed by measuring alkaline phosphatase activity, was highest for the control group on day 1, with significantly lower ALP activity detected on the
IGF-1 scaffold. ALP activity decreased over time between day 1 and day 14 for both scaffold groups.
3.3.2 Matrix Production
Matrix deposition was quantified at each timepoint in terms of GAG and collagen. (Fig. 3.3).
Significantly more GAG was measured on both scaffold types on day 14 compared to day 1. No differences were measured in terms of collagen deposition at any timepoint. Matrix distribution was visualized histologically, and staining revealed that both GAG and collagen were localized to the seeded surface of the scaffold.
3.4 Discussion
The goal of this chapter is to determine the response of chondrocytes on the fiber-based scaffolds in vitro. Chondrocytes are expected to interact with the scaffolds in vivo because they were shown to migrate into the scaffolds from cartilage tissue in chapter 1. To determine the response of chondrocytes 34 on the scaffolds over time, cells were isolated from full thickness articular cartilage and seeded onto the scaffolds. Cell number, matrix deposition, and mineralization potential were measured over three weeks of culture. The chondrocytes attached, remained viable, and produced matrix on both control and IGF-1 scaffolds. The results of this study demonstrate that the microfiber scaffolds support the attachment, proliferation and biosynthesis of chondrocytes.
In terms of cell number and matrix production, both scaffolds supported similar cellular activity in this study. While insulin-like growth factor-1 is known to enhance cartilage matrix deposition [147;165], these effects are reported for studies in which insulin-like growth factor-1 is introduced via continuous media supplementation at higher levels than the concentrations that are released from these scaffolds
[147;166;167]. Furthermore, for the studies presented here, the fiber-based scaffolds are soaked in serum media overnight prior to seeding to enhance cell attachment by surface protein coating. This pre- soak step occurs during the “burst release” phase of the insulin-like growth factor-1 release from the scaffolds, and therefore decreases the total growth factor to which cells are exposed. In an in vivo environment, insulin-like growth factor-1 is naturally present [167]. Therefore, the insulin-like growth factor-1 released from the scaffold will not act in isolation but will instead locally augment the native concentration. It is possible that, while the insulin-like growth factor-1 did not increase matrix deposition in this study over the control scaffold, it may have an effect when combined with the insulin-like growth factor-1 that is naturally present in an in vivo environment. In addition, Mauck et al. demonstrated synergy between insulin-like growth factor-1 stimulation and dynamic loading [166]. Applied to our system, this may suggest that the insulin-like growth factor-1 released from the fibers may enhance matrix production when combined with the loaded environment that is found in vivo. In order to further understand the potential additive effect of scaffold insulin-like growth factor-1, native insulin-like growth factor-1, and loading, future studies could evaluate the response of cells on the insulin-like growth factor-1 scaffolds in media containing insulin-like growth factor-1 levels similar to that found in vivo with and without loading.
In contrast to the similar trends for matrix deposition on scaffolds with and without insulin-like growth factor-1-free, the alkaline phosphatase activity was significantly modulated by scaffold type. On day 1, alkaline phosphatase activity of the chondrocytes was significantly reduced on the insulin-like growth factor-1 scaffolds. This finding is not surprising as others have reported the suppression of thyroid
35 hormone-induced alkaline phosphatase activity by insulin-like growth factor in rat epiphyseal chondrocytes [168]. While this study does not investigate the underlying mechanisms that drive the suppression of alkaline phosphatase activity, future studies that evaluate gene expression of hypertrophic markers may help to further elucidate the pathway by which insulin-like growth factor-1 affects mineralization behavior. The scaffold-mediated suppression of alkaline phosphatase activity, a marker of mineralization potential, may be advantageous for cartilage-cartilage healing because diseased cartilage tissue is particularly susceptible to pathological mineralization [169], and a scaffold which counteracts this natural tendency may reduce the susceptibility of the healing cartilage tissue to ectopic mineralization.
3.5 Conclusions
This study evaluates the response of chondrocytes on the microfiber scaffold that will comprise the walls of the integration cup system. The scaffolds were initially seeded with cells to mimic the ideal in vivo conditions, in which chondrocytes have migrated from the cartilage into the scaffold. The fiber-based scaffold that releases insulin-like growth factor-1 supported cell attachment and matrix elaboration over time, while reducing alkaline phosphatase activity. These findings suggest that the scaffold, optimized in chapter 1 to promote cell migration, will sustainably support chondrocyte attachment, growth, and biosynthesis over time.
36 PLGA: PCL control solution FTC +/- in vitro culture
IGF-1 100 ng/mg IGF-1
Figure 3.1 Study design. Polymer scaffold with and without insulin-like growth factor-1 (IGF-1) are generated via electrospinning using a 5:1 mixture of poly(lactide-co-glycolide) (PLGA) and polycaprolactone (PCL). Each scaffold type was seeded with full thickness chondrocytes (FTCs) and cultured in vitro for three weeks.
37 100 FTC
FTC+IGF 3 Control IGF - #
50
Cell No. x 10 x No. Cell Day 1 Day
0 D1 D14 D21 0.1 FTC
FTC+IGF Day 14 Day
0.05 *
200 µm
Day 21 Day # 200 µm
ALP Activity (pmoles/cell/min) Activity ALP # 0 D1 D14 D21
Figure 3.2 Cell viability, number, and alkaline phosphatase activity. Live cell imaging shows that chondrocytes are viable throughout the culture period. Cell number was similar between groups for all timepoints. ALP activity was significantly suppressed in the IGF scaffold on day 1, with significant decrease over time between day 1 and day 7 for both groups (n=5, *p<0.05 for difference between groups on a given timepoint, #p<0.05 for difference from corresponding group at previous timepoint).
38 FTC FTC+IGF 40 FTC FTC+IGF 30 GAG
20 # # Control IGF
GAG (µg) GAG 10 Collagen
0 200 μm D1 D14 D21 90 FTC FTC+IGF Collagen I 60
30
Collagen (µg) Collagen Collagen II 200 μm
0 200 μm Day 21 D1 D14 D21
Figure 3.3 Matrix deposition. Chondrocytes produced GAG over time, with significantly more GAG measured on day 14 compared to day 1 for both groups. Collagen content was not different over time or between groups (n=5, *p<0.05 for difference between groups on a given timepoint, #p<0.05 for difference from corresponding group at previous timepoint) GAG and collagen were observed on the seeded edge of the scaffold using alcian blue and picrosirius red staining, respectively (n=2). Scaffolds were negative for collagen I and positive for collagen II (n=2).
39 CHAPTER 4: THE EFFECT OF CERAMIC CRYSTALLINITY ON DEEP ZONE CHONDROCYTE RESPONSE AND CALCIFIED CARTILAGE FORMATION
40 4.1 Introduction
In chapters 2 and 3, the walls of the cup system were optimized to promote cartilage-cartilage integration. A microfiber scaffold, which released insulin-like growth factor-1, enhanced cell migration from cartilage tissue into the scaffold supported chondrocyte viability and matrix deposition. The focus of this chapter shifts to optimize a scaffold to promote cartilage-bone integration in the base of the cup system.
The ideal base will serve to integrate the cartilage to the underlying bone by promoting the formation of calcified cartilage tissue. Previous work demonstrated that hydrogel-ceramic scaffolds enhance the formation of calcified cartilage-like tissue by deep zone chondrocytes [11;15]; however, hydroxyapatite, a crystalline, relatively inert ceramic, was used and the effect of poorly crystalline bioactive ceramics on chondrocyte response have not yet been determined. In this chapter, the effect of ceramic crystallinity on chondrocyte response will be optimized for calcified cartilage formation to promote cartilage-bone integration in the base of the cup.
4.1.1 Background and Significance
Current cartilage repair products, such as chondrocyte implantation and minced cartilage transplantation, as well as many cartilage scaffolds in development, do not ensure calcified cartilage regeneration. As a result, the neo-cartilage-to-bone junction is vulnerable to poor integration. To address this limitation, osteochondral interface tissue engineering efforts began with cell-based approaches [113], followed by recent reports of a hydrogel-hydroxyapatite composite scaffold system for calcified cartilage formation [11;15]. Both alginate and agarose have been investigated as the hydrogel phase; however, the alginate system was limited by inherently low mechanical properties as well as slow gelation time, leading to ceramic particle aggregation [15]. Agarose exhibits a faster gelation time, allowing for homogenous distribution of mineral and cells within the constructs. Furthermore, the culturing of chondrocytes in agarose hydrogels resulted in higher mechanical properties than were measured for the alginate system
[11], further indicating that agarose is a promising material for calcified cartilage engineering.
While various hydrogels have been utilized for cartilage and calcified cartilage tissue engineering, relatively few studies have evaluated the effect of varying calcium phosphate parameters on chondrocyte response. To date, most studies have used crystalline hydroxyapatite, which is a relatively inert calcium phosphate; however, the mineral of the native osteochondral interface is poorly crystalline [59]. While the
41 effect of calcium phosphate crystallinity on chondrocyte response is unclear, reports indicate that ceramic crystallinity modulates cellular response, as it has been shown to control matrix elaboration in bone marrow cells [170;171]. In addition, studies show that bioactive calcium phosphate sources enhance mineralized tissue formation in bone tissue engineering applications [172]. Inspired by these findings, this study will investigate the effect of crystallinity on calcified cartilage formation using two bioactive calcium phosphate mineral sources.
4.1.2 Objectives
The objective of this study is to investigate the effect of calcium phosphate crystallinity on calcified cartilage formation by deep zone chondrocytes. Specifically, poorly crystalline calcium deficient apatite and crystalline beta tricalcium phosphate will be incorporated into agarose scaffolds with deep zone chondrocytes to evaluate formation of calcified cartilage. It is hypothesized that ceramic crystallinity will modulate chondrocyte response, with the less crystalline, more biomimetic calcium deficient apatite resulting in enhanced matrix production.
4.2 Materials and Methods
4.2.1 Ceramic Characterization
Calcium deficient apatite (CDA) was purchased from Sigma-Aldrich and sintered (Thermolyne,
Thermo Scientific) at 900° for three hours to produce β-tricalcium phosphate (TCP). Particles of both ceramics were imaged using scanning electron microscopy (SEM). In addition, calcium and phosphorus content (n=6), crystal structure (n=2), chemistry (n=2), and specific surface area (n=3) were determined.
Particle size and morphology was assessed using SEM (5kV, 1000x, Hitachi S-4700) after sputter-coating with gold-palladium for ten seconds (Cressington 108 Auto). Ceramic calcium and phosphorus content
(n=6) were determined using inductively coupled plasma analysis (ICP, Thermo Jarrell Ash, Trace Scan
Advantage) [173]. Briefly, 10 mg of ceramic was dissolved in several drops of 17% HCl and brought to
100 mL with double distilled water. The resulting solutions were pumped through argon plasma excited by a 2 kW/27.12 MHz radiofrequency generator. The concentration of each element was determined using its characteristic wavelength (Ca, 317.9 Å; P, 213.6 Å) [173]. The crystal structures of the ceramics were evaluated with X-ray diffraction (XRD, X-ray Diffractometer, Inel). The samples were scanned over a
42 range of 0-120°, with a step size of 0.029°. Ceramic chemistry was examined using Fourier transform infrared spectroscopy (FTIR, FTS 3000MX Excalibur Series, Digilab), wherein dehydrated samples were mixed with potassium bromide and scanned in absorbance mode (400 scans, 4 cm−1 resolution). The specific surface area of each ceramic was calculated using the Brunauer-Emmett-Teller (BET) method
(n=3, NOVA-win 2002 BET analyzer, Quantachrome Corporation) after overnight dehydration.
4.2.2 Cells and Cell Culture
Primary DZCs were isolated and pooled from the femoral articular cartilage of five immature calf knees (Green Village Packing Co.) following previously described protocols [79]. The bottom third of the cartilage was separated, and the calcified cartilage was removed by scraping. The cartilage was minced and digested with collagenase type 2 (310 U/mg, Worthington) for 16 hours in Dulbecco’s Modified
Eagle’s Medium (DMEM) supplemented with 10% fetal bovine serum (FBS, Atlanta Biologicals), 2% antibiotics (10,000 U/mL penicillin, 10 mg/mL streptomycin), and 0.2% antifungal (250 µg/mL amphotericin B). The deep zone chondrocyte suspension was filtered before plating (30 µm, Spectrum).
The isolated chondrocytes were maintained in high-density culture in fully-supplemented DMEM with 10%
FBS, 1% non-essential amino acids, 1% antibiotics, and 0.1% antifungal for three days prior to seeding in the hydrogel [15]. All media supplements were purchased from Cellgro-Mediatech, unless otherwise specified.
4.2.3 Scaffold Fabrication and Culture
Chondrocytes were encapsulated at a density of 10 million cells/mL in 2% low gelling agarose
(Agarose Type VII, Sigma), and a biopsy punch (Sklar Instruments) was used to core cylindrical scaffolds
(Ø = 5 mm, height = 2.4 mm) [11]. Acellular and cellular agarose scaffolds with 1.5% w/v ceramic, added at the time of fabrication, and corresponding controls without ceramic were fabricated (Fig. 4.2). All
⁰ samples were cultured under humidified conditions at 37 C and 5% CO2 and maintained in medium composed of DMEM supplemented with 1% ITS+ Premix (BD Biosciences), 1% penicillin-streptomycin,
0.1% gentamicin sulfate, 0.1% amphotericin B, and 40 µg/mL L-proline (Sigma). The media was changed every other day and freshly supplemented with 50 µg/mL ascorbic acid (Sigma). In order to induce
43 hypertrophy, DZCs were stimulated with 25 nM triiodothyronine (T3, Sigma) during the first three days of culture [174].
4.2.4 Cell Number and Distribution
Cell number (n=5) was determined using the Quanti-iT™ PicoGreen® dsDNA assay kit (Molecular
Probes) following sample digestion. The sample was rinsed with PBS and exposed to a freeze-thaw cycle in 500 μL of 0.1% Triton™-X solution (Sigma) in order to lyse the cells. A pestle was used to manually pulverize the samples. After desiccation for 12 hours in a CentriVap Concentrator (Labconco Co.), the samples were digested for 18 hours at 65°C with papain (8.3 activity units/mL) in 0.1 M sodium acetate
(Sigma), 10 mM cysteine HCl (Sigma), and 50 mM ethylenediaminetetraacetate (Sigma). For DNA content, a 25 μL aliquot of the sample was mixed with 175 μL of PicoGreen® working solution, and fluorescence was measured with a microplate reader (Tecan) at excitation and emission wavelengths of
485 and 535 nm, respectively. The conversion factor of 7.7 pg DNA/cell was used to determine cell number [158;159].
Cell distribution (n=2) was evaluated via histology. The samples were first fixed in neutral buffered formalin with 1% cetylpyridinium chloride (Sigma) for 24 hours, followed by dehydration with an ethanol series. The dehydrated samples were embedded in paraffin (Paraplast X-tra Tissue Embedding Medium,
Fisher Scientific), and 7 μm sections were obtained from the center of the scaffold. The paraffin was cleared with xylenes, and the samples were rehydrated and stained with hematoxylin for ten minutes and eosin Y for two minutes. Cover-slipped samples were imaged using a brightfield microscope (Zeiss,
Axiovert 25).
4.2.5 Matrix Deposition
Total collagen content (n=5) was quantified using a modified hydroxyproline assay [160] with bovine collagen I solution (Sigma) as the standard. Briefly, a 40 μL aliquot of sample digest was mixed with 10 μL of 10 N sodium hydroxide and heated to 250°C for 25 minutes in an autoclave to hydrolyze the collagen.
The hydrolyzate was oxidized at room temperature for 25 minutes with 450 μL of buffered chloramine-T reagent prior to incubation with 425 μL of Ehrlich’s reagent (15% p-dimethylaminobenzaldehyde in 2:1 isopropanol/percholoric acid) at 65°C. Absorbance was measured at 555 nm with a microplate reader
44 (Tecan). All matrix values were normalized by sample wet weight to account for differences in sample size.
Collagen distribution (n=2) was evaluated via histology. The samples were first fixed in neutral buffered formalin with 1% cetylpyridinium chloride (Sigma) for 24 hours, followed by dehydration with an ethanol series. The dehydrated samples were embedded in paraffin (Paraplast X-tra Tissue Embedding
Medium, Fisher Scientific), and 7 μm sections were obtained from the center of the scaffold. The paraffin was cleared with xylenes, and the samples were rehydrated, stained with picrosirius red for one hour, and exposed to 0.1 N hydrochloric acid for two minutes (n=2). Cover-slipped samples were imaged using a brightfield microscope (Zeiss, Axiovert 25). In addition, collagen I, II, and X were assessed using immunohistochemistry (n=2) [11]. Specifically, monoclonal antibodies for collagen I (1:00 dilution) and collagen II (1:100 dilution) were purchased from Abcam, and antibodies for collagen X were purchased from the Developmental Studies Hybridoma Bank (University of Iowa). After deparaffinization, sections were treated with 1% hyaluronidase for 30 minutes at 37°C, 1% acetic acid for four hours, and incubated with primary antibody overnight. Cell nuclei were stained with 4',6-diamidino-2-phenylindole (Sigma). A
FITC-conjugated secondary antibody (1:200 dilution, LSAB2 Abcam) was used, and samples were imaged under confocal microscopy (Olympus Fluoview IX70) at excitation and emission wavelengths of
488 nm and 568 nm, respectively.
Sample glycosaminoglycan content (GAG, n=5) was determined with a modified 1,9- dimethylmethylene blue (DMB) binding assay [161-163] with chondrotin-6-sulfate (Sigma) as the standard. The absorbance difference between 540 nm and 595 nm was used to improve the sensitivity in signal detection. In addition, histology was used to visualize GAG distribution. Deparaffinized sections were exposed to 3% acetic acid for three minutes, stained with alcian blue for 45 minutes, and rinsed twice with acid-alcohol (pH = 1) for one minute (n=2) [159]. Cover-slipped samples were imaged using a brightfield microscope (Zeiss, Axiovert 25).
4.2.6 Mineralization
Alkaline phosphatase (ALP) activity (n=5) was measured using a colorimetric assay based on the hydrolysis of p-nitrophenyl phosphate (pNP-PO4) to p-nitrophenol (pNP) [164]. The samples were lysed in 0.1% Triton™ X solution, exposed to a freeze-thaw cycle, and crushed with a pestle. A 25 μL aliquot
45 was added to pNP-PO4 solution (Sigma) and incubated for 10 minutes at 37°C. Absorbance was measured at 405 nm using a microplate reader (Tecan).
Calcium and phosphate distribution (n=2) were evaluated as an indicator of overall mineral distribution [175]. For calcium staining, deparafinized histology sections were stained with alizarin red for one hour, rinsed with tap water, followed by 0.01 N hydrochloric acid in 70% ethanol for two minutes. For phosphate staining, deparaffinized samples were stained with 5% silver nitrate and exposed to UV light
(365 nm) for 25 minutes before a tap water-rinse. Cover-slipped samples were imaged using a brightfield microscope (Zeiss, Axiovert 25).
4.2.7 Chondrocyte Hypertrophy
The expression (n=5) of collagen X, Indian Hedgehog (Ihh), matrix metalloproteinase-13 (MMP-13), parathyroid hormone-related protein (PTHrP), and alkaline phosphatase (ALP) were measured at day 1 and 14 using reverse transcription followed by real-time polymerase chain reaction (RT-PCR). Primer sequences are listed in Table 4.1. Total RNA was isolated using the TRIzol (Invitrogen) extraction method. The isolated RNA was reverse-transcribed into cDNA using the SuperScript III First-Strand
Synthesis System (Invitrogen). PCR reactions (25 ul) were carried out using SYBR GreenER qPCR
SuperMix (Invitrogen) on an iCycler instrument (Bio-Rad). Relative expression data were quantified using
2-(Ct sample−Ct GAPDH), where Ct is the cycle threshold normalized to the housekeeping gene, glyceraldehyde
3-phosphate dehydrogenase (GAPDH).
4.2.8 Media Ion Analysis
Media calcium concentration (n=6) was quantified using the Arsenazo III dye (Pointe Scientific). The media was diluted with water in a 1:10 ratio and allowed to react with the dye for five minutes.
Absorbance was measured at 620 nm using a microplate reader (Tecan). Media phosphate concentration
(n=6) was quantified using the BioVision Phosphate Assay Kit. The media was diluted with water in a 1:10 ratio and allowed to react with 30 µL of dye for 30 minutes. Absorbance was measured at 650 nm using a microplate reader (Tecan).
46 4.2.9 Statistical Analysis
Results are presented in the form of mean ± standard deviation, with n equal to the number of samples per group. A two-way analysis of variance (ANOVA) was performed to determine the effects of ceramic type and culturing time on cell response (proliferation, matrix deposition, ALP activity, solution ion concentration) as well as ceramic parameters (calcium and phosphorus weight content, Ca/P ratio and surface area). A one-way ANOVA was used to determine the effects of scaffold group on gene expression at each timepoint. The Tukey-Kramer post-hoc test was used for all pair-wise comparisons, and significance was attained at p<0.05. Statistical analyses were performed with JMP IN (4.0.4, SAS
Institute, Inc.).
4.3 Results
4.3.1 Ceramic Characterization
Ceramic particle shape, calcium and phosphate content, crystallite structure, chemistry, and specific surface area were characterized prior to scaffold fabrication and are summarized in Figure 4.1. Electron microscopy revealed irregular particle morphology for CDA and rhombic-shaped particles for TCP. Crystal planes characteristic of hydroxyapatite (002, 211, 300, and 202) (JCPDS 9-432) were identified in the
CDA XRD spectra (Fig. 4.1). Characteristic TCP crystal planes (214, 300, 0210, 128, and 220) were identified in the TCP XRD spectra (JCPDS 9-0169). The broader peaks present in the CDA spectra indicate smaller crystallite size compared to the sharper peaks of the TCP spectra. Analysis by FTIR confirmed phosphate presence via bending peaks for both ceramics, although a carbonate peak
[173;176;177] was observed only in the CDA spectra. In terms of molar calcium and phosphate content,
TCP had higher calcium and phosphorus content than the CDA (p<0.05); however, the two ceramics had similar calcium/phosphate molar ratios. The specific surface area of CDA was significantly higher than the specific surface area of the TCP.
4.3.2 Cell Distribution and Number
Deep zone chondrocytes were cultured in agarose scaffolds with CDA, TCP, or without ceramic , with and without T3 stimulation for two weeks (Fig 4.2). Cell distribution was assessed via hematoxylin and eosin staining for each group over time (Fig. 4.3). Chondrocytes were uniformly distributed throughout
47 every scaffold at all timepoints, with higher cell density observed by day 14 compared to day 1 for all groups. Quantitatively, cell number was similar in all groups on day 1, with an increase in cell number on day 7 and day 14. Significantly more cells were measured for the CDA group than the ceramic-free and
TCP groups on day 7 and day 14. Trends between groups and over time for the unstimulated and T3- stimulated cell populations were similar.
4.3.3 Matrix Deposition
Matrix deposition was measured in terms of collagen and GAG for each group over time. Collagen deposition increased over time for the unstimulated control group and all stimulated groups between day
1 and day 7 (Fig. 4.4). Between day 7 and day 14, collagen content increased for the unstimulated ceramic-containing groups and for the stimulated CDA group. For both unstimulated and stimulated groups, the CDA-containing scaffold resulted in the most collagen by day 14. This finding was qualitatively confirmed with picrosirius red staining. Immunohistochemistry was used to determine the collagen types present in the unstimulated groups on day 14 (Fig. 4.5). Positive collagen II staining was observed for all groups, with the most positive staining observed for the CDA group. Collagen I and X staining was observed only for the ceramic-free and the TCP groups on day 14. The most collagen I and
X was visualized in the TCP-containing scaffolds that were stimulated with T3.
Glycosaminoglycan content increased over time for all groups (Fig. 4.6). For the unstimulated constructs, the CDA-containing scaffold resulted in the highest GAG content by day 7 and day 14. In the presence of T3 stimulation, no difference in GAG was measured between groups on day 7; however, the
CDA group had significantly higher GAG content on day 14 than the control or TCP groups. Alcian blue staining showed pericellular GAG depositions for all groups on day 7, with positive staining observed throughout the scaffold for the unstimulated and stimulated CDA groups on day 14.
4.3.4 Mineralization Potential and Hypertrophy
Mineralization potential was assessed by measuring the alkaline phosphatase (ALP) activity for each group over time, and mineral distribution was evaluated via histology (Fig. 4.7). For the unstimulated chondrocytes, ALP activity was highest in the CDA group on day 1, with significantly lower ALP activity for all groups by day 7, which persisted to day 14. In the presence of T3, the ceramic-free and TCP groups
48 exhibited enhanced ALP activity on day 7 and day 14; however, low ALP activity was measured at every timepoint for the CDA group. Histological staining with alizarin red and von Kossa revealed uniformly distributed mineral for both ceramic-containing groups at all timepoints, with no phosphate or calcium staining observed in the ceramic-free control group at any timepoint.
The expression of hypertrophic markers were measured on day 1 and day 14 for each group (Fig.
4.8). On day 1, collagen X and Ihh expression were downregulated for ceramic-containing groups compared to the ceramic-free control. The CDA group exhibited upregulated PTHrP expression compared to the ceramic-free group and downregulated ALP expression compared to both the ceramic- free and TCP groups. On day 14, for the unstimulated groups, collagen X was downregulated in the CDA group compared to both the ceramic-free and TCP groups, and the MMP13 was upregulated for the TCP group with respect to the ceramic-free control group. In the presence of T3, Ihh was upregulated in the
TCP group, and MMP13 was downregulated for both ceramic-containing groups on day 14.
4.3.5 Media Ion Concentrations
Calcium and phosphate concentration in the media were measured for each group at each timepoint
(Fig. 4.9). For both stimulated and unstimulated groups, media calcium was significantly lower in the media of the CDA groups on day 1 than all other groups. Media phosphate concentration was significantly higher for the TCP group than all other groups on day 1. After day 1, there were no differences detected in either calcium or phosphate concentration between groups.
4.4 Discussion
The goal of this chapter is to determine the effect of ceramic crystallinity on deep zone chondrocyte response and to identify a bioactive calcium phosphate source that promotes calcified cartilage formation.
In this study, two composite scaffolds were generated by adding either poorly crystalline calcium deficient apatite or crystalline tricalcium phosphate to agarose. The response of deep zone chondrocytes cultured within the scaffold were evaluated in terms of cell viability and number, matrix deposition, mineralization potential, and gene expression over two weeks of in vitro culture. The results collectively demonstrate that, while chondrocytes survive and produce matrix in all constructs, the presence of poorly crystalline calcium deficient apatite enhances cell growth and matrix deposition both with and without the addition of
49 thyroid hormone stimulation, while the presence of crystalline tricalcium phosphate does not. Based on these findings, it is clear that ceramic crystallinity is an important design parameter for tissue engineering, and calcium deficient apatite is a promising calcium phosphate for calcified cartilage tissue engineering.
To limit confounding variables and differences derived from varying the ceramic manufacturer, the tricalcium phosphate was obtained by directly sintering the calcium deficient apatite. This experimental design ensured that the presence of trace minerals, such as zinc, which may affect cell response[178-
180], were not varied between the two calcium phosphates. While sintering the ceramic did produce crystalline tricalcium phosphate with a similar calcium phosphate ratio to the calcium deficient apatite, it also removed carbonate and decreased the specific surface area. Although this study does not specifically examine the effects of carbonate and particle shape, cellular response may be collectively influenced by a combination of crystallinity, carbonation, and ceramic surface area.
The agarose-calcium deficient apatite and agarose-tricalcium phosphate composite scaffolds both supported chondrocyte viability, growth, and matrix elaboration over time. While no differences were measured in terms of cell number, glycosaminoglycan content, or collagen content between the ceramic- free control and the tricalcium phosphate group, presence of calcium deficient apatite in the scaffold significantly enhanced cell number and matrix deposition. This finding expands on previously reported results demonstrating that calcium deficient carbonated apatite coatings enhanced matrix production by bovine chondrocytes seeded on cellulose scaffolds [181]. The agarose-calcium deficient apatite composite scaffold was transformed over the culture period by the cells into a mineralized matrix that was rich in collagen II and glycosaminoglycans.
Importantly, the response of cells to calcium deficient apatite was not altered by thyroid hormone stimulation. Thyroid hormone promotes the hypertrophic phenotype in deep zone chondrocytes [11;174] and will likely be introduced into the scaffold after in vivo implantation in a full thickness defect via the blood [182]. However, it is unknown if all cells in the defect will be exposed to the hormone; therefore, the ideal calcified cartilage scaffold will enhance matrix elaboration in both the presence and absence of thyroid hormone. Khanarian et al. has demonstrated that thyroid hormone stimulation affects deep zone chondrocyte growth, matrix deposition, and mineralization. In the agarose-hydroxyapatite system, enhanced matrix deposition was observed only in the presence of thyroid hormone. Thus, the ability of
50 the agarose-calcium deficient apatite scaffold to stimulate matrix deposition in the absence of thyroid hormone represents an improvement over the response observed for agarose-hydroxyapatite scaffolds
[11].
In addition to matrix elaboration, ceramic crystallinity modulated mineralization potential and hypertrophy of the chondrocytes. In the absence of thyroid hormone, on day 1, the calcium deficient apatite resulted in elevated alkaline phosphatase activity compared to the control and β-tricalcium phosphate groups; however, low alkaline phosphatase activity was measured thereafter. In the presence of hypertrophic stimulation, both the control and TCP groups exhibited elevated alkaline phosphatase activity on day 7 and day 14. Suppression of this activity was measured for the calcium deficient apatite group, with low alkaline phosphatase activity persisting throughout the two weeks of culture. In addition to decreased alkaline phosphatase activity, calcium deficient apatite presence also downregulated hypertrophic markers on day 1 and day 14, suggesting that the pre-incorporation of a biomimetic mineral may reduce the need for cell-mediated mineralization in the long term. Since cartilage tissue is particularly susceptible to mineralization in disease states [169], the ability to regulate hypertrophy and mineralization may offer a viable method to prevent the overgrowth of mineralized tissue in the cartilage defect over time.
While this study demonstrates that calcium deficient apatite and tricalcium phosphate elicit different deep zone chondrocyte responses, the effects of crystallinity cannot be decoupled from the effects of carbonate presence and surface area, which were also varied between the two calcium phosphates. The presence and dose of carbonate in calcium phosphates has been shown to modulate osteoblast response [183]. Specifically, hydroxyapatites with higher carbonate content resulted in lower alkaline phosphatase activity, consistent with the findings for the carbonated calcium deficient apatite in this study.
In addition, while the effect of calcium phosphate surface area on chondrocytes is unknown, differences in mineral surface area has been shown to modulate bone formation [184-186]. It has been postulated that differences in cell response related to surface area changes may be driven, in part, by the differences in proteins absorbed onto the ceramic surface [184] or by differences in calcium phosphate dissolution, as both processes are coupled to surface area [187]. While protein absorption was not measured in this study, differences between the two ceramics in terms of calcium and phosphorus dissolution were
51 detected, with decreased calcium in the media from the CDA group and increased phosphate in the media from the TCP group. Future studies that specifically evaluate varying carbonate content and calcium phosphate surface area may help to elucidate the mechanisms that underlie the differences in matrix deposition and mineralization potential driven by the ceramic type that were observed in this study.
4.5 Conclusions
This study focuses on optimization of the calcium phosphate phase of a hydrogel-ceramic composite scaffold for calcified cartilage regeneration. Two bioactive ceramics, calcium deficient apatite and tricalcium phosphate, were incorporated into deep zone chondrocyte-laden agarose hydrogels that were cultured in the presence or absence of thyroid hormone stimulation. Although the cells remained viable and deposited matrix in all scaffold groups, the most matrix deposition was measured for the scaffolds which contained calcium deficient apatite. Moreover, the promotion of matrix elaboration by calcium deficient apatite was robust and persisted in the presence of thyroid hormone stimulation. These findings suggest that calcium deficient apatite is a promising ceramic for calcified cartilage regeneration.
52 Table 4.1 Primer Sequences for Gene Expression.
Gene Sense Antisense Amplicon Size (bp)
ALP TGCGACTGACCCTTCACTCTC CACCAGCAGGAAGAAGCCTTT 84
Col X TGGATCCAAAGGCGATGTG GCCCAGTAGGTCCATTAAGGC 82
Ihh ATCTCGGTGATGAACCAGTG CCTTCGTAATGCAGCGACT 97
MMP13 ACATCCCAAAACGCCAGACAA GATGCAGCCGCCAGAAGAAT 109
PTHrP ACCTCGGAGGTGTCCCCTAA GCCCTCATCATCAGACCCAA 80
GAPDH GCTGGTGCTGAGTATGTGGT CAGAAGGTGCAGAGATGATGA 213
53 SEM XRD FTIR 15 0.6 v (PO )-3
CDA CDA 3 4 210
TCP 0 TCP
20 v (PO )-3
10 2 4 4 214
28 0.3
1 002 2 μm 300 v (CO )-2
5 2 3
Absorbance
211
02
300
0 202 -2 v3(CO3) (900 C) 0 0 20 25 30 35 40 2000 1600 1200 800 400 CDA TCP 2θ Wavenumber (cm-1)
CDA TCP Ca (wt%, n=6) 31.58 0.36* 33.54 0.40* P (wt%, n=6) 17.35 0.14* 18.56 0.18* Ca/P Molar Ratio (n=6) 1.41 0.02 1.40 0.02 BET SA (m2/g, n=3) 74.33 7.48* 2.069 0.78*
Figure 4.1 Ceramic characterization. Calcium deficient apatite (CDA) was sintered at 900°C for three hours to produce beta tricalcium phosphate (TCP). Scanning electron microscopy (SEM) images show irregular particle shape for the CDA and a rhombic particle shape for the TCP (10,000x, n=2). Crystalline peaks are detected for both ceramics using X-ray diffraction (XRD), with broader peak widths for the CDA (n=2). Using Fourier transform analysis (FTIR), phosphate bending peaks are observed for both ceramics; however, a carbonate peak is detected only for the CDA (n=2). TCP has higher calcium and phosphorus weight content than the TCP; however, both ceramics have similar calcium/phosphorus molar ratio (n=6, *p<0.05 for difference between ceramics). The specific surface area of the CDA is significantly larger than the surface area of the TCP (n=3, *p<0.05 for difference between ceramics).
54 No T3
Ceramic-free +T3
No T3
deep zone chondrocytes CDA CDA agarose +T3
No T3
TCP TCP +T3
Figure 4.2 Study design. Deep zone chondrocytes are mixed with agarose to make cellular constructs with either no particles (control), calcium deficient apatite (CDA) particles, or tricalcium phosphate (TCP) particles to generate three study groups. Each scaffold group was cultured in vitro with or without thyroid hormone (T3) stimulation.
55 Ceramic-free CDA TCP Ceramic-free +T3 CDA+T3 TCP+T3
D1
D7
D14 100 μm
16 16 Ceramic-free Ceramic-free+T3 CDA CDA+T3 TCP * * TCP+T3 12 # 12
* * * * # 8 * # 8 #
/ mg Wet Weight Wet mg / #
# # / mg Wet Weight Wet mg / 3 3 #
- # 3 3 # # - #
4 4 Cell No. x10 No. Cell 0 x10 CellNo. 0 D1 D7 D14 D1 D7 D14 Figure 4.3 Cell number and distribution. Hematoxylin and eosin staining reveals uniform cell distribution throughout all scaffolds at day 14 (10x, n=2). The calcium deficient apatite (CDA) group measures the highest cell number at day 14 both with and without thyroid hormone (T3) stimulation (n=5, *p<0.05 for difference between groups, #p<0.05 for difference from corresponding group at previous timepoint).
56 Ceramic-free CDA TCP Ceramic-free +T3 CDA+T3 TCP+T3
D1
D7
D14 100 μm
8 8 Ceramic-free Ceramic-free+T3 * * CDA * CDA+T3 # TCP * # TCP+T3 6 6
4 # 4
# # # #
2 2
μg Collagen/mg Wet Weight Collagen/mg μg Wet μg Collagen/mg Wet Weight Collagen/mg μg Wet 0 0 D1 D7 D14 D1 D7 D14 Figure 4.4 Collagen deposition. Picrosirius red staining shows positive collagen staining throughout all scaffolds, with the darkest staining observed for the calcium deficient apatite (CDA) group on day 14 (10x, n=2). The CDA group measures the highest collagen deposition on day 14 in the presence and absence of thyroid hormone (T3) stimulation (n=5, *p<0.05 for difference between groups, #p<0.05 for difference from corresponding group at previous timepoint).
57 Ceramic-free CDA TCP Ceramic-free +T3 CDA+T3 TCP+T3
Col I
Col II
Col X
200 μm
Figure 4.5 Day 14 collagen immunohistochemistry. Weak collagen I staining is detected for all groups except the calcium deficient apatite (CDA) group, with the brightest staining observed for the stimulated TCP group. Collagen II staining is detected for all groups with the most positive staining for the CDA group. Collagen X staining is positive for all groups except the CDA group, with the strongest staining observed in the stimulated control and TCP groups (10x, n=2).
58 Ceramic-free CDA TCP Ceramic-free +T3 CDA+T3 TCP+T3
D1
D7
D14
100 μm *
20 Cermaic-free * 20 * # Ceramic-free+T3 * CDA CDA+T3 # TCP TCP+T3 15 15
# 10 # 10 * * # # # # #
5 # # 5
μg GAG/mg Wet Weight GAG/mg μg Wet μg GAG/mg Wet Weight GAG/mg μg Wet
0 0 D1 D7 D14 D1 D7 D14
Figure 4.6 Glycosaminoglycan deposition. Alcian blue staining shows positive glycosaminoglycan staining throughout all scaffolds, with the darkest staining observed for the calcium deficient apatite (CDA) groups on day 14 (10x, n=2).The CDA group measures the highest proteoglycan deposition on day 14 in the presence and absence of thyroid hormone (T3) stimulation (n=5, *p<0.05 for difference between groups, #p<0.05 for difference from corresponding group at previous timepoint).
59 10 10 Ceramic-free Ceramic-free+T3 CDA CDA+T3 * TCP # 8 8 TCP+T3 * # 6 6 * # 4 4 #
* 2 * 2
ALP Activity (pmoles/cell/min) Activity ALP # # # (pmoles/cell/min) Activity ALP 0 0 D1 D7 D14 D1 D7 D14
Ceramic-free CDA TCP Ceramic-free +T3 CDA+T3 TCP+T3
D1 D1
Von Kossa Von
D14
D1 D1 Alizarin Red Alizarin
D14 100 μm
Figure 4.7 Alkaline phosphatase activity and mineral distribution. The calcium deficient apatite (CDA) group measures the highest ALP activity on day 1, with low ALP activity thereafter. The ceramic- free and TCP groups exhibit enhanced day 7 and day 14 ALP activity in the presence of thyroid hormone (T3), with the tricalcium phosphate (TCP) group measuring the highest ALP activity on day 14 (n=5, *p<0.05 for difference between groups, #p<0.05 for difference from corresponding group at previous timepoint). Von Kossa staining revealed similar staining on day 1 and day 14 for all groups, with black nodules present in the TCP and CDA groups. Alizarin red staining shows positive calcium staining throughout the CDA and TCP scaffolds at all timepoints, with negative staining for the ceramic-free scaffolds (10x, n=2).
60 DAY 1 DAY 14
10 10 Ceramic-free Ceramic-free CDA CDA * TCP TCP 8 8
6 6
4 * 4
2 2 *
* * * * *
Normalized Expression Gene Normalized GeneExpression 0 0 Col X Ihh MMP13 PTHrP ALP Col X Ihh MMP13 PTHrP ALP Figure 4.8 Gene expression. Calcium deficient 10 Ceramic-free apatite (CDA) and tricalcium phosphate (TCP) CDA+T3 groups exhibited downregulated col X and Ihh on TCP+T3 day 1 compared to the control. The CDA group 8 also exhibited the lowest ALP expression and the highest PTHrP. On day 14, the untreated CDA 6 group exhibited the lowest col X expression. * MMP13 was upregulated at day 14 for the 4 unstimulated TCP group. The stimulated TCP group exhibited upregulated Ihh, and both 2 stimulated ceramic groups MMP13 expression *
was downregulated with respect to the stimulated Normalized Expression Gene control group (n=5, *p<0.05 for difference between 0 groups). Col X Ihh MMP13 PTHrP ALP
61 120 Control Media Ceramic-free CDA TCP 120 Control Media Ceramic-free+T3 CDA+T3 TCP+T3
*
90 * * 90 *
g/ml)
g/ml)
μ μ
60 60
30 30
Media Media Calcium ( Media Media Calcium (
0 0 D1 D7 D14 D1 D7 D14
200 Control Media Ceramic-free CDA TCP 200 Control Media Ceramic-free+T3
CDA+T3 TCP+T3 g/ml)
150 g/ml) 150 μ * μ *
100 100
50 50
Media Phosphate Media Phosphate ( Media Phosphate (
0 0 D1 D7 D14 D1 D7 D14
Figure 4.9 Media ion concentrations. The calcium deficient apatite (CDA) group had lower media calcium concentration on day 1; however, no difference was noted between the CDA and control group after day 1. The tricalcium phosphate (TCP) group resulted in higher phosphate concentration in the media on day 1 (n=6, *p<0.05 for difference between groups).
62 CHAPTER 5: THE EFFECT OF CERAMIC DOSE ON DEEP ZONE CHONDROCYTE RESPONSE AND CALCIFIED CARTILAGE FORMATION IN AGAROSE SCAFFOLDS
63 5.1 Introduction
In chapter 4, the effect of ceramic crystallinity on calcified cartilage formation was investigated. It was determined that poorly crystalline calcium deficient apatite is promising for calcified cartilage regeneration because it promotes cell proliferation and matrix production, resulting in a calcified matrix that is rich in proteoglycans and collagen II. Reports have demonstrated, using an agarose-hydroxyapatite composite, that ceramic dose can influence cell response [11]; thus, in this chapter, the dose of calcium deficient apatite in agarose will be optimized to maximize matrix deposition and mechanical properties.
5.1.1 Background and Significance
This study evaluates the response of deep zone chondrocytes to varied doses of calcium deficient apatite in agarose scaffolds to promote calcified cartilage regeneration. While calcium deficient apatite enhances matrix deposition at a dose of 1.5% w/v, studies have shown that ceramic dose affects chondrocyte response and the resultant mechanical properties of the newly formed tissue [11]. Khanarian et al. evaluated the effect of the hydroxyapatite dose in agarose scaffolds on hypertrophy-stimulated deep zone chondrocytes. While 1.5% and 3% hydroxyapatite resulted in similar matrix deposition and cell number, the scaffold with 3% hydroxyapatite exhibited higher mechanical properties after two weeks of culture.
In this study, the response of deep zone chondrocytes in the presence and absence of thyroid hormone will be evaluated in agarose constructs with four doses of calcium deficient apatite: 0.75%,
1.5%, 3%, and 4.5%. In a full thickness defect, both hypertrophic and non-hypertrophic deep zone chondrocytes will be present near the cartilage-bone interface in the defect, as these are the cells located closest to this region. Studies have demonstrated that thyroid hormone (T3) stimulates the hypertrophic phenotype in deep zone chondrocytes [188], and the deep zone chondrocytes that reside near a full thickness defect will likely be exposed to thyroid hormone because it is present in the circulation [182].
5.1.2 Objectives
The objective of this study is to investigate the effect of calcium deficient apatite dose on chondrocyte response in order to determine the ceramic dose that is most conducive to regenerating
64 calcified cartilage. It is hypothesized that increasing ceramic content will increase hypertrophy and matrix deposition.
5.2 Materials and Methods
5.2.1 Cells and Cell Culture
Primary deep zone chondrocytes (DZCs) were isolated and pooled from the femoral articular cartilage of five immature calf knees (Green Village Packing Co.) following previously described protocols [79]. The bottom third of the cartilage was separated, and the calcified cartilage was removed by scraping. The cartilage was minced and digested with collagenase type 2 (310 u/mg, Worthington) for 16 hours in
Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 10% fetal bovine serum (FBS, Atlanta
Biologicals, Atlanta, GA), 2% antibiotics (10,000 U/mL penicillin, 10 mg/mL streptomycin), and 0.2% antifungal (250 µg/mL amphotericin B). The DZC suspension was filtered before plating (30 µm,
Spectrum). The isolated chondrocytes were maintained in high-density culture in fully-supplemented
DMEM with 10% FBS, 1% non-essential amino acids, 1% penicillin-streptomycin, 0.1% gentamicin sulfate, and 0.1% antifungal for three days prior to seeding in the hydrogel [15]. All media supplements were purchased from Cellgro-Mediatech unless otherwise specified.
5.2.2 Scaffold Fabrication, Characterization, and Culture
Chondrocytes were encapsulated at a density of 10 million cells/mL in 2% low gelling agarose
(Agarose Type VII, Sigma) and varying doses of CDA (Fig. 5.1). A biopsy punch (Sklar Instruments, West
Chester, PA) was used to core cylindrical scaffolds (Ø = 5 mm, height = 2.4 mm) [11]. Acellular and cellular agarose scaffolds with varying concentrations of ceramic, added at the time of fabrication, and corresponding controls without ceramic were fabricated.
⁰ All samples were cultured under humidified conditions at 37 C and 5% CO2 and maintained in ITS media composed of DMEM supplemented with 1% ITS+ Premix (BD Biosciences), 1% penicillin- streptomycin, 0.1% gentamicin sulfate, 0.1% antifungal (250 µg/mL amphotericin B), and 40 µg/mL L- proline (Sigma). The medium was changed every other day and freshly supplemented with 50 µg/mL ascorbic acid (Sigma). In order to induce hypertrophy, DZCs were stimulated with 25 nM triiodothyronine
(T3, Sigma) during the first three days of culture [174].
65 Gross morphology of acellular scaffolds was assessed with a stereoscope (Olympus SZ61). Scaffold wet weight (n=3) was measured (P114 Pinnacle Series Balance, Denver Instruments), and ash weight was determined by thermogravimetric analysis (TA Q-50, TA Instruments). Scaffold mechanical properties
(n=3) were determined on a shear-strain controlled rheometer (TA instruments) following published protocols [189]. Briefly, each sample was placed between two flat porous platens and immersed in DMEM to prevent dehydration. First, the equilibrium compressive Young’s modulus (Eeq) was calculated at 15% compressive strain, which is within the physiological range for articular cartilage [190]. A dynamic shear test was then performed (0.01 Hz to 10 Hz) with a logarithmic frequency sweep at a shear strain amplitude of 0.01 radian. Both the magnitude of the complex shear modulus (|G*|) and phase shift angle
(δ) between the applied strain and the resulting torque were determined at 1 Hz.
5.2.3 Cell Viability and Number
Cell viability and death (n = 2) were visualized using live/dead staining (Invitrogen). The samples were rinsed with phosphate buffered saline (PBS, Sigma), stained following the manufacturer’s suggested protocol, and imaged under confocal microscopy (Olympus Fluoview IX70) at wavelengths of 488 nm and
568 nm, respectively. Cell number (n=5) was determined using the Quanti-iT™ PicoGreen® dsDNA assay kit (Molecular Probes, Eugene, OR) following sample digestion. The sample was rinsed with PBS and exposed to a freeze-thaw cycle in 500 μL of 0.1% Triton™-X solution (Sigma) in order to lyse the cells.
After desiccation for 12 hours in a CentriVap Concentrator (Labconco Co.), the samples were digested for
18 hours at 65°C with papain (8.3 activity units/mL) in 0.1 M sodium acetate (Sigma), 10 mM cysteine- hydrochloric acid (Sigma), and 50 mM ethylenediaminetetraacetate (Sigma). For DNA content, a 25 μL aliquot of the sample was mixed with 175 μL of the PicoGreen® working solution, and fluorescence was measured with a microplate reader (Tecan) at excitation and emission wavelengths of 485 and 535 nm, respectively. The conversion factor of 7.7 pg DNA/cell was used to determine cell number [158;159].
5.2.4 Mineralization
Alkaline phosphatase (ALP) activity (n=5) was measured using a colorimetric assay based on the hydrolysis of p-nitrophenyl phosphate (pNP-PO4) to p-nitrophenol (pNP) [164]. The samples were lysed in
0.1% Triton™ X solution, exposed to a freeze-thaw cycle, and crushed with a pestle. A 25 μL aliquot was
66 added to pNP-PO4 solution (Sigma) and incubated for 10 minutes at 37°C. Absorbance was measured at
405 nm using a microplate reader (Tecan). In addition, phosphate distribution (n=2) was evaluated as an indicator of overall mineral distribution. For phosphate staining, depariffinized samples were stained with
5% silver nitrate and exposed to UV light (365 nm) for 25 minutes before a tap water rinse. Cover-slipped sections were imaged using a brightfield microscope (Zeiss, Axiovert 25).
5.2.5 Matrix Deposition
Total collagen content (n=5) was quantified using a modified hydroxyproline assay [160] with bovine collagen I solution (Biocolor, Carrickfergus, UK) as the standard. Briefly, a 40 μL aliquot of sample digest was mixed with 10 μL of 10 N sodium hydroxide and heated to 250°C for 25 minutes in order to hydrolyze the collagen. The hydrolyzate was then oxidized at room temperature for 25 minutes with 450 μL buffered chloramine-T reagent prior to the addition of 425 µL Ehrlich’s reagent (15% p- dimethylaminobenzaldehyde in 2:1 isopropanol/percholoric acid). Absorbance was measured at 555 nm with a microplate reader (Tecan). All matrix values were normalized by sample wet weight in order to account for any differences in sample size. Additionally, collagen distribution (n=2) was evaluated via histology. The samples were first fixed in neutral buffered formalin with 1% cetylpyridinium chloride
(Sigma) for 24 hours followed by dehydration with an ethanol series. The dehydrated samples were embedded in paraffin (Paraplast X-tra Tissue Embedding Medium, Fisher Scientific), and 7 μm sections were obtained from the center of the scaffold. Paraffin was cleared with xylenes, the samples were rehydrated, stained with picrosirius red staining for one hour, and exposed to 0.1 N hydrochloric acid for two minutes (n=2). In addition, collagen I and II were assessed using immunohistochemistry (n=2) [11].
Specifically, monoclonal antibodies for collagen I (1:00 dilution) and collagen II (1:100 dilution) were purchased from Abcam (Cambridge, MA). After fixation, samples were treated with 1% hyaluronidase for
30 minutes at 37°C and incubated with primary antibody overnight. Cell nuclei were stained with 4',6- diamidino-2-phenylindole (Sigma). A FITC-conjugated secondary antibody (1:200 dilution, LSAB2 Abcam) was used, and samples were imaged under confocal microscopy (Olympus Fluoview IX70, Center Valley,
PA) at excitation and emission wavelengths of 488 nm and 568 nm, respectively.
Sample glycosaminoglycan content (GAG, n=5) was determined with a modified 1,9- dimethylmethylene blue (DMB) binding assay [161-163], with chondrotin-6-sulfate (Sigma) as the
67 standard. The absorbance difference between 540 nm and 595 nm was used to improve the sensitivity in signal detection. In addition, histology was used to visualize glycosaminoglycan distribution.
Deparaffinized sections were exposed to 3% acetic acid for three minutes, stained with alcian blue for 45 minutes, and rinsed twice with acid-alcohol (pH=1) for one minute (n=2) [159].
5.2.6 Statistical Analysis
Results are presented as mean ± standard deviation, with n equal to the number of samples per group. A two-way analysis of variance (ANOVA) was performed to determine the effects of ceramic dose and culturing time on cell response (proliferation, matrix deposition, ALP activity, solution ion concentration, and mechanical properties). The Tukey-Kramer post-hoc test was used for all pair-wise comparisons, and significance was attained at p<0.05. Statistical analyses were performed with JMP IN
(4.0.4, SAS Institute, Inc.).
5.3 Results
5.3.1 Scaffold Characterization
Scaffolds with 0%, 0.75%, 1.5%, 3%, and 4.5% CDA were fabricated (Fig. 5.1) and characterized in terms of gross appearance, wet and ash weight, and mechanical properties (Table 5.1). The ceramic-free scaffold was translucent, whereas the ceramic-containing scaffolds were more opaque. While wet weights were similar between the control and the 0.75%, 1.5% and 3% CDA groups, the 4.5% CDA group was significantly heavier than the ceramic-free control. A dose-dependent increase in ash weight was detected for all ceramic-containing groups. Day 1 mechanical properties were measured for all groups in terms of compressive modulus, dynamic shear modulus, and phase shift angle. While there were no differences detected between groups for Young’s modulus or phase shift angle, the dynamic shear modulus was significantly higher than the control group for both the 3% and 4.5% CDA groups.
5.3.2 Cell Viability and Number
Cell number and viability were tracked over time for all groups (Fig. 5.2). Cell number was similar for all groups on day 1. For the unstimulated groups, cell number increased over time for all ceramic- containing groups and was maintained for the ceramic-free control group. On day 21, the highest cell
68 number was measured for the 4.5% group; however, there was a decrease in cell number in this group between day 21 and day 42. On day 42, the 0.75%, 1.5%, and 3% groups exhibited the highest cell number. These findings were confirmed with live/dead staining that showed viable cells in all groups. In the presence of T3 stimulation, an increase in cell number for all ceramic containing groups was measured by day 7, with higher cell number for ceramic-containing groups than the control regardless of dose. On day 21, the 3% and 4.5% groups exhibited the highest cell number; however, a significant decrease in cell number was measured for these groups between day 21 and day 42. On day 42, the highest cell number was detected for the 0.75% and 1.5% groups. Corresponding viability staining revealed many dead cells in the 3 and 4.5% CDA groups that had been stimulated with T3, with viable cells detected in all other groups.
5.3.3 Matrix Deposition
Collagen and glycosaminoglycan (GAG) deposition were measured for all groups over time (Fig. 5.3,
Fig. 5.4, and Fig. 5.5). Collagen deposition in the unstimulated groups increased over time between day 1 and day 14 for the ceramic-free, 0.75% and 1.5% groups. On day 14 and day 21, significantly higher collagen content was measured for the unstimulated 0%, 0.75% and 1.5% CDA groups compared to the unstimulated 3% and 4.5% groups and for the stimulated 0.75% and 1.5% CDA compared to the stimulated 3% and 4.5% CDA groups. On day 42, the highest collagen content was measured for the
0.75% group, with a dose-dependent decrease in collagen measured for the 1.5, 3, and 4.5% groups compared to the 0.75% group. For the groups stimulated with T3, collagen content was highest in the
0.75% and 1.5% CDA scaffolds on day 42 and lowest for the 3% and 4.5% groups throughout the study.
The histological analysis corroborated these findings, with strongly positive staining for the 0.75% and
1.5% groups both with and without T3 and weak staining observed in the unstimulated 4.5% group and the stimulated 3 and 4.5% groups. Collagen content was further analyzed for all groups on day 42 via immunohistochemical staining (Fig.5.4). All scaffolds exhibited weak or negative collagen I and collagen
X staining and positive collagen II staining.
Glycosaminoglycan content was measured for each group over time (Fig. 5.5). In the absence of T3, the highest GAG content was measured for the 0.75%, 1.5%, and 3% groups on day 42, and a significant loss of GAG was detected for the 4.5% group between day 21 and day 42. In the presence of T3, the
69 0.75% and 1.5% groups exhibited the highest GAG content on day 42, with no GAG detected in the 3% and 4.5% groups at this timepoint. In both cases, the control maintained constant GAG from day 14 to day 42. Histological analysis confirmed these findings, revealing strong staining of uniformly distributed
GAG for the 0.75%, 1.5%, and 3% unstimulated groups as well as the 0.75% and 1.5% stimulated groups.
5.3.4 Mechanical Properties
Mechanical properties were assessed on days 1, 21, and 42 for all groups (Fig. 5.6). Properties increased over time for all ceramic-containing groups, and the unstimulated 0.75%, 1.5% and 3% groups exhibited higher Young’s modulus and dynamic shear modulus by day 42 than the control. On day 42, the stimulated 0.75% and 1.5% exhibited higher mechanical properties than the control but lower properties than corresponding unstimulated groups. In the presence of T3, the 0%, 3%, and 4.5% group had the lowest mechanical properties; however, in the absence of T3, the 3% resulted in the strongest constructs.
5.3.5 Mineralization
Alkaline phosphatase (ALP) activity and von Kossa staining were used to assess mineralization potential and mineral distribution (Fig. 5.7). ALP activity was elevated on day 1 for the 3% and 4.5% groups, with low ALP activity after day 1 for all unstimulated groups. For T3-stimulated groups, a late peak in ALP activity on day 42 was observed for the 3% group. Positive von Kossa staining was observed for all mineral-containing groups at day 42.
5.4 Discussion
The goal of this study is to optimize the dose of ceramic in an agarose-calcium deficient apatite composite scaffold for calcified cartilage formation. To this end, the response of deep zone chondrocytes was evaluated over time in agarose scaffolds with 0%, 0.75%, 1.5%, 3%, and 4.5% calcium deficient apatite in the presence and absence of thyroid hormone. The results demonstrate that deep zone chondrocytes are sensitive to calcium deficient apatite dose within agarose gels, with the highest matrix deposition detected for 0.75%, 1.5%, and 3% unstimulated groups and 0.75% and 1.5% stimulated groups. While the 0.75% and 1.5% groups both promoted matrix elaboration in the presence and absence of thyroid hormone, the constructs with 1.5% calcium deficient apatite exhibited significantly
70 higher mechanical properties by day 42 compared to the 0.75% calcium deficient apatite. Based on these findings, a calcium deficient apatite dose of 1.5% is the optimal for calcified cartilage regeneration.
In this study, lower doses of calcium deficient apatite promoted the development of a matrix rich in glycosaminoglycan and collagen II, while higher doses inhibited matrix elaboration. This finding echoes previously reported results by Khanarian et al. [11], showing that matrix deposition was elevated for deep zone chondrocytes in the presence of 1.5% and 3% hydroxyapatite and thyroid hormone but was inhibited when the dose of hydroxyapatite was raised to 6%. In our study, we measured inhibition of matrix deposition in the presence of thyroid hormone beginning at a lower dose of 3%. This difference is likely a result of the increased bioactivity of calcium deficient apatite compared to hydroxyapatite, as bioactive ceramics at high doses have been shown to result in less matrix than more inert ceramics in vivo [172]. This effect may be related to surface chemistry and ion dissolution. In addition, in agarose- ceramic composites, high doses of calcium phosphate particles within the scaffold limit the space available for matrix elaboration relative to lower doses. The incorporation of a high dose of ceramic also influences the scaffold mechanics. Therefore, chondrocytes cultured within the high dose gels experience a stiffer environment, and the reduction in matrix deposition may be a result of these mechanical cues.
Schuh et al. has shown that chondrocytes cultured in stiff agarose scaffolds produce less GAG than chondrocytes cultured in softer agarose scaffolds, indicating that mechanosensing may play a central role in modulating matrix elaboration in three-dimensional constructs [191].
The trends for matrix/scaffold weight were similar to matrix/cell, with glycosaminoglycan and collagen production elevated per cell for the low dose groups and suppressed for the high dose groups. This suggests that low doses of calcium deficient apatite not only promote chondrocyte proliferation, resulting in more cells that are capable of producing glycosaminoglycans and collagen, but may also increase biosynthesis at a cellular level. Elevation of matrix production/cell over the control scaffold was significant for glycosaminoglycans for unstimulated 0.75%, 1.5%, and 3% groups and stimulated 0.75% and 1.5% groups. However, in terms of collagen/cell, a significant elevation in matrix/cell was measured only for the unstimulated 1.5% group. This finding is in agreement with previously reported results that indicate there is a tendency for chondrocytes to preferentially produce glycosaminoglycans relative to collagen in hydrogel-based tissue engineered constructs [11;192].
71 Mechanical properties of the gel, measured at day 1, 21, and 42 follow similar patterns to the matrix deposition. The highest compressive and shear moduli were measured for the 1.5% and 3% unstimulated groups. Despite the high glycosaminoglycan and collagen content found for the 0.75% group, the resulting construct was not as strong as the 1.5% and 3% groups, indicating that the mineral may play a synergistic role in reinforcing the glycosaminoglycan-collagen matrix. Similar findings were reported for agarose-hydroxyapatite scaffolds in which higher doses of mineral were found to result in higher mechanical properties despite no measurable differences in matrix content between the scaffold groups
[188].
This study demonstrated that, in addition to matrix production, chondrocyte mineralization potential was modulated by the dose of calcium deficient apatite. On day 1, alkaline phosphatase activity was upregulated for the 3% and 4.5% groups, with a significant decline in activity by day 7. In the stimulated groups, a second peak in ALP activity was measured for the 3% CDA group on day 42. Positive von
Kossa staining was observed for all groups, with no marked differences between corresponding stimulated or unstimulated groups. While the histological analysis indicates similar mineral presence, the nature of the mineral cannot be distilled from the stains, and it is possible that the cells have remodeled the mineral over time in the scaffolds. The early alkaline phosphatase activity is consistent with the results from chapter 4 and previous reports of deep zone chondrocytes cultured in agarose-ceramic constructs
[11]. Since the calcium deficient apatite causes early (day 1) calcium precipitation from the media
(chapter 4), it is possible that the precipitation of high doses of calcium into the scaffold enhance the alkaline phosphatase activity in the high dose scaffolds. In a study by Subramony et al., high doses of ceramic within polymer scaffolds were also reported to result in increased early alkaline phosphatase activity of mesenchymal stem cells, corroborating findings that alkaline phosphatase activity is a function of ceramic dose [193]. The late peak in alkaline phosphatase activity on day 42 for the 3% group mirrors a late peak previously reported for chondrocytes cultured on polymer scaffolds with 10% incorporated hydroxyapatite, in which a similar late peak was not observed for a higher dose of hydroxyapatite or for a ceramic-free control [154].
In summary, the results of this study demonstrate that deep zone chondrocyte response is modulated by calcium deficient apatite dose in agarose scaffolds in terms of growth, biosynthesis, and
72 mineralization. While low doses of bioactive mineral promote glycosaminoglycan and collagen II deposition, high doses of ceramic inhibit chondrocyte biosynthesis. Though both 0.75% and 1.5% ceramic enhanced matrix deposition in the presence and absence of thyroid hormone stimulation, the
1.5% dose resulted in the highest mechanical properties and was the only dose to result in increased glycosaminoglycan and collagen deposition per cell by day 42.
5.5 Conclusions
Collectively, the results of this study and chapter 4 provide the optimal design specifications to promote calcified cartilage-like tissue formation by the hydrogel-based scaffold of the multi-phased cup integration system. An agarose-calcium deficient apatite system incorporating 1.5% ceramic supports chondrocyte viability and promotes matrix deposition, resulting in increased mechanical properties.
Therefore, the hydrogel-ceramic scaffold in the base of the cup will consist of agarose with 1.5% calcium deficient apatite.
73 0% No T3
0.75% deep zone chondrocytes 1.5% CDA agarose
3% +T3
4.5%
Figure 5.1 Study design. Deep zone chondrocytes and calcium deficient apatite (CDA) were added to agarose to fabricate scaffolds with five different doses of CDA particles. Each scaffold group was cultured in vitro with or without thyroid hormone (T3) stimulation.
74 Table 5.1 Acellular scaffold characterization for agarose constructs with varied CDA content Day 1 (*p<0.05) 0% 0.75% 1.5% 3% 4.5% CDA CDA CDA CDA CDA Top View Scale Bar = 2 mm
Wet Weight (mg) 41.3 0.8 41.4 0.9 40.7 0.6 42.3 0.7 43.8 0.9* Ash Weight (mg) N/A 0.4 0.1 0.6 0.2 1.2 0.2 2.0 0.2
Compressive 5.1 1.6 5.8 1.4 4.7 0.9 8.4 1.7 7.8 1.5 Modulus (kPa)
Dynamic Shear 7.9 0.3 11.1 1.7 8.8 2.1 11.6 0.9* 13.4 1.1* Modulus (kPa) Phase Shift (deg) 1.3 0.7 2.7 0.7 1.9 1.3 1.8 0.7 2.7 0.8
*p<0.05 for difference from ceramic-free control
75 25000 25000 0% 0%+T3 0.75% * 0.75%+T3 20000 1.5% 1.5%+T3 # 20000 3% 3%+T3 # * # 4.5%+T3 * 4.5% * 15000 15000 * # * # # # # # 10000 10000
# #
Cells/mg Wet Weight Wet Cells/mg Cells/mg Wet Weight Wet Cells/mg 5000 5000
0 0 D1 D14 D21 D42 D1 D14 D21 D42 0% 0.75% 1.5% 3% 4.5%
-T3
+T3
Day 42, scale bar = 200 µm
Figure 5.2 Cell number, viability, and distribution. In the absence of thyroid hormone (T3) stimulation, cell number was highest for the 0.75, 1.5 and 3% calcium deficient apatite (CDA) groups by day 42. In the presence of thyroid hormone (T3), cell number was highest for the 0.75 and 1.5% groups on day 42 (n=6, *p<0.05 for difference between groups, #p<0.05 for difference from corresponding group at previous timepoint). Uniformly distributed, viable cells were observed in all groups, except the stimulated 3% group and both 4.5% groups (n=2).
76 20 0% 20 0%+T3 0.75% * 0.75%+T3 1.5% * 1.5%+T3 3% # 3%+T3 4.5% 4.5%+T3 * # * * * * 10 * * 10 # # # # * * # *# #
# #
ug Collagen/mg Wet Weight Wet Collagen/mg ug ug Collagen/mg Wet Weight Wet Collagen/mgug
0 0 D1 D14 D21 D42 D1 D14 D21 D42
Collagen/cell (ng/cell) on day 42 Groups 0% 0.75% 1.5% 3% 4.5% -T3 0.93 0.3 0.88 0.2 0.76 0.0 0.47 0.0 0.58 0.2 +T3 0.50 0.1 0.63 0.1 0.70 0.1 0.20 0.1 0.22 0.1
0% 0.75% 1.5% 3% 4.5%
-T3
+T3
Day 42, scale bar = 200 µm
Figure 5.3 Collagen production. Collagen production, normalized by wet weight, was highest for the unstimulated 0.75% and stimulated 0.75 and 1.5% calcium deficient apatite (CDA) groups by day 42 (n=6, *p<0.05 for difference between groups, #p<0.05 for difference from corresponding group at previous timepoint). Collagen content, normalized by cell number, was highest for the stimulated 1.5% CDA group, and lowest for the stimulated and unstimulated 3 and 4.5% CDA groups. Significant increases over the control are highlighted in green, and significant decreases are shown in red (n=6, *p<0.05). Picrosirius red staining confirms the quantitative findings and shows uniform collagen distribution throughout the agarose constructs (n=2).
77 0% 0.75% 1.5% 3% 4.5%
-T3 CollagenI
+T3
-T3 CollagenII
+T3
-T3 CollagenX
+T3
Day 42, scale bar = 400 µm
Figure 5.4 Collagen immunohistochemistry. Negative collagen I and X staining and positive collagen II staining were found for all groups (n=2).
78 30 0% * * 30 0%+T3 # 0.75% 0.75%+T3 * * 25 # # 25 1.5% 1.5%+T3 * 3% 3%T3 # 20 4.5% * 20 4.5%+T3 * # * # * 15 15 # * # # # 10 10 * # # # #
5 5
ug GAG/mg Wet Weght Wet GAG/mg ug ug GAG/mg Wet Weight Wet GAG/mg ug # 0 0 D1 D14 D21 D42 D1 D14 D21 D42 -5 -5
GAG/cell (ng/cell) on day 42 Groups 0% 0.75% 1.5% 3% 4.5% -T3 0.72 ± 0.1 1.46 ± 0.2 1.62 ± 0.2 1.30 ± 0.2 0.31 ± 0.3 +T3 0.32±0.3 1.0±0.3 1.45±0.4 -0.30±0.4 -0.18±0.3
0% 0.75% 1.5% 3% 4.5%
-T3
+T3
Day 42, scale bar = 200 µm
Figure 5.5 Glycosaminoglycan production. Glycosaminoglycan (GAG) content, normalized by wet weight, was highest for the unstimulated 0.75, 1.5 and 3% calcium deficient apatite (CDA) groups and for the stimulated 0.75 and 1.5% CDA groups by day 42 (n=6, *p<0.05 for difference between groups, #p<0.05 for difference from corresponding group at previous timepoint). GAG content, normalized by cell number, followed similar trends, with significant increases over the control highlighted in green and significant decreases shown in red (n=6, p<0.05). Alcian blue staining confirms the quantitative findings and shows uniform collagen distribution throughout the agarose constructs (n=2).
79 200 0% * 200 0% 0.75% # # 0.75% 1.5% 1.5% 3% 3% 150 150 4.5% 4.5% * # * # 100 * # #^ 100 (kPa) * * # *^ #^ #
# # (kPa) Modulus Young's 50 50 # * * * # ^ # Dynamic Shear Modulus lG*l at 1 Hz 1 at lG*l Modulus Shear Dynamic ^ ^ 0 0 D1 D21 D21+T3 D42 D42+T3 D1 D21 D21+T3 D42 D42+T3
20 0% Figure 5.6 Mechanical properties. Dynamic 0.75% shear modulus and Young’s modulus were 1.5% highest for the unstimulated 0.75 and 1.5% 3% 4.5% calcium deficient apatite (CDA) groups on day # * * 42 (n=3, *p<0.05 for difference between # groups, #p<0.05 for difference from 10 corresponding group at previous timepoint, # ^p<0.05 for difference from corresponding unstimulated group).
Modulus at 1 Hz (kPa) Hzat 1 Modulus #^ Phase Shift Angle of Dynamic Shear Dynamic of Angle Shift Phase
0 D1 D21 D21+T3 D42 D42+T3
80 6 * 0%+T3 6 0% * 0.75%+T3 0.75% 1.5%+T3 5 5 1.5% 3%+T3 3% 4.5%+T3 4.5% 4 4
3 * 3 *
2 2 # 1
1 (picomoles/cell/min) Activity ALP ALP Activity (picomoles/cell/min) Activity ALP # # 0 0 D1 D14 D21 D42 D1 D14 D21 D42
0% 0.75% 1.5% 3% 4.5%
-T3
+T3
Day 42, scale bar = 200 µm
Figure 5.7 Mineralization. ALP activity was highest for the 3 and 4.5% groups on day 1 with low ALP activity thereafter for all groups, except for the stimulated 3% group, which exhibited a peak in activity on day 42 (n=6, *p<0.05 for difference between groups, #p<0.05 for difference from corresponding group at previous timepoint). Von Kossa staining shows uniform mineral distribution through the ceramic- containing scaffolds at day 42 (n=2).
81 CHAPTER 6: EFFECT OF CERAMIC DOSE IN MICROFIBER SCAFFOLDS ON DEEP ZONE CHONDROCYTE RESPONSE AND CALCIFIED CARTILAGE FORMATION
82 6.1 Introduction
In chapters 4 and 5, ceramic composition and dose were optimized in an agarose-ceramic composite scaffold for interface regeneration, and it was determined that calcium deficient apatite promoted glycosaminoglycan and collagen production in a dose-dependent manner. In this chapter the dose of calcium deficient apatite in polymeric microfiber scaffolds will be optimized to enhance calcified cartilage formation in the base of the cup-shaped scaffold. Preliminary findings from an in vivo study in rabbits [17] suggest that the combination of fiber-based scaffolds with hydrogels containing ceramic increases the organization of the newly formed interface compared to either scaffold alone, indicating that a hybrid scaffold composed of polymer, hydrogel, and ceramic phases may be ideal for calcified cartilage regeneration. While agarose is well-suited for chondrocyte culture and regeneration of cartilage tissue, a hydrogel-based scaffold system may not provide a sufficient physical barrier between the cartilage and bone regions due to its high water content. Furthermore, although an interface scaffold made of hydrogel will integrate well with hydrogel-based cartilage grafts, an agarose-based scaffold is difficult to handle given the dimensions of the interface (<300 µm thick). Therefore, the agarose-ceramic scaffold will be combined with a fibrous scaffold to form the base of the cup to facilitate integration with cartilage grafts while improving structural integrity and ease of handling.
6.1.1 Background and Motivation
This study optimizes a microfiber-ceramic scaffold to serve as a temporary barrier to osseous upgrowth during the healing of full thickness defects and to promote calcified cartilage formation by chondrocytes. Fiber-based scaffolds composed of polymers and ceramic particles have been studied for osteochondral tissue engineering applications [125;137;194], and chondrocytes have been shown to attach, proliferate, and produce matrix on these scaffold systems. However, few studies have investigated the use of a poorly crystalline ceramic phase that mimics the native interface. Furthermore, though the response of articular, full thickness chondrocytes has been characterized on a variety of scaffolds, the response of deep zone chondrocytes, which reside directly above the interface on polymer-ceramic scaffolds, has not been studied in detail.
83 6.1.2 Objectives
The first objective of this study is to fabricate and characterize a scaffold for interface regeneration using the stable 5:1 PLGA:PCL microfiber composition, optimized in chapter 1, and varying concentrations of calcium deficient apatite. The second objective is to evaluate the dose of calcium deficient apatite that is most conducive to regenerating calcified cartilage in microfiber scaffolds.
Increasing the content of calcium deficient apatite is hypothesized to increase matrix deposition and mineralization potential, similar to the response of chondrocytes in composite hydrogel scaffolds.
6.2 Materials and Methods
6.2.1 Scaffold Fabrication and Characterization
Unaligned microfiber scaffolds were fabricated using the electrospinning process [20;149] (Fig. 6.1).
For control scaffolds, a 32% (w/v) polymer (5:1 PLGA (85:15, Lakeshore Biomaterials):PCL (Sigma-
Aldrich, Mw ≈ 70,000-90,000) solution in 60/40 DCM/DMF was vortexed continuously for one hour. To fabricate PLGA:PCL-calcium deficient apatite (CDA, Sigma-Aldrich) scaffolds, CDA nanoparticles were directly added to the PLGA:PCL polymer solution at varying concentrations. Each solution was loaded into a 5 mL syringe with a stainless steel blunt tip needle (26.5 gauge for PLGA:PCL blends, 23 gauge for polymer-ceramic blends) that was 13 cm from the collecting target, and electrospun at 8-10 kV using a custom electrospinning device. The polymer solution was deposited (1 mL/hour) onto a stationary collecting target using a syringe pump (Harvard Apparatus).
As-fabricated microfiber scaffolds were imaged with SEM (2 kV, Hitachi 4700, Hitachi Ltd.) to evaluate fiber morphology and Energy Dispersive X-ray Analysis (EDXA, 15 kV Princeton Gamma Tech) was utilized to determine scaffold elemental composition. Scaffolds were sputter-coated (Cressington
108auto) with gold-palladium to reduce charging effects. Microfiber diameter (n=3) was quantified via image analysis of SEM micrographs. The scaffolds were imaged in both secondary and backscatter modes and EDXA spectra were collected at randomly selected scaffold regions. Signal was collected at
100 seconds with a dead time of ~30% and 3000-4000 counts per second (CPS). The weight fraction of
CDA incorporated in the electrospun scaffolds (n=3) was validated using thermo-gravimetric analysis
(TGA-Q500, TA Instruments). The sample was loaded into a calibrated platinum pan, heated to 100°C in
84 nitrogen, and ramped at 20°C/minute to 700°C in oxygen. The ash weight was determined at the end of the temperature ramp. The degradation temperature of PLGA and PCL is approximately 400°C; therefore, the residual weight corresponded to the weight percent of mineral in the fibers.
Chemistry and crystallinity (n=2) of the as-fabricated polymer and polymer-ceramic microfiber scaffolds were analyzed using Fourier Transform Infrared Spectroscopy (FTIR, PerkinElmer Frontier) and
X-ray diffraction (XRD, X’Pert3 Powder, PANalytical), respectively. The FTIR spectra were collected in attenuated total reflectance (ATR) mode at a resolution of 4 cm-1. X-ray diffraction spectra were collected using a zero background plate over a range of 20-40° with a step size of 0.01 and a scan time of 120 seconds/step.
Polymer degradation was examined in vitro after 7, 14, 28, 42, 56, 70, and 84 days of culture in ITS- supplemented medium. At each timepoint, total weight loss (n=6) was determined for each sample. After incubation, the samples were rinsed with distilled water, dried in a vacuum (CentriVap, Labconco), and the dry weight was measured (SE2 balance, Sartorius).
6.2.2 Cells and Cell Culture
Primary deep zone chondrocytes were isolated and pooled from the femoral articular cartilage of five immature calf knees (Green Village Packing Co.) following published protocols [79]. The bottom third of the cartilage was separated and the calcified cartilage was removed by scraping. The cartilage was then minced and digested with collagenase type 2 (310 u/mg, Worthington) for 16 hours in Dulbecco’s Modified
Eagle’s Medium (DMEM) supplemented with 10% fetal bovine serum (FBS, Atlanta Biologicals, Atlanta,
GA), 2% antibiotics (10,000 U/mL penicillin, 10 mg/mL streptomycin), and 0.2% antifungal (250 µg/mL amphotericin B). The deep zone chondrocyte suspension was filtered before plating (30 µm, Spectrum,
Rancho Dominguez, CA). The isolated chondrocytes were maintained in high-density culture in fully- supplemented DMEM with 10% FBS, 1% non-essential amino acids, 1% penicillin-streptomycin, 0.1% gentamicin sulfate, and 0.1% antifungal for three days prior to seeding in the hydrogel [15]. All media supplements were purchased from Cellgro-Mediatech unless otherwise specified.
To seed the cells onto the scaffolds, 10 µL of concentrated cell suspension was pipetted onto each scaffold to achieve a seeding density of 100,000 cells/cm2 and incubated for 25 minutes at 37°C and 5%
85 CO2 before submersion in media. Cell-laden scaffolds were cultured at 37°C and 5% CO2 in 1.5 mL of media which was refreshed three times weekly.
6.2.3 Cell Viability and Number
Cell viability and death (n = 2) were visualized using live/dead staining (Invitrogen). The samples were rinsed with phosphate buffered saline (PBS, Sigma), stained following the manufacturer’s suggested protocol, and imaged under confocal microscopy (Olympus Fluoview IX70) at wavelengths of 488 nm and
568 nm, respectively. Cell proliferation (n=5) was determined using the Quanti-iT™ PicoGreen® dsDNA assay kit (Molecular Probes, Eugene, OR) following sample digestion. The sample was rinsed with PBS and exposed to a freeze-thaw cycle in 500 μL of 0.1% Triton-X solution (Sigma) in order to lyse the cells.
After desiccation for 12 hours in a CentriVap Concentrator (Labconco Co.), the samples were digested for
18 hours at 65°C with papain (8.3 activity units/mL) in 0.1 M sodium acetate (Sigma), 10 mM cysteine-
HCl (Sigma), and 50 mM ethylenediaminetetraacetate (Sigma). For DNA content, a 25 μL aliquot of the sample was mixed with 175 μL of the PicoGreen® working solution and fluorescence was measured with a microplate reader (Tecan, Research Triangle Park, NC), at excitation and emission wavelengths of 485 and 535 nm, respectively. The conversion factor of 7.7 pg DNA/cell was used to determine cell number
[158;159].
6.2.4 Matrix Deposition
Total collagen content (n=5) was quantified using a modified hydroxyproline assay [160] with bovine collagen I solution (Sigma) as the standard. Briefly, a 100 μL aliquot of sample digest was dehydrated in a vacuum oven (Isotemp Vacuum Oven Model 280A, Fisher Scientific) overnight and mixed with 25 μL 2 N sodium hydroxide and heated to 250°C for 25 minutes in order to hydrolyze the collagen. The hydrolyzate was then oxidized at room temperature for 25 minutes with 450 μL of buffered chloramine-T reagent prior to the addition of Ehrlich’s reagent (15% p-dimethylaminobenzaldehyde in 2:1 isopropanol/percholoric acid). Absorbance was measured at 555 nm with a microplate reader (Tecan). Additionally, collagen distribution (n=2) was evaluated via histology. The samples were first fixed in neutral buffered formalin with 1% cetylpyridinium chloride (Sigma) for one day and stored in 0.01 M cacodylic acid at 4°C. Prior to processing, samples were soaked overnight in 5% polyvinyl alcohol (PVA, Sigma-Aldrich), embedded in a
86 frozen block of PVA, and sectioned at -21 °C using a cryostat (Model OFT, Bright Instrument Company
Microtome). Sections were soaked in distilled water for one hour to remove residual PVA, stained with picrosirius red staining for one hour, and exposed to 0.1 N hydrochloric acid for two minutes (n=2). For collagen I and II immunostaining, sections were incubated with primary antibody overnight. Cell nuclei were stained with 4',6-diamidino-2-phenylindole (Sigma). A FITC-conjugated secondary antibody (1:200 dilution, Abcam) was used and samples were imaged under confocal microscopy (Olympus Fluoview
IX70) at excitation and emission wavelengths of 488 nm and 568 nm, respectively. Cover-slipped sections were imaged using a brightfield microscope (Zeiss, Axiovert 25).
Sample glycosaminogylcan content (GAG, n=5) was determined with a modified 1,9- dimethylmethylene blue (DMB) binding assay [161-163], with chondrotin-6-sulfate (Sigma) as the standard. The absorbance difference between 540 nm and 595 nm was used to improve the sensitivity in signal detection. In addition, histology was used to visualize GAG distribution. Sections were exposed to
3% acetic acid for three minutes, stained with alcian blue for 45 minutes, and rinsed twice with acid- alcohol (pH=1) for one minute (n=2) [159]. Cover-slipped sections were imaged using a brightfield microscope (Zeiss, Axiovert 25).
6.2.5 Mineralization
Alkaline phosphatase (ALP) activity (n=5) was measured using a colorimetric assay based on the hydrolysis of p-nitrophenyl phosphate (pNP-PO4) to p-nitrophenol (pNP) [164]. The samples were lysed in
0.1% Triton™ X solution, exposed to a freeze-thaw cycle, and crushed with a mortar. A 25 μL aliquot was added to pNP-PO4 solution (Sigma) and incubated for ten minutes at 37°C. Absorbance was measured at
405 nm using a microplate reader (Tecan). In addition, phosphate distribution (n=2) was evaluated as an indicator of overall mineral distribution. For phosphate staining, samples were stained with 5% silver nitrate and exposed to UV light (365 nm) for 25 minutes before a tap-water rinse. Cover-slipped sections were imaged using a brightfield microscope (Zeiss, Axiovert 25). Media calcium concentration (n=5) was quantified using the Arsenazo III dye (Pointe Scientific), The media was diluted with water in a 1:10 ratio and allowed to react with the dye for five minutes. Absorbance was measured at 620 nm using a microplate reader (Tecan). Media phosphate concentration (n=5) was quantified using the BioVision
Phosphate Assay Kit. The media was diluted with water in a 1:10 ratio and allowed to react with 30 µL of
87 dye for 30 minutes. Absorbance was measured at 650 nm using a microplate reader (Tecan).
6.2.6 Statistical Analysis
Results are presented in the form of mean ± standard deviation, with n equal to the number of samples per group. A two-way analysis of variance (ANOVA) was performed to determine the effects of ceramic dose and culturing time on cell response (proliferation, matrix deposition, ALP activity, and solution ion concentration). The Tukey-Kramer post-hoc test was used for all pair-wise comparisons and significance was attained at p<0.05. Statistical analyses were performed with JMP IN (4.0.4, SAS
Institute, Inc.).
6.3 Results
6.3.1 Scaffold Characterization
The structural and chemical properties of the scaffold were characterized post-fabrication (Fig. 6.2 and Fig. 6.3). The microfibers were randomly aligned when examined via SEM regardless of CDA dose
(Fig. 6.2). The presence of nodules along the fibers was noted for all ceramic-containing meshes. Fiber diameter was also measured for each composition, with no significant differences found between groups.
The elemental composition of the fibers was analyzed using energy-dispersive x-ray analysis (EDXA, Fig.
6.2). While carbon and oxygen were found in all groups, only the ceramic-containing groups exhibited peaks for phosphorus and calcium. Thermogravimetric analysis was used to verify mineral dose in the composite fibers. Residual weight of the 0% CDA microfibers was 0.36 ± 0.38% and 9.71 ± 0.46%,
14.55 ± 0.06% and 19.92 ± 0.44% for the 10%, 15% and 20% CDA microfibers, respectively.
The chemical composition of the fibers was also investigated via FTIR, which further confirmed the presence of ceramic within the fibers (Fig. 6.3). Typical infrared bands for the PCL and PLGA stretching modes were observed in the ceramic-free (0% HA) and ceramic-containing (10%, 15%, 20% CDA) groups. These bands include the ester carbonyl stretch from the PLGA (C=O) at 1746 cm-1 and from the
PCL at 1726 cm-1, the PLGA methyl group C–H stretch at 1452 cm-1, the PLGA C–O–C ether group at
1183 cm-1 and 1080 cm-1, and the PLGA C–O stretch at 1129 cm-1 [195]. The ceramic composite
3- microfibers showed characteristic peaks corresponding to the third vibrational PO4 band from 566-628 cm-1, as observed in the spectra reported in chapter 4 for the calcium deficient apatite. X-ray diffraction
88 was used to analyze the crystallinity of the scaffolds. One peak, at 2Θ =23.83 was present in the ceramic- free, blended polymer scaffold which is characteristic of PCL [196]. In addition to the characteristic PCL peak, crystal planes 002, 211, 300, and 202, characteristic of the CDA (as reported in chapter 4), were identified in the XRD spectra of all ceramic containing scaffolds. Composition-dependent degradation was assessed by measuring dry weight over the course of a three month in vitro culture period (Fig. 6.4). The ceramic-free fibers exhibited the greatest degradation, with 87.76 ± 0.89% remaining after 84 days.
Greater mass was retained with increasing ceramic content as 20% CDA fibers retained 91.08 ± 0.74% of their initial mass after 84 days.
6.3.2 Cell Viability and Number
The attachment, viability, and distribution of deep zone chondrocytes on the microfiber meshes were visualized using confocal microscopy (Fig. 6.5). The chondrocytes attached uniformly to the fiber surface on day 1 and assumed similar morphologies on all compositions. The cells maintained a rounded morphology throughout culture. Dead cells were rarely observed on any scaffold group.
Cell number was similar on all scaffold compositions on days 1, 7, 14, and 21 (Fig. 6.5). A significant increase in the number of cells was measured on all compositions between day 1 and day 7 with no significant increase over time noted thereafter for any scaffold type. A significantly greater number of cells were measured on the 20% CDA group on day 42 as compared to the ceramic-free control group.
6.3.3 Matrix Deposition
Matrix deposition in terms of collagen and GAG were measured for all compositions over the duration of the study (Fig. 6.6 and Fig. 6.7). Significantly greater collagen was measured in the 10% CDA group as compared to the ceramic-free control and the 15% CDA group on day 14. By day 21, similar collagen deposition was measured for all groups, with more collagen measured for each group as compared to the corresponding value from day 1. On day 42, more collagen was measured in the ceramic-free control as compared to the corresponding value from day 21. Additionally, the 20% CDA group exhibited higher collagen content than the 10% CDA group on day 42. Collagen distribution was visualized by picrosirius red staining, which showed positive staining for all groups. While collagen was localized to the edge of
89 the ceramic-free control scaffold, it was distributed throughout the depth of the ceramic-containing scaffolds.
In terms of GAG content, significantly more GAG was detected in the 10% CDA scaffolds on day 7 compared to day 1. By day 14, GAG content was similar between all scaffold compositions, with more
GAG measured at day 14 for all groups compared to corresponding day 1 values. On day 42, higher GAG content was measured in the 15% CDA and 20% CDA scaffolds as compared to corresponding day 21 values. Significantly more GAG was measured in the 15% CDA group as compared to the ceramic-free control and 10% CDA group on day 42; however, the most GAG was measured for the 20% CDA group on day 42. Media GAG content was also measured for all compositions over the duration of the study, and GAG was detected in the media of all scaffold groups during the culture period. The distribution of
GAG in the scaffolds was visualized by alcian blue staining. On day 42, the GAG was concentrated at the edge of the scaffold for all groups, with the greatest penetration observed for the 20% CDA scaffold.
6.3.4 Mineralization
Alkaline phosphatase activity was quantified to measure chondrocyte mineralization potential over time (Fig. 6.8). The highest ALP activity was found for the 10% group on day 1, with a significant decrease in ALP activity measured for all compositions between day 1 and day 7. No significant differences between groups were measured between groups after for any timepoint after day 1. Mineral distribution visualized via von Kossa staining showed similar dose-dependent mineral distribution on the composite microfibers and no mineral in the 0% HA group (Fig. 6.8).
Media calcium and phosphate concentration was also measured over the duration of the study to better understand mineralization mechanics on the microfibers (Fig. 6.8). Significantly more calcium was measured in the media of the ceramic-free control scaffolds on day 1 compared to the control media and the media from the ceramic-containing scaffold groups. After day 1, no difference was detected between groups in terms of calcium concentration. Measurement of media phosphate showed significantly lower media phosphate in the all scaffold groups on day 1 as compared to the control media.
90 6.4 Discussion
The overarching goal of thesis aim 2 is to engineer an integrative scaffold that can be placed between the healing cartilage and host subchondral bone to promote calcified cartilage formation and support osteointegration of the cartilage graft with the native bone tissue. The regeneration of the calcified cartilage will improve fixation of the grafts to the bone and protect the cartilage from osseous invasion.
This study specifically addresses the characterization and in vitro optimization of a microfiber-calcium deficient apatite composite to form the base of the cup-shaped integration scaffold. In this study, microfiber scaffolds with varying concentrations of calcium deficient apatite are fabricated using electrospinning, and deep zone chondrocyte response is investigated in terms of cell viability, cell number, mineralization potential, and matrix production over time. The cell response measured on the scaffolds collectively demonstrate that, while all scaffold groups support chondrocyte attachment and proliferation, increasing ceramic dose results in increased matrix deposition. Based on these findings, it is apparent that 20% CDA is the optimal dose of ceramic for the base of the integration scaffold.
Composite meshes that approximate the native osteochondral interface in thickness (~150 µm [39]) are reproducibly generated using electrospinning and can be handled with ease. By adjusting electrospinning parameters, scaffolds containing up to 20% calcium deficient apatite can be electrospun with similar fiber diameters. While the fiber diameter is controlled, it is noted that fiber nodularity appears to increase with mineral content, similar to other reported findings [154]. Therefore, although this study did not specifically investigate cellular response with respect to varying surface nodularity, chondrocyte response may be influenced collectively by the change in mineral content and scaffold texture.
In this study, ceramic incorporation increased glycosaminoglycan production at both 15% and 20% doses compared to the ceramic-free control and 10% groups and increased collagen production at the
20% dose. This finding is similar to results reported by Chuang et al. [150], in which deep zone chondrocytes cultured on poly(lactide-co-glycolide) microfiber scaffolds with 15% incorporated hydroxyapatite particles resulted in significantly more glycosaminoglycan and collagen deposition by day
21 compared to ceramic-free controls. In a study by Moffat et al. [154], matrix deposition of full thickness chondrocytes on aligned polymer nanofibers was not dependent on the dose of hydroxyapatite.
Specifically, no differences were found in glycosaminoglycan or collagen deposition on scaffolds
91 containing 10% or 15% hydroxyapatite particles. The difference observed in our study, in which 15% calcium deficient apatite resulted in more glycosaminoglycan deposition than 10%, may be due to the increased bioactivity of the calcium deficient apatite with respect to hydroxyapatite, as increased bioactivity has been shown to promote matrix deposition in other scaffolding systems [172]. Alternatively, the enhanced glycosaminoglycan production may be attributed to the zonal cell population used here, as chondrocytes isolated from different zones have varying biosynthetic activity [37].
In addition to promoting matrix production, the scaffold should ideally retain the deposited matrix to enable the formation of a dense, calcified matrix. In this study, measured collagen and glycosaminoglycan increased over time for all groups, and day 42 histological analysis revealed dense regions of matrix on the scaffold edge initially seeded with cells as well as penetration of the matrix into the scaffold for ceramic-containing groups. By day 42, collagen and glycosaminoglycans completely penetrated the 20% CDA scaffold. While glycosaminoglycans were detected in the media at each timepoint for all scaffold groups, the amount detected in the media was small in comparison to the glycosaminoglycans measured on the scaffolds (<10% for the 20% group on days 21 and 42), indicating that the majority of produced glycosaminoglycans were retained in the scaffold. This finding shows an improvement from the poly(lactide-co-glycolide)-hydroxyapatite scaffold used in studies by Moffat et al.
[154] in which GAG and collagen were found to decrease over time, but is similar to findings by Chuang et al. [143]. In this study and the study by Chuang et al., larger fibers and deep zone chondrocytes were used, as compared to the smaller fibers and full thickness chondrocytes used by Moffat et al. These differences may account for the increased matrix deposition and retention.
It is envisioned that this scaffold will serve three important functions in vivo. First, during early tissue formation, it is anticipated that the scaffold will serve as a temporary barrier against vascular invasion and ectopic mineralization, degrading over time as cell-mediated tissue formation occurs. Second, it will promote elaboration and support retention of matrix at the cartilage-bone interface, leading to the formation of a calcified cartilage layer. Finally, the ceramic within the fiber mesh is expected to promote integration with underlying bone. To this end, unpublished data from our laboratory demonstrates the osteointegrative potential of polymer-ceramic fiber meshes in vivo using a bone core model.
92 6.5 Conclusions
Collectively, the results of this study provide the optimal design specifications for the microfiber- ceramic composite that will compose the base of the cup integration system. A microfiber-based system incorporating 20% calcium deficient apatite results in enhanced cell growth and matrix elaboration by deep zone chondrocytes. Therefore, the base of the cup will consist of composite polymer-ceramic microfibers containing 20% calcium deficient apatite.
93 deep zone chondrocytes CDA +
Ceramic 10% 15% 20% in vitro -free CDA CDA CDA culture PLGA:PCL solution Figure 6.1 Study design. Scaffolds were generated by electrospinning PLGA:PCL polymer with varying amounts of calcium deficient apatite (CDA) nanoparticles and cultured in vitro.
94 Control 10% CDA 15% CDA 20% CDA
CDA CDA CDA 5 µm
1.2 µm 0.2 1.4 µm 0.1 1.1 µm 0.1 1.3 µm 0.03
P Ca 20% CDA 100 20% CDA 15% CDA 80 10% CDA 15% CDA Ceramic-free 60
10% CDA Counts 40
% Initial Weight Initial % 20 Ceramic-free 0 0 3 6 9 keV 100 300 500 700 Temperature (ºC)
Figure 6.2 Incorporation of calcium deficient apatite in microfiber scaffolds. Scanning electron microscopy (SEM) was used to evaluate fiber morphology (n=3, white arrows indicate CDA). Fiber diameter, shown below each image, was measured using ImageJ. Energy dispersive x-ray analysis (n=3, bottom left) confirms calcium and phosphorus incorporation in scaffolds with ceramic. Thermogravimetric analysis verifies that ceramic is incorporated at the target dose (n=3).
95
*
Ʌ
v (PO )-3 Ʌ
4 4 Ʌ
20% CDA Ʌ *
Ʌ
Ʌ
Ʌ
Ʌ
15% CDA *
Ʌ
Ʌ
* Ʌ Ʌ
10% CDA Counts Absorbance (a.u.) Absorbance Ceramic-free
2000 1600 1200 800 400 20 25 30 35 40 -1 wavenumber (cm ) 2θ Figure 6.3 Characterization of composite scaffolds composed of microfibers and calcium deficient apatite (CDA). Fourier transform infrared analysis shows similar spectra for all scaffolds, with additional phosphate bending peaks present in all ceramic-containing scaffolds. Crystalline peaks that correspond to polycaprolactone (*) were observed in all spectra, and peaks corresponding to calcium deficient apatite (v) were observed only in ceramic-containing scaffolds.
96 105 Ceramic-free 10% CDA 15% CDA 20% CDA
95
85
Scaffold Remaining (%) Remaining Scaffold 75 0 30 60 90 Time (days)
Figure 6.4 Scaffold degradation over time. Polymer scaffolds with and without calcium deficient apatite degrade over time in culture media. At day 84, the remaining weight is 87.8% ± 0.89, 89.6% ± 1.40, 88.37% ± 1.06, and 91.1% ±0.74 for the 0%, 10%, 15%, and 20% CDA scaffolds, respectively (n=6).
97 Control 10% CDA 15% CDA 20% CDA
Day 1
Day 42
200 µm
300 Ceramic-free Figure 6.5 Cell viability and number. 10% CDA Chondrocytes are viable on all scaffolds
15% CDA throughout the 42 day culture period. Cell number 3 - * 200 20% CDA increases for all groups by day 7, but by day 42 there are significantly more cells on the 20% ## calcium deficient apatite (CDA) scaffolds than the # # control scaffolds (n=5, *p<0.05 for difference 100 between groups on a given timepoint, #p<0.05 for
Cell No. x 10 x No. Cell difference from corresponding group at previous timepoint).
0 D1 D7 D14 D21 D42
98
Control 10% CDA 15% CDA 20% CDA
Total CollagenTotal
Collagen II Collagen Collagen I Collagen
100 µm
180 Ceramic-free * * 10% CDA * # # g) 15% CDA ^ μ 120 20% CDA
60 Collagen ( Collagen
0 D1 D7 D14 D21 D42
Figure 6.6 Collagen deposition. Picrosirius red staining (top) shows collagen deposition in all scaffold groups on day 42 (n=2, scale bar = 100 um). Collagen II (middle panel) and collagen I (bottom panel) show positive collagen II staining for all groups and negative staining for collagen I (n=2, scale bar = 100 um). Significant collagen deposition is measured for all groups by day 21, with the highest collagen content detected for the 20% group on day 42. (n=5, *p<0.05 for difference between groups on a given timepoint, #p<0.05 for difference from corresponding group at previous timepoint, ^p<0.05 for difference from corresponding group at day 1).
99 120 Ceramic-free * 120 Ceramic-free 10% CDA * 10% CDA 15% CDA # 90 20% CDA 90 15% CDA # 20% CDA
60 60 ^ #
30 30
GAG in media (µg) media in GAG GAG in scaffold (µg) scaffold in GAG * * * * * * * 0 0 D1 D7 D14 D21 D42 D1 D7 D14 D21 D42
Control 10% CDA 15% CDA 20% CDA
100 µm
Figure 6.7 Glycosaminoglycan (GAG) content. Glycosaminoglycan content on the scaffolds increased for all groups by day 14, with the highest GAG content measured for the 20% calcium deficient apatite (CDA) group on day 42 (n=5, *p<0.05 for difference between groups on a given timepoint, #p<0.05 for difference from corresponding group at previous timepoint, ^p<0.05 for difference from corresponding group at day 1). GAG was measured in the media for all groups at day 7; however by day 42, GAG is only detected in the media of the 20% CDA group (n=6, *p<0.05 for difference from control media). Alcian blue staining on day 42 (bottom) confirmed the quantitative findings (n=2, scale bar = 100 µm).
100 0.6 200 Control media Ceramic-free Ceramic-free 10% CDA 10% CDA
g/ml) 15% CDA 15% CDA µ 20% CDA 20% CDA 150 0.4 * * * * * 100 * 0.2 50
# ALP Activity (pmoles/cell/min) Activity ALP 0 Media Ion Concentration ( 0 D1 D7 D14 D21 D42 Phosphate Calcium
Control 10% CDA 15% CDA 20% CDA
100 µm
Figure 6.8 Mineralization. Black nodules are visible in von Kossa-stained sections for mineral-containing groups on day 42 (top left, n=2). Alkaline phosphatase (ALP) activity is highest for the 10% group on day 1 with a significant decrease measured for all groups between day 1 and day 7. Media calcium is significantly higher for the control group on day 1, with no difference between groups thereafter. Media phosphate is lower than the control media for the control and all ceramic groups on day 1, with no difference between scaffold groups thereafter (n=5, *p<0.05 for difference between groups on a given timepoint, #p<0.05 for difference from corresponding group at previous timepoint).
101 CHAPTER 7: IN VITRO EVALUATION OF INTERFACE FORMATION IN AN OSTEOCHONDRAL EXPLANT MODEL
102 7.1 Introduction
In the previous chapters, the phases of the scaffold that will form the cup walls (chapters 2 and 3) and the cup base (chapters 4, 5, and 6) were optimized independently to maximize cell migration and calcified cartilage formation by deep zone chondrocytes. This chapter evaluates the translational potential of the assembled cup integration system. The scaffold phases will be joined and tested in a full thickness defect model to investigate the effect of the scaffold system on cartilage repair.
7.1.1 Background and Significance
This study evaluates the translational potential of the cup integration system for use with current cartilage repair strategies. As the previous in vitro studies evaluated cell response on each scaffold phase, the ability of the multi-phased scaffold system to improve integration during cartilage repair must be assessed at the tissue level. The cup is designed to be versatile and compatible with a variety of available techniques, such as tissue transplantation, tissue engineered grafts, and cell implantation.
Although rabbit models are frequently used to study cartilage repair, it is particularly challenging to study the implementation of a complex scaffold system using this model because the cartilage depth is limited to approximately 400 µm [2]. Due to the shallow depth of the rabbit cartilage, defects made surgically often penetrate into the marrow space [188]. Since the base of the cup scaffold is designed to be placed directly at the cartilage-bone interface, it is important that the defect depth is precisely controlled.
Therefore, in this study, a full thickness defect model will be characterized and used to test the cup for use with clinically-relevant repair techniques in vitro.
Organ culture models, derived from either large animal [112;197-201] or human [202;203] tissue, can assess integration and the effect of a scaffold on the maintenance of the surrounding native tissue with greater control and lower cost than in vivo studies. Specifically, the use of bovine tissue in this study affords a cartilage layer which is sufficiently thick to study scaffold design and evaluate the performance of the cup with various repair strategies. Importantly, the ability to generate defects with precisely defined depth in organ culture models has been previously demonstrated [200].
103 7.1.2 Objectives
There are two objectives of this study. The first objective is to test if the cup scaffold is compatible with clinically relevant cartilage repair strategies. The second objective of this study is to evaluate the ability of the cup system to improve integration between a cartilage graft and the surrounding host tissue.
It is hypothesized that (1) the cup can be implemented with autograft procedures, cell implantation, and hydrogel grafts, (2) the cup will promote integration with surrounding cartilage and bone, and (3) the combined agarose-fiber-ceramic base will result in more matrix formation at the cartilage-bone interface than the fiber-ceramic base without agarose.
7.2 Materials and Methods
7.2.1 Explant Harvest and Culture
Fresh, immature metacarpophalangeal bovine joints (Green Village Packing Co.) were skinned, de- gloved, and soaked in soapy water followed by 70% ethanol for 20 minutes. The joint was opened in a sterile environment, and the articular cartilage rinsed with sterile phosphate buffered saline (PBS).
Osteochondral explants were extracted using a 1/2" Milwaukee Pistol Grip Electric Drill (Model 0300-20) with a 7/16” diamond tipped cylindrical drill bit (Model number 102080 with a 3/8 inch stem mount,
Starlite) while the joint surface was continuously irrigated with sterile PBS. Three or four plugs were harvested from each joint and stored in culture medium. The bone of each plug was trimmed (h = 5 mm) using a high profile histology blade (Shandon Blade, ThermoFisher). Defects were created using a biopsy punch (Sklar, 5 mm), and loose debris was removed using a high velocity water jet (Waterpik) (Fig. 7.1).
Full thickness cartilage explants (Ø = 8 mm) were excised from the flat surface of bovine metacarpophalangeal joints with a biopsy punch (Sklar), and the bottom was gently scraped to remove osseous tissue. Explants were cultured in 8 mL Dulbecco’s Modified Eagle’s Medium (DMEM) with 1% insulin, transferrin, selenous acid (ITS), 50 μg/mL proline, 0.1 μM dexamethasone, 0.9 mM sodium pyruvate, and 50 μg/mL ascorbic acid in a six-well plate.
7.2.2 Cells and Cell Culture
Primary articular chondrocytes were isolated from neonatal calf wrists obtained from a local abattoir
(Green Village Packing Co.) according to published protocols [79]. Briefly, cartilage tissue was extracted
104 from the surface of the metacarpophalangeal joint, minced, and incubated for 16 hours with 0.1% (w/v) collagenase (Worthington) in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS, Atlanta Biologicals), 2% antibiotics (10,000 U/mL penicillin, 10 mg/mL streptomycin), and 0.1% antifungal (250 µg/mL amphotericin B). The cell suspension was filtered to separate the cells from extracellular matrix debris before plating (30 μm, Spectrum). The isolated chondrocytes were maintained in high-density culture in fully-supplemented DMEM with 10% FBS, 1% non-essential amino acids, 1% antibiotics, and 0.1% antifungal for three days before seeding. All media supplements are purchased from Cellgro-Mediatech unless otherwise specified.
7.2.3 Scaffold Fabrication
Unaligned microfiber scaffolds were fabricated by electrospinning [20;149]. For ceramic-free polymer fiber fabrication, a 32% (w/v) polymer (5:1 PLGA (85:15, Lakeshore Biomaterials):PCL (Sigma-Aldrich,
Mw ≈ 70,000-90,000)) solution in 60/40 DCM/DMF was vortexed continuously for one hour. For fibers containing insulin-like growth factor-1 (IGF-1, Invitrogen), finely ground BSA (5% w/w, Sigma Aldrich) was added directly to a solution of 32% polymer (5:1 PLGA:PCL) in 60/40 DCM/DMF and vortexed continuously. After one hour, IGF-1, suspended in distilled water at a concentration of 5 mg/mL, was added to the polymer melt, and the solution was vortexed for an additional hour. To fabricate PLGA:PCL- calcium deficient apatite (CDA, Sigma-Aldrich) scaffolds, 20% w/w CDA nanoparticles were directly added to the PLGA:PCL polymer solution and vortexed for an additional hour. Each solution was loaded into a 5 mL syringe with a stainless steel blunt tip needle (26.5 gauge for PLGA:PCL blends, 23 gauge for polymer-ceramic blends, 18 gauge for IGF-1-polymer blends) that was 13 cm from the collecting target and electrospun at 8-10 kV using a custom electrospinning device. The polymer solution was deposited
(1 mL/hour) onto a stationary collecting target using a syringe pump (Harvard Apparatus).
To fabricate polymer cups, ceramic-free mesh (with or without IGF-1) was cut into squares (1.6 cm x
1.6 cm, 90-120µm thick), wetted with 39 µL sol-gel solution (12:1 tetamethylorthosilicate (Sigma):water), compressed in a custom mold, and allowed to dry for five hours in ambient conditions (Fig. 7.3). Next, the bottom portion of the cup was removed with a biopsy punch (Sklar, 4 mm), and 20% CDA-polymer composite scaffolds (Ø = 5 mm) were wetted with 10 µL sol-gel solution, compressed onto the walls of the
105 cup in the custom-designed mold to form the base, and allowed to dry for five hours. The cup walls were trimmed (h = 3 mm) and sterilized with ultraviolet exposure for 15 minutes.
7.2.4 Defect Repair and Culture
Defects were immediately repaired following generation (Fig. 7.4). For autograft control repair, the excised plug was press-fit back into the defect site. For autograft+cup repair, the autograft was removed, placed into a scaffold cup (with or without IGF-1), and the autograft+cup was press-fit back into the defect site. For hydrogel control repair, full thickness chondrocytes, which were isolated from immature bovine wrist joints, were mixed at a density of 10 million cells/mL in 2% low gelling agarose (Agarose Type VII,
Sigma) and pipetted into the defect site (~100 µL). The hydrogel was allowed to set in the defect for 20 minutes before the repaired explant was submersed in media. For hydrogel+cup repair, a scaffold cup was press-fit into the defect, and the chondrocyte-agarose was added directly into the cup in the defect until the hydrogel was flush with the top of the defect (~100 µL). The hydrogel was allowed to set in the defect for 20 minutes before the repaired explant was submersed in media. For the hydrogel+cup+agarose-ceramic base repair, acellular agarose with 1.5% CDA was pipetted into the bottom of the cup prior to the addition of the chondrocyte-agarose gel. For chondrocyte implantation repair, Bio-Gide® (Ø = 5 mm, Geistlich) was saturated with 25 µL chondrocyte solution (12 million full thickness chondrocytes/mL in sterile saline) and sutured over the defect with four or five stiches (6-0
Vicyrl, Ethicon). A needle was inserted under one edge of the flap, and the patch-defect border was sealed with 1-2 drops Evicel® Fibrin Sealant (Ethicon), which was allowed to set for 90 seconds.
Chondrocytes suspended in sterile saline (100 µL, 12 million cells/mL) were injected into the defect under the patch using a syringe attached to the needle. The needle was removed, an additional suture stich closed the hole, and 1-2 drops Evicel® Fibrin Sealant (Ethicon) were administered to the top of the flap to completely seal the patch-defect border. After 90 seconds, the repaired defect was submersed in media.
For the chondrocyte implantation+cup repair, a scaffold cup was inserted into the defect prior to suturing the patch, and the chondrocyte implantation procedure described above was employed.
All repaired defects were cultured in a six-well plate in 9 mL DMEM with 1% insulin, transferrin, selenous acid (ITS), 50 μg/mL proline, 0.1 μM dexamethasone, 0.9 mM sodium pyruvate, and 50 μg/mL ascorbic acid refreshed three times weekly.
106 7.2.5 Histology Analysis
Explants were rinsed in PBS and fixed in neutral buffered formalin with 1% cetylpyridinium chloride
(Sigma) for three days. Following fixation, the samples were decalcified with 10% ethylenediaminetetraacetic acid (EDTA, Sigma) in tris-buffer at a pH of 7.3 for five weeks, followed by dehydration with an ethanol series. The dehydrated samples were embedded in paraffin (Paraplast X-tra
Tissue Embedding Medium, Fisher Scientific), and 7 μm sections were obtained (Reichert-Jung RM 2030
Microtome, Leica) from the center of the scaffold. To visualize collagen (n=3), samples were deparaffinized and stained in 0.1% direct red for one hour before rinsing in 0.01 N hydrochloric acid and cover-slipping. Glycosaminoglycans and cell nuclei were visualized (n=2) via staining with safranin-O for
20 minutes, Weigert’s Hematoxylin for 7 minutes, and Fast Green for 12 minutes. Cover-slipped (Cytoseal
XYL) samples were imaged with a brightfield microscope (Olympus DP72).
7.2.6 Integration Strength
Full thickness cartilage explants were cultured in DMEM with 1% ITS, 50 μg/mL proline, 0.1 μM dexamethasone, 0.9 mM sodium pyruvate, and 50 μg/mL ascorbic acid in a six-well plate for three days to confirm sterility. Explants were bisected, using a histology blade and custom blade guide. The blade guide ensured that the cut was made in the center of the explant perpendicular to the surface. After the cut was made, the explants were immediately rejoined with suture (6-0 Vicryl, Ethicon) with or without a scaffold placed between the two halves (n=3/group). Re-joined cartilage explants were cultured for four weeks with media refreshed three times weekly.
Prior to testing, the suture was cut and removed from the cartilage explant, and the sample was loaded into a custom-designed shearing device (Fig. 7.9). The sample was positioned onto a platform that supported half of the cartilage explant, and a clamp was used to hold the explant in place. A platen in contact with the unsupported half was lowered at a rate of 0.5 mm/minute (DynaMight 8840, Instron) to shear the two halves apart. Force was recorded and converted to stress by dividing the surface area of the cartilage explant cross-section, which was measured for each explant following testing (n=3/group).
The peak stress was averaged across three samples per group (Fig. 7.9).
107 7.2.7 Statistical Analysis
Results are presented as mean ± standard deviation, with n equal to the number of samples per group. One-way ANOVA was used to determine the effects of scaffold presence on cartilage-cartilage integration strength. The Tukey-Kramer post-hoc test was used for all pair-wise comparisons, and significance was attained at p<0.05. Statistical analyses were performed with JMP IN (4.0.4, SAS
Institute, Inc.).
7.3 Results
7.3.1 Full Thickness Defect Model Characterization
On average, four osteochondral plugs were harvested per joint, and full thickness defects were reproducibly generated in each. Cells within both tissue regions (bone and cartilage) remained viable throughout the culture period, although a zone of death was observed at the cut edges of the cartilage for all samples by day 1 (Fig. 7.2). The zone of death produced by the drill appeared to penetrate further into the tissue compared to the zone created by the biopsy punch. Consistent with the live/dead cell imaging, the cell number measured in both the cartilage and bone regions remained unchanged over 14 days of culture. As expected, ALP activity was higher in the bone region than in the cartilage region, but no change was observed over time for either tissue type. Similarly, no changes were measured for the matrix content over time for either region.
7.3.2 Gross Morphology of Repaired Full Thickness Defects
After in vitro culture, top and side view images of all samples were recorded, and the gross morphology of the samples was assessed (Fig. 7.5). The cartilage and bone showed no macroscopic signs of deterioration, and the cartilage remained shiny and white on all samples. For the hydrogel groups, the hydrogel-based graft in the tissue remained translucent for all groups, although the groups repaired with a cup appeared more opaque than the cup-free control. For the autograft groups, the graft was proud to the defect in some samples, but there was no trend observed between groups for position of the graft. The samples repaired with allogenic cells remained intact, with the membrane covering the defect.
108 7.3.3 Histological Analysis
Histological analysis was performed to evaluate collagen and GAG content at the graft-cartilage interface and at the graft-bone interface. For the groups repaired with an autograft (Fig. 7.6), evaluation of the cartilage-cartilage interface using safranin-o and picrosirius red revealed no evidence of cartilage- cartilage connection; however, residual scaffold was visible in both groups repaired with a cup, and it appeared to be bonded to both the graft and host cartilage. No integration was observed at the graft-bone interface.
For the groups repaired with a hydrogel graft (Fig. 7.7), evaluation of the cartilage-cartilage interface using safranin-o and picrosirius red showed evidence of cartilage-cartilage connection, with the hydrogel congruent to the cartilage edge in portions of all repairs. Residual scaffold was visible in the groups repaired with a polymer cup, and cells were noted within the fibrous matrix in both groups. At the hydrogel-bone interface, the hydrogel was interdigitated with the bone in the control group and in portions of the groups repaired with the integration cup. Residual scaffold was present at this interface as well, and cell nuclei were identified within the scaffold matrix.
For the groups repaired with chondrocyte implantation (Fig. 7.8), graft-cartilage connection was observed in sections stained with safranin-o and picrosirius red. The graft tissue observed at the cartilage-cartilage edge in the control group stained weakly for GAG and collagen. Stronger positive staining for both GAG and collagen was observed in the group repaired with the integration cup at the graft-host cartilage interface. The graft-bone interface was similar for both control and scaffold groups, with new tissue observed above the bone region that stained weakly for GAG and collagen.
7.3.4 Integration Strength
The integration strength at the cartilage-cartilage interface was determined for cartilage repaired with and without a scaffold containing IGF after four weeks of culture (Fig.7.9). All samples exhibited integration that enabled handling of the cartilage after the suture was removed to load and test the interface strength. Significantly stronger integration was measured for the cartilage repaired with a scaffold compared to the cartilage repaired without a scaffold, with an average measured maximum stress of 39.9 ± 6.4 kPa and 15.3 ± 4.4 kPa for the scaffold and control, respectively.
109 7.4 Discussion
This study evaluates a degradable integration scaffold doped with insulin-like growth factor-1 and calcium phosphate nanoparticles for use with three clinically relevant cartilage repair techniques. Full thickness chondral defects in bovine osteochondral explants were repaired with an autograft, tissue- engineered graft, or chondrocyte implantation in the presence and absence of the integration scaffold and cultured for four weeks in vitro. The ability of the scaffold to improve the strength of cartilage integration was measured by mechanical testing after four weeks of in vitro culture. The results of this study demonstrate that the integration cup can be implemented with all three repair techniques with minimal change to the standard repair process. Furthermore, the scaffold can enhance the integration strength between two full thickness cartilage explants over four weeks of in vitro culture.
The defect model used in this study disrupts the cartilage, with limited damage to the surrounding tissue. Each sample was repaired immediately following defect creation and cultured statically for four weeks. All grafts remained seated in the defect throughout the culture period, with no macroscopic degradation or growth noted on any of the explants after four weeks. Histologically, changes in the matrix of the native tissue were minimal; several samples exhibited reduced glycosaminoglycan staining in regions of cartilage and increased collagen staining near the outer border of the host cartilage tissue.
In the autograft repair samples, minimal integration was noted. In the control repair, the edge of both the graft and host tissue remained smooth and appeared unchanged. While the histological processing may have caused the samples to separate, no evidence of new tissue formation was observed at the interface. In the cup-repaired groups, the tissue looked similar to the control; however, residual scaffold material was observed in the gap between the graft and host cartilage. While the majority of scaffold was destroyed during processing, the remaining scaffold was bonded to the edge, indicating that the scaffold may have integrated with the cartilage. There was no evidence of integration at the cartilage-bone interface in the autograft samples, suggesting that blood and marrow that are usually present in the bone may be needed to provide a source of stem cells and growth factors in vivo [204] for cartilage-bone integration; however, in this study the bone was cleaned of marrow and blood.
In the groups repaired with a hydrogel graft, evidence of integration at the cartilage-cartilage junction was observed in all three groups. In the control repair, the hydrogel graft was congruent with the host
110 cartilage, and positive collagen staining was observed at the interface. In the groups repaired with a cup, residual fiber was observed at the interface, with nuclei present in the scaffold, indicating that cells had migrated into the cup walls during culture. Other investigators have shown that hydrogel constructs more readily integrate with host cartilage than autograft cartilage tissue [201], which was supported by our histological findings in this study. In addition, the hydrogel graft remained connected to the bone, with positive staining for glycosaminoglycans and cells evident at the interface. In the group with the hydrogel- ceramic composite in the base of the repair, cell nuclei were observed at the interface, suggesting that cells may have migrated into the thin acellular hydrogel-fiber-ceramic layer.
In the groups repaired using chondrocyte implantation, new tissue formation was observed at the host-cartilage interface. In the presence of the integration scaffold, the tissue stained more positively for glycosaminoglycan content and collagen than the control repair. New tissue was also detected at the cartilage-bone interface in both groups; however, it did not stain strongly for glycosaminoglycans or collagen. The repair tissue was fibrous and did not resemble the structural organization of native cartilage, consistent with previous reports [200;203].
In addition to histology, the integration of two pieces of full thickness cartilage was measured using a shear test when repaired with and without an IGF scaffold. This experimental design represents the autograft-repair technique, in which a healthy cartilage graft is placed next to the healthy host cartilage that shoulders a defect. The model employed here was used to ensure congruency between the two cartilage pieces, which is not always achieved in push-out tests that measure the integration of an inner disk and an annular ring. After four weeks of culture, the cartilage pieces integrated with each other in the control case; however, higher integration strength was achieved when a scaffold doped with insulin-like growth factor-1 was placed between the cartilage during repair. While the histological comparison of the cartilage-cartilage interface showed little difference in the autograft repair with and without the cup scaffold, this quantitative measurement suggests that the scaffold does promote integrative repair. Due to the unique testing procedure employed here and the use of full thickness tissue, it is difficult to compare our mechanical evaluation to other published studies; however, the integration strengths measured were the same order of magnitude to previously reported values (56.7 kPa and 28 kPa, respectively, after four weeks of in vitro culture with serum) determined using push-out tests to measure the integration of full
111 thickness bovine cartilage with a tissue engineered construct [201] and an autograft [205]. Unlike histology, the mechanical testing required only minor manipulation of the sample prior to analysis. This is important because it is likely that the polymer scaffold was destroyed during histological processing, limiting the observable differences between the control and scaffold-repaired group in the stained sections. In the histology images, remnants of the scaffold were detected on the edge of both the graft and host cartilage; this observation, combined with the higher integration strength that was measured in the scaffold group, suggests that the scaffold may improve integration by bonding to both graft and host cartilage. Importantly, the bonding measured here does not restore native strength, which has been measured to be 8.8 MPa using a push-out test [198].
The explants used in this study were cultured in serum-free, chemically defined media that was previously optimized for osteochondral explant culture [206]. The use of this media facilitates study of the scaffold in isolation from confounding cellular effects caused by serum. However, because fetal bovine serum has been shown to enhance cartilage-cartilage integration [207], the use of serum-free media limits the comparisons that may be made to other studies that used media enriched with serum
[197;198;200;201].
Although this study provides valuable information regarding the interaction of the scaffold on native osteochondral tissue and integration of cartilage grafts, it is limited by several inherent shortcomings of the model. Primarily, because the explant tissue was isolated from the marrow, synovium, and mechanical loading that exist in the native joint, the in vitro explant model eliminates many of the factors that may provide additional healing or immunologic response in vivo. For example, after an intra-articular defect is formed, migrating cells from the synovial tissues may enhance scaffold-guided repair [208].
Furthermore, since mechanical loading and growth factors have a synergistic effect on cartilage formation
[166], the growth-factor releasing scaffold may result in more cartilage formation in a loaded environment.
The lack of marrow in the bone compartment reduces exposure to cells, such as mesenchymal stem cells, which may populate the base of the cup and contribute to enhanced calcified cartilage formation.
Finally, in vivo, the healing cartilage would be perfused with a multitude of growth factors and immune cells that are present in the blood. Implanting the full thickness defect model in vivo in a subcutaneous
112 pouch, will more closely mimic this aspect of the complex native healing environment, and will be discussed in chapter 8.
7.5 Conclusions
Collectively, the results of this study indicate the cup integration scaffold can be utilized with a variety of cartilage repair techniques, including autografting, tissue engineering, and cell-based repair approaches. This study demonstrates that placement of the scaffold in a cartilage defect causes cells from the surrounding tissues to migrate into the scaffold, resulting in new tissue formation around the scaffold over time. Moreover, the scaffold increases the integration strength in a cartilage explant repair model.
113 osteochondral full thickness bone is trimmed bone is cleaned defect is created plug is harvested defect model
Figure 7.1 Osteochondral tissue harvest and full thickness defect generation. Osteochondral tissue plugs were harvested using a diamond-tipped drill bit. The bone was trimmed using a histology blade and remaining debris was removed with a high velocity jet of phosphate buffered saline. A biopsy punch was used to create a defect in the center of the plug.
114 Overview Side View Top View Cartilage Bone Collagen GAG Cartilage Viability Viability
Defect
Day 1 Day 1 cm 1 cm Bone 1 cm 250 μm 250 μm
180 20 Day 1 Day 1 Day 7 Day 14 Day 7 15 Day 14 *
120
/ mg mg / DW
3 3 - 10 *
60
5 14 Day Cell Cell 10 x No.
ALP Activity Activity ALP (pmoles/cell/min) 0 0 Cartilage Bone Cartilage Bone
Figure 7.2 Characterization of the full thickness defect model. Full thickness defect models were generated from bovine cartilage. Cell number and alkaline phosphatase (ALP) activity was maintained in culture over time for the bone and cartilage tissue, with higher ALP activity in the bone compared to the cartilage (n=5, * p<0.05 for difference between cartilage and bone at the same timepoint). Cells were viable in the bone and cartilage tissues overtime; however, a zone of death was observed on the cut edges of the cartilage. Collagen and GAG content was similar on day 1 and day 14 (picrosirus red and alcian blue, respectively).
115 top of mold is lowered
2 mm
top view fiber mesh
bottom of mold
side view cup scaffold in defect
Figure 7.3 Cup fabrication. To fabricate the cup scaffolds, fiber mesh is cut into squares, wetted with sol gel solution, and placed onto the bottom of a custom mold with protruding pegs. The top of the mold, with slots for the pegs, is lowered onto the base and a weight is applied. After drying, cup scaffolds are removed from the pegs with tweezers. The cup is designed to fit tightly in a 5 mm defect (right).
116 Hydrogel Cartilage Graft Polymer Nanofiber Cup CaP nanoparticles IGF-1 Chondrocyte
Hydrogel+IGF(IGF-1+cup) + Cell Suspension+(IFG-1+cup)+ IGF Autograft ChHydrogel+hydrogel Cell Suspension Cup+Autograft (IFGAutograft-1+cup)++ (IFGHydrogel+-1+cup)+ Cup+(Ag-MF Ag base)-CDA + (Ch+hydrogelCup ) + cells (cartilageIGF Cup graft) (Ch+hydrogelIGF Cup ) (Ch+hydrogelBase )
Control Groups Experimental Groups
Full thickness defect model
In Vitro Culture In vitro culture
Figure 7.4 Study design. Full thickness defects were repaired using three clinically relevant repair methods with varied scaffolds and cultured in vitro for four weeks.
117 Autograft + Autograft + Autograft Control Cup IGF Cup
Hydrogel + Hydrogel + IGF Cup Hydrogel IGF Cup + Ag-CDA Base
Figure 7.5 Images of repaired explants after in vitro culture. Repaired explants were fixed after four weeks of culture. The cartilage remained white and shiny, and no differences were observed in the bone region. In all groups, the defect was filled with repair tissue. Cells Cells + IGF Cup
118
cartilage-cartilage cartilage-bone GAG GAG
Autograft
Collagen GAG GAG
Autograft +
Control Cup
Collagen GAG GAG
Autograft +
IGF Cup Collagen
Figure 7.6 Autograft-based repair. For each group, the top row shows safranin-O staining, and the bottom row shows picrosirius red staining (n=2, 20x, bar = 100 µm).
119
cartilage-cartilage cartilage-bone GAG GAG
Hydrogel
Collagen GAG GAG
Hydrogel +
IGF Cup
Collagen GAG GAG
Hydrogel + IGF Cup + Ag-CDA
Base Collagen
Figure 7.7 Hydrogel-based repair. For each group, the top row shows safranin-O staining, and the bottom row shows picrosirius red staining (n=2, 20x, bar = 100 µm).
120
cartilage-cartilage cartilage-bone GAG GAG
Cells
Collagen Collagen GAG GAG
Cells + IGF Cup Collagen
Figure 7.8 Cell implantation-based repair. For each group, the top row shows safranin-O staining, and the bottom row shows picrosirius red staining for each group (n=2, 20x, bar = 100 µm).
121 80 Control IGF-1 Scaffold platen platform 60 * Control 40
20 (kPa) Stress Max
IGF-1 Scaffold 0 Week 4 Figure 7.9 Integration strength. A custom device (left) was used to test the integration strength of the repaired cartilage (middle). Full thickness cartilage explants (d = 8 mm) were repaired using suture with either no scaffold (n=3, left) or an IGF-1 microfiber scaffold (100 ng/mg), and the integration strength (n=3) of the two halves was measured after four weeks in vitro culture.
122 CHAPTER 8: IN VIVO EVALUATION OF INTERFACE FORMATION IN AN OSTEOCHONDRAL EXPLANT MODEL
123 8.1 Introduction
In chapter 7, the translational potential of the fully assembled cup integration system was assessed in vitro. The scaffold phases were joined and tested in a full thickness defect model to investigate the effect of the scaffold system on integration of cartilage grafts with the surrounding host tissue. The scaffold was compatible with three clinically relevant repair strategies and improved the integration between full thickness cartilage explants. In this chapter, histological analysis of the scaffold in the full thickness explant model will be extended to in vivo subcutaneous culture.
8.1.1 Background and Significance
Building on the in vitro testing of each scaffold phase and the evaluation of the assembled cup in a full thickness defect model in vitro, this study evaluates the translational potential of the cup integration system for use with cartilage repair strategies in vivo. Subcutaneous culture of osteochondral organ models, which provides a nutrient-rich environment, has been previously used to assess cartilage- cartilage integration with explant tissue models [106;198;203;209;210].
8.1.2 Objectives
The objective of this study is to evaluate the clinical translational potential of the total integration scaffold in vivo. It is hypothesized that the cup will improve integration with surrounding cartilage and bone and that the combined agarose-fiber-ceramic base will result in calcified cartilage formation that is more organized than the fiber-ceramic base without agarose.
8.2 Materials and Methods
8.2.1 Explant Harvest and Culture
Fresh, immature metacarpophalangeal bovine joints (Green Village Packing Co.) were skinned, de- gloved, and soaked in soapy water followed by 70% ethanol for 20 minutes. The joint was opened in a sterile environment, and the articular cartilage rinsed with sterile phosphate buffered saline (PBS). Full thickness cartilage explants (Ø = 8 mm) were excised from the flat surface of bovine metacarpophalangeal joints with a biopsy punch (Sklar), and the bottom was gently scraped to remove osseous tissue. Osteochondral explants were extracted using a 1/2" Milwaukee Pistol Grip Electric Drill
124 (model 0300-20) with a 7/16” diamond tipped cylindrical drill bit (Model number 102080 with a 3/8” stem mount, Starlite) while the joint surface was continuously irrigated with sterile PBS solution. Three or four plugs were harvested from each joint and stored in culture medium. The bone of each plug was trimmed
(h = 5 mm) using a high profile histology blade (Shandon Blade, ThermoFisher). Defects were created using a biopsy punch (Sklar, 5 mm), and loose debris was removed using a high velocity water jet
(Waterpik). Explants were cultured in 8 mL Dulbecco’s Modified Eagle’s Medium (DMEM) with 1% insulin, transferrin, selenous acid (ITS), 50 μg/mL proline, 0.1 μM dexamethasone, 0.9 mM sodium pyruvate, and
50 μg/mL ascorbic acid in a six-well plate.
8.2.2 Cells and Cell Culture
Primary articular chondrocytes were isolated from neonatal calf wrists obtained from a local abattoir
(Green Village Packing Co.) according to published protocols [79]. Briefly, cartilage tissue was extracted from the surface of the metacarpophalangeal joint, minced, and incubated for 16 hours with 0.1 w/v% collagenase (Worthington) in DMEM, supplemented with 10% fetal bovine serum (FBS, Atlanta
Biologicals), 2% antibiotics (10,000 U/mL penicillin, 10 mg/mL streptomycin), and 0.2% antifungal
(250 µg/mL amphotericin B). The cell suspension was filtered to separate the cells from extracellular matrix debris before plating (30 μm, Spectrum). The isolated chondrocytes were maintained in high- density culture in fully-supplemented DMEM with 10% FBS, 1% non-essential amino acids, 1% antibiotics, and 0.1% antifungal for three days before seeding. All media supplements are purchased from
Cellgro-Mediatech unless otherwise specified.
8.2.3 Scaffold Fabrication
Unaligned microfiber scaffolds were fabricated by electrospinning [20;149]. For ceramic-free polymer fiber fabrication, a 32% (w/v) polymer (5:1 PLGA (85:15, Lakeshore Biomaterials):PCL (Sigma-Aldrich,
Mw ≈ 70,000-90,000) solution in 60/40 DCM/DMF was vortexed continuously for one hour. For fibers containing insulin-like growth factor-1 (IGF-1, Invitrogen), finely ground bovine serum albumin (5% w/w,
Sigma Aldrich) was added directly to a solution of 32% polymer (5:1 PLGA:PCL) in 60/40 DCM/DMF and vortexed continuously. After one hour, IGF-1, suspended in distilled water at a concentration of 5 mg/mL, was added to the polymer melt, and the solution was vortexed for an additional hour. To fabricate
125 PLGA:PCL-calcium deficient apatite (CDA, Sigma-Aldrich) scaffolds, 20% (w/w) CDA nanoparticles were directly added to the PLGA:PCL polymer solution and vortexed for an additional hour. Each solution was loaded into a 5 mL syringe with a stainless steel blunt tip needle (26.5 gauge for PLGA:PCL blends, 23 gauge for polymer-ceramic blends, 18 gauge for IGF-1-polymer blends), placed 13 cm from the collecting target, and electrospun at 8-10 kV using a custom electrospinning device. The polymer solution was deposited (1 mL/hour) onto a stationary collecting target using a syringe pump (Harvard Apparatus).
To fabricate polymer cups, ceramic-free mesh (with or without IGF-1) was cut into squares (1.6 cm x
1.6 cm, 90-120µm thick), wetted with 39 µL sol-gel solution (12:1 tetamethylorthosilicate (Sigma):water), compressed in a custom mold, and allowed to dry for five hours in ambient conditions. Next, the bottom portion of the cup was removed with a 4 mm biopsy punch (Sklar), and 20% CDA-polymer composite scaffolds (Ø = 5 mm, t = 90-120µm) were wetted with 10 µL sol-gel solution, compressed onto the walls of the cup in the custom-designed mold to form the base, and allowed to dry for five hours. The cup walls were then trimmed (h = 3 mm) and sterilized with ultraviolet exposure for 15 minutes.
8.2.4 Sample Preparation
Defects were generated in osteochondral plugs with a biopsy punch (Sklar, 5 mm) and immediately repaired. For autograft control repair, the removed plug was press-fit back into the defect site. For autograft+cup repair, the autograft was removed, placed into a scaffold cup (with or without IGF-1), and the autograft+cup was press-fit back into the defect site. For chondrocyte implantation control repair, Bio-
Gide® (Ø = 5 mm, Geistlich) was saturated with 25 µL chondrocyte solution (12 million full thickness chondrocytes/mL in sterile saline) and sutured over the defect with four or five stiches (6-0 Vicyrl,
Ethicon). A needle was inserted under one edge of the flap, and the patch-defect border was sealed using 1-2 drops Evicel® Fibrin Sealant (Ethicon), which was allowed to set for 90 seconds. Chondrocytes suspended in sterile saline (100 µL, 12 million cells/mL) were injected into the defect under the patch using a syringe attached to the needle. The needle was removed, an additional suture stitch was used to close the hole, and 1-2 drops Evicel® Fibrin Sealant (Ethicon) was administered to the top of the flap to completely seal the patch-defect border. After 90 seconds, the repaired defect was submersed in media.
For the chondrocyte implantation+IGF cup repair, a scaffold cup was inserted into the defect prior to suturing the patch, and the chondrocyte implantation procedure described above was employed. For
126 hydrogel control repair, full thickness chondrocytes, isolated from immature bovine wrist joints, were mixed at a density of 10 million cells/mL in 2% low gelling agarose (Agarose Type VII, Sigma) and pipetted into the defect site (~100 µL). The hydrogel was allowed to set in the defect for 20 minutes before the repaired explant was submersed in medium. For hydrogel+IGF cup repair, a scaffold cup was press-fit into the defect, and the chondrocyte-agarose solution was added directly into the cup in the defect until the hydrogel was flush with the surface (~100 µL). The hydrogel was allowed to set in the defect for 20 minutes before the repaired explant was submersed in medium. For the hydrogel+IGF- cup+hydrogel-ceramic base repair, acellular agarose with 1.5% CDA was pipetted into the bottom of the cup prior to the addition of the chondrocyte-agarose gel.
8.2.5 Subcutaneous Implantation and Culture
All surgical procedures were performed in accordance with a protocol approved by the Institutional
Animal Care and Use Committee (IACUC) at the Columbia University Medical Center. Athymic rats (NIH- rnu, 175-200 g) were used for this study. Animals were anesthetized in an inhalation chamber with 1–5% isoflurane in high flow oxygen and maintained under a surgical plane of anesthesia using isoflurane (1–
2%) administered through an oxygen mask. The surgical area was shaved, draped, and prepared using an alternating isopropanol/betadine scrubbing technique. Sustained release buprenorphine (1.2 mg/kg
SQ), carprofen (5 mg/kg SQ q24 for 72 hours), and marcaine (2 mg/kg at the incision site) were administered to alleviate pain due to the operation. For prophylaxis against infection, an injection of baytril
(5 mg/kg SQ) was administered immediately prior to surgery. Using aseptic technique, four individual subcutaneous pouches were surgically created in the dorsum of each rat by incisions approximately 1.5 cm in length made with a scalpel (#15 blade, Feather). One repaired full thickness defect model (Fig. 8.1 and Fig. 8.2) was placed in each pouch, and the incision was closed with wound clips (9 mm stainless steel autoclip, BD Diagnostic Systems). After four weeks, animals were sacrificed by carbon dioxide inhalation, and the scaffolds were excised and prepared for analysis.
8.2.6 Histological Analysis
Explants were rinsed in PBS and fixed in neutral buffered formalin with 1% cetylpyridinium chloride
(Sigma) for three days. Following fixation, half the samples were decalcified with 10%
127 ethylenediaminetetraacetic acid (EDTA, Sigma) in tris-buffer at a pH of 7.3 for five weeks, followed by dehydration with an ethanol series. The dehydrated samples were embedded in paraffin (Paraplast X-tra
Tissue Embedding Medium, Fisher Scientific), and 7 μm sections were obtained (Reichert-Jung RM 2030
Microtome, Leica) from the center of the scaffold. Calcified samples were fixed in 80% ethanol for three days and embedded in poly-methylmethacrylate (PMMA, Sigma). The samples were sectioned (7 μm) with a Leica sliding microtome (SM2500S, Leica Microsystems Inc.) using a tungsten carbide blade
(Delaware Diamond Knives Inc.).
Sections were stained for cell nuclei, collagen, GAG, and mineral content (n=3. To visualize cell nuclei, samples were deparaffinized and stained with Weigert’s Hematoxylin for five minutes followed by eosin for two minutes. To visualize collagen content, deparaffinized samples were stained in 0.1% direct red for one hour before rinsing in 0.01 N hydrochloric acid and cover-slipping. Collagen alignment was evaluated by imaging picrosirius red-staining samples with polarized light. To distinguish collagen II, immunohistochemical staining was performed. Antigen retrieval was achieved via incubation with testicular hyaluronidase at 37ºC, and samples were subsequently incubated with collagen II antibody
(Abcam, 1:100 dilution) overnight at room temperature. To visualize the bound antibodies, the sections were incubated with 3,3'-diaminobenzidine (DAB peroxidase kit, Vector Laboratories) and counterstained with Fast Green for three minutes.
Glycosaminoglycans were visualized via staining with safranin-O for twenty minutes, Weigert’s
Hematoxylin for seven minutes and Fast Green for 12 minutes. To visualize mineral content, calcified sections were stained with von Kossa (5% silver nitrate) and exposed to ultraviolet light (365 nm) for 15 minutes. Following radiation, the plastic was removed by soaking in a 1:1 xylenes:chloroform mixture for
45 minutes and the sample was stained with hematoxylin for seven minutes followed by safranin-o for 20 minutes. Cover-slipped (Cytoseal XYL) samples were imaged with a brightfield microscope (Olympus
DP72). The images were stitched together in Photoshop (Adobe) using the auto-align and auto-blend tools. After stitching, the area surrounding the stained section was digitally cleaned.
8.2.7 Statistical Analysis
Results are presented as mean ± standard deviation, with n equal to the number of samples per group. One-way ANOVA was used to determine the effects of scaffold presence on the strength of
128 cartilage integration. The Tukey-Kramer post-hoc test was used for all pair-wise comparisons, and significance was attained at p<0.05. Statistical analyses were performed with JMP IN (4.0.4, SAS
Institute, Inc.).
8.3 Results
8.3.1 Animal Surgery
Four implants were placed into the back of each rat (Fig. 8.1 and Fig. 8.2). All animals survived the surgery and gained weight over time (data not shown). No infections were noted, and no rats required additional antibiotics or analgesics. Immediately following surgery, nine rats reopened their incisions by removing the wound clips. The incisions were cleaned and closed within 12 hours using a buried stitch
(Vicryl 4-0, Ethicon) under the supervision of a veterinarian.
8.3.2 Gross Morphology of Implants
Each repaired full thickness defect was removed from the subcutaneous pouch after four weeks and imaged prior to fixation (Fig. 8.3). All grafts remained well-seated in the defect, and no gross deterioration was noted for the cartilage or bone regions. Furthermore, no nodular growth on either tissue type was observed; all samples were similar in appearance to before implantation. All explants were surrounded by a thin sheath of fibrous and vascularized tissue, which bound the sample to the skin of the rat. While removal of the sheath was possible in some cases, the tissue surrounding the cartilage region was not removed to avoid manipulation that could interfere with the repair tissue.
8.3.3 Autograft Control
The autograft repair without a scaffold served as a control for the autograft groups that were augmented with the control (IGF-1-free) and optimized (with IGF-1) cups. The cartilage tissue maintained a glossy white appearance after implantation (Fig. 8.3). Histological analysis of the autograft control repair demonstrated that the graft maintained a matrix rich in GAGs and collagen, resembling healthy cartilage tissue (Fig. 8.4 and Fig. 8.5). The repair and native cartilage tissue were connected at the base of the graft, but the surface and middle regions of the graft were not congruent with the host tissue. Collagen alignment at the graft-host interface was observed in polarized images. Cells were not observed in the
129 cartilage-cartilage gap, and, despite the host and graft tissue remaining close together, there was little evidence of new cartilaginous tissue. Staining with von Kossa (Fig. 8.13) revealed mineralization in the cartilage compartment at the host-graft cartilage junction, although the presence of mineral in the cartilage tissue was not uniform across samples.
8.3.4 Autograft with Control Cup
The cartilage tissue maintained a glossy white appearance after implantation of the autograft group that was augmented with a cup without IGF-1 (Fig. 8.3). Histological analysis of the autograft repair with the IGF-1-free cup demonstrated that the graft maintained a matrix rich in GAGs and collagen, with noticeable depletion of the GAGs at the surface of the host tissue and loss of collagen evident in the center of the host and grafted cartilage (Fig. 8.6). The cartilage-cartilage interface was connected at the bottom of the defect, with separation between the graft and host tissue observed in the surface region.
Evidence of newly formed, cellular cartilage tissue at the host-graft interface that stains positively for both
GAG and collagen was observed. In addition, collagen alignment was increased at the interface, with the interface region appearing bright yellow when viewed using polarized light. The region just above the cartilage-bone interface stained strongly for collagen, which was aligned. Just below this region, where the base of the scaffold was placed, there was a region void of GAGs and collagen. Staining with von
Kossa (Fig. 8.13) revealed mineralization in the cartilage compartment at the graft-bone junction, although the presence of mineral in the cartilage tissue was not uniform across samples.
8.3.5 Autograft with IGF Cup
Similar to the other autograft groups, the cartilage tissue maintained a glossy white appearance after implantation (Fig. 8.3). Histological analysis of the autograft repair demonstrated that the graft maintained a matrix rich in GAGs and collagen that resembles healthy cartilage tissue, although loss of
GAGs was noted in the surface of the host tissue on one side of the cross section (Fig. 8.7). At the cartilage-cartilage interface, close contact was observed at the base of the grafts, with a gap observed in the region between the host and graft tissue in the middle and surface zone where the scaffold was placed. There was evidence of the scaffold in the gap, with cells budding from the graft tissue in the direction of the gap. At the graft-host junction, aligned collagen was observed using polarized light. The
130 cartilage-bone interface was uniform, with a thin region void of GAG and collagen where the base of the cup was placed, and a region directly above this which stained darkly for aligned collagen and GAG.
Similar to the control cup, staining with von Kossa (Fig. 8.13) revealed mineralization in the cartilage compartment at the host-graft cartilage junction, although the presence of mineral in the cartilage tissue varied between samples.
8.3.6 Chondrocyte Implantation Control
For the chondrocyte implantation control group, the host cartilage maintained a glossy white appearance, and the defect was filled with translucent tissue (Fig. 8.3). Histological analysis revealed that the defect was filled with a collagen-rich, GAG-poor fibrous tissue (Fig. 8.8). The host tissue consisted of a matrix rich in GAG and collagen, although noticeable GAG depletion was observed in the surface region of the host cartilage. Inconsistent formation of new cartilage tissue at the cartilage-cartilage interface was also observed. While several cartilage-cartilage interfaces in the samples from this group were populated by a GAG-rich tissue deposit, others were void of cartilaginous tissue. Rounded cells were not observed in the middle of the defect, although chondrocyte-like cells were found in the newly formed tissue at the cartilage-cartilage interface. Positive GAG staining was also observed in the bone compartment throughout the depth of the explant. Evidence of ectopic mineralization was not detected
(Fig. 8.13).
8.3.7 Chondrocyte Implantation with IGF Cup
For the cell-based repair with the IGF cup, the host cartilage maintained a glossy white appearance, and the defect was filled with a translucent tissue (Fig. 8.3). Histological analysis revealed a fibrous, collagen-rich, GAG-deficient tissue filling the defect (Fig. 8.9). The host cartilage tissue exhibited GAG loss from the surface region, although the collagen matrix was maintained. New cartilaginous tissue was observed at the cartilage-cartilage interfaces, although this tissue was observed only near the deep and middle cartilage zones. The tissue at the interface was cellular and stained positively for both GAG and collagen. Similar to the control chondrocyte implantation group, positive GAG staining was observed in the bone compartment throughout the depth of the explant. Positive mineral staining at the defect-bone interface was observed in the location where the mineralized cup base was placed (Fig. 8.13).
131 8.3.8 Hydrogel Graft Control
The hydrogel graft served as a control for the hydrogel groups that were augmented with the IGF cup and the IGF cup with the agarose-calcium deficient apatite base. For this group, the host cartilage maintained a glossy white appearance, and the defect was filled with translucent tissue (Fig. 8.3).
Histological analysis demonstrated that the host tissue maintained a matrix rich in GAG and collagen (Fig.
8.10). Collagen and GAG were also observed in the hydrogel graft, with similar staining throughout, except for more positive GAG staining near the cartilage-bone interface. Inconsistent integration was noted at the cartilage-cartilage interface between samples, witch fibrous tissue observed between the hydrogel and native tissue. The cartilage-bone interface was interdigitated, and collagen and GAGs were observed directly above the bone tissue. There was no evidence of ectopic mineralization (Fig. 8.13).
8.3.9 Hydrogel Graft with IGF Cup
The host cartilage maintained a glossy white appearance, and the defect was filled with translucent tissue for the hydrogel group that was augmented with an optimized cup scaffold (Fig. 8.3). The cup had not degraded after four weeks and was visible in the cross-section of the defect prior to histological processing. Histological analysis demonstrated that the graft consisted of a matrix with GAGs and collagen, although there was noticeable depletion of the GAGs at the surface of the host tissue (Fig.
8.11). New cellular tissue that stained positively for GAGs and collagen was observed at the cartilage- cartilage interface throughout the depth of the cartilage tissue. The collagen in the new tissue was aligned. The cells in the new tissue that were closer to the surface of the tissue were small and similar in size to the cells in the adjacent tissue; however, cells near the deep zone were larger, matching the cells in the adjacent deep zone tissue. The newly formed tissue appeared to be on the host side of the fiber scaffold, although the fibrous scaffold was not clearly visible in the processed sections. The cartilaginous tissue regeneration observed in the hydrogel was confined to the cartilage defect. Positive mineral staining at the defect-bone interface was observed in the location where the mineralized cup base was placed (Fig. 8.13).
132 8.3.10 Hydrogel Graft with IGF Cup and Agarose-CDA Composite Base
For the hydrogel group with the IGF cup and the agarose-CDA base, the host cartilage maintained a glossy white appearance, and the defect was filled with translucent tissue (Fig. 8.3). Similar to the hydrogel group with the IGF cup, the fiber scaffold had not degraded after four weeks and was visible in the cross-section of the defect prior to histological processing. Histological analysis revealed that the host tissue maintained a matrix rich in GAGs and collagen (Fig. 8.12), and the grafts consisted of a GAG- and collagen-rich matrix with uniformly distributed chondrocytes. A region void of matrix was observed in the center of the scaffold in one sample, likely the result of an air bubble that was introduced during in situ scaffold formation. Similar to the hydrogel repair with the IGF cup, new tissue was observed at the cartilage-cartilage interface in all depths of the cartilage tissue. The cells in the new tissue at the interface were consistent with the cell size in the adjacent host tissue. The bottom of the graft had a GAG-rich area in the region where the agarose-CDA base had been placed. The collagen that was deposited in the hydrogel at the cartilage-bone interface was aligned and was visible under polarized light. Positive mineral staining at the defect-bone interface was observed in the location where the mineralized cup base was placed (Fig. 8.13).
8.4 Discussion
The overarching goal of this research is to improve cartilage repair outcomes by engineering a scaffold system that improves integration of cartilage grafts with the host tissue. Specifically, this study evaluates the efficacy of a degradable polymer cup scaffold to promote cartilage-cartilage integration and cartilage-bone integration with three clinically relevant repair techniques. Full thickness chondral defects in bovine osteochondral explants were repaired with an autograft, tissue-engineered graft, or allogenic cell implantation in the presence and absence of an integration scaffold and cultured for four weeks subcutaneously in an athymic rat. The integration cup was implemented with all three repair techniques with minimal change to the standard repair process. Furthermore, the cup demonstrated potential to enhance the formation of new cartilage tissue at the cartilage-cartilage interface in the hydrogel repair methods, and led to aligned collagen at the cartilage-bone interface when combined with an agarose-
CDA base and used with a hydrogel cartilage graft.
133 In this study, the grafts were repaired ex vivo and transferred to a subcutaneous pouch in an athymic rat within one day. Despite the lack of a membrane or flap in the autograft and hydrogel repair groups, the grafts remained well-seated in the defect during culture, suggesting that the press-fit technique for the autograft placement and the in situ gelation for the hydrogel grafts are viable methods to achieve a tight fit of the graft within the defect. In addition, the fibrous sheath that surrounded the explant in the subcutaneous pouch likely contributes to holding the graft in place.
This defect model disrupts the cartilage, with limited damage to the surrounding tissue. After the subcutaneous culture period, however, depletion of the host glycosaminoglycans was observed in several samples, which has been previously reported for similar models [210]. The loss of glycosaminoglycans occurred primarily in the region of the cartilage closest to the surface, which may be a result of the interaction of the cartilage with the rat subcutaneous environment. Alternatively, it could also be caused by degradation of the matrix by the resident chondrocytes, as surface zone cells have been shown to degrade proteoglycans more rapidly than deep zone chondrocytes [37]. In our study, depletion of the glycosaminoglycan matrix was inconsistent across groups, with no clear link between repair type and host tissue maintenance.
For the defects repaired with an autologous graft, more consistent cartilage-cartilage integration was observed in the lower regions of the graft for all three groups. This finding may be an artifact of the model, as the surface zone was closer to the fibrous capsule, which may have interfered with the healing process in these regions. In several of the cross-sectional images, the fibrous capsule (most likely originating from the rat) extended from the outside of the explant into the region between the graft and host tissue. In addition, some of the grafts were proud to the surface of the defect, preventing the host and graft cartilage from healing together. This effect may be reduced in an intra-articular model in which the surface of the defect experiences loading.
Differences in integration may also be due to varying between the biosynthetic capacities of the cells that reside in the distinct zones of articular cartilage. It was previously demonstrated that cells isolated from the deep zone of articular cartilage are more active than cells from the surface zone, producing more glycosaminoglycans when cultured in agarose [38]. Additionally, the chondrocytes residing in the deeper regions of immature bovine cartilage are more synthetically active than those in the surface zone when
134 surrounded by their native matrix, with a ten-fold increase in synthetic activity for the cells in the radial zone compared to the surface zone [211]. Generally, clear differences between groups in terms of integration at the cartilage-cartilage interface were not observed histologically, and quantification of the integration across all three samples did not reveal differences between groups (data not shown). The integration in the cup groups may improve as the fibrous scaffold degrades. In the IGF cup samples, a gap between the host and graft cartilage was observed where the scaffold was placed. Cells from the surrounding cartilage tissue are budding off the surface, and it is possible that, given more time, these cells will bridge the gap, connecting the graft to the host tissue. To further understand how the cup affects autograft repair, later timepoints may provide insightful data. In addition, to further encourage integration and cell migration into the cup, a digestion agent which has been shown to facilitate integration [104-108], such as chondroitinase ABC, hyaluronidase, or collagenase, could be employed. Future studies should evaluate the compatibility of the integration cup with digestive washing protocols and assess the effects of combining the two integration approaches.
For the defects repaired via allogenic cell implantation, the bulk repair tissue that filled the defects was fibrous in nature, staining positive for collagen and negative for glycosaminoglycans. While the repair tissue was congruent with the host tissue, the formation of new cartilage tissue was inconsistent at the graft-host interface with and without the addition of the integration scaffold. The repair tissue at the interface was cellular and rich in glycosaminoglycans and collagen. After implantation, positive staining for glycosaminoglycans was observed in the bone tissue for all of the cell implantation-repaired samples, indicating that the injected chondrocytes may have traveled beyond the defect site, lodging in the bone tissue. Differences between groups were not observed with regard to this ectopic glycosaminoglycan-rich tissue formation. Because it is unclear if the final location of the injected cells recapitulates the scenario of cell implantation in an intact joint, future studies should be performed to evaluate the utility of the integration scaffold with cell-based repair techniques using an intra-articular model.
For the defects repaired with a hydrogel graft, a matrix consisting of glycosaminoglycans and collagen was deposited within the bulk of the graft for all samples. No clear differences in bulk matrix were observed between groups. More consistent cartilage-cartilage integration, however, was observed in the groups that were augmented with an integration scaffold. New tissue formation in these samples
135 appeared to align with the location of the fibrous scaffold and was rich in glycosaminoglycans and collagen. Integration in the cup-repaired hydrogel samples was observed in the deep, middle, and surface zones of the cartilage, and the cells in the repair tissue were similar in appearance to the cells in the adjacent host tissue. The integration observed for the hydrogel groups is not surprising because the hydrogel graft used in this study is an immature tissue engineered graft, and immature grafts integrate with host tissue better than more mature grafts [199;201]. The addition of the agarose-calcium deficient apatite base resulted in dense glycosaminoglycan deposition in the base of the graft and aligned collagen, visible under polarized light. Interestingly, alignment of the collagen was not observed at the base of any other hydrogel groups, suggesting that that addition of agarose-calcium deficient apatite leads to more organized tissue formation at the cartilage-bone interface. This finding parallels results from the rabbit study completed by Kharanrian et al. [17], in which the combination of a polymer mesh and agarose-ceramic scaffold resulted in calcified cartilage formation that was more organized than in cases in which the defect was repaired with either an agarose-ceramic scaffold or polymer-ceramic scaffold alone. The mechanism by which the hydrogel-ceramic base drives tissue organization, however, remains unknown.
This study enables the comparison between in vitro culture and subcutaneous culture of the full thickness defect model. In chapter 7, the model was cultured in static conditions in media that was replaced three times weekly, whereas in this chapter, the explant tissue was transferred to a subcutaneous pouch within one day of defect creation. As expected, the additional nutrients and the dynamic environment of the subcutaneous pouch result in more robust tissue growth, observed histologically. While differences between in vitro and in vivo culture are influenced by the available nutrients, they may also have been, in part, due to the difference in scaffold degradation rates in vitro and in vivo, as polymers degrade faster in vivo [212;213]. Therefore, although the same timepoint was used for both studies (four weeks), they may have been at different stages in the healing process.
In this study, the use of explant tissue provided a model to study the interaction of repair scaffolds with host and graft tissue. The data described here indicate that subcutaneous culture of the full thickness explant model can be used to study a variety of cartilage repair methods. Although this model provides a valuable platform to study scaffold-host interaction and integration, it has several limitations which should
136 be addressed in subsequent studies. Since the explant tissue was isolated from the marrow, synovium, and mechanical loading that are found in the joint, the spontaneous repair that is observed in intra- articular cartilage defects [214] is not recapitulated in this model. For example, after an intra-articular defect is formed, migrating cells from the synovial tissues may enhance scaffold-guided repair
Furthermore, since mechanical loading and growth factors have a synergistic effect on cartilage formation
[166], the growth-factor releasing scaffold may result in more cartilage formation in a loaded environment.
The lack of marrow in the bone compartment also reduces exposure to cells, such as mesenchymal stem cells, which may populate the base of the cup and contribute to calcified cartilage formation in vivo. In this study, a glycosaminoglycan-rich tissue was observed in the bone compartment of the cell implantation- repaired groups, suggesting that chondrocytes injected into the defect penetrated into the bone compartment. In an intra-articular defect, this would be less likely, as the creation of a defect results in blood that travels upward into the defect. The upward motion of the blood may counteract the downward trajectory of the cells injected into the defect. Another limitation of this model is the immaturity of the neonatal tissue used, as immature cartilage has higher cell density than older cartilage and possesses a greater innate healing capacity [74]. To elucidate the importance of these key differences, a full thickness defect model derived from mature tissue or in an intra-articular model of a mature animal could be used.
8.5 Conclusions
The results of this study collectively demonstrate that the integration scaffold system can be used to augment autografts, tissue engineered grafts, and cell implantation procedures. Furthermore, the optimized integration cup promotes cartilaginous tissue deposition at the graft-host cartilage junction in hydrogel and cell-based repairs.
137 Hydrogel Cartilage Graft Polymer Nanofiber Cup CaP nanoparticles IGF-1 Chondrocyte
Hydrogel+IGF(IGF-1+cup) + Autograft+ Cell Suspension+(IFG-1+cup)+ IGF Autograft ChHydrogel+hydrogel Cell Suspension Cup+Autograft (IFG-1+cup)+ (IFGHydrogel+-1+cup)+ Cup+(Ag-MF Ag base)-CDA + IGF Cup (Ch+hydrogelCup ) + cells (cartilage graft) (Ch+hydrogelIGF Cup ) (Ch+hydrogelBase )
Control Groups Experimental Groups
Full thickness defect model
Figure 8.1 Study design. Eight groups were generated by repairing the full thickness defect using different techniques. The repaired samples were cultured subcutaneously in an athymic rat for four weeks.
138
Figure 8.2 Subcutaneous rat study surgical procedure. An incision is made by holding the skin taut while a second surgeon creates a lateral cut using a #15 blade. The skin is then released and Metzenbaum scissors are used to create a subcutaneous pouch along the side of the animal. The repaired explant sample is placed into the pouch, the incision is closed using wound clips, and the samples are harvested after four weeks.
139 Autograft Control
Autograft + Control Cup
Autograft + IGF Cup
Chondrocyte Implantation Control
Chondrocyte Implantation + IGF Cup
Hydrogel Control
Hydrogel + IGF Cup
Hydrogel + IGF Cup + Hydrogel- CDA Base
Figure 8.3 Samples after in vivo implantation. All samples maintained similar dimensions and appearance through the implantation period, with a thin surrounding sheath of fibrous tissue.
140
Control + Control Cup + IGF-1 Cup AUTOGRAFT REPAIR AUTOGRAFT
Control + IGF Cup CHONDROCYTE REPAIR CHONDROCYTE
Control + IGF-1 Cup + IGF-1 Cup + Ag-CDA Base HYDROGEL REPAIR HYDROGEL
Figure 8.4 Histology overview of repair. All sample defects were filled with repair tissue after implantation (hematoxylin and eosin, scale bar = 1 mm).
141 Safranin-O Picrosirius Red Collagen II
C C B B A A D D
A. Cartilage-cartilage B. Cartilage-cartilage C. Bulk repair D. Cartilage-bone
H&E
O
-
Safranin
PicrosiriusRed PolarizedLight
Figure 8.5 Autograft control repair. The top three images show a cross-section stained with safranin-O (left) picrosirius red (middle) and collagen II (right) staining (4x, bar = 1 mm). Regions of interest are imaged (10x, bar = 200 µm) from areas indicated in the low magnification image with and without a polarizer. The hematoxylin and eosin (H&E) images in the top panel represent regions similar to the areas identified in the safranin-o overview image.
142 Safranin-O Picrosirius Red Collagen II
C B A C A B D D
A. Cartilage-cartilage B. Cartilage-cartilage C. Bulk repair D. Cartilage-bone
H&E
O
-
Safranin
PicrosiriusRed PolarizedLight
Figure 8.6 Autograft with control cup repair. The top three images show a cross-section stained with safranin-O (left) picrosirius red (middle) and collagen II (right) staining (4x, bar = 1 mm). Regions of interest are imaged (10x, bar = 200 µm) from areas indicated in the low magnification image with and without a polarizer. The hematoxylin and eosin (H&E) images in the top panel represent regions similar to the areas identified in the safranin-o overview image.
143 Safranin-O Picrosirius Red Collagen II
C
A C B A B D D
A. Cartilage-cartilage B. Cartilage-cartilage C. Bulk repair D. Cartilage-bone
H&E
O
-
Safranin
PicrosiriusRed PolarizedLight
Figure 8.7 Autograft with IGF cup repair. The top three images show a cross-section stained with safranin-O (left) picrosirius red (middle) and collagen II (right) staining (4x, bar = 1 mm). Regions of interest are imaged (10x, bar = 200 µm) from areas indicated in the low magnification image with and without a polarizer. The hematoxylin and eosin (H&E) images in the top panel represent regions similar to the areas identified in the safranin-o overview image.
144 Safranin-O Picrosirius Red Collagen II
B C C B A A D D
A. Cartilage-cartilage B. Cartilage-cartilage C. Bulk repair D. Cartilage-bone
H&E
O
-
Safranin
PicrosiriusRed PolarizedLight
Figure 8.8 Chondrocyte implantation control repair. The top three images show a cross-section stained with safranin-O (left) picrosirius red (middle) and collagen II (right) staining (4x, bar = 1 mm). Regions of interest are imaged (10x, bar = 200 µm) from areas indicated in the low magnification image with and without a polarizer. The hematoxylin and eosin (H&E) images in the top panel represent regions similar to the areas identified in the safranin-o overview image.
145 Safranin-O Picrosirius Red Collagen II
C C B B A A D D
A. Cartilage-cartilage B. Cartilage-cartilage C. Bulk repair D. Cartilage-bone
H&E
O
-
Safranin
PicrosiriusRed PolarizedLight
Figure 8.9 Chondrocyte implantation with IGF cup repair. The top three images show a cross-section stained with safranin-O (left) picrosirius red (middle) and collagen II (right) staining (4x, bar = 1 mm). Regions of interest are imaged (10x, bar = 200 µm) from areas indicated in the low magnification image with and without a polarizer. The hematoxylin and eosin (H&E) images in the top panel represent regions similar to the areas identified in the safranin-o overview image.
146 Safranin-O Picrosirius Red Collagen II
C B A C A B D D
A. Cartilage-cartilage B. Cartilage-cartilage C. Bulk repair D. Cartilage-bone
H&E
O
-
Safranin
PicrosiriusRed PolarizedLight
Figure 8.10 Hydrogel graft control repair. The top three images show a cross-section stained with safranin-O (left) picrosirius red (middle) and collagen II (right) staining (4x, bar = 1 mm). Regions of interest are imaged (10x, bar = 200 µm) from areas indicated in the low magnification image with and without a polarizer. The hematoxylin and eosin (H&E) images in the top panel represent regions similar to the areas identified in the safranin-o overview image.
147 Safranin-O Picrosirius Red Collagen II
C C A B D A B D
A. Cartilage-cartilage B. Cartilage-cartilage C. Bulk repair D. Cartilage-bone
H&E
O
-
Safranin
PicrosiriusRed PolarizedLight
Figure 8.11 Hydrogel graft with IGF cup repair. The top three images show a cross-section stained with safranin-O (left) picrosirius red (middle) and collagen II (right) staining (4x, bar = 1 mm). Regions of interest are imaged (10x, bar = 200 µm) from areas indicated in the low magnification image with and without a polarizer. The hematoxylin and eosin (H&E) images in the top panel represent regions similar to the areas identified in the safranin-o overview image.
148 Safranin-O Picrosirius Red Collagen II
C
A B B A C D D
A. Cartilage-cartilage B. Cartilage-cartilage C. Bulk repair D. Cartilage-bone
H&E
O
-
Safranin
PicrosiriusRed PolarizedLight
Figure 8.12 Hydrogel graft with IGF cup and agarose-calcium deficient apatite base repair. The top three images show a cross-section stained with safranin-O (left) picrosirius red (middle) and collagen II (right) staining (4x, bar = 1 mm). Regions of interest are imaged (10x, bar = 200 µm) from areas indicated in the low magnification image with and without a polarizer. The hematoxylin and eosin (H&E) images in the top panel represent regions similar to the areas identified in the safranin-o overview image.
149
Control + Control Cup + IGF-1 Cup AUTOGRAFT REPAIR AUTOGRAFT
Control + IGF Cup CHONDROCYTE REPAIR CHONDROCYTE
Control + IGF-1 Cup + IGF-1 Cup + Ag-CDA Base HYDROGEL REPAIR HYDROGEL
Figure 8.13 Mineralization. The bone and the bottom of the cup scaffold stain positively for mineral (von Kossa and safranin-O, scale bar = 1 mm).
150 CHAPTER 9: SUMMARY AND FUTURE DIRECTIONS
151 9.1 Summary
The objective of this thesis was to design a multi-phased integration scaffold for cartilage repair to improve healing of joint injuries. A scaffold was rationally designed and tested to address shortcomings of tissue repair methods that currently limit long-term clinical outcomes. The ideal scaffold system should promote graft integration to the host tissue at both the cartilage-cartilage and the cartilage-bone interfaces. The scaffold should increase cellularity at the cartilage-cartilage interface to promote cell- mediated biological fixation of the graft and enhance calcified cartilage formation at the graft-subchondral bone junction. Finally, the ideal scaffold should be compatible with current clinical repair strategies. It was hypothesized that a cup-shaped scaffold, with insulin-like growth factor-1 in the walls to home cells and bioactive ceramic in the base to promote calcified cartilage formation, would promote integration of cartilage grafts at both the cartilage-cartilage and cartilage-bone interfaces.
To evaluate these hypotheses, studies were designed to address three specific aims. Aim 1 focused on the design and optimization of the cup walls to address cartilage-cartilage integration. To this end, insulin-like growth factor-1, a chemical homing agent for chondrocytes and stem cells, was incorporated into electrospun fibrous scaffolds to attract cells from cartilage tissue into the scaffold. The dose of insulin-like growth factor-1 was optimized to promote the homing of cells into the scaffold (chapter
2). It was demonstrated that the insulin-like growth factor-1, incorporated at 100 ng/mg, significantly increased cell number on scaffolds cultured adjacent to cartilage tissue compared to growth factor-free control scaffolds. Next, the response of chondrocytes on the scaffold was assessed and it was shown that the scaffold supports cell attachment, viability, and matrix elaboration (chapter 3).
In Aim 2, cartilage-bone integration was addressed. Bioactive ceramics were evaluated with varying crystallinity to optimize the ceramic phase of the scaffold (Chapter 4). Calcium deficient apatite, a biomimetic, bioactive calcium phosphate, was identified as an optimal ceramic for osteochondral interface regeneration because it significantly enhanced collagen II and glycosaminoglycan deposition. Since preliminary in vivo data had suggested that a polymer fiber-hydrogel-ceramic composite may be ideal for calcified cartilage regeneration, the ceramic dose was optimized in both agarose (Chapter 5) and polymeric fiber scaffolds (Chapter 6).
152 After optimizing both the walls and base of the cup for cartilage-cartilage and cartilage-bone integration, Aim 3 evaluated the potential for clinical translation of the complete scaffold. Using bovine tissue, a full thickness defect model was developed, characterized, and used to test the scaffold system.
The cup system was used to repair the defect in conjunction with autologous tissue grafts, chondrocyte implantation, and hydrogel-based tissue engineered cartilage grafts. The effect of the cup on healing and integration was evaluated in vitro (chapter 7) and in vivo (chapter 8) with a subcutaneous athymic rat model. The major findings of each chapter are briefly highlighted below.
9.1.1 Scaffold Design and Optimization for Cartilage-Cartilage Integration
The first aim developed and optimized the walls of the cup for placement at the cartilage-cartilage junction to promote integration. The walls were designed to be thin, easy-to-handle, degradable, and able to release homing agents. To meet these criteria, a polymeric fiber-based scaffold was developed by electrospinning a blend of poly(lactide-co-glycolide) and polycaprolactone with incorporated insulin-like growth factor-1. In chapter 2, the number of cells on scaffolds that were cultured adjacent to cartilage tissue was measured as a function of the concentration of insulin-like growth factor-1 in the fibers
(chapter 2). Growth factor incorporation at 100 ng/mg led to increased cell number on scaffolds after two weeks of in vitro culture. This finding demonstrated that the release of a homing agent from a fibrous scaffold increases cellularity, and this dose of insulin-like growth factor-1 was used in subsequent studies.
Chapter 3 determined the response of chondrocytes to the growth factor-releasing scaffold. Isolated from bovine tissue and seeded on the scaffolds, the chondrocytes demonstrated cell attachment, viability, and matrix elaboration. Lower mineralization potential was measured for scaffolds containing growth factors compared to those without. Collectively, these results informed the design of the walls of the cup, the cartilage-cartilage integration phase of the scaffold system.
9.1.2 Scaffold Design and Optimization for Cartilage-Bone Integration
The second aim developed the base of the cup, designed to be placed at the cartilage-bone interface and optimized to promote integration by the formation of calcified cartilage. Previous reports showed that ceramics within a polymer scaffold promote cell-mediated formation of calcified cartilage
[11;15]. Thus, bioactive calcium phosphates were incorporated into agarose hydrogels, and the response
153 of deep zone chondrocytes was evaluated with varying ceramic crystallinity (chapter 4). Poorly crystalline calcium deficient apatite resulted in significantly more glycosaminoglycan and collagen production over time compared to scaffolds without ceramic or with incorporated tricalcium phosphate, a crystalline ceramic. Next, the dose of calcium deficient apatite was optimized in agarose hydrogel scaffolds and polymer fiber scaffolds. For both scaffold platforms, a dose of ceramic was identified that promoted cell growth and biosynthesis, resulting in a calcified cartilage-like tissue.
9.1.3 Evaluating the Clinical Potential of the Integrative Scaffold System
The third aim evaluated the clinical potential of the total integrative scaffold system. As tissue grafts and cell-based therapies are used in the clinic to treat cartilage lesions, the cup was tested with both approaches. In addition, since tissue-engineered cartilage grafts have been approved in Europe and are in the pipeline for FDA approval, the cup was also tested with an agarose-based, tissue-engineered cartilage graft. To evaluate clinical implementation, bovine osteochondral tissue was harvested, and a full thickness defect was made in the cartilage tissue. This organ culture model, derived from a large animal with cartilage sufficiently thick for scaffold evaluation, avoids the high costs associated with large animal in vivo studies. In vitro testing of the cup in the defect model demonstrated that the scaffold system was compatible with clinical repair techniques and led to stronger cartilage-cartilage integration (chapter 7).
Subsequently, the full thickness defect model, repaired with the integration scaffold, was implanted subcutaneously into athymic rats, and cells migrated into the cup, depositing new tissue at the graft-host interface. When combined with the agarose-calcium deficient apatite base and used with the hydrogel graft, the cup scaffold resulted in organized collagen formation at the cartilage-bone interface.
9.2 Future Directions
The findings of this thesis demonstrate the promise of using a multi-phased scaffold to address cartilage-graft integration. However, to realize the clinical translation of the scaffold system, several areas of further study are needed, as described below.
9.2.1 Human Cartilage Organ Model Testing
The bovine full thickness defect model enabled the evaluation of the clinical implementation of the integration scaffold system in conjunction with techniques similar to those used in the clinic. Nevertheless,
154 the immature cartilage tissue is inherently different from the tissue that would interact with the scaffold in a human surgery. To address this, the full thickness defect model could be extended to human osteochondral tissue. While human tissue is less available, excess osteochondral tissue can be harvested for this purpose from human grafts after allografting procedures.
9.2.2 Intra-Articular in Vivo Testing
Although the full thickness defect model enables evaluation of the scaffold with host cartilage and bone tissue, it is necessary to evaluate the scaffold in the intra-articular environment. Compared to the organ culture model, an intra-articular model, such as the rabbit, would enable assessment of the scaffold in an environment with mechanical loading and the chemical cues from synovial fluid.
9.2.3 Stem Cell-Seeded Scaffold
While an acellular approach offers advantages in cost, packaging, and clinical implementation, pre-seeding the scaffold with stem cells may improve the integration capacity of the system. Future studies will investigate the effects of scaffolds pre-seeded with synovium derived stem cells, which are well-suited for cartilage repair. In addition, the recent development of induced pluripotent stem cells
(iPSC) may enable cell-based approaches that do not require an additional surgical procedure [215].
Preliminary studies with bovine synovium-derived stem cells show that the scaffolds support stem cell attachment and viability (Fig. 9.1), live stem cell tracking shown in red); however culture conditions will need to be optimized to promote matrix elaboration and differentiation in future studies.
Day 1 Day 14 Day 21
Figure 9.1 Synovium Derived Stem Cells on SDSC Microfiber Scaffolds with and without IGF. Synovium derived stem cells are shown in red on microfiber scaffolds. The cells attached and 250 µm uniformly on the scaffolds SDSC+IGF over time (n=3, DiI dye).
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