Biomimetic Scaffold Design for Tendon Tissue Engineering

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BIOMIMETIC SCAFFOLD DESIGN FOR TENDON TISSUE ENGINEERING

TO ENHANCE MECHANORESPONSE AND TENOGENESIS

ANOWARUL ISLAM

Submitted in partial fulfillment of the requirements

For the degree of Doctor of Philosophy

Dissertation Adviser: Dr. Ozan Akkus

Department of Mechanical and Aerospace Engineering

CASE WESTERN RESERVE UNIVERSITY

January, 2017

CASE WESTERN RESERVE UNIVERSITY SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis/dissertation1 of

ANOWARUL ISLAM

for the degree of

Doctor of Philosophy

Dr. Ozan Akkus

Committee Chair, Adviser 07/25/2016 Department of Mechanical and Aerospace Engineering

Dr. Joseph Mansour

Committee Member 07/25/2016 Department of Mechanical and Aerospace Engineering

Dr. Umut A. Gurkan

Committee Member 07/25/2016 Department of Mechanical and Aerospace Engineering

Dr. Horst Von Recum

Committee Member 07/25/2016 Department of Biomedical Engineering

1We certify that written approval has been obtained for any proprietary material contained therein. i

TABLE OF CONTENTS

Table of Tables ...... viii

Table of Figures ...... ix

Acknowledgements ...... xiii

ABSTRACT ...... xv

1. Chapter 1: Background ...... 1

1.1 Introduction ...... 1

1.2 Tendon Structure ...... 1

1.3 Tendon Composition ...... 3

1.4 Tendon Mechanical Properties ...... 5

1.5 Tendon Injuries ...... 6

1.6 Current Repair Techniques...... 6

1.6.1. Suture based surgical repair ...... 6

1.6.2. Augmentation strategy in suture based technique ...... 8

1.6.3. Biological graft based technique ...... 10

1.7 Tendon Tissue Engineering ...... 11

1.7.1. Synthetic Scaffolds ...... 12

1.7.2. ECM Derived Scaffolds ...... 13

1.7.3. Cells Used in Tendon Tissue Engineering: ...... 14 ii

1.7.4. Growth Factors (GFs) Used in Tendon Tissue Engineering: ...... 15

1.8 Collagen as a Bioscaffold ...... 16

1.9 Electrochemically Aligned Collagen (ELAC) for Tendon Repair 126-133 ...... 17

1.9.1. Fabrication and physical characteristics of ELAC 126: ...... 18

1.9.2. Mechanical properties of ELAC threads 131: ...... 19

1.9.3. Differentiation and in vitro response of MSCs to ELAC 127 128: ...... 20

1.9.4. In vivo biocompatibility and degradation of ELAC in a Rabbit patellar tendon model

130: 21

1.10 Matrix Stiffness and Topography:...... 22

1.11 Ideal Scaffold for Tendon Tissue Engineering ...... 23

2. Chapter 2: Research Objectives ...... 24

2.1 Introduction ...... 24

2.2 Aim 1: Assess the effect of stiffness anisotropy of ELAC substrate on the cytoskeletal behavior and tenogenesis ...... 24

2.3 Aim 2: Increase tenogenic differentiation on electrocompacted collagen substrates. .... 25

2.4 Aim 3: Construct a biomimetic tendon repair scaffold ...... 26

3. Chapter 3: Collagen Substrate Stiffness Anisotropy Affects Cellular Elongation, Nuclear

Shape and Stem Cell Fate towards Anisotropic Tissue Lineage ...... 28

3.1 Abstract ...... 28

3.2 Introduction ...... 29

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3.3 Materials and Methods: ...... 30

3.3.1. Fabrication of Collagen Sheets by Electrochemical Compaction [Figure 11A]: ...... 30

3.3.2. Resolving the effects of topography from the effects of SA by cellulose acetate replication of surface topography ...... 31

3.3.3. Tuning of Molecular Alignment via Mechanical Stretch: ...... 31

3.3.4. Imaging of Molecular Alignment in Compacted Collagen Sheets ...... 33

3.3.5. Assessment of Mechanical Properties of Stretched Sheets (SA): ...... 33

3.3.6. Surface Roughness Measurement by Atomic Force Microscopy (AFM): ...... 34

3.3.7. Cell Viability after day 1 ...... 34

3.3.8. Effect of SA on Cytoskeletal and Nuclear Morphologies ...... 35

3.3.9. Cell seeding on the replicated cellulose acetate film ...... 36

3.3.10. Measurement of Cytoskeletal and Nucleus Morphologies ...... 36

3.3.11. Effect of SA on Tenogenic Differentiation of Human MSCs ...... 36

3.3.12. Effect of SA on Tendon Related Matrix Synthesis ...... 37

3.3.13. Statistical Analysis: ...... 39

3.4 Results ...... 40

3.5 Discussion ...... 50

3.6 Conclusions ...... 54

4. Chapter 4: Tenoinduction by Mimicry of Tendon Topography on Pure Collagen Substrate

4.1 Abstract ...... 56

4.2 Introduction ...... 57

4.3 Materials and Methods: ...... 58

4.3.1. Collagen Sheet Fabrication by Electrochemical Compaction [Figure 21]: ...... 58

4.3.2. Synthesis of Collagen Gel: ...... 59

4.3.3. Experimental Groups: ...... 61

4.3.4. Imaging of Molecular Alignment in Compacted Collagen Sheets ...... 61

4.3.5. Assessment of Mechanical Properties of Collagen Sheets: ...... 62

4.3.6. Effect of Matrix Modulus and Alignment on Tenogenic Differentiation of Human

MSCs…… ...... 62

4.3.7. Effect of Matrix Modulus and Alignment on Tendon Related Matrix Synthesis ...... 63

4.3.8. Statistical Analysis: ...... 65

4.4 Results: ...... 65

4.5 Discussions: ...... 71

4.6 Conclusions ...... 74

5. Chapter 5: Assess effects of dermatan sulfate incorporation on the differentiation of

MSCs...... 75

5.1 Abstract ...... 75

5.2 Introduction ...... 76

5.3 Materials and Methods: ...... 76

5.3.1. DS Incorporated Collagen Sheet Fabrication: ...... 76

5.3.2. Assessment of Mechanical Properties of Collagen Sheets: ...... 78

5.3.3. Effect of DS Incorporation On Cell Proliferation On Compact Sheets:...... 78

5.3.4. Effect of DS Incorporation on Tenogenic Differentiation of Human MSCs ...... 78

5.3.5. Statistical Analysis: ...... 79

5.4 Results: ...... 79

5.5 Discussions: ...... 82

5.6 Conclusion:...... 84

6. Chapter 6: Computer Aided Biomanufacturing of Mechanically Robust Pure Collagen

Meshes with Controlled Macroporosity ...... 85

6.1 Abstract ...... 85

6.2 Introduction ...... 87

6.3 Materials and Method: ...... 88

6.3.1. Overview of the Patterned Electrocompaction Process (Figure 30): ...... 88

6.3.2. Manufacturing an individual patterned layer...... 89

6.3.3. Fabrication of 3-D Scaffolds with Controlled Interconnected Porosity: ...... 92

6.3.4. Imaging of Molecular Alignment within Patterned Channels: ...... 93

6.3.5. Assessment of Mechanical Properties of Patterned Bioscaffolds: ...... 93

6.3.6. Adhesion and Morphology of Human Mesenchymal Stem Cells Seeded on Patterned

Bioscaffolds ...... 94

6.3.7. Benchmark for Mechanical Performance of Scaffolds: ...... 95

6.3.8. Computational Screening of Scaffold Morphologies: ...... 96

6.3.9. Statistical Analysis: ...... 98

6.4 Results: ...... 98

6.5 Discussion: ...... 103

6.6 Conclusion ...... 108

7. Chapter 7: Summary and Future Directions ...... 110

Appendix A. Patterned Polycarbonate-Cathode Bilayer Manufacturing Steps ...... 116

Appendix A1. Preparation of Plastic Cathode Bilayer ...... 116

Appendix A2. CAD/CAM Design of Pattern Mold ...... 117

Appendix A3. G-Code for CNC Machining of Negative Replica of Patterned Scaffold...... 117

Appendix A4. Procedure for using Meshcam ...... 122

Appendix A5. Machining Pattern Mold ...... 127

Bibliography ...... 128

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Table of Tables

Table 1: Scaffold devices with FDA clearance for rotator cuff repair 43 ...... 12

Table 2: Experimental groups for different level of modulus range and unidirectionally aligned substrate ...... 61

Table 3: Comparative mechanical properties of collagen based scaffolds for tissue engineering application ...... 107

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Table of Figures

Figure 1: Schematic illustration of the hierarchical structure of tendon...... 3

Figure 2: Using suture anchor in rotator cuff repair ...... 7

Figure 3: Using bone tunnel during surgery 51 ...... 7

Figure 4: Knotless fixation during rotator cuff surgery ...... 8

Figure 5: A and B, Intraoperative photographs showing augmentation patch 45 ...... 9

Figure 6: Enthesis in tendon bone interface ...... 9

Figure 7: A) Schematic representation of the monomeric collagen to solid ELAC B)

Compensated polarized images of tendon and ELAC fibers...... 18

Figure 8: Mechanical Properties of ELAC threads optimized using different crosslinking treatments...... 19

Figure 9: A) In vitro cell-matrix layer (black arrows) deposition on ELAC threads (white arrows) by MSCs. B) The orientational anisotropy of ELAC is dictated on the cellular cytoskeleton. Also note that nuclei are elongated along the ELAC fiber axis...... 20

Figure 10: Tenogenic differentiation of human MSCs on ELAC (fold change with respect to randomly oriented collagen)...... 21

Figure 11: A) Overview of electrochemical compaction of collagen sheets, B) Overview of molecular alignment by stretching...... 32

Figure 12: A) CPI images of collagen sheets at different levels of stretch. B) SEM images of collagen sheets stretched to different strain levels indicate an aligned topography along the stretching direction (arrows; scale bar 2μm). C) Replication of the surface topography of highly stretched (SA=8) collagen sheet on cellulose acetate film...... 41

Figure 13: Top right figure shows the mechanical testing direction of the longitudinal and transverse samples with the stretching direction. A) Transverse and longitudinal stiffness of collagen sheets at different stretch levels. B) Stiffness (left) and failure stress (right) anisotropy in collagen sheets increased with stretch...... 42

Figure 14: Surface Roughness measurement by AFM. The Mean RMS surface measurement showed that SA incorporation did not affect the surface roughness...... 43

Figure 15: Cell viability on SA=1 and SA=8 groups...... 44

Figure 16: A) MSCs’ cystoskeletal morphology at SA=1, SA=3 and SA=8 (Scale bar 50 μm) after 6 and 12 hours of seeding. Stretching direction indicated by arrows. B) Morphology of cells seeded on cellulose acetate with and without surface replication shows lack of cellular elongation and alignment in both cases. (Scale bar: 100µm)...... 45

Figure 17: Cell aspect ratio (A), major axis (B) and minor axis (C) length after 6 and 12 hours

(Left column). Nuclear aspect ratio (D), major axis (E) and minor axis (F) length after 6 and 12 hours (Right column). Significance was set at p < 0.025 using Bonferroni correction. (G)The schema is drawn to scale from the reported average measurements to illustrate cell and nucleus dimensions at different values of anisotropy. (H) Cells seeded on surface replicated cellulose acetate sheet showed that there was no significant difference in aspect ratios of cells seeded on surfaces replicated with the surface topography of collagen vs. control group...... 46

Figure 18: Cumulative Relative Histogram Plot of Cell Alignment Angle...... 47

Figure 19: Effect of SA on tenogenic differentiation of human MSCs...... 48

Figure 20: Effect of SA on long term matrix synthesis...... 49

Figure 21: Overview of fabricating collagen sheets with different modulus levels...... 60

Figure 22: Manufacturing aligned sheet with mechanical stretching. Molecular alignment is evident by CPI image (Emergence of blue color after stretching indicates molecular alignment).

Collagen fibril alignment is evident by SEM images...... 66

Figure 23: Effect of collagen matrix modulus and anisotropy on tenogenic differentiation of human MSCs...... 68

Figure 24: Effect of Modulus on long term matrix synthesis...... 69

Figure 25: After 21 days of hMSc culture, 100 MPa and 10 MPa aligned groups showed 3-fold increase in Type-I collagen synthesis than 0.1 MPa group; cells on 100 MPa group showed 4- fold increase in TSP-4 production than 0.1 MPa group indicated by corrected total cell fluorescence (CCTF); and type-III collagen showed 2-fold upregulation from 0.1 MPa to 100

MPa with a slightly increasing trend in aligned group compared to unaligned group...... 70

Figure 26: A) Dermatan Sulfate (DS) incorporated electrocompacted collagen sheet. B)

Concentration of DS by DMMB assay...... 77

Figure 27: Mechanical test result of the groups ...... 79

Figure 28: Cell proliferation did not affect by DS incorporation...... 80

Figure 29: Effect of DS incorporation on tenogenic differentiation of human MSCs...... 81

Figure 30: Process of fabricating individual patterned layer...... 89

Figure 32: Scaffolds with 4 different porosity shapes. A) Diamond-shaped pore; B) Square- shaped pore; C) Rectangle-shaped pore 4) Parallel channel pore...... 92

Figure 33: a) Finite element result of scaffold geometries b) Stiffness plot for different geometries from FEM c) Porosity and stiffness from FEM comparison for different geometries d) Rectangular porosity gives maximum porosity and stiffness ...... 97

Figure 34: A) Branching in the diamond-shaped patterned lattice displayed lack of alignment of collagen molecules as manifested by the magenta in the compensated polarize image (CPI).

B) Parallel channels introduce alignment in the patterned layer (emergence of blue color in CPI indicates alignment)...... 99

Figure 35: Mechanical assessment of patterned scaffolds with three different layer numbers.

...... 100

Figure 36: Mechanical assessment of patterned scaffolds with four different pore shapes. ... 101

Figure 37: DAPI (blue) and F-Actin (green) staining images revealed that cells covered the entire scaffold through thickness. Cells and nucleus became elongated along the length of the collagen filament...... 102

Figure 38: Cytoskeletal arrangement and tenogenic differentiation of MSCs ...... 114

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Acknowledgements

First and foremost, I would like to thank my Ph.D. advisor, Dr. Ozan Akkus, for giving me the opportunity to work under his supervision. He was always there when I had questions either regarding the direction of the projects or required a training to use any equipment. He trained me in person on how to use some of the equipment in the lab and for this extraordinary quality, I am really grateful to him. Overall, under his wing, I shaped up as an experimental researcher and developed skills to become an independent researcher which will help me in my future career. Besides academic advising, during a very tough time in my personal life, he extensively supported me and gave me space in a professional manner so that I could get through the time. He has been an excellent role model as a mentor and in my future career, I wish to be a mentor like him.

I would like to acknowledge my Ph.D. committee members: Dr. Joseph Mansour,

Dr. Umut Gurkan and Dr. Horst von Recum for their insightful feedback. I would like to specially thank Dr. Mansour for his helpful suggestion regarding mechanical testing and

Dr. Gurkan for developing a wonderful training process for using the Olympus IX83 microscope in his lab.

I would like to thank current lab members Mousa Younesi, Dr. Thomas Mbimba and previous lab member Dr. Vipuil Kishore as a collaborator in different projects as well as partner in brainstorm for different projects and experiments. I would also like to acknowledge the help from other current and previous lab members: Katherine Chapin,

Bolan Li, Hyung Jin Jung, Mustafa Unal, Yunus Alapan, Hakan Celik, Ping He, Greg

xiii

Learn, Andrew Tsai, Michael Bohl, Joel Ford and Emily Moore

I would like to thank Cheryl King and India Bowie from purchasing department for their help regarding purchasing necessary supplies of research in a timely manner.

I would also like to thank my wife who gave me mental support throughout my

PhD education. I would like to acknowledge all my friends for their suggestion, support and encouragement. I am thankful to my parents and other family members for motivating me to start and continue the PhD education.

Finally, I would like to thank Musculoskeletal Transplant Foundation (MTF) for providing support of my graduate study for first couple of years. I would also like to thank

National Scientific Foundation (NSF) and National Institute of Health (NIH) for providing the funding of this study.

xiv

Biomimetic Scaffold Design for Tendon Tissue Engineering to Enhance Mechanoresponse and Tenogenesis

ANOWARUL ISLAM

ABSTRACT

Tendon disorders are some of the most common musculoskeletal problems. Despite improvements in surgical techniques, revision of tendon tear surgery is significantly high. There is a recognized need for suitable bioscaffold that is porous enough for cell migration and proliferation, can promote tenogenic differentiation, and is simultaneously mechanically strong enough for tissue regeneration after implantation.

The inability to incorporate high strength and high porosity in a structure has been one of the major barriers in the engineering of load-bearing tissue. There is a general lack of biofabrication methods that will provide macroporosity (0.5 mm or greater) with mechanical robustness in delicate protein-based biomaterials such as collagen. The current study developed a CAD/CAM based electrocompaction method to manufacture highly porous patterned scaffolds using pure collagen which has a potential to combine these characteristics. The lattice structures are made up of anisotropically aligned sheet like lattice structures. Therefore, the dissertation also develops a material model which considers stiffness as well as stiffness anisotropy (SA) to mimic two key features of natural tendon.

Most natural tissues are substantially stronger along the load bearing direction than the direction transverse to the longer axis (such as tendons, muscle etc.). Notably, the cells of such tissues are elongated along the stiffer direction. There are not many biomaterials to emulate and xv study the effects of stiffness anisotropy on cellular response. In addition to the SA, extracellular matrix stiffness or mechanics also plays an important role to regulate cell morphology, proliferation, differentiation during regular and diseased states. There has been relatively little investigation on the roles of topographical factors (matrix anisotropy vs. matrix stiffness) in inducing tenogenic differentiation and the roles are still unknown. Elucidation of the mechanism of topographically induced tenogenesis in human MSCs will assist in optimizing the material to produce scaffolds for tendon repair and may provide insight into the differentiation mechanisms of MSCs and other stem cells in vivo, where type I collagen is the major substrate protein.

Effect of SA and stiffness study demonstrated that stem cell fate is affected by not only the magnitude of stiffness but also by the directional anisotropy of the substrate stiffness. The study suggests that increasing stiffness anisotropy has a positive effect on tenogenesis. However, highly stiff substrate without SA can generate similar tenogenic differentiation as substrate with

SA. This indicates, may be stiffness can overcome the effect of stiffness anisotropy on tenogenesis. Overall, SA has a positive effect on tenogenesis as anisotropy can be a key determinant in driving cell morphology and differentiation during development and maintenance of anistropically stiff tissues. In the stiffness study, a material profile was developed where the modulus can be changed from several hundred kPa, to single digit MPa, to hundred MPa. The tenoinductive collagen material model developed in the study can be used in the research and development of tissue engineering tendon repair constructs in future.

The patterned lattice structure fabrication method demonstrated the capabilities of manufacturing mechanically robust ‘scaffolds with controlled porosity’. In this method, by changing the number of layers and shape of the structure, mechanical properties can be modulated for different tissue engineering application such as tendon, hernia, SUI, thoracic wall xvi reconstruction. Pure collagen based scaffolds has not been used in clinical mesh application due to their weak mechanics and usually collagen rich decellularized tissue based xenografts are used clinically when mechanical robustness is required. The fabrication method developed in this study generates pure collagen scaffold mesh within the mechanical strength range of decellularized tissue.

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1. Chapter 1: Background

1.1 Introduction

Tendons are collagenous tissue that bear and transfer load from muscle to bone. Tendon disorders are some of the most common musculoskeletal problems, which occur by acute tendon tears due to trauma, injury, or by age-related degeneration. People with damaged tendon face complications which are related to the function of tendons such as instability, loss of partial or complete range of motion and pain in a particular joint. Moreover, the aforementioned problems result in progression of degenerative diseases. Tendon-repair procedures occur in excess of 100,000 annually in the

U.S. alone costing approximately $30 billion dollars 1, 2. Despite improvements in tendon repair techniques, revision of tendon tear surgery is significantly high. This necessitates more study to a more effective approach of tendon repair. A novel tissue engineering approach of tendon repair is the focus of this dissertation.

1.2 Tendon Structure

Tendons are mainly composed of type I collagen fibril. These collagen fibrils are arranged axially in load bearing direction3. Their ECM is arranged in hierarchical structure with different dimensions. The most basic building block of tendons are collagen molecules or tropocollagen, which is composed of three polypeptide strands, known as alpha chains. The alpha chains are basically composed of amino acid sequence, Glycine-X-Y (Where X and Y positions are commonly filled with proline and hydroxyproline4). Alpha chains are individually left handed helix and as a result twisted around each other and form a right handed coils. Tropocollagens then self-assemble into collagen fibrils with a stagger distance (D) of 67nm5. The fibril is the smallest tendon structural unit which has diameter varies from 10 to 500 nm, depending on different biological factors such as species, age and sample location6. 1

The fibrils are then self-assembled in collagen fibers which is the next level of tendon structure. Collagen fibrils are bound by endotenons in collagen fiber6. Endotenons are connective tissue sheaths which supply vasculature, lymphatics, and nerves 7-10. Collagen fiber diameters range from 5 – 30 μm in rat tail tendons up to 300μm in human tendons8, 11. Fibers are then packed into both primary and secondary bundles (fascicles). Similar to collagen fibers, primary fiber bundles and secondary fiber bundles (fascicles) are also surrounded by endotenon. Fascicles are then packaged into tertiary fiber bundles and form tendon unit by surrounded by epitenon 9. The purpose of epitenon is same as endotenon. Endotenon is a reticular network of thin collagen fibrils in a crisscross pattern while epitenon is a relatively dense network of collagen fibrils 9. In human tendons, fascicle diameters range from 150 – 1000μm and tertiary bundle diameters range from

1000 – 3000μm (the diameters of both fascicles and tertiary bundles are directly related to the macroscopic size of the tendon structure) 8. Tendons are also bound by the paratenon (synovial sheath for some tendons) 6. Paratenon is a loose, fatty, areolar connective tissue that functions as an elastic sleeve.

The hierarchical structure appears as a ‘‘crimp pattern’’ when longitudinal sections of the tendon are viewed in a polarized microscope 12, 13. The initial 2% of tendon strain corresponds to the elongation of the crimp and the crimped configuration is removed when the tendon is strained

14, 15 . The crimp pattern facilitates a buffer against tendon injury 14, 15. The hierarchical structure aligns of fiber bundles in load bearing direction or the long axis of tendon which facilitates tendon’s high tensile strength.

Figure 1: Schematic illustration of the hierarchical structure of tendon. The tendon has a multi- unit hierarchical structure composed of collagen molecules, fibrils, fibre bundles, fascicles and tendon units that run parallel to the tendon’s long axis. This hierarchical structure contributes to the mechanical competence of the tendon 6

1.3 Tendon Composition

Tendons consist of collagen fibers, ground substances, water and cells. Water is about 50-

75% of tendon’s mass. 30% of dry mass of tendon are fibers and ground substances. Fibers and ground substances are collectively called extracellular matrix (ECM). Whereas Cells maintain the structural and functional integrity, vasculature provide nutrients to the cells.

The main fiber in tendon ECM is collagen fiber. 85% of tendon’s dry mass is collagen and of the collagen, about 95% is type I collagen and 5% is type III and V collagens 6. Type III collagen is found in endotenon and epitenon 16. Type III collagen is synthesized quickly during early stages of tendon wound healing and stabilizes the wound site until the beginning of tissue remodeling and synthesis of type I collagen 17, 18. Collagen type V aids to regulate fibril diameter and found intercalated into the core of collagen type I 6, 19, 20. Other types of collagen such as types II, VI, IX,

X, and XI, are also evident in tendons in very small quantities 21. These collagens are found at the bone insertion site of fibrocartilage. They are believed to strengthen the connection by reducing stress concentration at the hard tissue interface 21, 22.

Ground substances of tendon are glycoproteins, glycosaminoglycans (GAGs), and proteoglycans (PGs) 6, 23. Glycoproteins commonly found in tendon are fibronectin, tenascin-C, undulin, the thrombospondin (TSP) family, cartilage oligomeric matrix protein (COMP) 8, 24-27.

While Tenascin-C contributes to the mechanical stability of the ECM, Fibronectin is located on the collagen surfaces and plays a key role in ECM-cell interactions such as adhesion, migration, growth, and differentiation 18, 28, 29. TSP family are thought to function as adaptors and modulators of cell-matrix interactions26. COMP plays a major role in tendon development which binds collagen type I and may increase the rate of type I collagen fibril formation 30, 31.

Proteoglycans are composed of a protein core and one or more covalently bound glycosaminoglycan (GAG) chains. They are basically a class of glycoproteins. The primary PG in tendon is decorin with some other in trace quantity such as lumican, biglycan, fibromodulin, hyaluronan and others 32-34. Decorin has a protein core and one bound GAG chain. In bone, the bound GAG is chondroitin sulfate and in tendon it is dermatan sulfate 35, 36. Proteoglycan content in mature tendon is about 1-2% of the dry mass and varies depending on mechanical loading condition of tendon region 37. For example, for bovine flexor tendon, while the compression- 4 bearing region have 3-5% proteoglycan content, at tension-bearing region that content is 0.2 –

0.5% 6, 38, 39.

1.4 Tendon Mechanical Properties

Parallel collagen fiber orientation along the load bearing direction as well as the hierarchical fiber structure provides tendon’s high tensile strength capability. Along with the fiber pattern, the viscoelastic characteristics of tendon facilitate the in vivo dynamic mechanical forces exerted on tendon. A typical tendon stress–strain curve shows an initial “toe region” 14. The toe region is usually 2% of the strain where the tendon flattens the crimp pattern of a tendon 6. The crimp pattern is basically a nature’s safety factor in tendon against injury and it affects the mechanical properties of tendon. Fibers with small crimp angle fails before those with larger crimp angle. After 2% strain, if the tendon is elongated further, the stress-strain curve forms a linear region. The linear region is up to 4% strain where tendon behaves elastically. Beyond 4% strain, microscopic failure begins, even if the tendon is unloaded, it will not return to its original length. At the end of linear region tendon begins to yield macroscopic failures occur at 8-10% 40. Typical maximum tendon strain is

12-15%, stress is 50-150 MPa and Young’s Modulus is 500-1200 MPa 14, 41.

Due to the viscoelasticity tendons sensitive to strain rate. At low strain rates, tendons are more deformable, absorbs less energy and less effective in load transfer. During high strain rate, tendon becomes stiff with less deformation and more efficient in transferring load. Therefore, tendons are at high risk of failure if tension is applied quickly and obliquely.

1.5 Tendon Injuries

Tendon injury is one of the most common musculoskeletal problems in the US. For example, an estimated 30,000 to 75,000 rotator cuff tendon repairs are performed each year in the United

States, and 40% or more of patients older than 60 years old are affected by rotator cuff tendon related disorders 42, 43. Initial research in tendon tear includes improvement of surgical technique, augmentation of surgical technique by mechanically reinforcing the repair with various graft materials 42,

44-48. Despite the improvements of surgical techniques, revision of tendon tear surgery is high ranging from 11% to 94% depending on the size of the tear, patient age, tendon quality and the level of tendon degeneration49, 50.

1.6 Current Repair Techniques

Tendon does not heal naturally like regular tissue. Therefore, patient with damaged tendon initially lose range of motion, feel pain and in progression get degenerative diseases. Treatment is necessary to reduce pain and prevent against degenerative diseases. Initial treatment for tendon injury includes suture based surgical repair (direct repair), augmentation of surgical technique by mechanically reinforcing the repair with various graft materials and tissue engineering constructs and recently regenerative healing by tissue engineering.

1.6.1. Suture based surgical repair

Surgery of rotator cuff tear repair involves suturing the torn tendon to the head of the humerus either by bone tunnel, knotless fixture or anchor screws48.

Suture anchors have been introduced in 1993 by Stefen J.Snyder 46. Suture anchor surgery techniques involves suturing the torn tendon with an absorbable screw and tension with a push lock (Figure 2).

Figure 2: Using suture anchor in rotator cuff repair

Suture anchors are disadvantageous due to poor tendon-bone contact, suture breakage and insufficient pullout strength44. Anchors must be abandoned if the suture breakage occurs, which cause extra surgical complication with additional tissue damage. So to avoid anchor screw suturing

Burkhart, et al. in 1997 invented utilizing bone tunnels for suture attachment 44.

Figure 3: Using bone tunnel during surgery 51

In this method, a straight, cannulated drill guide and a drill hook is used for drilling a tunnel in the proximal of humerus. A suture hook is then pierced through the rotator cuff into the position of the drill hook slot. The suture is then pulled through the tunnel.

Suture breakage also occurs due to knot tying 44. Adjusting Knot tie is also important as it determines how much tension the repaired tendon will have after the repair. It is very hard for conventional methodology to adjust the knot tying. Also knot tie is almost impossible in arthroscopic surgery. To address this issue, a knotless fixation was invented by Ritchart et al. in

1999 47. Usually knotless suture anchors are used in this method. These are special types of screws so that suture can pass through them and a special device is used to adjust the knot tying.

Figure 4: Knotless fixation during rotator cuff surgery

1.6.2. Augmentation strategy in suture based technique

Sometimes suture based techniques are augmented with surgical mesh. Surgical mesh is used to reinforce the repair site once the repair is completed. (e.g. Polycarbonate polyurethane patch, poly-L-lactic acid-reinforced fascia patch ) 42, 45 . Some augmentation patch also supports cell repopulation, revascularization as well as act as a load sharing matrix at the repair site (e.g. 8

Conexa). Augmentation with a surgical mesh may permanently reinforce the repair and decrease failure rates.

Figure 5: A and B, Intraoperative photographs showing augmentation patch 45

In spite of all the advancement of suture based surgical technique, the revision of rotator cuff tear is still unacceptably high. This high rate of repair failure is due to the poor means of attaching tendon to bone.

Figure 6: Enthesis in tendon bone interface

The natural interface where tendon attaches to bone is called the enthesis. Enthesis is composed of mineralized and unmineralized fibrocartilage separated by a tidemark as one progresses from bone to the tendon. So naturally tendon-bone junction adjusts its change in

9 modulus gradually by enthesis as opposed to suddenly. The suture based technique doesn’t address the enthesis. So, for sudden mismatch in modulus of suture-tendon-bone is the main reason for high re-tearing rate of rotator cuff repair.

1.6.3. Biological graft based technique

Uses of biological grafts have also recently been started. They are available in three forms: autografts, allografts and xenografts. Each of these forms has its own advantages and disadvantages.

Autograft tendon is transplanted from persons own body and used for reducing repair tension between tendon-bone. This also starts natural biological process of tendon-bone junction healing by providing growth factors and growth scaffold for cell proliferation 49. However, autograft can induce donor site morbidity in addition to surgical complication and longer healing time.

Allograft tendon transplants are also performed 52. Massive rotator cuff tears are repaired with allografts of cadaveric Achilles, patellar, or quadriceps tendon. Moore et al. showed that although patient shows improved function but not as good as the usual suture based technique without the allograft 52. Out of 28 patients one patient showed infection and one patient showed acute rejection. Therefore, the problems with the allograft transplants are tissue rejection, disease transmission and availability in limited amounts.

Xenografts are usually used as an augmentation strategy in rotator cuff repair 49. As allografts have potential threat of tissue rejection, extracellular matrices from xenogenic material with no cellular component act as tissue bridge between tendon and bone. This finally acts as a

10 scaffold for aligned cellular growth and collagen assembly. 20% of patients showed inflammatory reaction with xenograft augmentation and currently not recommended 52-54.

1.7 Tendon Tissue Engineering

Although due to the advancement of research and technology, the understanding of this disease process and surgical technique has improved, the failure rates of these repairs are significantly high. Therefore, there is a recognized need to improve the current techniques or to find new strategies that can promote tendon regeneration and at the same time can augment the repair by mechanical reinforcement.

Tendon tissue engineering approach of rotator cuff repair is a more recent and promising approach. Not only it augments the repair by mechanical reinforcement, but also at the same time biologically enhances the intrinsic healing potential of the tendon by tendon regeneration rather than replacing tendons with foreign partially functionalized substitute. Tissue engineering based technique involves the use of scaffolds, growth factors and cell seeding. Scaffolds act as a temporary structure to support initial three dimensional tissue growths. It also facilitates cell proliferation then promotes matrix production and finally organize matrix into functional tendon tissues. Growth factor, cell seeding, mechanical stimulation, contact guidance is also used to further facilitate tendogenesis55.

Table 1: Scaffold devices with FDA clearance for rotator cuff repair 43

There has been a significant research and development for suitable scaffolds for rotator cuff in last decade. Extracellular matrix (ECM), synthetic polymer derived or combinations of these two scaffolds are currently available. Scaffold devices that are approved by the U.S. Food and Drug Administration (FDA) till date for rotator cuff repair in humans are shown in Table 1.

1.7.1. Synthetic Scaffolds

Synthetic scaffolds are mostly polysters like polyglycolic acid (PGA), polylactic acid

(PLA) and their copolymer polylactic-coglycolic acid (PLGA). The main advantage of synthetic scaffold is that they have good mechanical properties and hence better processability. Synthetic scaffolds that are used in tendon tissue engineering are biodegradable. Scaffolds from PLA, PGA,

PLGA, polycaprolactone, polyhydroxybutyrate has degradation products glycolic, lactic acid, acetyl coenzyme A and fatty acids are normally present in the body. Therefore these degradation end products easily metabolized and eliminated without any toxicity 56.

There a scarcity of data on the host response of synthetic scaffold and only two synthetic scaffolds in the market has been tested for host cell response 43. Derwin et al used X-Repair poly-

Llactide scaffold (Synthasome Inc, San Diego, CA, USA) for rotator cuff repair in canine model and found foreign body giant cells 57. Cole et al implanted the Biomerix RCR Patch, a polycarbonate polyurethane scaffold (Biomerix Corp, Fremont, CA, USA) into a rat model of acute rotator cuff repair and found there was an 80% average infiltration with host connective tissue with almost no evidence of inflammation 58. But in general synthetic scaffolds shows acute inflammation followed by chronic inflammation if the material is not biodegradable 56, 59.

Several studies have been done on the mechanical properties and tendenogenesis capability of synthetic scaffold. Wei et al. showed unwoven PGA scaffold degrade more slowly and have better mechanical property than the woven scaffolds 60. Ouyang et al. showed that knitted PLGA scaffolds augmented tendenogenesis and contributed to better mechanical properties 61.

Although synthetic polymers show better mechanical properties, can be tailored to be biodegradable and promote tendon regeneration, it has some disadvantages. Due to the hydrophobic nature, they do not show good cell adhesion 62, 63. The degradation materials are acidic hence high presence of this end product can cause systematic or local reactions 64, 65.

1.7.2. ECM Derived Scaffolds

ECM derived scaffolds are mainly collagen derivatives. As the tendon ECMs are mainly composed of type I collagen, these scaffolds are highly biocompatible and show better cell adhesion and cell proliferation.

Host response is one of the most important criteria for scaffolds. Presence of cell has high impact on the scaffold host response. If the scaffolds are from xenogenic sources, the cells and cellular remnants should be removed. Human dermal matrix with cells and cell remnants showed immune cell infiltration, tumor necrosis factor-α deposition and macrophage activation 66.

Acellular human dermal matrix showed minimal or no inflammation with active host response 66.

Laboratory made pure Collagen gel, Collagen sponge and collagen fiber don’t have the issue with cell removal but they possess insufficient mechanical strength. Collagen gels in combination with polyglyconpolyglyconate suture showed better biomechanical properties but inferior to uninjured tendons 67. Study showed that gel–collagen sponge constructs could greatly enhance the development of functional tendon tissue 68, 69.

Although collagen gel with collagen fiber or collagen sponge shows promising strategy in terms of superior cell seeding efficiency and biofunctionality, ECM derived and pure collagen based scaffolds have some inherent limitations. They have limited process ability with less mechanical properties which make them vulnerable to suture retention properties. Also batch-to- batch variation in collagen constructs makes reliable reproduction of these scaffolds difficult.

1.7.3. Cells Used in Tendon Tissue Engineering:

Tendon derived fibroblasts (TDFs) 70, 71, skin derived fibroblasts(DFs) 72-74, MSCs (tendon- adipose- or marrow-derived) 61, 67, 69, 71, 74-78 and embryonic stem cells 79 have been assessed for tissue engineering based repair of tendons. TDFs obtained from autologous tendon biopsies seem to be obvious choice. However, tenoblast or tenocytes are not available in abundance and they may de-differentiate 72, 80-83. Furthermore, it increases tissue morbidity at the harvesting site. The multipotency and common origin of progenitor cell make them vulnerable to form ectopic bone 14 and cartilage in the repair site 84. Bone marrow stem cells (BMSC) have been mostly used in tendon tissue engineering. Comparative study between TDFs,DFs and BMSCs showed that BMSC was most promising followed by DFs 74, 85. When seeded at a high density, MSCs may result in ectopic calcification 67, 86; however, reducing the cell-seeding density mitigates the problem 87. Embryonic stem cells (ESCs) have not been assessed rigorously for tendon repair. One study reported tendon- like tissue formation when ESC-seeded scaffolds were implanted in nude mice 79. However, some scaffolds displayed ossification. The optimal conditions to attain tenocytic differentiation of ESCs are unknown. Another limitation with ESCs is the risk of teratoma formation. Recently adipose derived stem cells (ASCs) has been started to use 88-90 . ASCs with or without growth factor has been shown to differentiate in tenocyte 89, 90. ASCs can be collected at large amount with less hassle and donor site morbidity. Therefore, ASCs has high potential in tendon tissue engineering.

Overall, MSCs seem to be the most feasible cell-populations for tissue engineering based repair of tendon.

1.7.4. Growth Factors (GFs) Used in Tendon Tissue Engineering:

GFs are biological agents which can help to increase differentiation of MSCs to tenocytes as well as matrix synthesis by cells. They also prevent de-differentiation of tenocytes/fibroblasts.

As there is no GF which showed ultimate efficacy in tendon tissue engineering, researchers have been trying different GFs in tendon repair strategies. GFs involve in tendon tissue engineering are

BMP-12/GDF-7, BMP-13/GDF-6, BMP-14/GDF-5, FGF-2, CTGF, TGF-ß, PDGFbb and IGF-1

91-98. Bone marrow stromal cells showed tenomodulin expression or in some case formation in the presence of GDF6, GDF7, BMP 2, BMP 12, PD98059 99-106. Adipose-derived mesenchymal stem cells (ADMSCs) also showed tenomodulin expression in the presence of GDF589, 107. However, there are several significant drawbacks associated with growth factors: 1) risk of non-tenocytic

15 differentiation 108-110, 2) risk of tumor formation in some cases 111, 3) short half-life 112, 113, 4) manufacturing costs (e.g. recombinant production followed by purification). All these concerns make it lengthy and very costly to get regulatory approval for growth factors and to this date none of the growth factors stated in the above are approved by the FDA for tendon regeneration.

Therefore, a regenerative solution which circumvents growth factors would be highly significant.

1.8 Collagen as a Bioscaffold

Tendons are primarily consisting of type I and type III collagenous fibril. Out of these collagenous fibril 95 % are type I collagen and the biomechanical function of type I collagen in tendon is to resist tensile load. Therefore, type I collagen rich scaffolds are the finest candidates for tendon repair. Laboratory made pure Collagen gel, Collagen sponge and collagen fiber are promising strategies in terms of superior cell seeding efficiency and biofunctionality, but they possess insufficient mechanical strength and due to this, limited process ability. The inferior mechanical strength of collagenous scaffold is due to two reasons. First, in native tendon, type I collagen fibrils are uniformly oriented along the longer axis114 whereas in collagen scaffolds formerly describe has randomly oriented collagen fibrils115, 116. Secondly, in native tendon, collagen is densely packed giving the higher stiffness but in collagen gel or sponge, there is limited packing density of collagen molecules. Overall, in native tendon collagen fibrils are densely packed and uniformly aligned but current collagen derived scaffolds lack both of these criteria.

Therefore, there is a rationale need to align collagen molecule in the venue of collagenous scaffolds.

Several means such as magnetic fields, mechanical stretching, extrusion and electrospinning have been employed to align collagen molecules. Magnetic fields transversely

16 orient collagen molecules 117-121. However, this requires expensive super conductive magnets on the tesla-order strength. Mechanical means of orienting collagen by stretching provides some degree of alignment, but spatially inhomogeneous structures are still present 122. Collagen gels extruded through orifices generates thread like structure. However, it has recently been shown that the alignment of molecules during extrusion is limited to the surface and the core of extruded fibers lack alignment 123. Recently electrospinning has been used to generate aligned collagen scaffolds

124, 125. However, electrospinning denatures collagen molecules 125. Therefore, none of the aforementioned techniques was able to produce mechanically strong, aligned and densely packed collagen scaffold in a cost effective way. Dr. Akkus’ lab developed a method of aligning collagen molecules by electrochemical compaction (ELAC) which generates a biomaterial that meets the needs for an ideal biomaterial of tendon tissue engineering scaffold.

1.9 Electrochemically Aligned Collagen (ELAC) for Tendon Repair 126-133

Summary of the studies on Electrochemically Aligned Collagen (ELAC) technology:

Monomeric collagen solutions were converted to electrochemically aligned collagen

(ELAC) threads by a novel biofabrication method. Fabric orientation, mechanical properties and packing density of ELAC matches those of the native tendon 126. Proper crosslinking of ELAC threads improved the failure strength (110 MPa) and modulus (0.9 GPa) to the level of tendon 131.

MSCs seeded on ELAC thread goes under tenogenic differentiation due to ELACs topographical cues 128. In vivo study of cross linked ELAC showed limited inflammation after 4 months and

ELAC was mostly absorbed after 8 months130. New preliminary data demonstrates our ability to deposit patterned lattice structures by employing computer aided design and manufacturing.

1.9.1. Fabrication and physical characteristics of ELAC 126:

Figure 7: A) Schematic representation of the monomeric collagen to solid ELAC B) Compensated polarized images of tendon and ELAC fibers. Molecular alignment is manifested by blue color when alignment is parallel to fast axis of the retardation plate (double headed arrow) (scale bar 200 microns).

At first acid solubilized monomeric collagen solution was dialyzed. The dialyzed solution was applied between two parallel electrodes using a syringe. After the application of electrical current, collagen molecules become aligned and packed along the longer axis of the electrode and

ELAC is formed. ELAC is then treated with PBS and genipin crosslinking to induce d-banding and elevate strength respectively. The mechanism behind ELAC formation can be described as: a pH gradient electrodes (acidic at anode to basic at cathode) is formed between the electrodes as soon as the voltage applied. As Collagen is an ampholytic molecule, the net charge of it becomes negative near cathode and positive near anode which causes repulsing collagen molecule from both of the electrodes. The repulsed collagen molecules migrate and accumulate towards the isoelectric point (pI) between the electrodes where they have zero net charge (Figure 7). ELAC formed in this way are in the form of thread around 200-800 micron diameter and has packing density 1030 mg/mL (one of the highest that has been reported to date 134) whereas the solution it made from has collagen density 3 mg/mL. Compensated polarized microscopy, high resolution scanning electron microscopy (SEM), transmission electron microscopy (TEM) and quantitative

18 small-angle X-ray spectroscopy (SAXS) showed that the degree of collagen alignment within

ELAC threads was comparable to the native tendon 126, 132. The advantages associated with the electrochemical process are: (1) occurs in aqueous environment (no toxic solvents as compared to electrospinning); (2) low cost (stainless steel or carbon electrodes, power supply); (3) capability to fabricate intricate patterns by improvising on electrodes as arrays, (4) Electrochemically aligned threads can be induced by D-banding by PBS treatment 126, 132, indicating that molecules are not denatured during processing, 5) ability to accommodate other matrix molecules as additives (such as glycosaminoglycans) 129.

1.9.2. Mechanical properties of ELAC threads 131:

Tendon, depending on anatomical location and species, has a failure stress of 60-100 MPa and elastic modulus of 500-1000 MPa 67, 135-138. Genipin crosslinking conditions can be optimized by modulating the type of solvent, genipin concentration and crosslinking duration, to reach a failure stress range of 80-110 MPa (Figure 8A) and modulus range of 600-900 MPa (Figure 8C)131.

Crosslinked ELAC threads fail in the range of 10% - 15% strain which is also in accordance with tendons (Figure 8B) 131. Therefore, ELAC provides the cells with a substrate that is not only topographically similar to tendon, but also mechanically similar to tendon.

Figure 8: Mechanical Properties of ELAC threads optimized using different crosslinking treatments. (A) Wet Stress, (B) Wet Strain, and (C) Wet Modulus. Black bar: 0.625% genipin in 1x PBS; White bar: 0.625% genipin in 90% ethanol; Gray bar: 2% genipin in 90% ethanol. 19

1.9.3. Differentiation and in vitro response of MSCs to ELAC 127 128:

In vitro studies showed that MSCs were able to attach, migrate and invaginate between braided ELAC threads. Human MSCs were seeded on ELAC and cultured under conditions conducive to matrix production by inclusion of ascorbic acid (50 µg/mL) in the culture medium for four weeks. MSCs formed about 100 micron thick white colored cell-matrix layer over the

ELAC threads (Figure 9A) where the cell-matrix layer was oriented along the longer axis of ELAC threads. Importantly, cellular cytoskeleton was oriented along the longer axis of ELAC fibers

(Figure 9B).

Differentiation of human MSCs on ELAC threads in the absence growth factors was also assessed. Human MSCs (Lonza, MD) were seeded on ELAC threads and random collagen strips and the effect of collagen orientation on tenogenic differentiation was investigated using real-time

PCR. The expression of an early stage tendon-specific marker, scleraxis, and a later stage tendon- specific marker, tenomodulin, were significantly greater on ELAC on days 3 and 14, respectively

(Figure 10A and 10B). Tendon-related markers (tenascin-C and collagen-III) were significantly higher on ELAC threads compared to random collagen threads. Osteocalcin, a specific marker for bone differentiation was significantly suppressed on ELAC threads (Figure 10C). These results

20 demonstrate that ELAC threads stimulate MSC differentiation specifically towards the tendon lineage without growth factors. Implicit in these results is the relative effects of matrix stiffness and matrix alignment. Proposed studies will elucidate the effect of these variables on differentiation further.

Figure 10: Tenogenic differentiation of human MSCs on ELAC (fold change with respect to randomly oriented collagen). (A, B) Tendon specific genes – Scleraxis (A) and Tenomodulin (B), (C) Bone specific genes (Runx2 and Osteocalcin). Black bar: Random; White bar: ELAC. (* indicates statistical significance, p < 0.05)

1.9.4. In vivo biocompatibility and degradation of ELAC in a Rabbit patellar tendon model 130:

ELAC bioscaffolds were inlaid into the patellar tendon of rabbits Implant degradation and local response were assessed. ELAC was mostly absorbed by 8 months. A mild inflammation was observed around the ELAC threads at 4 months (comparable to that around standard sutures, no giant cells, foreign body encapsulation or presence of neutrophils) which was resolved by 8 months and replaced by de novo fibrous tissue that appeared to be indistinguishable from the tendon proper histologically 130. Continued presence of ELAC up to 4 months is advantageous as tendon is a slowly healing tissue 67, 139 and prolonged support of ELAC would therefore be desirable. Baseline in vivo assessment indicated that ELAC is biocompatible and biodegradable.

1.10 Matrix Stiffness and Topography:

Tendons consist of parallel collagenous fibril along the longer axis of the tendon. This gives tendon tissue excellent tensile force bearing capacity along the direction of the fibrils. The elongated size of tenoblast ( L= 20 – 60μm; W= 8 – 20μm) may be due to the anisotropic nature of tendon. It has been showed that matrix stiffness and topography guides cellular morphology, phenotype and differentiation pathway of mesenchymal stem cells (MSCs). This phenomena is due to the fact that cell can sense the rigidity of the matrix by focal adhesion force140. Cell senses rigidity or anisotropy (if there is any) of the matrix and this synergistically affects the mechano chemical signaling between cell and the matrix141. This results in the change of cell morphology and the commitment of cell fate. Cell generates nanoscale level forces within single mature integrin-based focal adhesions (FAs). Individual FAs apply repeatedly tugging force to locally sense the ECM stiffness. FAs in soft substrate/soft direction give softer feedback to the cells and cells fail to assemble a cytoskeleton rich in polymerized actin. On the other hand FAs in stiff substrate/ stiff direction offer rigid feed back to the cell and this result in actin polymerization140,

142.

Aligned topography of matrix resulted elongated and aligned cells in the direction of alignment. For example, aligned electrochemically compacted collagen fiber, electrospun polyurethane (PU), silk fibroin, poly (l-lactic acid) and two-dimensional (2D) microgrooved surface showed elongated cell morphology and tendon related ECM formation128, 143-149. Soft substrate favors MSC differentiation into neuronal-like cells, moderate elasticity promoted myogenic differentiation, and a rigid matrix stimulated osteogenic differentiation 150. Substrate stiffness ~ 0.5-1kPa (Collagen I gel, Collagen-Glycosaminoglycan) showed chodrogenic differentiation 151, 152. Substrate stiffness 40 kPa (Collagen I coated PMA) showed tenogenic differentiation 104.While there are disparate report on the effects of alignment and matrix stiffness 22 separately, the synergy between the two in terms of inducing tenogenic differentiation is not studied.

1.11 Ideal Scaffold for Tendon Tissue Engineering

From the above literature review, it is clear that there is a scarcity of methods which can yield strong, anisotropic and densely packed ideal collagen scaffolds in a cost effective fashion.

Studies by our laboratory on electrochemical processing of collagen molecules during the past 8 years resulted in a biomaterial that meets these needs.

An ideal scaffold would require certain characteristics to work auspiciously for tendon regeneration. It should be biocompatible, biodegradable as well as biofunctional. For cell proliferation and migration, it should be porous enough and should be mechanically strong enough for tissue regeneration after implantation. Finally, to acknowledge the issue of batch-to-batch variation, it also should have reliable reproducibility. Moreover, the material should also consider stiffness as well as stiffness anisotropy to mimic two key features of natural tendon.

An altered method similar to ELAC fabrication was developed in this study to fabricate patterned lattice structures with controlled pore size and morphology. The inability to incorporate high strength and high porosity in a structure has been one of the major barriers in the engineering of load-bearing tissue 153. This method utilizes computerized scaffold design and fabrication which allows the integration of ‘scaffolds with controlled macroporosity’, an essential feature for populating a scaffold with cells and vasculature. There is a general lack of biofabrication methods that will provide controlled interconnected macroporosity. The cutting edge patterned electrochemical deposition method will address this challenge at the fundamental level.

2. Chapter 2: Research Objectives

2.1 Introduction

This dissertation describes a novel biomimetic scaffold design for tendon tissue engineering via micropatterned deposition of collagen. As a part of the scaffold design, this study also investigated the possibility of a suitable material model which mimic the tendon topography as well as alignment and hence favors tenogenesis.

Previous studies by this laboratory demonstrated that ELAC is biocompatible and favors tenogenesis. However, the micropatterned deposition uses planar electrode instead of liner electrode as in ELAC and generates highly dense collagen sheet like structure 132, 154. It can be speculated that this compact collagen substrate would be biocompatible as ELAC as similar technique, material and chemical post processing was used to fabricate them. However, the capability of tenogenesis needs to be evaluated for this highly compact collagen substrate. While doing so, this dissertation investigated two fundamental research questions such as effect of stiffness anisotropy and stiffness on tenogenesis.

2.2 Aim 1: Assess the effect of stiffness anisotropy of ELAC substrate on the cytoskeletal

behavior and tenogenesis

Sub-Aim 1.1: Assess the effect of stiffness anisotropy on cellular and nuclear morphology

Most natural tissues are substantially stronger along the load bearing direction than the direction transverse to the longer axis (such as tendons, muscle etc.). Notably, the cells of such tissues are elongated along the stiffer direction. In a single cell context, each cell will sense different stiffness in longitudinal and transverse direction. This will eventually effect the cell

24 behavior e.g. cell alignment or even may be on cell fate. Therefore, it is important to assess the effect of stiffness anisotropy of ELAC sheet on cytoskeletal behavior.

Sub-Aim 1.2: Assess the effect of stiffness anisotropy on stem cell fate towards tenogenesis

There are not many biomaterials to emulate and study the effects of stiffness anisotropy on cellular response. MSCs seeded on fully aligned ELAC undergo tenogenic differentiation via topographic cues and gets elongated along the length of ELAC 128. However, the roles of topographical factors (matrix anisotropy vs. matrix stiffness) in inducing the observed tenogenic differentiation are unknown. Therefore, a controlled method of generating a biomaterial with tunable stiffness anisotropy is needed to better understand the cellular fate in anisotropic tissues.

This study will report the effects of a controlled unidirectional stretch process on the stiffness/strength anisotropy of collagen substrates. In these activities, MSCs will be seeded on aligned collagen sheets at increasing stiffness anisotropy values. The tenocytic differentiation and matrix synthesis by MSCs on different stiffness anisotropy collagen sheets will be assessed.

2.3 Aim 2: Increase tenogenic differentiation on electrocompacted collagen substrates.

Sub-Aim 2.1: Assess effects of matrix topography (matrix stiffness magnitude and matrix alignment) on the differentiation of MSCs

Extracellular matrix stiffness or mechanics plays an important role to regulate cell morphology, proliferation, differentiation during regular and diseased states. There has been relatively little investigation of this phenomenon with respect to tenogenesis. Morever, the roles of topographical factors (matrix anisotropy vs. matrix stiffness) in inducing the observed tenogenic differentiation are unknown. In this study, MSCs will be seeded on random collagen sheet at

25 increasing stiffness values and aligned collagen sheets a fixed stiffness values. Notably, collagen sheets can be crosslinked to encompass four orders of magnitude of Young’s modulus (0.1 MPa to 100 MPa). The tenocytic differentiation and matrix synthesis by MSCs on aligned collagen sheets will be assessed to identify the conditions which maximize the differentiation.

Sub-Aim 2.2: Assess effects of dermatan sulfate incorporation on the differentiation of MSCs.

Based on the result of Sub-Aim 1.1, a potent topography will be supplemented with dermatan sulfate (DS), a glycosaminoglycan richly present in tendon to enhance the topographical differentiation cues with compositional cues.

2.4 Aim 3: Construct a biomimetic tendon repair scaffold

A novel micropatterned deposition method will be employed to construct a 3-D scaffold from the biomaterial formulation developed in Aim 1. Computer Aided Design (CAD) and fabrication will be used to generate patterned electrode pairs to transform collagen solutions to solid patterned lattice layers.

Sub-Aim 3.1: Improve mechanical function of scaffolds.

The studies will vary the microarchitecture of lattice patterns to obtain mechanically competent scaffolds with interconnected porosity. These variables will include filament area, pore size, filament angles and deposition pattern (linear filaments vs. crimp like sinusoidal filaments).

Due to the high number of variables, these combinations will be assessed computationally in the first step via finite element analysis to determine variables which provides closer stiffness (N/mm) values of rabbit infraspinatus tendon with maximum porosity. The deposition patterns favored by computational analysis will then be fabricated to confirm the outcome by mechanical tests.

Sub-Aim 3.2: Assess in vitro response of cells in 3D-scaffolds.

The interconnected porosity allows populating scaffold with cells expeditiously and also facilitates nutrient diffusion. The second phase will address the extent to which these benefits occur. In these experiments, micropatterned 3D-scaffolds will be populated with MSCs and the cell population will be assessed.

3. Chapter 3: Collagen Substrate Stiffness Anisotropy Affects Cellular Elongation, Nuclear Shape and Stem Cell Fate towards Anisotropic Tissue Lineage

3.1 Abstract

Rigidity of substrates plays an important role in stem cell fate. Studies were commonly carried out on isotropically stiff substrate or substrates with unidirectional stiffness gradients.

However, many native tissues are anisotropically stiff and it is unknown whether controlled presentation of stiff and compliant material axes on the same substrate governs cytoskeletal and nuclear morphology, as well as stem cell differentiation. In this study, electrocompacted collagen sheets were stretched to varying degrees to tune the stiffness anisotropy (SA) in the range of 1 to

8, resulting in stiff and compliant material axes orthogonal to each other. The cytoskeletal aspect ratio increased with increasing SA by about 4-fold. Such elongation was absent on cellulose acetate replicas of aligned collagen surfaces indicating that the elongation was not driven by surface topography. Mesenchymal stem cells (MSCs) seeded on varying anisotropy sheets displayed a dose-dependent upregulation of tendon-related markers such as Mohawk and

Scleraxis. After 21 days of culture, highly anisotropic sheets induced greater levels of production of type-I, type-III collagen and thrombospondin-4. Therefore, SA has direct effects on MSC differentiation. These findings may also have ramifications of stem cell fate on other anisotropically stiff tissues, such as skeletal/cardiac muscles, ligaments and bone.

3.2 Introduction

A number of natural tissues such as tendons, blood vessels and muscles are substantially stiffer along their load bearing direction than other tissue axes. Notably, the cytoskeletons and nuclei of cells in such tissues are generally elongated along the stiffer direction. In such a fundamental context, the effects of extracellular matrix (ECM) stiffness anisotropy (SA) on cellular morphology and response have not been investigated. Prior research has demonstrated the effects of extracellular matrix rigidity on morphology, proliferation and differentiation of mesenchymal stem cells (MSCs) 150, 155-160. Such studies were commonly carried out on environments with isotropic stiffness. Depending on matrix rigidity, MSCs may undergo neurogenesis, myogenesis or osteogenesis 150. Matrix rigidity has also shown to affect the degree to which a cell spreads 161-163.

It has also been shown that cell migration can be controlled by stiffness gradients in a process termed as durotaxis 140.

Alignment and elongation of the cytoskeleton have been achieved predominantly by topographical cues such as micropatterned ridges, micropillars, gratings, wells or microimprinted cell-adhesive patterns 155-160, 164-166. There is evidence that cells seeded on nanofibrous topographies align along the longer axes of fibers 167-169. This outcome is mainly due to the defined topography of the substrates. It was demonstrated that topography induced cytoskeletal alignment differentiated MSCs to tenogenic and neuronal differentiation 128, 170. It is yet unknown whether controlled presentation of substrate SA governs cytoskeletal and nuclear morphology, as well as stem cell fate.

Type I collagen is the most abundant and major structural protein in connective tissues 171,

172. Therefore, it is the center of many tissue engineering strategies for anisotropic tissues such as tendon, ligament, etc. To the best of our knowledge there is no study which introduces SA directly

29 to the pure collagen substrate by incorporating different stiffness values in different material directions and studies the effect on cytoskeletal response and differentiation toward anisotropic tissue lineage. Our research group demonstrated transformation of monomeric collagen solutions to solid phase by electrochemical gradients induced by linear electrodes 126, 173, 174. The method generates highly dense and aligned collagen threads with a packing density (1030 mg/ml) one of the highest reported to date 132, 154. The molecular alignment lacks when planar electrodes are used to fabricate sheets. In the current study, SA was introduced in electrocompacted sheets by controlled unidirectional stretch of the sheets to various levels. Variations in the cytoskeletal and nuclear morphology of cells seeded on collagen sheets of varying SA were studied. Effects of collagen SA on the expression of tendon-related transcription factors and tendon-related extracellular matrix synthesis were also investigated.

3.3 Materials and Methods:

3.3.1. Fabrication of Collagen Sheets by Electrochemical Compaction [Figure 11A]:

Bovine dermis derived acid soluble monomeric type-I collagen solution (Advanced

Biomatrix, San Diego, CA; 6 mg/ml) was diluted two-fold using RNAase/DNAase free water. The pH of the collagen solution was adjusted to 8-10 using 1 N NaOH and dialyzed against ultrapure water for 18 hours to prepare the collagen solution for electrocompaction.

Electrocompaction of collagen in the sheet form was carried out as described before 173, 174.

Briefly, a rectangular window of 30x10x1.5 mm was cut in a plastic piece. The plastic window was placed on planar carbon electrode and filled with the dialyzed collagen solution. Another plane carbon electrode was placed above the plastic boundary, sandwiching the collagen solution between the electrodes between which 30 VDC was applied for 2 min. Electric current electrophoretically mobilizes collagen molecules and compacts them under the effects of

30 mechanisms detailed in an earlier publication 175. This electrocompaction generates 100 microns thick rectangular sheets of 30x10 mm dimensions between the two planar electrodes.

3.3.2.Resolving the effects of topography from the effects of SA by cellulose acetate replication of surface topography

The topography of a highly stretched collagen sheet (SA=8) was replicated using a cellulose acetate sheet to assess whether stretch induced surface topographical alignment induce cell alignment and elongation. Briefly, a thin film of cellulose acetate sheet is wetted with acetone, placed on top of the highly stretched collagen sheet and kept for 5 minutes. The surface topographical pattern of the collagen sheet is replicated through the flow of solubilized the cellulose molecules. The replicated cellulose acetate samples were then peeled off and washed with deionized water to remove any trace of acetone.

3.3.3. Tuning of Molecular Alignment via Mechanical Stretch:

A motorized mechanical device was built to align collagen molecules by stretching (Figure

11B). Randomly oriented collagen sheet was gripped in the device and stretched at a translational speed of 35µ/s to 20%, 40% and 60 % of their initial length of 20 mm to induce different levels of

SA.

(A)

(B)

Figure 11: A) Overview of electrochemical compaction of collagen sheets, B) Overview of molecular alignment by stretching.

3.3.4. Imaging of Molecular Alignment in Compacted Collagen Sheets

The degree of alignment of collagen molecules at different levels of stretching was assessed by a polarized optical microscope equipped with a first order wavelength gypsum plate (Olympus

BX51, Melville, NY, USA) and by using SEM.

Collagen is a positive birefringent material and the aligned molecules along the slower axis of the gypsum plate shows blue interference color and the molecules which are perpendicular to the slow axis appear yellow 176. Magenta color indicates lack of alignment and emergence of blue in the CPI image indicates molecular alignment in the SW-NE direction or along the slower axis or perpendicular to the long dimension of the gypsum plate.

The surface morphology of collagen and cellulose acetate samples were observed by a scanning electron microscope (FEI Nova Nanolab 200, Hillsboro, Oregon) at a voltage of 3 kV and a beam current of 0.12 nA. Before analyzing the samples with SEM, all the samples were sputter coated with 5 nm thick layer of palladium (Denton Desk IV Coater DCH240).

3.3.5. Assessment of Mechanical Properties of Stretched Sheets (SA):

Treatment groups were: i) Unstretched, ii) 20% stretched, iii) 40% stretch, iv) 60% stretch.

Collagen sheets were cut into 20 x 2 mm strips along the stretch direction (Longitudinal) or transversely to the stretch direction (Transverse) (n = 6-8 samples per group). Samples were hydrated in deionized water for 30 minutes and then they were tested under monotonic tension

(Rheometrics Inc., NJ) until failure at a strain rate of 10 mm/min. Cross sectional areas of sheet samples were measured with a multi-photon confocal microscope (Leica TCS SP2, Wetzlar,

Germany) in the hydrated state. Stress-strain curves were constructed from the load-displacement data using sample geometry. Moduli along the longitudinal (EL) and transverse (ET) directions 33 were calculated from the slopes of the linear regions of stress strain curves. SA was defined by dividing the modulus in the longitudinal by the modulus in the transverse directions:

EL SA  (1) ET

Similarly, stress anisotropy (STA) was defined as:

 L STA  (2) T

Where, σL and σT is defined as the maximum failure stress.

3.3.6. Surface Roughness Measurement by Atomic Force Microscopy (AFM):

To check the effect of SA on surface roughness, AFM was performed on SA=1 and

SA=8. Surface roughness was measured using Atomic Force Microscopy (AFM). AFM was performed using Veeco Dimension 3100 (Bruker, MA, USA) at room temperature in tapping mode with typical anisotropic AFM probes. Images were taken in several locations of each samples (N= 6-8 repeat measurements/group) with 20 X 20 µm image size. The quantitative measurement was performed by Nanoscope 6.5.3 software accompanied by the AFM system.

3.3.7. Cell Viability after day 1

Cell viability study was performed at day 1 to check whether introducing stiffness anisotropy in electrocompacted collagen sheet affects the cell viability. LIVE/DEAD®

Viability/Cytotoxicity Kit (ThermoFisher Scientific, MA, USA) was used according to manufacturer’s instruction. In brief, 10 µL of 2mM ethidium bromide and 2.5 µL of 4mM Calcein was mixed in 5mL sterile 1X PBS and added on the cell cultured samples. Samples were incubated 34 in these reagents for 30 min at room temperature. Following incubation, samples were examined and photographed. Cell viability counting was performed using ImageJ.

3.3.8. Effect of SA on Cytoskeletal and Nuclear Morphologies

Cells were seeded on sheets stretched to anisotropy levels of SA = 1 (isotropic), SA = 3 and SA = 8. Prior to cell seeding, samples were disinfected in 70% ethanol for 4 hours and washed in 1X PBS. Samples (n = 3 wells/group) were placed into ultralow attachment 24 well plates

(Corning). Human MSCs (Lonza, Walkersville, MD) at passage 5 were seeded at a density of 5000 cells/cm2. The culture medium composed of alpha MEM (Invitrogen) supplemented with 10%

MSC-Qualified FBS (Invitrogen), 1% penicillin/streptomycin and 50 μg/mL ascorbic acid.

Cell morphology was visualized by staining cytoskeletal actin filaments with AlexaFluor

488 Phalloidin (Life Technologies, Grand Island, NY, USA) at 6 and 12 hour time points. Cells were fixed with 3% formaldehyde (with 0.1% TritonX-100) for 10 min followed 1x PBS wash.

The actin filaments were stained by incubating the cells in AlexaFluor 488 Phalloidin at 37 °C for

20 min. The stain was washed with 1x PBS and images of stained cells were taken using an

Olympus Microscope (Olympus IX83) with a 20X objective lens. Images were taken from randomly selected fields of view (N=3-5 repeat measurements/sample). Cell nuclei were visualized by DAPI nucleic acid stain (Invitrogen). Briefly the stock solution was diluted to 300 nM in 1x PBS and added to the sample wells. Samples were incubated for 3 minutes for nucleic acid staining. The staining was washed with 1x PBS and images were taken as described for actin staining within the same field of view as actin staining. For actin staining imaging, EGFP filter

(486/509 nm wave length) and for DAPI staining image DAPI filter (358/461 nm wavelength) was used.

3.3.9. Cell seeding on the replicated cellulose acetate film

Cells were seeded on the cellulose acetate film with the topography replication and without the topography replication (control) in similar method as described above. Cells were also seeded on collagen coated cellulose acetate sheet. Cellulose acetate samples were functionalized with

Type I collagen with a stock solution of 50 µg/ml applied at 6 µg/cm2 area. Collagen-I surface coating was verified by Picro Sirius Red staining. Cell morphology was visualized by staining cytoskeletal actin filaments using AlexaFluor 488 Phalloidin (Life Technologies, Grand Island,

NY, USA) after 6 hours.

3.3.10. Measurement of Cytoskeletal and Nucleus Morphologies

Cellular and nuclear morphologies were measured by ImageJ (NIH, MD, USA). An ellipse was fitted around each cell. The diameters of the major and the minor axes of the ellipse were recorded. The angle between the stretching direction and the major axis was defined as the cellular alignment angle Ɵ. Cell aspect ratio (r) was defined by the ratio of the diameters of the major axis to the minor axis.

3.3.11. Effect of SA on Tenogenic Differentiation of Human MSCs

Cells were seeded on electrocompacted collagen sheets which were stretched to anisotropy levels of SA = 1 (isotropic), SA = 3 and SA = 8 as elucidated earlier. Additionally, a random collagen gel group was included for this experiment which provides an SA of 1; however, collagen gel is less compliant than unstretched electrocompacted collagen sheet. Therefore, collagen gel enabled to sort out the effects of substrate stiffness. Collagen gel was synthesized by mixing acid soluble monomeric collagen solution with 10x PBS at a ratio of 8:1 parts and the pH was adjusted to 7.4 using 0.1 N NaOH. The neutral collagen solution was poured into the wells of a 6 well plate

36 and gelled by incubating at 37 °C for 2h. At time points 1, 3 and 5 days total RNA was extracted by lysing the sells using TRIZOL reagent (Invitrogen) following manufacturer’s protocol. Briefly, chloroform was added to the trizol homogenized samples and the phase separation was performed by centrifuging the samples at 12,000 g for 15 min at 4 ºC. The RNA was collected in a separate tube from the supernatant aqueous phase, precipitated by adding isopropanol and pelleted by centrifuging at 12,000 g for 10 min at 4 °C. 70% ethanol was used to wash the RNA pellet. The pellet was then dried and resuspended in RNAse/DNAse free water (Invitrogen) and stored at -80

°C. 2 µg of total RNA was reversed transcripted to synthesize cDNA using the cDNA Reverse

Transcription Kit (Applied Biosystems). Taqman real time PCR mastermix (Applied Biosystems) and Taqman gene expression assays (Applied Biosystems) for early tendon specific or related genes (Scleraxis 89, 99, 177-181 and Mohawk 182-188) were used to evaluate the expression of the genes by quantitative real time PCR (Applied Biosystems 7500 Real Time PCR System). The relative fold change in the target gene expression was quantified using 2 deltadeltaCt method by normalizing the target gene expression to RPLP0 169 as a housekeeping gene and relative to the expression on the random collagen gel at each time point.

3.3.12. Effect of SA on Tendon Related Matrix Synthesis

Cells were seeded on electrocompacted collagen sheets at anisotropy levels of SA = 1

(isotropic), SA = 3 and SA = 8 as described above. Cells were cultured for 21 days as described earlier. Three tendon-related extracellular matrix molecules were investigated: i) type-I collagen, ii) type-III collagen and iii) Thrombospondin-4 (TSP-4). For type-I and type-III collagen, immunohistochemistry was performed and for TSP-4 green immunofluorescence was performed.

Collagen sheet samples were fixed in 10% neutral buffered formalin, rinsed in 3 washes of

PBS for 5 min each, permeabilized for 15 min in PBS containing 0.25% Triton X-100 followed 37 by 3 washes in PBS for 5 min each. After blocking the samples in PBST (Phosphate Buffered

Saline with Tween) containing 5% BSA, 22.52 mg/ml glycine and 0.1% Tween 20 for 30 min, the samples were incubated with primary antibody overnight at 4°C. The following antibodies were adopted for immunohistochemistry: mouse Anti-Col1A1 (Abcam), rabbit anti-Col III (Abcam),

Thrombospondin 4 (Santa-Cruz). Samples incubated with blocking solution without primary antibody were used as negative control for the secondary antibody, and collagen sheet without cells were used as background negative control. Staining was performed by Alkaline phosphate substrate-chromogen using StayRed/AP kit (Abcam) according to manufacturer’s recommendations. Briefly, the samples were washed 3 times in PBS for 5 min each after overnight incubation with primary antibodies. Following the washes, the samples were incubated with a secondary ALP-conjugated antibody for an hour at room temperature. The samples were washed

3 times in PBS for 5 min each and incubated with StayRed/AP working solution (3 ml of

StayRed/AP Substrate buffer containing one drop of StayRed/AP chromogen) for 15 min at room temperature. Samples were rinsed in PBS and imaged (N=3-5 repeat measurements/sample) using a microscope (Olympus IX83 digital microscope, Olympus Life Science). Images were processed and quantified by Cellsens Dimension software (Olympus Life Science).

For immunofluorescence, the anti-Rabbit Alexa-488 conjugated specific secondary antibody (Pierce Protein Biology, Thermo fisher Scientific) was used. The samples were visualized

(N=3-5 repeat measurements/sample) under Olympus IX83 digital microscope (Olympus Life

Science) using Cellsens Dimension software (Olympus Life Science)

Quantification of the protein staining was performed by ImageJ (NIH, Maryland, USA).

The amount of type-I and type-III collagen staining was calculated by ImageJ colour thresholding

189. The regions of interest were chosen randomly across the samples. Amount of TSP-4 was

38 measured by corrected total cell fluorescence (CCTF) of each cell seeded on different group 190-

193. Briefly, the CCTF of a cell was measured by deducting the average fluorescence of the background around the cell from the average fluorescence of the cell area. Regions of interest were chosen randomly across the samples.

3.3.13. Statistical Analysis:

Analysis of covariance (ANCOVA) was performed for mechanical data to check whether longitudinal modulus is significantly different from transverse modulus. Stretching level was taken as the covariant. Q-Q (Quantile-Quantile) plot of the residuals was plotted to check the normality of the data. A post hoc analysis using the Tukey’s test was conducted to compare the pair wise differences between different groups. The significance was set at p<0.05.

The cell study data were not normal according to the Q-Q plot, a two-way ANOVA equivalent of nonparametric test, ordered logistic regression (OLR) was used to check whether there is an effect of time point as well as anisotropy on cellular response. Non parametric Mann-

Whitney U test was conducted to compare pairwise difference between groups (Figure 17) as a post hoc analysis. A Bonferroni correction was applied to the cell study statistical analysis to compensate for large data set and statistical significance was set at p<0.025. For protein quantification staining Mann-Whitney U test was conducted to compare difference between groups.

In the case of CCTF quantification of TSP-4, as cell data are involved, a Bonferroni correction was applied and statistical significance was set at p<0.025. Statistical computing tool

“R” (R-Project, Vienna, Austria) was used to do all the statistical analysis.

3.4 Results

Electrochemically compact collagen sheets were fabricated by using planar electrodes

(Figure 11A) and stretched by a mechanical stretching device to incorporate SA in isotropic collagen sheets (Figure 11B) (see Methods). Collagen molecules are randomly (i.e. isotropically) oriented in compact sheets as evidenced by CPI (Compensated Polarized Imaging) image

(magenta indicates randomness 126, 194, 195 in Figure 12A, 0% stretch). Molecular alignment increased gradually with stretch as indicated by the emergence of blue color in the polarized images (Figure 12A). 0% stretch group was isotropic and 60% stretched group had the highest alignment as indicated by the uniform blue color over the aligned compact sheet. CPI results were backed up by the SEM (scanning electron microscopy) images (Figure 12B) which revealed collagen fibrils of 0% stretch group were randomly oriented. A gradual increase (Figure 12B) in fibrillary alignment was observed along the stretching direction. SEM images indicated a full replication of the aligned collagen surface topography by the cellulose acetate (Figure 12C).

(A)

(B)

(C)

Figure 12: A) CPI images of collagen sheets at different levels of stretch. The arrow indicates the slow axis of the gypsum plate and molecular alignment along the direction of the slow axis is manifested by blue which emerges gradually with stretch. At 60% stretch, full alignment is present over the field of view. Scale bar 100 µm, B) SEM images of collagen sheets stretched to different strain levels indicate an aligned topography along the stretching direction (arrows; scale bar 2μm). C) Replication of the surface topography of highly stretched (SA=8) collagen sheet on cellulose acetate film. The replicated surface was used to determine whether the surface topography played a role in cellular alignment and elongation. Scale bar: 20µm

The electro compacted collagen sheets were tested mechanically both in longitudinal and

transverse to the stretching direction. The mechanical test result of different SA collagen sheet

indicated that longitudinal direction modulus (EL) and maximum failure stress (σL) increased about

퐸퐿 4-fold when SA (= ) increased from 1 to 8 (Figure 13A). On the other hand, ET and σT declined 퐸푇

by 2-fold from SA=1 group to SA=8 group (“T” indicates transverse direction property).

Electrocompacted random collagen sheet (0% stretch) was isotropically stiff and strong

(SA=1.11 ± 0.51, STA=1.09 ± 0.55, respectively), in agreement with the random fibrillar

휎 orientation observed by CPI and SEM. SA and STA (stress anisotropy = 퐿) increased gradually 휎푇

by up to 8-fold with increasing stretching level (Figure 13B).

(A)

(B)

The Root Mean Square (RMS) surface roughness, Rq for SA=1 and SA=8 was 50.71 ±

22.67 nm and 53.03 ± 16.45 nm. There was no significant difference between the groups which indicates that incorporation of SA does not affect the surface roughness of the samples. (Figure

14).

Figure 14: Surface Roughness measurement by AFM. The Mean RMS surface measurement showed that SA incorporation did not affect the surface roughness.

The cell viability result indicates that electrocompacted collagen sheets are not cytotoxic and incorporation of SA has no effect on cell viability (Figure 15).

Figure 15: Cell viability on SA=1 and SA=8 groups.

Human mesenchymal stem cells (hMSC) seeded on collagen sheets of varying SA showed a 4-fold increase in the cytoskeletal aspect ratio with increasing SA (Figure 16A, 17A) both at 6 and 12 hours. This increase in aspect ratio was mainly due to continual increase in the length of major axis from 40 microns at SA=1 to 110 microns at SA=8 at 12-hour time point (Figure 16A,

17B). On the other hand, cell minor axis length was stagnant at about 30 micrometers by 12-hour time point regardless of the SA (Figure 17C). Both major axis length (p =0.003) and minor axis length (p = 0.000) increased between 6 and 12 hours indicating continual spreading of cells over time (Figure 16A, 17B, 17C). There was no significant difference of cell aspect ratio between cells seeded on surface replicated cellulose acetate sheet and control cellulose acetate sheet

(Figure 16B & 17H). Cells on cellulose acetate with functionalized Collagen-I coating also did not show any significant difference like the uncoated cellulose acetate sample experiment.

Nuclear aspect ratio (Figure 17D) increased moderately but significantly within increasing

SA (p<0.025). In parallel with the changes observed for cytoskeletal dimensions, major axis length

(Figure 17E) of nuclei increased with increasing SA whereas the minor axis (Figure 17F) did not change with increasing SA.

(G) (H)

Figure 17: Cell aspect ratio (A), major axis (B) and minor axis (C) length after 6 and 12 hours (Left column). Nuclear aspect ratio (D), major axis (E) and minor axis (F) length after 6 and 12 hours (Right column). Significance was set at p < 0.025 using Bonferroni correction. (G)The schema is drawn to scale from the reported average measurements to illustrate cell and nucleus dimensions at different values of anisotropy. (H) Cells seeded on surface replicated cellulose acetate sheet showed that there was no significant difference in aspect ratios of cells seeded on surfaces replicated with the surface topography of collagen vs. control group.

The cumulative histograms (Figure 18) indicated that a greater fraction of cells were aligned along the stretch axis with increasing amount of stretch. Such that, at 12 hours, 24%,47% and 78.8% aligned cells were within 20° angle of stretching direction for SA=1, SA=3 and SA=8 groups, respectively. For SA=1 group, after 6 hours, 65% of cells had r < 1.2 and after 12 hours

33.33% of cells had r < 1.2. Cells with aspect ratios of r < 1.2 were considered to be round and therefore, not included in the histogram.

SA=1 SA=3 SA=8 Figure 18: Cumulative Relative Histogram Plot of Cell Alignment Angle. After 6 hours, for SA=1, SA=3 and SA=8 group, 22.4%, 41.0% and 77.0% aligned cells are within 200 angle of the stretching direction respectively. After 12 hours, for SA=1, SA=3 and SA=8 group, 24.0%, 47.0% and 78.8.0% of aligned cells are within 200 angle of the stretching direction respectively. For SA=1 group, after 6 hours, 65% cell has r <1.2 and after 12 hours 33.33% of cells have r<1.2 are considered to be round and therefore, not included in the cell alignment angle histogram.

Gene expression results on days 1 and 3 (Figure 19) showed that increasing the stiffness by compaction while maintaining isotropy (gel vs. unstretched sheet) did not affect the expression levels of MKX and SCX. On day 1, induction of SA resulted in significant increases in the expressions of MKX and SCX. On day 3, expressions of MKX and SCX were greater only for the highest level of stretch. By day 5, expressions of all electrocompacted groups (stretched or not) were comparable, and, greater than those of the collagen gel.

Figure 19: Effect of SA on tenogenic differentiation of human MSCs. After day 1, 3 and 5 SCX showed gradual increase of expression level. MKX showed increased expression for SA=3 and SA=8 group after day 1 and 3. Within the same marker and day, significance difference between gel and compact group is noted by “*” and between non anisotropic compact with other anisotropic group is denoted by “**” and SA=3 vs SA=8 is denoted by “***”.

Figure 20: Effect of SA on long term matrix synthesis. The collagen laid by cells on anisotropically stiff substrates was oriented parallel to the stretch direction. After day 21 days of hMSc culture, highly anisotropic (SA=8) showed 2-fold increase in type-I collagen synthesis than non-anisotropic sheet (SA=1); cells on SA=8 sheet showed 2- fold increase in TSP-4 production than SA=1 indicated by corrected total cell fluorescence (CCTF); and gradual increase in type-III collagen synthesis by 2-fold increase from SA=8 to SA=1 sheet; Single head white arrow indicates collagen synthesis; for type-I and type-III collagen, scale bar 100 µm and for TSP-4 scale bar 50 µm. Double head white arrow indicate stretching direction.

After 21 days of culture, type-I and type-III collagen production increased by 2-fold in highly anisotropic group (SA=8) than SA= 1 and SA=3 (Figure 20). For type-I collagen, there was no significant difference between SA=1 and SA=3. For type-III collagen, SA=3 samples showed significantly greater matrix synthesis than SA=1. In case of TSP-4, cells on SA=8 sheet showed 2.5- fold increase in TSP-4 expression than SA=1 indicated by corrected total cell fluorescence (CCTF) level (Figure 20). There was no significant difference in TSP-4 immunofluorescence level of cells between SA=1 and SA=3.

3.5 Discussion

The planar stretch induced SA in collagen sheet affected cellular and nuclear morphology.

Furthermore, cell fate, specifically tenogenic differentiation of MSCs were inducible by substrate

SA. The observed outcome was attributable to SA rather than aligned topography as demonstrated by the absence of cellular elongation on isotopically stiff cellulose acetate sheets which replicated the topography of aligned collagen sheets.

The increase in anisotropy was mostly due to an increase in the modulus along the stretch axis whereas the modulus transverse to the stretch axis varied less prominently (4-fold vs 2-fold) with the stretching. This outcome implies that the lateral interactions between collagen molecules define the stiffness in the transverse direction, regardless the molecules are oblique (as in the isotropic state) or fully parallel (as in the aligned state) to each other. Overall, molecular alignment predominantly benefits the modulus along the longer axes of molecules.

Preferential alignment towards the stiffer material axis as we report in this study is an agreement with the preferential migration of cells towards higher rigidity regions of materials with rigidity gradients 140. Plotnikov et al. reported that individual focal adhesions (FAs) sense ECM

50 locally by repeatedly applying tugging forces and that soft ECMs promote tugging traction dynamics in FAs whereas rigid ECMs promote stable traction in FAs 140. Therefore, cells migrate towards the rigid ECM with stable traction of FAs by directed migration. In this study cells showed elongation along the stiff direction. This indicates that the FAs of cells were able to sense and differentiate between the stiff and softer directions of the anisotropic sheets and responded accordingly.

The response of nuclear aspect ratio to substrate anisotropy (22% increase) was significant but less pronounced than the response of cytoskeletal aspect ratio (201% increase). The change in nuclear morphology occurred largely by an increase in the major axis length of the nucleus while there was no change in the minor axis length of the nucleus. In highly anisotropic tendon tissue, cell aspect ratio varies from 3-8 196-198 whereas nucleus aspect ratio varies from 2.5-6 100, 196, 199.

The nucleus aspect ratio in isotropic tissue types (adipocyte, cornea) varies from 1.1-1.8 200-204.

Cells seeded on the highly anisotropic sheets in this study had a nucleus aspect ratio of 2, a ratio that is on the higher end of isotropic tissues and lower end of anisotropic tissues. Experiments with longer durations of cell-seeding on anisotropic sheets may show greater elongation of the nuclei which remains to be determined. Also, cells are investigated in 2D context whereas the 3D nature of native tissues may be promoting a greater degree of increase in the nuclear aspect ratio.

The nucleus had an increasing major axes length with increasing SA whereas the minor axes remained static. It is likely that this behavior is determined by preferential recruitment of actin fibers along the stiffer direction pulling on the nucleus along the major axis predominantly.

Therefore, substrate anisotropy may be transmitted to the nucleus via mediation of actin filaments along the major axis. Future experiments targeting actin anchorage to the nuclei would prove this point conclusively.

For the isotropic (SA=1) group, 65% and 34% of cells were round after 6 and 12h. Whereas for medium (SA=3) and highly anisotropic (SA=8) sheet close to 100% cells were elongated. The average alignment direction gradually comes closer to the stretching direction as the anisotropy of the sheet increases. This indicates that cell sensed the stiffer direction of the anisotropic sheet and showed direct response by gradually increasing cell aspect ratio and by gradually extending along the stretching direction. In this context, arguably, SA is a form of durotaxis where the cell translates locally by expanding along the stiffer axis at a higher rate than the compliant axis to assume both an elongated form while aligning along the stiffer direction of the material.

It has been well established that biophysical cues such as substrate stiffness induce MSCs to commit to different lineages 150, 205. Younesi et al. showed that anisotropic alignment of the electrocompacted collagen fibers yields tenogenic differentiation of MSCs as demonstrated by gene expression and matrix production 169. Mature tendon tissues are highly anisotropic in nature and during tendon development in embryos, the environment where cells are undergoing tenogenesis are observed to be formed of highly anisotropic parallel bundle of collagen fibrils 206,

207. The anisotropic collagen sheets in this study partially mimic the anisotropic nature of the tendon ECM by presenting parallel collagen molecular alignment. Therefore, it is likely that the substrate SA induced cellular elongation is a contributor to the upregulation of the tendon markers as reported in this study. Therefore, we propose anisotropy of the stiffness as one of the determinants of MSC fate.

Scleraxis is an early progenitor marker which regulates tendon formation 177, 179, 181. It has been shown that scleraxis was expressed in day 14.5 embryonic mouse tendon 180. Mechanical force and scleraxis incorporation in the hESCs induced hESCs commitment to tenocytes 99.

Tendons of scleraxis null mutant mice showed severe defects. All these studies suggest that

52 expression of scleraxis is important in tendon development. Previous studies of MSCs seeded on highly anisotropic collagen fibers 128, 169 and knitted silk collagen scaffold 178 showed early increase in scleraxis expression and later the expression was decreased. Growth differentiation factor-5

(GDF-5)89 treated MSCs seeded on culture plates are reported to induce early increase in scleraxis expression followed by a decrease. The current study similarly showed early increase in the scleraxis expression and interestingly, at day 1 and day 3 highly anisotropic sheet showed higher expression than gel, non-anisotropic and intermediate anisotropic groups (Figure 19). At day 5 scleraxis expression decreases and levels between groups (Figure 19). This early increase in the scleraxis expression in highly anisotropic sheet indicates that, the anisotropic nature of the collagen sheet promotes the genes involved in early phases tenogenic differentiation.

Mohawk is expressed in developing tendons of mouse embryo and Mohawk knockout mice showed hypoplastic tendons and down regulation of type I collagen182, 184, 187. These results indicate that Mohawk is another critical regulator for tendon differentiation. Liu et al. demonstrated that the transcription factor Mohawk upregulates scleraxis in murine MSCs. Otabe et al. demonstrated that Mohawk was upregulated by 1.8 fold when rat MSCs were seeded on a collagen scaffold to repair a tendon defect185, 188. MSCs 186 or tendon-derived cells 183 seeded on collagen gels exhibited 1.5- and 3-fold upregulation of Mohawk after 7 days and 2 days of culture period, respectively, without the use of any growth factor. The intermediate and highly anisotropic sheet in this study showed 4-fold upregulation of Mohawk expression at day 1 without the use of any growth factor. At day 3, the expression increases from 2- to 4-fold from isotropic to highly anisotropic sheet. At day 5, there is further increase in Mohawk expression. Overall, Mohawk displayed the most robust response to SA among the markers investigated in this study.

Long term tendon related matrix production study indicated that highly anisotropic samples favored tendon related matrix production. Type-I collagen is the main tendon collagen and type-

III collagen is one of the major tendon-associated collagen which is crucial for type-I collagen fibrilogenesis 208, 209. Studies demonstrated that transcription factor SCX and MKX is involved in tendon formation by regulating type-I collagen production 184, 210-212. This study also conforms with the previous studies by showing that SCX and MKX upregulation in the first five days and increased type-I collagen production by day 21 (Figure 20) occurred for highly anisotropic group

(SA=8). Type-III collagen mediate attachments of tendon and helps during healing of tendon injury 17, 213, 214. The response of the body of tendon injury is to produce type-III collagen to quickly repair the damage 27, 215. After long periods type-III collagen is remodeled to type-I collagen 214,

215. In this study type-III collagen production was higher in anisotropic groups than the isotropic group (Figure 20). TSP-4 is one of the main tendon-related genes 216. Tendon extracellular matrix contains TSP-4 which displays the highest tendon selective expression compared to other tissue types 216-218. TSP-4 is also believed to bind to collagen and form complexes with COMP in tendon

218, 219. TSP-4 was shown to be upregulated in both engineered scaffold-free tendon tissue and in collagen matrix 220, 221. Recently, Ning et al. showed that TSP-4 was upregulated when cells were seeded on decellularized tendon slices 222. The highly anisotropic group (SA=8) in this study showed higher level of TSP-4 immunofluorescence (Figure 20) than the SA=1 and SA=3 groups indicating that the anisotropy of the collagen sheet mimics the anisotropy of the tendon and increase TSP-4 level as in Ning et al.’s study.

3.6 Conclusions

To the best of our knowledge, this study is the first which demonstrated that stem cell fate is affected by not only the magnitude of stiffness but also by the directional anisotropy of the

54 substrate stiffness. Such anisotropy can be a key determinant in driving cell morphology and differentiation during development and maintenance of anistropically stiff tissues. We have demonstrated a case for tendon differentiation and future studies will investigate genotypes and phenotypes associated with other anisotropic tissues such as skeletal or cardiac muscles.

4. Chapter 4: Tenoinduction by Mimicry of Tendon Topography on Pure Collagen Substrate

4.1 Abstract

Extracellular matrix modulus or mechanics plays an important role to regulate cell morphology, proliferation, differentiation during regular and diseased states. Although the effects of substrate topography and modulus on MSC differentiation are well known with respect to osteogenesis and adipogenesis, there has been relatively little investigation of this phenomenon with respect to tenogenesis. Overall, the roles of topographical factors (matrix alignment vs. matrix modulus) in inducing tenogenic differentiation are unknown. In this study we investigated both the effect of modulus and alignment of type I collagen substrate for tendon differentiation, which are two of the major factors of tenogenic differentiation. Moreover, seminal studies in the literature have reported the effect of modulus in the kPa range and there is a lack of studies that assess cellular response on materials in the MPa range of modulus. In this study, type I collagen sheet substrate in range of 0.1-100 MPa was generated by tuning the modulus value of 1000-fold by using ectrocompaction and crosslinking processes. Matrix alignment was introduced by controlled unidirectional stretching of the sheet. This study showed that mimicking the tendon topography, i.e. increasing the substrate modulus as well as alignment increased the tenogenic differentiation.

Therefore, the tenoinductive collagen material model developed in this study can be used in the research and development of tissue engineering tendon repair constructs in future.

4.2 Introduction

Modulus of the extracellular matrix plays an important role in regulating cell morphology, proliferation, differentiation during regular and diseased states. For example, 1 kPa matrices favor differentiation of MSCs into neuronal-like cells, 10 kPa elasticity promotes myogenic differentiation, and a 100 kPa matrix stimulates osteogenic differentiation 150. The effects of substrate topography and modulus on MSC differentiation are well known with respect to osteogenesis and adipogenesis104, 150, 223-226. However, there has been relatively little investigation of this phenomenon with respect to tenogenesis. For example, Sharma et al studied the effect of tenogenesis on collagen coated polyacrylamide substrate with modulus gradient and modulus range less than 100 kPa104, 105. Therefore, the studies investigate tenogenesis are few and vary widely in design and materials used, and none of them specifically investigated tenogenesis in human MSC104, 105. The studies to date have investigated the effects of a modulus range of .01 to

.08 MPa104. On the other hand, it is known that the native modulus of the tendon tissue to be as high as 1200 MPa 14, 41. Therefore, studies to date do not cover a modulus range that is more compatible with the tendon tissue environment.

The literature also reports that topography, specifically unidirectionally aligned fibrous topography as another tenogenic differentiation cue. Tong et al. reported that tendon cell differentiation of hMSCs occurs on collagen coated PDMS substrates which replicate native tendon surface227. However, they did not investigate the effects of modulus on MSC to tenocyte differentiation227. Yin found that aligned substrate promote tenogenesis in tendon/stem progenitor cells (TPSC) but did not evaluate MSCs 228. Furthermore, aligned electrochemically compacted collagen fiber, electrospun polyurethane (PU), silk fibroin, poly (l-lactic acid) and two-

57 dimensional (2D) microgrooved surface showed elongated cell morphology and tendon related

ECM formation128, 143-149.

The effects of matrix alignment and matrix modulus on tenogenesis have been investigated separately and the synergy between the two variables in terms of inducing tenogenic differentiation is not studied. Our research group created a collagen based material platform to investigate the effect of modulus in a broad range of 0.1 MPa to 100 MPa while controlling the topographical alignment random or unidirectional. In this method, collagen solutions are transformed to highly dense randomly oriented collagen sheets by electrochemical compaction induced by planar electrodes169, 173, 229. Matrix alignment is introduced in these sheets by controlled unidirectional stretch of the sheets as molecular alignment lacks when planar electrodes are used to fabricate sheets229. The aim of the current study is to study the effects of matrix modulus and alignment on tenogenic differentiation of human MSCs. Importantly, a broad modulus range of 0.1 MPa to 100 MPa is covered logarithmically in this study for the first time in the literature. Elucidation of the mechanism of topographically induced tenogenesis in human

MSCs as such will assist in optimizing materials to produce scaffolds for tendon repair and may provide insight into the differentiation mechanisms of MSCs and other stem cells in vivo.

4.3 Materials and Methods:

4.3.1. Collagen Sheet Fabrication by Electrochemical Compaction [Figure 21]:

Acid soluble monomeric Type-I collagen solution (Collagen Solutions, San Jose, CA; 6 mg/ml) was diluted two-fold with RNAase/DNAase free water. The pH of the collagen solution was adjusted to 8-10 using 1 N NaOH. The collagen solution was then dialyzed against ultrapure water for 18 hours to prepare the collagen solution for electrocompaction.

Collagen sheet was fabricated in sheet form by electrocompaction method described before

173, 174. Briefly, a rectangular window of 30x10x1.5 mm was cut in a plastic piece. The plastic piece was placed on planar carbon electrode and the window was filled with the dialyzed collagen solution. Another plane carbon electrode was placed above the plastic spacer to sandwich the collagen solution between the electrodes. A 30 VDC electrical potential was applied across the electrodes for 2 min. Collagen molecules are electrophoretically mobilized and compacted due to pH gradients established between the two electrodes under the mechanisms detailed in an earlier publication 175. 100-200 microns thick collagen sheets of 30x10 mm dimensions were generated by the electrocompaction. Collagen molecules are randomly oriented within the plane of the sheet.

In one of the experimental groups, collagen molecules were unidirectionally aligned by stretching the sheet using a motorized mechanical device as described earlier229 (Figure 21 & 22). Collagen sheet samples were incubated in phosphate buffered saline (PBS) for six hours at 37 °C to induce fibril formation and treated with 2-propanol solution for 12 hours. Different levels of modulus were attained by crosslinking the sheets as will be described.

4.3.2. Synthesis of Collagen Gel:

Elastically most compliant group was the standard collagen gel which was not subjected to electrocompaction. Collagen gel was synthesized by mixing acid soluble monomeric collagen solution with 10x PBS at a ratio of 8:1 parts and the pH was adjusted to 7.4 using 0.1 N NaOH.

The collagen solution was then poured into the glass petri dish. The petri dish was placed in at 37

°C for 1h to form collagen gel. The gel was then crosslinked with genipin in 95% ethanol for 3- days as detailed in Table 2.

Figure 21: Overview of fabricating collagen sheets with different modulus levels.

4.3.3. Experimental Groups:

Four levels of modulus were targeted (0.1, 1, 10 and 100 MPa) for randomly aligned collagen sheets to investigate the effect of modulus on tenoinduction. These modulus levels were attained by changing the crosslinking protocols [Table 2]. Crosslinking was carried out by genipin

(Wako Chemical, Japan) in 90% v/v ethanol solution at 37 °C. The fifth group was a unidirectionally aligned collagen sheet group at 10 MPa along the axis of alignment to assess the effects of alignment. Samples were treated with per acetic acid (Sigma Aldrich, USA) ethanol solution (2% Acetic acid+96% Ethanol) after the cross linking to bleach out extra genipin which may keep crosslinking the samples.

Table 2: Experimental groups for different level of modulus range and unidirectionally aligned substrate

Target Experimental No Groups Modulus Modulus (MPa) (MPa)

1 Randomly oriented gel (3day, 0.6% genipin crosslinked) 0.1 0.09 ± 0.04

Randomly oriented compact sheet (1hr, 0.1% genipin 2 1 1.12 ± 0.18 crosslinked) Randomly oriented compact sheet (6hr, 0.6% genipin 3 10 10.39 ± 2.63 crosslinked)

4 Aligned compact sheet (1hr, 0.3% genipin crosslinked) 10 11.78 ± 2.44

Randomly oriented compact sheet (3 day, 2% genipin 5 100 92.93 ± 15.24 crosslinked)

4.3.4. Imaging of Molecular Alignment in Compacted Collagen Sheets

The alignment of collagen molecules at 10 MPa aligned group was assessed by a polarized optical microscope equipped with a first order wavelength gypsum plate (Olympus BX51, 61

Melville, NY, USA) and by using SEM. The aligned molecules along the slower axis of the gypsum plate shows blue interference color and the molecules which are perpendicular to the slow axis appear yellow 176 as collagen is a positive birefringent material . Magenta color indicates lack of alignment and emergence of blue in the Compensated Polarized Imaging (CPI) indicates molecular alignment.

The surface morphology of collagen samples was observed by a scanning electron microscope (FEI Nova Nanolab 200, Hillsboro, Oregon).

4.3.5. Assessment of Mechanical Properties of Collagen Sheets:

Collagen sheet of treatment groups described in Table 2 were cut into 20 x 2 mm strips (n

= 6-8 samples per group). Samples were hydrated in deionized water for 30 minutes and tested

under monotonic tension (Rheometrics Inc., NJ) until failure at a strain rate of 10 mm/min to

assess that attainment of targeted modulus values. The thickness of the sheet samples was

measured by a custom-made micrometer where the micrometer closes an electrical circuit upon

contact with the surface of the sample. Stress-strain curves were constructed using the load-

displacement data and sample geometry. Modulus was calculated from the slopes of the linear

regions of stress strain curves.

4.3.6. Effect of Matrix Modulus and Alignment on Tenogenic Differentiation of Human MSCs

The culture medium composed of alpha MEM (Invitrogen) supplemented with 10% MSC-

Qualified FBS (Invitrogen), 1% penicillin/streptomycin and 50 μg/mL ascorbic acid. Cells were cultured for 21 days with medium changes every 3 days. Tenogenic differentiation was assessed by RT-PCR at day 3, 14 and 21. At time points 3, 14 and 21 days total RNA was extracted by lysing the sells using TRIZOL reagent (Invitrogen) following manufacturer’s protocol. Briefly, chloroform was added to the trizol homogenized samples and the phase separation was performed by centrifuging the samples at 12,000 g for 15 min at 4 ºC. The RNA was collected in a separate tube from the supernatant aqueous phase, precipitated by adding isopropanol and pelleted by centrifuging at 12,000 g for 10 min at 4 °C. 70% ethanol was used to wash the RNA pellet. The pellet was then dried and resuspended in RNAse/DNAse free water (Invitrogen) and stored at -80

°C. 2 µg of total RNA was reversed transcripted to synthesize cDNA using the cDNA Reverse

Transcription Kit (Applied Biosystems). Taqman real time PCR mastermix (Applied Biosystems) and Taqman gene expression assays (Applied Biosystems) for tendon related markers (Collagen

I, Collagen III, and COMP) and tendon specific markers (Scleraxis, Thrombospondin) were used to evaluate the expression of the genes by quantitative real time PCR (Applied Biosystems 7500

Real Time PCR System). The relative fold change in the target gene expression was quantified using 2 deltadeltaCt method by normalizing the target gene expression to RPLP0 169 as a housekeeping gene and relative to the expression on 0.1 MPa group at day 3.

4.3.7. Effect of Matrix Modulus and Alignment on Tendon Related Matrix Synthesis

Based on the results of RT-PCR, 1 MPa group was not included in matrix synthesis study.

Cells were seeded on 0.1 MPa, 10 MPa, 10 MPa aligned and 100 MPa groups as described above.

Cells were cultured for 21 days as described earlier. Three long term tendon-related extracellular matrix molecules were investigated: i) type-I collagen, ii) type-III collagen and iii)

Thrombospondin-4 (TSP-4). For type-I and type-III collagen, immunohistochemistry was performed and for TSP-4 green immunofluorescence was performed.

Collagen sheet samples were fixed in 10% neutral buffered formalin, rinsed in 3 washes of

PBS for 5 min each, permeabilized for 15 min in PBS containing 0.25% Triton X-100 followed by 3 washes in PBS for 5 min each. After blocking the samples in PBST (Phosphate Buffered

3 times in PBS for 5 min each and incubated with StayRed/AP working solution (3 ml of

StayRed/AP Substrate buffer containing one drop of StayRed/AP chromogen) for 15 min at room temperature. Samples were rinsed in PBS and imaged using a microscope (Olympus IX83 digital microscope, Olympus Life Science). Images were processed and quantified by Cellsens Dimension software (Olympus Life Science).

For immunofluorescence, the anti-Rabbit Alexa-488 conjugated specific secondary antibody (Pierce Protein Biology, Thermo fisher Scientific) was used. The samples were visualized

64 under Olympus IX83 digital microscope (Olympus Life Science) using Cellsens Dimension software (Olympus Life Science)

Quantification of the protein staining was performed by ImageJ (NIH, Maryland, USA).

The amount of type-I and type-III collagen staining was calculated by ImageJ colour thresholding

189. The regions of interest were chosen randomly across the samples. Amount of TSP-4 was measured by corrected total cell fluorescence (CCTF) of each cell seeded on different group 190-

4.3.8. Statistical Analysis:

One-way analysis of variance (ANOVA) was performed for RT-PCR data and Tuckey’s post hoc analysis was performed for pairwise comparison. Significance was set at p<0.05.

For COL I and COL III quantification Mann-Whitney U test was conducted to compare difference between groups (p<0.05). In case of CCTF quantification of TSP-4, as cell data are involved, a Bonferroni correction was applied and statistical significance was set at p<0.025.

Minitab Statistical package (Minitab Inc., State College, PA, USA) was used to perform the statistical analysis.

4.4 Results:

The mechanical test results [Table 2] showed that the targeted modulus levels were attained over 4 order of magnitudes.

Planar stretching introduced molecular as well as fibrillar alignment evident by CPI and

SEM image respectively [Figure 22].

Figure 22: Manufacturing aligned sheet with mechanical stretching. Molecular alignment is evident by CPI image (Emergence of blue color after stretching indicates molecular alignment). Collagen fibril alignment is evident by SEM images.

SCX expression at day 3 was greater for 100 MPa than for 0.1 MPa (p = 0.02) [Figure 23A].

There was no significant difference between 10 MPa aligned and 10 MPa random groups indicating that alignment doesn’t have any effect at 10 MPa.

COL I expression at day 3 was the lowest for 0.1 MPa level and the highest for 100 MPa

[Figure 23B]. Expression at 1-10 MPa range modulus values were positioned intermediately.

Between each modulus level, there was about 2-fold upregulation in COL I expression. At day 14, the expression was grouped over two levels, 0.1-1 and 10-100 MPa and there was about 2-fold upregulation between these two modulus levels. By day 21, there was no significant difference

66 between any of the groups. Alignment did not have any effect on COL I expression at 10 MPa groups.

COL III expression was the highest for 100 MPa modulus and it showed more than 2-fold upregulation than 0.1 MPa at different time points [Figure 23C]. Alignment favored COL III expression such that the expression for aligned 10 MPa group at day 14 was two-fold greater than that for random 10 MPa group.

COMP expression for 100 MPa group showed 2 and 3-fold greater upregulation than that of random 0.1-10 MPa groups at day 14 and 21 respectively [Figure 23D]. Alignment increased

COMP expression such that 10 MPa aligned group’s expression was greater than 10 MPa random group’s expression. Furthermore, aligned 10 MPa group’s COMP expression did not differ from that of 100 MPa group.

There was a gradual increase in TSP-4 expression with increasing modulus [Figure 23E].

Alignment also significantly favored TSP-4 expression (p = 0.04).

After 21 days of culture, Type-I collagen production increased by 2-fold in 10 MPa and 3-fold in 100 MPa group in comparison to the production by the 0.1 MPa group [Figure 24&25].

Alignment increased COL I synthesis significantl (p = .031). In case of COL III, there was a gradual increase in synthesis with increasing modulus with a 2-fold increase from 0.1 to 100 MPa group [Figure 24&25]. In this case also, alignment increased COL III synthesis (p = .034). In case of TSP-4, there was no significant difference between 0.1-10 MPa while 100 MPa showed 4- fold greater TSP-4 production than that of the0.1 MPa group [Figure 24&25]. There was no significant difference in TSP-4 immunofluorescence level of cells between aligned and random group.

Figure 23: Effect of collagen matrix modulus and anisotropy on tenogenic differentiation of human MSCs. (A) Scleraxis, (B) Collagen-I, (C) Collagen-III, (D) COMP, (E) Thrombospondin- 4. Anisotropy and increasing modulus improved tenoinduction. Statistical differences are highlighted by horizontal lines with the corresponding p values. Statistical significance was set at p <0.05.

Figure 24: Effect of Modulus on long term matrix synthesis. After day 21 days of hMSc culture, higher modulus favored matrix synthesis. For collagen I, stiffer groups showed thick collagen fiber formation. For TSP-4, 100 MPA group showed more production. For Collagen III also 100 MPa showed maximum synthesis and alignment (10MPa aligned) increase matrix synthesis. Scale bar 50 µm. Alignment direction of 10 MPa aligned group is shown by white arrow.

160000 p = 0.000

/ROI) p = 0.005

2 140000 p = 0.000 p = 0.031 µm 120000 p = 0.001 100000 80000

60000 40000 20000

( Synthesis Collagen I Type 0 0.1 Mpa 10 Mpa 10 Mpa 100 MPa Aligned 20 p = 0.000 p = 0.000

16 p = 0.000

4 -

12 TSP

8 CCTF for for CCTF

0 0.1 MPa 10 Mpa 10 MPa 100 MPa Aligned 450000

/ROI) 400000 p = 0.003 2 p = 0.010

µm 350000 p = 0.003 p = 0.034 300000 p = 0.003 250000 200000

150000 100000 50000 ( Synthesis Collagen III Type 0 0.1 Mpa 10 Mpa 10 Mpa 100 Mpa Aligned

4.5 Discussions:

Natural tendon is composed of highly stiff and aligned type I collagen fibrils. In this study we investigated both the effect of modulus and alignment of type I collagen substrate for tendon differentiation. This study showed that mimicking the tendon topography, i.e. increasing the substrate modulus as well as alignment increased the differentiation of human MSCs to a tenogenic lineage.

Past literature reported the effect of modulus on differentiation (e.g. muscle, neuron) in the kPa range which was then followed by glass which has a modulus around 90 GPa 150, 230.

Therefore, there is a gap in the literature on substrate modulus effects in the range of 100 kPa to

100 MPa range in terms of cellular response. This study not only fills the gap of the modulus ranges that had been studied to date, but also incorporates matrix alignment to investigate tendon differentiation104, 105.

Previous studies demonstrated that increasing modulus generally increases cell adhesion,

proliferation and multilineage differentiation104, 105, 150, 231-237. Very few of these studies

investigated the effect of modulus on tenogenic differentiation on collagen substrate which is

the major tendon ECM104, 105. Moreover, the studies that investigated tenogenic differentiation

used substrate modulus (10-80 kPa) levels which are orders of magnitude softer than tendon and

did not consider synergistic effect of alignment 104, 105. This is the first study to our knowledge,

which considers modulus of collagen substrate to MPa level as well as investigates effect of

substrate alignment on tendon differentiation.

The current study fabricated type I collagen substrates with modulus values encompassing

three orders of magnitudes of change beginning from hundred kPa, to hundred MPa. Moreover,

71 by introducing planar stretch, we were able to generate aligned substrates. Therefore, the collagen material model developed in this study allowed for systematic investigation of the effects of matrix modulus and anisotropy tenogenic differentiation.

Scleraxis is an early progenitor marker which is important in tendon development177, 179, 181

180 99. Sharma et al. showed increased SCX expression with increasing collagen coated substrate modulus in their studies related to tendon differentiation 104, 105. Recently, Chen at al showed arterial stiffening via SCX upregulation238. This indicates that, substrate modulus is associated with SCX expression. The current study showed early increase in the scleraxis expression and interestingly, at day 3 stiffer groups showed higher expression. Previous studies of MSCs seeded on highly anisotropic collagen fibers 128, 169, knitted silk collagen scaffold 178 and on culture plates 89 reported to induce early increase in scleraxis expression followed by a decrease.

However, the modulus was not varied systematically in these studies. This study also showed that scleraxis expression levels between groups by day 21 (Figure 23A). Therefore, stiffer substrates seem to benefit tenogenesis by expediting the inception of MSC to tendon differentiation cascade.

Tendon is mostly composed of type-I collagen. Studies showed that matrix stiffening increases collagen I synthesis239 and with aging collagen turnover decreases in tendon with decreasing modulus240. The current study showed increased COL I expression with increasing modulus such as 100 > 1-10> 0.1 MPa at day 3 and 100 > 10> 0.1-1 MPa at day 14. This indicates that stiffer matrices induce earlier and greater levels of COL I expression than compliant matrices [Figure 23B]. From the protein production data, at day 21, there was a gradual increase in COL I synthesis from 0.1 MPa to 100 MPa. At the production level, alignment showed upregulation of COL I synthesis and synthesis in 10 MPa aligned group was similar to 100 MPa

72 group. This increase in COL I synthesis may be due to the alignment of the matrix as indicated by previous studies which showed that aligned matrix increased tenogenic differentiation 128, 169,

241.

Type-III collagen is one of the major tendon-associated collagen and crucial for type-I collagen fibrilogenesis 208, 209. During tendon injury type-III collagen is produced to quickly repair the damage 27, 215 and after long periods it is remodeled to type-I collagen 214, 215 In the current study, the trend in COL III expression as well as production was similar to that of COL

I. Moreover, in the expression level, 10 MPa aligned group showed similar expression as 100

MPa group both at day 3 and 14. In the production level, 10 MPa aligned group showed higher expression than 10 MPa unaligned at day 21. This indicates that alignment also helps in tenogenesis. Previous studies also showed similar outcome as stiffer matrix104 and aligned matrix128 increased COL III expression. It can be noticed that the amount of COL III synthesis is higher than COL I. Whether or not COL III is remodeled to COL I as is the case in tendon injury repair sites would require longer duration studies.

TSP-4 is present in tendon ECM, where it shows the highest tendon selective expression compared to other tissue types 216-218. Therefore, TSP-4 is considered to be one of the main tendon-related genes 216. Cells seeded on decellularized tendon slices, engineered scaffold-free tendon tissue and collagen matrix showed upregulation of TSP-4 222 220, 221. This study also showed upregulation of TSP-4 expression both with modulus and alignment after day 21 [Figure

23E]. In matrix synthesis study, after day 21, only 100 MPa group showed higher level of TSP-

4 immunofluorescence (Figure 24&25) than the other groups. In tendon, TSP-4 binds to collagen and form complexes with COMP 218, 219. Smith et. al suggested that COMP has an organizational role in tendon formation as well as COMP is necessary for tendon to resist load

242, 243. The current study shows, upregulation of COMP expression with stiffer (100 MPa) and

aligned (10 MPa aligned) matrix [Figure 23D]. This indicates that, higher modulus and

alignment mimicked tendon topography to an extent and had an influence on tenogenic

differentiation.

4.6 Conclusions

The current study demonstrates tenogenic differentiation is influenced by both modulus and alignment of the substrate. This study tried to mimic both the modulus anisotropy as well as the modulus of the natural tendon and investigated the synergistic effect of these two key factors of tenogenic differentiation. Therefore, the tenoinductive collagen material model developed in this study can be used in the research and development of tissue engineering tendon repair constructs in future.

5. Chapter 5: Assess effects of dermatan sulfate incorporation on the differentiation of MSCs.

5.1 Abstract

The tension bearing regions of tendon has abundant Dermatan Sulfate (DS) content which is one of the major glycosaminoglycans (GAG) in tendon. Therefore, it was hypothesized that incorporating DS in compact collagen sheet would render composition that is more convergent to that of tendon’s and may synergistically add to the topographical cues in terms of promoting differentiation. Additionally, while there are extensive research studies on chondroitin sulfate loaded scaffolds in tendon repair and regeneration, there is a lack of studies related to DS scaffolds for tendon repair and regeneration. In this study, electrocompacted collagen sheets with composite mixtures of collagen and DS at three different molar ratios (collagen only (no DS) and 30:1 and

10:1, collagen: DS) were fabricated and the effect of DS incorporation in compact sheets on tenogenic differentiation was investigated. This study showed that high concentration (10:1) of DS inclusion increased the failure strain by 3% and DS incorporation did not have any adverse effect on cell proliferation. However, DS incorporation did not affect tenogenic differentiation also and it may require other parameter such as mechanical loading, to investigate the effect of DS inclusion.

5.2 Introduction

Dermatan Sulfate (DS) is one of the major glycosaminoglycans (GAG) in tendon. The tension bearing regions of tendon has abundant DS content 244. DS assist collagen fibril assembly process during tendon development. DS chains cause parallel alignment of collagen fibril and helps in bridging collagen phase and cell signaling 245. Therefore, incorporating DS in compact collagen sheet would possibly give more alignment to the sheet as well as would render composition that is more convergent to that of tendon’s and may synergistically add to the topographical cues in terms of promoting differentiation. In our earlier work, we demonstrated that when collagen-DS mixtures are subjected to electrochemical gradients, DS is mobilized with collagen and becomes trapped in the final ELAC 129 . The experiments of this aim will include DS in compact collagen sheet at various concentrations and assess the effects on mechanical function and cell differentiation. Our earlier experience in inclusion of DS mimics indicated that such molecules do not compromise mechanical properties of ELAC threads and extend the failure strain in uncrosslinked fibers 129.

Composite mixtures of collagen and DS were prepared at three different molar ratios

(collagen only (no DS) and 30:1 and 10:1 collagen:DS) prior to loading into the electrochemical process. Based on the result of previous chapter, electrocompacted sheet with a moderate crosslinking protocol was chosen for this study. The effect of DS incorporation on the mechanical properties and cell differentiation was assessed.

5.3 Materials and Methods:

5.3.1. DS Incorporated Collagen Sheet Fabrication:

Collagen solution was dialyzed according to the method described in section 4.3.1.

Composite mixtures of collagen and DS (Celsus, Cincinnati, Ohio) was be prepared at three different molar ratios i) collagen only (no DS)-Control, ii) 30:1 collagen: DS and iii) 10:1 collagen:

DS. The mixture was used as a stock solution to form compact collagen sheet. Compact collagen sheet was fabricated by electrochemical process as described in section 4.3.1. Collagen sheet samples were incubated in phosphate buffered saline (PBS) for six hours at 37 °C to induce fibril formation and treated with 2-propanol solution for 12 hours. Finally, collagen sheets were crosslinked in .625% genipin (Wako Chemical, Japan) for 3 days. Samples were treated with per acetic acid (Sigma Aldrich, USA) ethanol solution (2% Acetic acid+96% Ethanol) after the cross linking to bleach out extra genipin which may keep crosslinking the samples. DS incorporated compact sheets were dissolved in 1 N HCl to confirm the amount of DS in the DS mixed groups.

Dimethylmethylene Blue (DMMB) assay was used to measure the concentration of DS in the solution which indicates the amount of DS incorporated in the compact collagen sheets.

Figure 26: A) Dermatan Sulfate (DS) incorporated electrocompacted collagen sheet. DS is uniformly distributed along the volume of the sheet confirmed by dimethylmethylene blue (DMMB) staining (control sheets absent in DS are transparent). B) Concentration of DS by DMMB assay.

5.3.2. Assessment of Mechanical Properties of Collagen Sheets:

Mechanical testing of different groups collagen sheet was performed as described in section

4.3.5

5.3.3. Effect of DS Incorporation On Cell Proliferation On Compact Sheets:

Samples were disinfected in 70% ethanol for 4 hours and washed in 1X PBS and placed into ultralow attachment 24 well plates (Corning) (n = 3 wells/group). Human mesenchymal stem cells (MSCs) (Lonza, Walkersville, MD) at passage 5 were seeded at a density of 20000 cells/cm2 on different sample groups. The culture medium composed of alpha MEM (Invitrogen) supplemented with 10% MSC-Qualified FBS (Invitrogen), 1% penicillin/streptomycin and 50

μg/mL ascorbic acid. After 4 h, non-attached cells were removed by changing the media and the attached cells were cultured for 7 days. After day 1 and day 7, cell proliferation was quantified using alamar blue assay by following the manufacturer’s instructions. Briefly, 0.5 ml of alamar blue mix (culture medium þ 10% alamar blue) was added to each well and incubated for 2 h at 37

°C. Following this,100 ml of alamar blue mix from each wellwas transferred to a 96 well plate in triplicate and the absorbance was recorded at 570 nm and 600 nm. Cell number was quantified by calculating the percentage reduction in alamar blue and comparing the values to a standard curve generated using known number of cells.

5.3.4. Effect of DS Incorporation on Tenogenic Differentiation of Human MSCs

This study was performed on the three groups as described in section 4.3.6 except in this case cells were cultured for 14 days and PCR was performed for single time point (14 days).

Tendon related markers -Collagen I, Collagen III, COMP, Decorin (DCN) and tendon specific markers (Scleraxis, Thrombospondin-4 (TSP-4) were used to evaluate the expression of the genes by quantitative real time PCR (Applied Biosystems 7500 Real Time PCR System). The relative

78 fold change in the target gene expression was quantified using 2 deltadeltaCt method by normalizing the target gene expression to RPLP0 169 as a housekeeping gene and relative to the expression on control group at day 14.

5.3.5. Statistical Analysis:

Statistical analysis was performed similarly as described in section 4.3.8.

5.4 Results:

DMMB assay result indicates that DS was incorporated in desired concentration in compacted collagen sheet (Figure 26).

The mechanical test result (Figure 27) shows that there is a significant decrease in elastic modulus in 10:1 group than control group (p = .02). However, there was no significant difference of elastic modulus between DS groups as well as between control and 30:1 groups. Strain increased significantly in higher DS concentration (10:1) group than control (p = .02) and lower DS (30:1) concentration group (p = .01).

Figure 27: Mechanical test result of the groups

Alamar blue assay (Figure 28) revealed that there was a moderate but significant decrease in cell numbers in higher DS group than control group at day 1. However, after day 7, there was no significant difference in cell numbers between groups. From day 1 to day 7 there was about 3.5- fold increase in all groups which indicates DS incorporation does not affect cell proliferation.

Figure 28: Cell proliferation did not affect by DS incorporation.

Gene expression results after day 14 (Figure 29) showed that DS incorporation did not affect the expression levels of COLIII, DCN, COMP and THP-4. However, DS incorporated group showed lower expression of COLI, indicating that DS incorporation has an adverse effect on tenogenesis. There was no DS dose dependent (10:1 vs 30:1) variation in any gene expression levels.

Figure 29: Effect of DS incorporation on tenogenic differentiation of human MSCs. After day 14, DS incorporation did not affect the expression levels of COLIII, DCN, COMP and TSP-4. DS incorporated group showed lower expression of COLI showed lower expression level in DS incorporated group. Horizontal lines indicate significant difference between groups.

5.5 Discussions:

This study investigated the effect of DS incorporation in compact sheets on tenogenic differentiation. This study showed that DS incorporation did not have adverse effect on cell proliferation. However, DS incorporation did not affect tenogenic differentiation also. Even in some case (e.g. COL I gene expression level) DS incorporation had unfavorable effect on tenogenesis.

The mechanical test result showed that low DS concentration (30:1 group) did not affect the elastic modulus or failure strain of the compact sheet. However, high concentration of DS (10:1 group) decrease the elastic modulus by 1.5 fold and increase failure strain by about 3%. The higher concentration of DS allowed the collagen fibril slipping within the sheet. This slipping of the collagen fibril increased failure strain which in turn decreased elastic modulus of the high DS concentration group. Scott, in his sliding proteoglycan-filament model hypothesized that GAG chains take part in sliding filament and converts local compressions into disseminated tensile strains246. Previous studies showed DS decreased collagen fibril diameter, retards the rate of fibril diameter increase and decrease mechanical strength of reconstituted collagen fibril 247-249 . In this study also, addition of higher concentration of DS may produce thinner collagen fibril which leaded to lower elastic modulus.

DS incorporation either did not affect (COLIII, DCN, COMP and THP-4) or negatively affect (COLI) gene expression levels. DS is abundant in tension bearing region of tendon. It has been shown that substrate tension increased tendon related gene expression 250, 251. Also overuse of tendon increased DS concentration and GAG deposit significantly increase in later time points

82 in tendon development while greater level of mechanical loading is involved, indicating that GAG content may be associated with the mechanical loading 252, 253. Studies have also shown that GAG content is related to the physical activity. For example, GAG content decreases with less physical activity and this effect can be reversed with increasing physical activity 35, 254, 255. All of these studies suggest that only incorporation of DS may not enough to effect tenogenic differentiation and may be mechanical loading also necessary which is an important factor associated with DS.

The biological effect of DS and DS containing PGS (Proteoglycans) depends on their complex structure 256. The variable DS chain length, disaccharide composition, and sulfation control functional interactions with potential protein associates and determines binding affinity.

The function of DS depends on both the GAG structure and the core protein that makes up a dermatan sulfate proteoglycan (DSPG) as it has been shown that, in specific developmental and physiologic conditions, DS sequence is influenced by variable expression through the PG core protein 257-259.

One of the most important DSPG is decorin which is a small leucine-rich proteoglycans

(SLRP)32, 256. SLRPs constitute a network of extracellular signal regulation and crucial for multiple signaling cascades260. Intracellular phosphorylation is a major channel of information for cellular response which is affected by SLRPs256, 260. They are believed to direct the functionality of tendons by binding to collagen fibrils and regulating collagen fibrillogenesis36. The leucine rich protein core of decorin binds non-covalently to the surface of the collagen fibrils261 and the GAG chain

(DS) binds to tenascin-X which colocalizes the neighboring collagen fibrils with one another to form inter- fibrillar crosslinks in connective tissue28, 256, 262-264. Therefore, the protein core is necessary to bind DSPG on Collagen fibril along with the DS chain for ECM assembly in connective tissue. In this study, we have used pure DS instead of DSPG which lacks the protein

83 core. This may be the reason for reduced expression of tendon related markers. DS in humans are composed of repeating disaccharide units of iduronic acid (IdoA) and GalNAc N-acetyl galactosamine. The presence IdoA in DS distinguishes it from chondroitin sulfates-A. Westergren-

Thorsson et al. showed that GAG chains containing high amounts of IdoA inhibit the proliferation of normal fibroblasts265. Adding DS instead of DSPG created only a IdoA rich composition which may lead to the down regulation of tendon related marker as shown by Westergren-Thorsson et al. showed.

5.6 Conclusion:

While there are extensive research studies on chondroitin sulfate loaded scaffolds in tendon repair and regeneration, there is a lack of studies related to DS scaffolds for tendon repair and regeneration. The current study investigates the effect of DS incorporation in tendon differentiation and reveals that DS does not affect tenogenic differentiation.

6. Chapter 6: Computer Aided Biomanufacturing of Mechanically Robust Pure Collagen Meshes with Controlled Macroporosity

6.1 Abstract

Reconciliation of high strength and high porosity in pure collagen based structures is a major barrier in collagen’s use in load-bearing applications. The current study developed a CAD/CAM based electrocompaction method to manufacture highly porous patterned scaffolds using pure collagen. Utilization of computerized scaffold design and fabrication allows the integration of mesh-scaffolds with controlled pore size, shape and spacing. Mechanical properties of fabricated collagen meshes were investigated as a function of number of patterned layers, and with different pore geometries. The tensile stiffness, tensile strength and modulus ranges from 10-50 N/cm, 1-

6 MPa and 5-40 MPa respectively for all the scaffold groups. These results are within the range of practical usability of different tissue engineering application such as tendon, hernia, stress urinary incontinence or thoracic wall reconstruction. Moreover, 3-fold increase in the layer number resulted in more than 5-fold increases in failure load, toughness and stiffness which suggests that by changing the number of layers and shape of the structure, mechanical properties can be modulated for the aforementioned tissue engineering application. These patterned scaffolds offer a porosity ranging from 0.8-1.5 mm in size, a range that is commensurate with pore sizes of repair meshes in the market. The connected macroporosity of the scaffolds facilitated cell-seeding such that cells populated the entire scaffold at the time of seeding. After 3 days of culture, cell nuclei became elongated. These results indicate that the patterned electrochemical deposition method in this study was able to develop mechanically robust, highly porous collagen scaffolds with controlled porosity which not only tries to solve one of the major tissue engineering

85 problems in a fundamental level but also has a significant potential to be used in different tissue engineering applications.

6.2 Introduction

Collagen is at the center of many tissue engineering strategies. It is a ubiquitous molecule that can be extracted from animal tissues. Generally, collagen is well tolerated in vivo 266-268. It presents cell adhesion sites and it can be digested enzymatically by cellular action. Owing to such favorable properties, collagen is used in many clinical applications 269-274. While pure collagen scaffolds present advantages in terms of cell response, they are weak and their use is mostly limited to non- load bearing applications such as barrier sheets, hemostatic chips and sponges to fill cavities 275-

282. Therefore, biofabrication modalities that would increase the mechanical robustness of collagen-based materials would expand the spectrum of applications for collagen.

Another challenge associated with collagen biofabrication is the introduction of porosity.

Porosity is essential for tissue integration and neo-vascularization. However, at the same time, porosity intrinsically reduces the mechanical strength of scaffolds. Classically, porosity is induced by freeze drying or salt-leaching283-285. While these methods are useful, porosity is random, has limiting interconnectedness and degree of control on the uniformity of pore size and shape is low.

Furthermore, most applications which require tissue integration and vascularization require macroscale porosity (0.5 mm or greater)286. Therefore, it is a major challenge to reconcile macroporosity with mechanical robustness in delicate protein-based biomaterials such as collagen.

Recent years have seen the introduction of the electrochemical compaction of collagen molecules to manufacture condensed tissue-analog forms126, 287, 288. In this approach, electrical currents are applied to collagen solutions. The resulting pH gradient in the solution results in the repulsion of collagen molecules by both electrodes and molecules become compacted between the electrodes. The electrocompaction method has been used for high strength collagen threads by

87 using parallel wire electrodes169, 289. In this study we developed a novel biofabrication method to manufacture patterned pore-lattice structures with controlled size and shape by computer-aided electrocompaction of collagen molecules using electrical currents. The aims of this study are: 1) to demonstrate the capabilities of patterned electrocompaction method in manufacturing scaffolds with different pore size and shapes; 2) report on mechanical characteristics of resulting scaffolds,

3) demonstrate the propensity of the macroporous scaffold for cell-seeding.

6.3 Materials and Method:

6.3.1. Overview of the Patterned Electrocompaction Process (Figure 30):

The method requires two planar carbon electrodes (each approximately 25x25 mm) for patterned deposition. A polycarbonate sheet of 2 mm in thickness is glued on the cathode using cyanoacrylate. This bilayered structure is then mounted on a computer controlled micromill

(Sherline CNC, Haverford CA). The negative replica of the desired pattern is designed in CAD environment (Solidworks) and it is machined on the plastic sheet in full depth to expose the underlying cathode’s surface. The channels machined within the polycarbonate layer as such are later filled with the collagen solution to be electrocompacted.

Figure 30: Process of fabricating individual patterned layer. Patterned electrochemical compaction of monomeric collagen solutions as mechanically robust lattice layers. The layers can be stacked to obtain thicker scaffolds.

6.3.2. Manufacturing an individual patterned layer

Acid soluble monomeric collagen solution (bovine dermis, Advanced Biomatrix, CA; 6 mg/ml) was diluted two-fold, pH was adjusted between 8-10 using 1N NaOH and dialyzed against ultrapure water for 18 hours. Dialyzed collagen was loaded in the patterned groves of the machined plastic-cathode bilayer by applying with a syringe. The carbon anode layer was placed on the top of the plastic-cathode bilayer. 30 VDC was applied for 2 min. across the electrodes. This results in electrophoretic compaction of collagen molecules within the patterned groove. The width of the groove was 1 mm for all of the scaffold groups. The biophysical mechanisms causing the electrocompaction of molecules were explained and discussed in detail before126, 287. Briefly,

89 electrical current generates acidic conditions near the anode and basic pH near the cathode.

Collagen has positive and negative net charge under acidic and basic conditions, respectively.

These charges are similar to the charges of the electrodes to which the molecules are close to, resulting in the repulsion of molecules from both electrodes. The net effect of repulsion is the compaction of molecules as a dense layer. Following electrocompaction, the plastic-cathode bilayer with the collagen deposit is recovered.

Three different pore geometries were manufactured to demonstrate the control on pore morphology: i) rectangular ii) square and iii) diamond. The average pore sizes for rectangular, square and diamond-shaped scaffolds are 1.5, 0.8 and 1.2 mm with corresponding porosities of

55%, 43% and 61%, respectively. To assess the effect of pore geometries on the mechanical properties of the scaffold, another pore architecture with parallel pore channels (group iv) with intermittent connections was manufactured. The parallel channel pore scaffold group has 53% porosity which is comparable to rectangular and diamond shape pore scaffold.

The computerization of the process allowed for fabrication of multiple layers in one electrode platform as shown here (Figure 31A&31B). The lattice structure can be designed to control the width of the channels and the spacing between them. An increased spacing or reduced channel width resulted in greater pore volume. The lattice was cast in a fashion to include a rectangular frame at the perimeter which to allow for recovering and handling the patterned deposit. The frame was also helpful in registering multiple patterned sheets to make multilayered scaffolds.

Figure 31: 3D-scaffolds can be obtained by overlaying the individual lattice layers. The pore network is staggered between each consecutive layer to attain interconnected porosity. The registration of the stagger pattern is accounted for during the CNC based machining of the electrode system. A) 3D-interconnected porosity can be attained by overlaying multiple layers while staggering the pore structure. Blue and black ‘ ’ denote the two separate scaffold layers. B) Layers can be patterned as parallel channel to induce collagen molecule alignment and make desired pore shaped scaffolds by staggering the individual layers.

Figure 32: Scaffolds with 4 different porosity shapes. A) Diamond-shaped pore; B) Square-shaped pore; C) Rectangle-shaped pore 4) Parallel channel pore. Black color of the scaffolds are due to genipin cross linking. Black arrow indicates porosity through the thickness of the scaffold and blue arrow indicates spaces between the filaments of the layers of the scaffold.

6.3.3. Fabrication of 3-D Scaffolds with Controlled Interconnected Porosity:

The individual patterned layers were stacked on top of each other in register to obtain the final scaffold as shown in Figure 31A&31B. Collagen solution (6 mg/mL, pH = 6) was brushed between layers as a binder prior to stacking the layers. The layered structure was kept under load

92 under a deadweight for an hour, placed in 1x PBS at 37 °C for 6 hours to induce fibrillogenesis as we have shown before132. Scaffolds were stored in 2-propanol solution when not used. Scaffolds were crosslinked in 0.625% genipin (Wako Chemical, Japan) in 90% v/v ethanol solution at 37 °C for 3 days289.

To evaluate the effect of number of layers on the overall mechanical properties of the scaffold, square pore scaffolds with 3 different layer numbers: v) 2 layers vi) 4 layers and vii) 6 layers were manufactured. The lateral dimensions of the individual layer as well as the final scaffolds were 20 X 12 mm.

6.3.4. Imaging of Molecular Alignment within Patterned Channels:

After manufacturing the patterned layer, the molecular alignment of collagen molecules within the electrocompacted pattern was examined by a polarized optical microscope (Olympus

BX51, Melville, NY, USA) and a first order wavelength gypsum plate. Collagen is a positive birefringent material, where the molecules which are aligned along the slower axis of this plate shows blue interference color and the molecules which are perpendicular to the slow axis appear yellow176. Magenta color indicates lack of alignment and blue color indicates alignment of the collagen molecule along the slower axis or perpendicular to the long dimension of the gypsum plate.

6.3.5. Assessment of Mechanical Properties of Patterned Bioscaffolds:

Patterned scaffolds with different pore geometries and different layer numbers (N =

6/group) were tested under monotonic tension until failure using a displacement rate of 10 mm/min

(RSA II, Rheometrics Inc., Piscataway, NJ, USA). Scaffolds were hydrated in water for 30 min prior to testing. The ends of the scaffolds were gripped by tensile fixture at a gauge length of 10

93 mm. A 10 N load-cell was used to measure the load. The thickness of the scaffolds was measured by a custom-made micrometer where the micrometer closes an electrical circuit and makes an audible sound in a multimeter when touches the surface of wet scaffolds. Load values were normalized with the wet cross sectional area and displacement values were normalized by the gage length to obtain stress-strain plots. Area is calculated as the product of number of collagen filament in the tensile direction, width of the filaments and average thickness of the filaments in the tensile direction of the scaffolds. Failure load, stiffness, elastic modulus, failure stress and toughness were calculated from stress-strain curves. Elastic modulus and stiffness were calculated at the steepest region of the stress strain curves. Toughness was calculated as the area under the stress-strain curves.

6.3.6. Adhesion and Morphology of Human Mesenchymal Stem Cells Seeded on Patterned Bioscaffolds

To demonstrate that the 3D-controlled porosity enables cell-loading throughout the continuum of the patterned scaffold, cells were seeded on 6-layered scaffolds (10 mm X 5 mm X

0.5 mm, rectangular porosity with 1.5 mm pore size) following sterilization in 70% ethanol overnight. Scaffolds were placed in an ultralow attachment 24-well plate (1 scaffold/well). Human mesenchymal stem cells (Lonza, Walkersville, MD) were seeded at a density of 250,000 cells/well.

The culture medium was composed of alpha MEM (Invitrogen) supplemented with 10% MSC-

FBS (Invitrogen), 1% penicillin/streptomycin and 50 μg/mL ascorbic acid. Twelve hours after seeding, the unattached cells were removed by replacing the growth medium and the cells adherent on the scaffolds were cultured for 3 days. Cell morphology was visualized by staining the cell actin filaments using AlexaFluor 488 Phalloidin (Life Technologies, Grand Island, NY, USA) at day 3.

Briefly, cells were fixed with 3% formaldehyde (with 0.1% TritonX-100) for 10 min and washed with 1x PBS. The actin filaments were stained by incubating the cells in AlexaFluor 488 Phalloidin

94 at 37 0C for 20 min. The stain was washed with 1x PBS and images of actin stained cells were taken using an Olympus Microscope. To visualize the cell nucleus, DAPI nucleic acid staining

(Invitrogen) was also performed after day 3.

6.3.7. Benchmark for Mechanical Performance of Scaffolds:

The long term goal of this project is to apply these scaffolds to rabbit infraspinatus tendon

(RIT) repair. Therefore, the structural load displacement behavior of RIT was chosen as the benchmark. This tendon is about 10 mm x 5 mm by 10 mm. Mechanical tests of Intact Bone-

Tendon-Muscle Complex was performed to Identify Target Strength/Stiffness Values for Scaffold.

Six (6) rabbit (New Zealand White, 1 yr. old, female) shoulders were dissected from fresh rabbit carcasses obtained from the Animal Resource Center (ARC) of Case Western Reserve University.

Since the tissue were collected post-mortem from another study which did not involve the shoulder region, IACUC approval was not applicable. The shoulders were dissected such that all the soft tissues were removed except the humerus–infraspinatus–muscle unit. The humeri were potted in

(poly) methylmethacrylate cement (Millennium Pour Denture Acrylic, Cherry Hill, NJ) up to 20 mm distant from the humeral head inside a hollow rectangular aluminum pipe which was fixed to the loading frame (Figure 32). The tendon muscle complex was gripped at the muscle by fixtures at 15 mm distance from the tendon-bone insertion. The muscle was frozen locally at the grip site by a piece of dry ice. The tensile fixture was also cooled by dry ice, all facilitating a solid grip at the fixture. The intact infraspinatus tendons were loaded in a physiologically relevant direction of the infraspinatus tendon perpendicular to the longitudinal axis of the humerus (Figure 32). The samples were loaded monotonically at a rate of 10 mm/min until failure (Test Resource 800LE3-

2, Test Resources Inc., MN, USA). A 220 N load cell was used to measure the load. All the samples were kept hydrated at all stages of suturing and testing. Stiffness was calculated at the steepest

95 region of the load-displacement curves. Following simulations and experiments was aimed to emulate this benchmark.

6.3.8. Computational Screening of Scaffold Morphologies:

The numbers of variables for designing the patterned scaffold morphology are quite large.

A preliminary computational screening of mechanical behavior was employed in four scaffold geometries. They are: a) sinusoidal filament with zigzag shape pore b) linear filament with square pore c) linear filament with rectangular pore and d) linear angular filament with diamond shape pore [Figure 33A]. The model will simulate load-displacement behavior of the multilayered lattice structure to determine morphologies which converges to the stiffness of equivalent tendon structure with highest porosity. The morphological variables of the individual lattice layers are filament width and cross sectional area of porosity. The lattice morphology considered linear filaments and sinusoidal filaments (to emulate the crimp pattern inherent in tendon). For any given lattice morphology, the simulations were executed at a constant filament width (500 µm) with four types of pore shapes. Scaffold geometries were designed in Solidworks and simulation was performed in Ansys workbench 12.0. As different geometry of the same material was compared, for simplicity, the material was modeled as a simple linear solid and nonlinearity wasn’t considered. Tensile test data (RSAII, Rheometrics Inc., Piscataway, NJ) of collagen sheet was employed to describe the material properties in FEM analysis. The load-displacement profile that is the most convergent to the load displacement profile of RIT can be considered suitable for tendon repair scaffold.

The preliminary goal was to investigate which geometries give maximum porosity with maximum stiffness. Finite element result shows that linear filament with rectangular porosity gives

maximum stiffness close to native tendon [Figure 33B]. A relative comparison between porosity

and stiffness [Figure 32C&D] showed that with similar porosity level, diamond shape pores give

more stiffness whereas with a high porosity than diamond shape pore, rectangular pore gives more

stiffness. This y result indicates that patterned scaffold with rectangular porosity has higher

potential to meet the requirement benchmark of 3-D scaffold for tendon repair.

d) Linear angular filament- a) Sinusoidal filament- b) Linear filament- c) Linear filament- diamond shape pores zigzag pores square pores rectangular pores

6.3.9. Statistical Analysis:

A one-way analysis of variance (ANOVA) test was performed to evaluate for significant differences between groups. Two sets of groups were analyzed separately for significant difference within the groups. The first set was different pore geometries groups: i) rectangular ii) square iii) diamond-shaped pore and iv) parallel channel pore. The second set was square pore scaffolds with: v) 2 layers vi) 4 layers and vii) 6 layers. Significant differences between means were calculated and significance level was set at p < 0.05. A post-hoc analysis using the Tukey’s test was conducted to compare pairwise differences between groups.

6.4 Results:

Four types of patterned lattice structures were fabricated (Figure 32). The collagen filaments of the scaffold with diamond-shaped pores lacked alignment (magenta in CPI image in

Figure 34A). In contrast, the scaffolds which are fabricated with parallel filaments had collagen molecules aligned parallel to the longer axes of the filaments as evidenced by blue coloration

(Figure 34B).

5 mm

Figure 34: A) Branching in the diamond-shaped patterned lattice displayed lack of alignment of collagen molecules as manifested by the magenta in the compensated polarize image (CPI). B) Parallel channels introduce alignment in the patterned layer (emergence of blue color in CPI indicates alignment).

The typical stress-strain plots of scaffolds with increasing number of layers (group v, vi and vii) showed that 2-layered scaffold failed abruptly with limited plastic deformation. 4-layered and 6-layered groups showed prominent post-yield deformability (Figure 35a).

There was a non-linear increase in the structural mechanical properties of scaffolds with increasing number of layers. Such that, 3-fold increase (2 layer to 6 layer) in the number of layers resulted in increases in more than 3-fold increase in failure load (7-fold), toughness (5.5-fold) and stiffness (7- fold) (Fig. 35b-f).

Figure 35: Mechanical assessment of patterned scaffolds with three different layer numbers. (a) Typical stress-strain curves for different layer scaffolds, (b) Failure load, (c) Elastic modulus, (d) Toughness (e) Failure stress and (f) Stiffness of 6 layer scaffold is more than 3 fold greater than the 2 layer scaffolds. The horizontal line indicates significant difference (p <0.05).

The stress-strain plots of the different porosity geometry shape showed that the scaffolds with diamond-shaped pores failed more abruptly than square/rectangular shape pore and parallel channel pore scaffolds (Figure 36a). Failure stress, elastic modulus, stiffness and toughness were highest for rectangular/square-shaped pore scaffolds, intermediate for parallel channel pore

100 scaffolds and lowest for diamond-shaped pore scaffolds and these properties increased about 6- fold, 7-fold, 2.5-fold and 8-fold respectively from diamond-shaped pore scaffold to square shape pore scaffold (Figure 36b-f). There were no significant differences between the material properties of rectangular and square-shaped pore scaffolds (Figure 36b-f).

Figure 36: Mechanical assessment of patterned scaffolds with four different pore shapes. (a) Typical stress-strain curves for different porosity scaffolds, (b) Failure stress, (c) Elastic modulus, (d) Toughness (e) Failure load and (f) Stiffness of rectangular/square-shaped pore scaffolds were highest, parallel channel pore scaffolds were intermediate and diamond-shaped pore scaffolds were lowest. Asterisks indicate significant differences between groups (p <0.05).

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F-actin and DAPI stained images taken from fields of view located in the deeper layers of scaffolds (Figure 37) revealed that cells were uniformly seeded over the filaments. Actin cytoskeleton of cells was elongated along the longer axis of the threads. The nuclei also became elongated along the length of the collagen filament in the scaffolds (Figure 37) with a nuclear aspect ratio of 2.15 ± 0.34 after 3 days of culture.

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6.5 Discussion:

A novel method of manufacturing, patterned planar electrochemical compaction of pure collagen was developed. The method fabricated mechanically promising patterned scaffolds with controlled porosity. Different pattern types were generated to show the versatility of the fabrication. The process involved the design of desired pore configuration via computer aided design and the machining of the pattern by computer controlled numerical machine tool. These computational modalities provided the control over 3D scaffold morphology.

There are several methods to prepare porous three-dimensional biodegradable scaffolds, including gas foaming290, 291, phase separation292-294, porogen leaching295, 296, fiber extrusion and bonding297, emulsion freeze-drying,298 and solid free-form fabrication (SFF) such as 3-D printing299, 300, bioplotting301-303, selective laser sintering304. Gas foaming process may produce structure with largely unconnected pores and a non-porous external surface. Particulate leaching is limited for thicker scaffolds, phase separation is limited by the number of materials included in the formulation, and SFF of collagen generally requires the inclusion of secondary polymers to provide consistency. SFF also require expensive specialized instruments and mass production via

SFF may present challenges. Electrospinning and fiber meshes uses fibers to make porous structure. However, electrospinning uses toxic solvents and control over pore size and shape is not possible. Moreover electrospinning and other techniques usually utilize a secondary synthetic polymer besides the collagen to enable manufacturability 305-309. Freeze drying provides porous collagen scaffolds and pore size and elongation can be controlled to some extent 283, 285, 310, 311. On the other hand, freeze drying process generally produces weak structures with elastic modulus and failure stress values in the kPa range62. There is no single technique which can provide mechanically robust pure collagen based 3-D scaffolds with controlled porosity. The demonstrated

103 electrocompaction method addresses this limitation to a significant extent. Importantly, the method lends itself to scaled-up production by using arrays of electrodes over large surface areas.

Mechanical test results of different pattern types and layer numbers suggest that the patterned scaffold manufactured by the method in this study has a potential to be used in the biomaterials and tissue engineering fields where strength and controlled porosity is required. The enhancement in mechanical properties is largely associated with the ability of the method to compact the pure monomeric collagen to solid phase within the channels of the cathode (Figure

1). Prior studies from our group demonstrated up to 300 fold increase in packing density of collagen (3 mg/mL to 1030 mg/mL) 154 molecules. The alignment of collagen molecules along the length of the channels also contributes to mechanical robustness.

Increasing the layer number has a disproportionately greater benefit on the mechanical properties of meshes. From 2 layer to 4 layer scaffold the failure load increased about 2.5 fold whereas from 2 layer to 6 layer scaffold it increased about 7 fold. Similarly, stiffness and toughness increased 7 fold and 5.5 fold respectively from 2 layer to 6 layer. Two-layer scaffolds failed abruptly at the end of the linear elastic region while as the layer number increased, the 4-layered and 6-layered scaffolds showed prominent post yield deformability. This increase is mechanical properties and prominent post yield deformability in increasing layer numbers may be a result of additional layers complementing the weak points in other layers. This support, in turn, results in a nonlinear increase in the mechanical properties with increasing number of layers. This result also indicates that layer numbers can be increased as an option to match the strength of native tissues.

There were variations between mechanical properties for variation of pattern type (e.g. rectangular/square- shaped pore, parallel channel pore and diamond-shaped pore). Diamond- shaped pore scaffold showed significantly inferior mechanical performance than the other types of

104 scaffold. This outcome was associated with lack of molecular alignment in the diamond-shaped patterned pores. In case of rectangular/square and parallel channel pore scaffold, the direction of loading is same as the direction of the collagen filament in the patterned scaffold which helps the scaffold to withstand greater load. Conversely, collagen filaments are obliquely oriented to the loading direction in diamond-shaped pore which may have reduced the strength and the stiffness.

Although the diamond-shaped pore may be weak in tensile direction, they may perform better where uniform load distribution is required due to the angular filament distribution. Parallel channel pore showed intermediate failure load due to the weak points which arose from lack of alignment in the interconnections between filaments.

The connected macroporosity of the scaffolds facilitated cell-seeding. Cells populated the entire scaffold at the time of seeding. After 3 days of culture, cell nuclei became elongated with an aspect ratio of 2.15. Native tendon, ligament, cardio myocyte and muscle cell has elongated nucleus with nucleus aspect ratio ranging from 2- 6 which is important in promoting a tenogenic, myogenic and ligament phenotypes196, 312-316. The observed elongation of cell morphology may be beneficial in engineering such tissues using the proposed scaffold concept.

Unification of mechanical robustness with ample amount of macroporosity renders the electrocompacted collagen scaffolds for mesh-based applications which require rapid tissue ingrowth and neovascularization. Such applications include hernia repair, stress urinary incontinence (SUI), vaginal prolapse, thoracic wall reconstruction and tendon tissue engineering.

At the present, such applications use synthetic polymers (e.g. polypropylene, polytetrafluoroethylene, polyester, etc.) autografts or decellularized allo/xenografts. Synthetic polymers provide acceptable mechanical properties. However they may present issues regarding

105 cell adhesion62, 63, systemic or local reactions 64, 65. Biodegradable synthetic polymer scaffolds may be associated with foreign body giant cells57.

To our knowledge pure collagen based scaffolds has not been used in clinical mesh application due to their weak mechanics [Table 3]. For clinical mesh application, to achieve mechanical robustness in collagen based scaffold, mostly collagen rich decellularized tissue such as small intestine submucosa (SIS), porcine acellular dermal matrix, and abdominal fascia

(decellularized xenografts or allografts) were used317, 318. Decellularization may not always be fully effective and cell remnants may impact the scaffold host response, immune cell infiltration, tumor necrosis factor-α expression and macrophage activation66. Chemicals used in decellularization and radiation used for antigen deactivation results in damage to the extracellular matrix which in turn may drive premature degradation of the implant before tissue integration takes place. The scaffold presented in this paper is a bottom-up fabrication sequence which utilizes pure collagen stock.

Prior animal studies using electrocompacted threads demonstrated a high level of biocompatibility267. Therefore, incomplete removal of antigens is not a limitation for electrocompacted collagen. Moreover previous in vitro study of electrocompacted collagen showed favorable cell proliferation128, 289 and differentiation128 to tenogenic lineage. To the best of our knowledge, this is the first time a pure collagen scaffold mesh is reported within the mechanical strength range of decellularized tissue based xenografts [Table 3].

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Table 3: Comparative mechanical properties of collagen based scaffolds for tissue engineering application

Market Compressive Compressive Tensile Elastic Collagen Based Scaffold Name Modulus Strength Strength Modulus

Collagen rich xenografts (decellularized collagen matrix) Permacol317 38 MPa 210 MPa CollaMend317 10MPa 30 MPa Porcine dermis Strattice317 15 MPa 50 MPa XenMatrix317 12 MPa 40MPa Pelvitex319 1.2 MPa 5 MPa FlexHD317 10MPa 30 MPa Human dermis AlloMax317 22 MPa 60 MPa Veritas317 6MPa 20 MPa Bovine pericardium PeriGuard317 16 MPa 110 MPa Porcine SIS Surgisis317, 319 4 MPa 15 MPa Human abdominal fascia320 2 MPa 10 MPa 0.2 -3.5 Porcine dermis321 MPa Animal SIS Strasis322 3 MPa Human cadaveric dermis Alloderm323 7.2 MPa Human cadaveric fascia lata Faslata323 10.85 MPa Decellularized tendon section 0.5- 6 324 0.2-0.7 MPa (Sheet form) MPa Electrocompacted

Pure collagen (reconstituted collagen) patterned scaffolds in this 1-6 MPa 5-40 MPa study Gel form (dogbone shape) 324 0.1 MPa 0.2 MPa Electrospun Collagen325 0.3 MPa 0.4 MPa Collagen, Collagen+ Elastin 350-200 25 kPa 12 kPa 80 kPa (freeze dry) 326 kPa Collagen freeze dry327 7.8 kPa 81 kPa Collagen–glycosaminoglycan 5.1 kPa 30 kPa (freeze dry) 310 Gel form (dog bone shape) 1.54-25 0.5-9 kPa 328 kPa Collagen (freeze dry) 329 20 kPa Collagen-chitosan (freeze 10-20 kPa dry) 329

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All of the aforementioned mesh based application requires mechanical robustness to a certain extent. Hernia repair meshes requires flexibility and optimized tensile strength to reduce discomfort. Polymer based meshes for hernia repair may be too strong and result in movement restriction and pain330. The tensile strength of a mesh required to withstand the maximum abdominal pressure is 32 N/cm330 which is only a tenth of that of most meshes available now.

High tensile strength polymers may be associated with erosion or extrusion in SUI repair331. The range of apparent tensile strength of existing xenograft and polypropylene sling products are 2-12

MPa and the apparent modulus range is 5-15 MPa332. Abdominal fascia used in autograft procedures in SUI application have a strength of up to 2 MPa and modulus of 10 MPa320. Polymer based meshes for thoracic diaphragm reconstruction has mechanical strength 25-40 N/cm333.

However, thoracic reconstruction with synthetic materials are more prone to infection which necessitates costly removal procedure334. In the current study, six-layered scaffolds with different pore geometries have mechanical properties such as tensile stiffness, tensile strength and modulus ranges from 10-50 N/cm, 1-6 MPa and 5-40 MPa respectively which are within the range of practical usability of these applications. These patterned scaffolds also have porosity size ranging from 0.8-1.5 mm which is also within the range of most of the present repair meshes in the market.

Moreover, the tensile strength can be optimized by changing the layer number of the scaffold, whenever optimization of tensile strength is required.

6.6 Conclusion

In summary, the current study developed a CAD/CAM based method to manufacture a pure collagen based highly porous patterned scaffold. High compaction and alignment of the collagen molecules rendered the construct mechanically robust. The results suggest that by changing the number of layers and shape of the structure, mechanical properties can be modulated

108 for different tissue engineering application such as tendon, hernia, SUI, thoracic wall reconstruction.

The inability to incorporate high strength and high porosity in a structure is one of the major barriers in the engineering of load-bearing tissue, and the fabricated structure in this method addresses this limitation to some extent. This method utilizes computerized scaffold design and fabrication which allows the integration of ‘scaffolds with controlled porosity’. There is a general lack of biofabrication methods that will provide controlled porosity and the patterned electrochemical deposition method in this study has a potential to address this challenge at the fundamental level.

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7. Chapter 7: Summary and Future Directions

The inability to incorporate high strength and high porosity in a structure has been one of the major barriers in the engineering of load-bearing tissue. There is a general lack of biofabrication methods that will provide macroporosity with mechanical robustness in delicate protein-based biomaterials such as collagen. The patterned lattice structure fabrication method, in this dissertation has a potential to combine these characteristics. This method utilizes computerized scaffold design and fabrication which allows the integration of macroscale porosity (0.5 mm or greater), an essential feature for populating a scaffold with cells and vasculature. The lattice structures are made up of anisotropically aligned sheet like lattice structures. Therefore, the dissertation also develops a material model which also considers stiffness as well as stiffness anisotropy to mimic two key features of natural tendon.

Most natural tissues are substantially stronger along the load bearing direction than the direction transverse to the longer axis (such as tendons, muscle etc.). Notably, the cells of such tissues are elongated along the stiffer direction. There are not many biomaterials to emulate and study the effects of stiffness anisotropy on cellular response. In addition to the stiffness anisotropy, extracellular matrix stiffness or mechanics also plays an important role to regulate cell morphology, proliferation, differentiation during regular and diseased states. Although the effects of substrate topography and stiffness on MSC differentiation are well known with respect to osteogenesis and adipogenesis, there has been relatively little investigation of this phenomenon with respect to tenogenesis. Overall, the roles of topographical factors (matrix anisotropy vs. matrix stiffness) in inducing tenogenic differentiation are unknown. Elucidation of the mechanism of topographically induced tenogenesis in human MSCs will assist in optimizing the material to

110 produce scaffolds for tendon repair and may provide insight into the differentiation mechanisms of MSCs and other stem cells in vivo, where type I collagen is the major substrate protein.

Two key features of natural tendon such as effect of stiffness and stiffness anisotropy on tenogenesis were studied in this study. Stiffness anisotropy was introduced in electrocompacted sheets by controlled unidirectional stretch of the sheets to various levels and Stiffness was.

Variations in the cytoskeletal and nuclear morphology of cells seeded on collagen sheets, the expression of tendon-related transcription factors and tendon-related extracellular matrix synthesis of varying SA were investigated. The stiffness value of type I collagen substrate were tuned in range of 0.1-100 MPa by using ectrocompaction and crosslinking processes. A systematic investigation of the effects of matrix stiffness on tenocytic differentiation and matrix synthesis by

MSCs on collagen sheets was investigated. The merit to inclusion of dermatan sulfate in the stiffer matrix to increase tenogenic differentiation was also studied. Finally, the current study developed a CAD/CAM based method to manufacture a pure collagen based highly porous patterned scaffold.

Effect of stiffness anisotropy and stiffness study demonstrated that stem cell fate is affected by not only the magnitude of stiffness but also by the directional anisotropy of the substrate stiffness.

The study suggests that increasing stiffness anisotropy has a positive effect on tenogenesis.

However, highly stiff substrate without SA can generate similar tenogenic differentiation as substrate with SA. This indicates, may be stiffness can overcome the effect of stiffness anisotropy on tenogenesis. Overall, SA has a positive effect on tenogenesis as anisotropy can be a key determinant in driving cell morphology and differentiation during development and maintenance of anistropically stiff tissues. In the SA study demonstrated a case for tendon differentiation and future studies will investigate genotypes and phenotypes associated with other anisotropic tissues such as skeletal or cardiac muscles. In the stiffness study, we developed a material profile where

111 the modulus can be changed from several hundred kPa, to single digit MPa, to hundred MPa. The tenoinductive collagen material model developed in the study can be used in the research and development of tissue engineering tendon repair constructs in future. The DS incorporation study revealed that only DS inclusion does not affect the tenogenic differentiation of the substrate and it may require other parameter such as mechanical loading, to investigate the effect of DS inclusion.

This patterned lattice structure fabrication method utilizes computerized scaffold design and fabrication which allows the integration of ‘scaffolds with controlled porosity’. The method demonstrates the capabilities of manufacturing mechanically robust scaffolds with different pore size and shapes. In this method, by changing the number of layers and shape of the structure, mechanical properties can be modulated for different tissue engineering application such as tendon, hernia, SUI, thoracic wall reconstruction. Pure collagen based scaffolds has not been used in clinical mesh application due to their weak mechanics. For clinical mesh application, to achieve mechanical robustness in collagen based scaffold, mostly collagen rich decellularized tissue based xenografts are used. The fabrication method developed in this study generates pure collagen scaffold mesh within the mechanical strength range of decellularized tissue.

In future studies, highly anisotropic sheet with stiffness ratio 8 along with the isotropically stiff substrate can be employed to elucidate the role of cytoskeletal re-organization of MSCs in inducing tenocytic differentiation. It has been shown that Rho-associated kinase (ROCK) and non- muscle myosin II effects focal adhesion formation and focal adhesion kinase which are crucial in topography-mediated differentiation.

It has been shown that MSCs on soft substrates usually have smaller focal adhesion, poorly aligned stress fibers which in turn produce less cell spreading than MSCs on stiff substrates225, 234.

Soft substrates inhibit Rho-induced stress fiber formation and α-actin assembly. Therefore, soft

112 substrate shows unspread and round cells234. Moreover, inhibition of RhoA/Rock results in reduced focal adhesion formation and changes fibroblast like cells to stellate like cells 226, 335-337. .

On the other hand, in stiff substrate, RhoA and ROCK activation promotes acto-myosin stress fiber assembly and demonstrates long parallel actin stress fiber with elongated cellular shape338.

Therefore, RhoA/ROCK may be major molecular pathway that promotes matured focal adhesion, organized stress fibers and elongated cells in stiff substrate

Previous studies showed that RhoA pathway was involved in MSC differentiation.

Inhibition of RhoA resulted in adipogenic differentiation339, 340. Rho GTPase was also involved in chondrogenic, myogenic and neurogenic differentiation234, 341, 342 The result of this dissertation showed a predominant cytoskeletal alignment of MSCs along the stiffer axis of anisotropically stiff collagen substrates343 which promotes tenogenesis. RhoA signaling may governed the onset of this topography mediated differentiation by affecting SMAD signaling in the downstream as it has been shown that SMAD manipulation promoted tenoenesis336 (Figure 38). It has been also showed that SMAD 2/3 signalling increased upregulation of tendon related markers344 . The involvement of RhoA and other downstream molecules such ROCKII, Myosin II, Actin can be probed by the corresponding inhibitors such as Y-27632, Blebbistatin, Cytochalasin D or actin polymerization stabilizer such as Jasplakinolide (Figure 38). MSCs seeded both on highly anisotropic aligned sheet as well as isotropic sheet can be employed to probe these molecules.

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Figure 38: Cytoskeletal arrangement and tenogenic differentiation of MSCs

The patterned scaffold in this study can be designed to generate scaffolds with sinusoidal filament [As in Figure 33A] to mimic the crimp pattern in natural tendon. This crimp patterned scaffold may result similar stress-strain plot like tendon.

The patterned lattice scaffolds developed in this dissertation has a potential to use for other tissue engineering application such as hernia, SUI, thoracic wall reconstruction. Therefore, in future expressions of osteogenic (Runx2, osteopontin and osteocalcin), chondrogenic (SOX9, aggrecan and collagen Type II) and adipogenic (PPARγ, LPL and adiponectin) markers can be assessed on the developed material model to pursue differentiation other than tenogenesis.

In future studies, the anisotropic and stiff electrocompacted pure collagen sheet can be used to develop 3D cell-laden collagen scaffolds by sandwiching cells between two sheets. Cells can help adhering the adjacent sheets. A preliminary study, showed that, cells were alive between two sandwiching sheets after day 1 and 3 which indicates that cells got enough nutrient between the two sheets to survive. Cells in natural tendons are in a 3D aligned micro environment. Therefore, 114 this method has a potential to investigate tenogenic differentiation of MSCs in similar 3D environment as natural tendon. A tube shaped 3D cell laden scaffold can also be fabricated by rolling the aligned sheets. Intermittent holes along the periphery towards the central core hole of the rolled scaffold can introduce interconnected porosity. The scaffold can be populated with cells along the core hole as well as the peripheral porosity. This type of scaffold can also be used in ligament and tendon tissue engineering

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Appendix A. Patterned Polycarbonate-Cathode Bilayer Manufacturing Steps

Appendix A1. Preparation of Polycarbonate-Cathode Bilayer

Cyanoacrylate glue (Loctite Instant Adhesive 495, Henkel, Australia) is applied uniformly on one of the carbon electrodes (cathode). The polycarbonate sheet is then glued on the cathode. During gluing, press down the plastic, if there is any bubble. The plastic-cathode bilayer is then kept under load for 30 minutes before using it for next step.

Steps for Making Polycarbonate-Cathode Bilayer

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Appendix A2. CAD/CAM Design of Pattern Mold

The negative replica of the desired pattern is designed in CAD environment (Solidworks). For simple pattern, a G-Code is written to machine the intended pattern on the plastic-cathode bilayer. For complicated pattern a CAM software (Meshcam, GRZ Software) is used to cut the desired pattern. An example of complicated pattern is given below:

A Complicated Pattern Suitable for Using in MeshCAM

Some Sample G-Code is provided in the following section.

Procedure for using Meshcam is given in the following section

Appendix A3. G-Code for CNC Machining of Negative Replica of Patterned Scaffold.

Square Pore:

(Line Program) g01 g21 g40 g49 g90 x0 y0 z0 f2 g01 z120.53 f10 g01 x0 f15 s1250 m3 g01 z121.03 f5 g01 z121.94 f5 g00 x0 y2.6 f150 (Pass1) g01 x-24 f15 g01 x-24 f15 g00 z120.5 f60 g01 z121.48 f5 g01 z122.41 f5

117 g01 x0 f15 g01 z120.53 f10 g01 y7.1 f15 g01 z122.68 f5 g01 z121.03 f5 g01 z122.68 f5 g01 x-24 f15 g01 y2.1 f15 g01 y10.1 f15 g01 z122.88 f5 g01 z121.48 f5 g01 z122.88 f5 g01 x0 f15 g01 y1 f15 g01 y7.1 f15 g00 z115 f60 g01 z121.94 f5 g00 z115 f60 g00 x0 y6.6 f150 (Pass2) g01 y2.1 f15 g00 x-6.25 y11.1 f150 (Pass5) g00 z120.5 f60 g01 z122.41 f5 g00 z120.5 f60 g01 z120.53 f10 g01 y1 f15 g01 z120.53 f10 g01 z121.03 f5 g01 z122.68 f5 g01 z121.03 f5 g01 x-24 f15 g01 y2.1 f15 g01 y14.1 f15 g01 z121.48 f5 g01 z122.88 f5 g01 z121.48 f5 g01 x0 f15 g01 y1 f15 g01 y11.1 f15 g01 z121.94 f5 g00 z115 f60 g01 z121.94 f5 g01 x-24 f15 g00 x-6.25 y3.1 f150 (Pass2) g01 y14.1 f15 g01 z122.41 f5 g00 z120.5 f60 g01 z122.41 f5 g01 x0 f15 g01 z120.53 f10 g01 y11.1 f15 g01 z122.68 f5 g01 z121.03 f5 g01 z122.68 f5 g01 x-24 f15 g01 y6.1 f15 g01 y14.1 f15 g01 z122.88 f5 g01 z121.48 f5 g01 z122.88 f5 g01 x0 f15 g01 y3.1 f15 g01 y11.1 f15 g00 z115 f60 g01 z121.94 f5 g00 z115 f60 g00 x0 y10.6 f150 (Pass3) g01 y6.1 f15 g00 x-17.75 y11.1 f150 (Pass6) g00 z120.5 f60 g01 z122.41 f5 g00 z120.5 f60 g01 z120.53 f10 g01 y3.1 f15 g01 z120.53 f10 g01 z121.03 f5 g01 z122.68 f5 g01 z121.03 f5 g01 x-24 f15 g01 y6.1 f15 g01 y14.1 f15 g01 z121.48 f5 g01 z122.88 f5 g01 z121.48 f5 g01 x0 f15 g01 y3.1 f15 g01 y11.1 f15 g01 z121.94 f5 g00 z115 f60 g01 z121.94 f5 g01 x-24 f15 g00 x-17.75 y3.1 f150 (Pass3) g01 y14.1 f15 g01 z122.41 f5 g00 z120.5 f60 g01 z122.41 f5 g01 x0 f15 g01 z120.53 f10 g01 y11.1 f15 g01 z122.68 f5 g01 z121.03 f5 g01 z122.68 f5 g01 x-24 f15 g01 y6.1 f15 g01 y14.1 f15 g01 z122.88 f5 g01 z121.48 f5 g01 z122.88 f5 g01 x0 f15 g01 y3.1 f15 g01 y11.1 f15 g00 z115 f60 g01 z121.94 f5 g00 z115 f60 g00 x0 y14.6 f150 (Pass4) g01 y6.1 f15 g00 x-12 y15.1 f150 (Pass7) g00 z120.5 f60 g01 z122.41 f5 g00 z120.5 f60 g01 z120.53 f10 g01 y3.1 f15 g01 z120.53 f10 g01 z121.03 f5 g01 z122.68 f5 g01 z121.03 f5 g01 x-24 f15 g01 y6.1 f15 g01 y16.2 f15 g01 z121.48 f5 g01 z122.88 f5 g01 z121.48 f5 g01 x0 f15 g01 y3.1 f15 g01 y15.1 f15 g01 z121.94 f5 g00 z115 f60 g01 z121.94 f5 g01 x-24 f15 g00 x-12 y7.1 f150 (Pass4) g01 y16.2 f15 g01 z122.41 f5 g00 z120.5 f60 g01 z122.41 f5 g01 x0 f15 g01 z120.53 f10 g01 y15.1 f15 g01 z122.68 f5 g01 z121.03 f5 g01 z122.68 f5 g01 x-24 f15 g01 y10.1 f15 g01 y16.2 f15 g01 z122.88 f5 g01 z121.48 f5 g01 z122.88 f5 g01 x0 f15 g01 y7.1 f15 g01 y15.1 f15 g00 z115 f60 g01 z121.94 f5 g00 z115 f60 g00 x-12 y1 f150 (Pass1) g01 y10.1 f15 (end program) g00 z120.5 f60 g01 z122.41 f5 m5

118 g00 z0 f150 g00 x0 f150 % g00 y0 f150 m2

Rectangular Pore:

% g00 z115 f60 g01 z120.53 f10 (Line Program) g00 x-6.25 y1 f150 (Pass1) g01 z121.03 f5 g01 g21 g40 g49 g90 x0 y0 z0 f2 g00 z120.5 f60 g01 y12.1 f15 s1250 m3 g01 z120.53 f10 g01 z121.48 f5 g00 x0 y4.6 f150 (Pass1) g01 z121.03 f5 g01 y9.1 f15 g00 z120.5 f60 g01 y4.1 f15 g01 z121.94 f5 g01 z120.53 f10 g01 z121.48 f5 g01 y12.1 f15 g01 z121.03 f5 g01 y1 f15 g01 z122.41 f5 g01 x-24 f15 g01 z121.94 f5 g01 y9.1 f15 g01 z121.48 f5 g01 y4.1 f15 g01 z122.68 f5 g01 x0 f15 g01 z122.41 f5 g01 y12.1 f15 g01 z121.94 f5 g01 y1 f15 g01 z122.88 f5 g01 x-24 f15 g01 z122.68 f5 g01 y9.1 f15 g01 z122.41 f5 g01 y4.1 f15 g00 z115 f60 g01 x0 f15 g01 z122.88 f5 g00 x-17.75 y9.1 f150 (Pass5) g01 z122.68 f5 g01 y1 f15 g00 z120.5 f60 g01 x-24 f15 g00 z115 f60 g01 z120.53 f10 g01 z122.88 f5 g00 x-17.75 y1 f150 (Pass2) g01 z121.03 f5 g01 x0 f15 g00 z120.5 f60 g01 y12.1 f15 g00 z115 f60 g01 z120.53 f10 g01 z121.48 f5 g00 x0 y8.6 f150 (Pass2) g01 z121.03 f5 g01 y9.1 f15 g00 z120.5 f60 g01 y4.1 f15 g01 z121.94 f5 g01 z120.53 f10 g01 z121.48 f5 g01 y12.1 f15 g01 z121.03 f5 g01 y1 f15 g01 z122.41 f5 g01 x-24 f15 g01 z121.94 f5 g01 y9.1 f15 g01 z121.48 f5 g01 y4.1 f15 g01 z122.68 f5 g01 x0 f15 g01 z122.41 f5 g01 y12.1 f15 g01 z121.94 f5 g01 y1 f15 g01 z122.88 f5 g01 x-24 f15 g01 z122.68 f5 g01 y9.1 f15 g01 z122.41 f5 g01 y4.1 f15 g00 z115 f60 g01 x0 f15 g01 z122.88 f5 g00 x-12 y13.1 f150 (Pass6) g01 z122.68 f5 g01 y1 f15 g00 z120.5 f60 g01 x-24 f15 g00 z115 f60 g01 z120.53 f10 g01 z122.88 f5 g00 x-12 y5.1 f150 (Pass3) g01 z121.03 f5 g01 x0 f15 g00 z120.5 f60 g01 y16.2 f15 g00 z115 f60 g01 z120.53 f10 g01 z121.48 f5 g00 x0 y12.6 f150 (Pass3) g01 z121.03 f5 g01 y13.1 f15 g00 z120.5 f60 g01 y8.1 f15 g01 z121.94 f5 g01 z120.53 f10 g01 z121.48 f5 g01 y16.2 f15 g01 z121.03 f5 g01 y5.1 f15 g01 z122.41 f5 g01 x-24 f15 g01 z121.94 f5 g01 y13.1 f15 g01 z121.48 f5 g01 y8.1 f15 g01 z122.68 f5 g01 x0 f15 g01 z122.41 f5 g01 y16.2 f15 g01 z121.94 f5 g01 y5.1 f15 g01 z122.88 f5 g01 x-24 f15 g01 z122.68 f5 g01 y13.1 f15 g01 z122.41 f5 g01 y8.1 f15 g00 z115 f60 g01 x0 f15 g01 z122.88 f5 (end program) g01 z122.68 f5 g01 y5.1 f15 m5 g01 x-24 f15 g00 z115 f60 g00 z0 f150 g01 z122.88 f5 g00 x-6.25 y9.1 f150 (Pass4) g00 y0 f150 g01 x0 f15 g00 z120.5 f60 g00 x0 f150

119 m2 %

Parallel Pore:

% g01 y0 f15 g01 x0 f15 (Line Program) g01 x0 f15 g00 z116 f60 g01 g21 g40 g49 g90 x0 y0 z0 f2 g00 z116 f60 g00 x0 y0 f150 (rectangle Pass5) s1250 m3 g00 x0 y0 f150 (rectangle Pass3) g00 z120.45 f60 g00 x0 y0 f150 (rectangle Pass1) g00 z120.45 f60 g01 z122.41 f10 g00 z120.45 f60 g01 z121.48 f10 g01 z122.88 f5 g01 z120.53 f10 g01 z121.94 f5 g01 y17.2 f15 g01 z121.03 f5 g01 y17.2 f15 g01 x-24 f15 g01 y17.2 f15 g01 x-24 f15 g01 y0 f15 g01 x-24 f15 g01 y0 f15 g01 x0 f15 g01 y0 f15 g01 x0 f15 g00 z116 f60 g01 x0 f15 g00 z116 f60 (finishing) g00 z116 f60 g00 x0 y0 f150 (rectangle Pass4) m05 (spindle stop) g00 x0 y0 f150 (rectangle Pass2) g00 z120.45 f60 g00 z0 f150 g00 z120.53 f60 g01 z121.94 f10 g00 y0 f150 g01 z121.03 f10 g01 z122.41 f5 g00 x0 f150 g01 z121.48 f5 g01 y17.2 f15 m2 g01 y17.2 f15 g01 x-24 f15 % g01 x-24 f15 g01 y0 f15

Diamond Pore

% g01 z102.5335 f10 g00 Z102 f60 (Line Program) g01 z102.9835 f5 g00 x156 y9.413 f150 (Pass6) g01 g21 g40 g49 g90 x0 y0 z0 g01 x134 y10.6015 f15 g00 Z102 f60 f2 g01 z103.4435 f5 g01 z102.5335 f10 s1250 m3 g01 x156 y13.9652 f15 g01 z102.9835 f5 g00 x156 y17 f150 (Pass1) g01 z103.98 f5 g01 x134 y6.0493 f15 g00 Z102 f60 g01 x134 y10.6015 f15 g01 z103.4435 f5 g01 z102.5335 f10 g00 Z102 f60 g01 x156 y9.413 f15 g01 z102.9835 f5 g00 x156 y12.4478 f150 (Pass4) g01 z103.98 f5 g01 x134 y13.6363 f15 g00 Z102 f60 g01 x134 y6.0493 f15 g01 z103.4435 f5 g01 z102.5335 f10 g00 Z102 f60 g01 x156 y17 f15 g01 z102.9835 f5 g00 x156 y7.8956 f150 (Pass7) g01 z103.98 f5 g01 x134 y9.0841 f15 g00 Z102 f60 g01 x134 y13.6363 f15 g01 z103.4435 f5 g01 z102.5335 f10 g00 Z102 f60 g01 x156 y12.4478 f15 g01 z102.9835 f5 g00 x156 y15.4826 f150 (Pass2) g01 z103.98 f5 g01 x134 y4.5319 f15 g00 Z102 f60 g01 x134 y9.0841 f15 g01 z103.4435 f5 g01 z102.5335 f10 g00 Z102 f60 g01 x156 y7.8956 f15 g01 z102.9835 f5 g00 x156 y10.9304 f150 (Pass5) g01 z103.98 f5 g01 x134 y12.1189 f15 g00 Z102 f60 g01 x134 y4.5319 f15 g01 z103.4435 f5 g01 z102.5335 f10 g00 Z102 f60 g01 x156 y15.4826 f15 g01 z102.9835 f5 g00 x156 y6.3782 f150 (Pass8) g01 z103.98 f5 g01 x134 y7.5667 f15 g00 Z102 f60 g01 x134 y12.1189 f15 g01 z103.4435 f5 g01 z102.5335 f10 g00 Z102 f60 g01 x156 y10.9304 f15 g01 z102.9835 f5 g00 x156 y13.9652 f150 (Pass3) g01 z103.98 f5 g01 x134 y3.0145 f15 g00 Z102 f60 g01 x134 y7.5667 f15 g01 z103.4435 f5

120 g01 x156 y6.3782 f15 g00 Z102 f60 g01 z103.98 f5 g01 z103.98 f5 g01 z102.5335 f10 g01 x134 y15.1538 f15 g01 x134 y3.0145 f15 g01 z102.9835 f5 g00 Z102 f60 g00 Z102 f60 g01 x155.1468 y17 f15 g00 x156 y3.244 f150 (Pass7) g00 x156 y4.8608 f150 (Pass9) g01 z103.4435 f5 g00 Z102 f60 g00 Z102 f60 g01 x156 y16.6385 f15 g01 z102.5335 f10 g01 z102.5335 f10 g01 z103.98 f5 g01 z102.9835 f5 g01 z102.9835 f5 g01 x155.1468 y17 f15 g01 x134 y12.9273 f15 g01 x134 y1.4971 f15 g00 Z102 f60 g01 z103.4435 f5 g01 z103.4435 f5 g00 x156 y14.1049 f150 (Pass2) g01 x156 y3.244 f15 g01 x156 y4.8608 f15 g00 Z102 f60 g01 z103.98 f5 g01 z103.98 f5 g01 z102.5335 f10 g01 x134 y12.9273 f15 g01 x134 y1.4971 f15 g01 z102.9835 f5 g00 Z102 f60 g00 Z102 f60 g01 x150.0203 y17 f15 g00 x156 y1.0723 f150 (Pass8) g00 x156 y3.3434 f150 (Pass10) g01 z103.4435 f5 g00 Z102 f60 g00 Z102 f60 g01 x156 y14.1049 f15 g01 z102.5335 f10 g01 z102.5335 f10 g01 z103.98 f5 g01 z102.9835 f5 g01 z102.9835 f5 g01 x150.0203 y17 f15 g01 x134 y10.7552 f15 g01 x134 y-0.0203 f15 g00 Z102 f60 g01 z103.4435 f5 g01 z103.4435 f5 g00 x156 y11.9328 f150 (Pass3) g01 x156 y1.0723 f15 g01 x156 y3.3434 f15 g00 Z102 f60 g01 z103.98 f5 g01 z103.98 f5 g01 z102.5335 f10 g01 x134 y-0.0203 f15 g01 z102.9835 f5 g01 x134 y10.7552 f15 g00 Z102 f60 g01 x144.8938 y17 f15 g00 Z102 f60 g00 x156 y1.826 f150 (Pass11) g01 z103.4435 f5 g00 x155.4373 y-0.5 f150 g00 Z102 f60 g01 x156 y11.9328 f15 (Pass9) g01 z102.5335 f10 g01 z103.98 f5 g00 Z102 f60 g01 z102.9835 f5 g01 x144.8938 y17 f15 g01 z102.5335 f10 g01 x140.7889 y-0.5 f15 g00 Z102 f60 g01 z102.9835 f5 g01 z103.4435 f5 g00 x156 y9.7607 f150 (Pass4) g01 x134 y8.5831 f15 g01 x156 y1.826 f15 g00 Z102 f60 g01 z103.4435 f5 g01 z103.98 f5 g01 z102.5335 f10 g01 x155.4373 y-0.5 f15 g01 x140.7889 y-0.5 f15 g01 z102.9835 f5 g01 z103.98 f5 g00 Z102 f60 g01 x139.7673 y17 f15 g01 x134 y8.5831 f15 g00 x156 y0.3086 f150 (Pass12) g01 z103.4435 f5 g00 Z102 f60 g00 Z102 f60 g01 x156 y9.7607 f15 g00 x150.3108 y-0.5 f150 g01 z102.5335 f10 g01 z103.98 f5 (Pass10) g01 z102.9835 f5 g01 x139.7673 y17 f15 g00 Z102 f60 g01 x150.7136 y-0.5 f15 g00 Z102 f60 g01 z102.5335 f10 g01 z103.4435 f5 g00 x156 y7.5886 f150 (Pass5) g01 z102.9835 f5 g01 x156 y0.3086 f15 g00 Z102 f60 g01 x134 y6.411 f15 g01 z103.98 f5 g01 z102.5335 f10 g01 z103.4435 f5 g01 x150.7136 y-0.5 f15 g01 z102.9835 f5 g01 x150.3108 y-0.5 f15 g00 Z102 f60 g01 x134.6408 y17 f15 g01 z103.98 f5 g00 x146.0753 y17 f150 g01 z103.4435 f5 g01 x134 y6.411 f15 (Pass13) g01 x156 y7.5886 f15 g00 Z102 f60 g00 Z102 f60 g01 z103.98 f5 g00 x145.1842 y-0.5 f150 g01 z102.5335 f10 g01 x134.6408 y17 f15 (Pass11) g01 z102.9835 f5 g00 Z102 f60 g00 Z102 f60 g01 x134 y15.1538 f15 g00 x156 y5.4165 f150 (Pass6) g01 z102.5335 f10 g01 z103.4435 f5 g00 Z102 f60 g01 z102.9835 f5 g01 x146.0753 y17 f15 g01 z102.5335 f10 g01 x134 y4.2389 f15 g01 z103.98 f5 g01 z102.9835 f5 g01 z103.4435 f5 g01 x134 y15.1538 f15 g01 x134 y15.1538 f15 g01 x145.1842 y-0.5 f15 g00 Z102 f60 g01 z103.4435 f5 g01 z103.98 f5 g00 x156 y16.6385 f150 (Pass1) g01 x156 y5.4165 f15 g01 x134 y4.2389 f15

121 g00 Z102 f60 g01 x134 y-0.1053 f15 g00 x140.0577 y-0.5 f150 g01 z103.4435 f5 (Pass12) g01 x134.9312 y-0.5 f15 g00 Z102 f60 g01 z103.98 f5 g01 z102.5335 f10 g01 x134 y-0.1053 f15 g01 z102.9835 f5 g00 Z102 f60 g01 x134 y2.0668 f15 m5 g01 z103.4435 f5 g00 z0 f150 g01 x140.0577 y-0.5 f15 g00 y0 f150 g01 z103.98 f5 g00 x0 f150 g01 x134 y2.0668 f15 m2 g00 Z102 f60 % g00 x134.9312 y-0.5 f150 (Pass13) g00 Z102 f60 g01 z102.5335 f10 g01 z102.9835 f5

Appendix A4. Procedure for using Meshcam

1. Export the pattern CAD model as an STL file

2. Open the STL file in MeshCAM

3. Chose 3-axis machining for Sherline CNC

4. Select “Rotate Geometry” Chose Z-axis to be the spindle axis, and the X-Y plane to

match the table orientation.

5. Click the “Translate Geometry” button, and select “Make geometry positive X and Y”

and “Make top of part zero as shown in the following image.

122

6. Click “Define Stock.” Measure the plastic-cathode bilayer size and enter the size in

the “Stock Size” panel. MeshCAM will automatically populate the “XY Position” and “Z

Position” panels, centering the part in the stock. These values can be changed to position

the part relative to the stock if necessary. Please refer to the following picture.

123

7. Select “Retract Height.” This sets the distance that the cutter will retract to above

the material during rapid moves. Use judgment. Usually 10 mm.

8. Set “Program Zero.” This is an important step. Usually one of the corner of the material

stock. Set the Z position to be the top of the stock. Set the XY position to the bottom left

corner

124

9. Click “Set Max Depth.” This will be the maximum depth of cut on your part. In this case,

it will be the thickness of the plastic.

125

10. Select “Generate Toolpath.” In the “Global Parameters” section, set the tolerance to 0.001-

0.005 inches. Select “Machine geometry only,” which will just cut to the geometry of your

original STL file, plus whatever “Machining Margin” was added. In “Roughing,” select

“Enable Roughing Pass” and click “Select Tool.”

11. You can add different types of cutters or edit existing ones, with options to set the

measurements of the cutter as well as the different associated rates. Chose the default Step

over, 0.5 inch Federate and default plunge rate.

12. Click OK. MeshCAM will calculate the toolpath

13. Click “Estimate Machining Time.” for machining time estimation.

14. “Save Toolpath.” Select the “Sherline EMC-MM(*.nc)” and save the .nc file which is the

G-Code file.

15. Take the .nc file and open in Sherline CNC software to machine the plastic-cathode bilayer.

126

Appendix A5. Machining Pattern Mold

The G-Code is loaded into the CNC software and the negative replica of the desired pattern is machined on the plastic sheet in full depth to expose the underlying cathode’s surface.

CNC Machining of a Plastic-cathode bilayer

Patterns after CNC Machining

127

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