A Dissertation
entitled
Mechanically-Conditioned Biphasic Composite Scaffolds to Augment Healing of
Tendon-Bone Interface
by
Gayathri Subramanian
Submitted to the Graduate Faculty as partial fulfillment of the requirements for the
Doctor of Philosophy Degree in Biomedical Engineering
______Dr. Eda Yildirim-Ayan, Committee Chair
______Dr. Beata Lecka-Czernik, Committee Member
______Dr. Yakov Lapitsky, Committee Member
______Dr. Ronald Fournier, Committee Member
______Dr. Arunan Nadarajah, Committee Member
______Dr. Amanda Bryant-Friedrich, Dean College of Graduate Studies
The University of Toledo
August 2017
Copyright 2017, Gayathri Subramanian
This document is copyrighted material. Under copyright law, no parts of this document may be reproduced without the expressed permission of the author. An Abstract of
Mechanically-Conditioned Biphasic Composite Scaffolds to Augment Healing of Tendon-Bone Interface
by
Gayathri Subramanian
Submitted to the Graduate Faculty as partial fulfillment of the requirements for the Doctor of Philosophy Degree in Biomedical Engineering
The University of Toledo
August 2017
Rotator cuff injuries are very common among people over the age of 60, with more than
600,000 surgeries performed annually in the United States for rotator cuff repairs.
However, in 20-80% of the cases, the repair fails due to re-rupture of the tendon at the tendon-bone insertion site. The complexity of the tendon tissue in terms of their structure, composition, and function at the tendon-to-bone interface demands for a combinatorial tissue-engineering approach in which cell maturation and function can be directed using bioactive proteins encapsulated within a biomaterial with appropriate material stiffness.
Further, since tendons experience routine mechanical strains in their native environment, providing suitable mechanical cues to the engineered scaffold was considered important for the success of rotator cuff repair strategies.
The objective of this dissertation was to synthesize and characterize a mechanically- conditioned biphasic composite collagen scaffold to enhance rotator cuff regeneration with
(1) controlled delivery of adipose-derived stem cells (ASCs) and platelet-derived growth factor (PDGF) to augment and accelerate tendon healing, and (2) spatial material stiffness to promote gradient mineralization and matrix directionality at the tendon-bone interface. iii
To this end, a mechanical loading bioreactor consisting of unique silicone loading chambers was designed that was capable of applying homogenous uniaxial tensile strains over 60% of the length of cell-encapsulated 3D collagen scaffolds. Uniaxial tensile mechanical loading at 2% strain with 0.1 Hz frequency was identified to be the appropriate loading modality to induce pure ASC tenogenic differentiation, along with enhanced matrix directionality and ECM gene expression within ASC-encapsulated 3D collagen scaffolds. Next, the poor protein retention and matrix stiffness properties of collagen were improved by synthesizing a composite collagen scaffold (PNCOL) interspersed with functionalized polycaprolactone (PCL) nanofibers. The PDGF-conjugated PNCOL scaffolds demonstrated controlled release of bioactive proteins (0.5% per day) under uniaxial tensile mechanical loading. Finally, the PDGF/ASC-encapsulated COLPNCOL biphasic composite scaffold with gradient matrix stiffness was engineered and subjected to uniaxial tensile mechanical loading to mimic the tissue at the tendon-bone insertion point.
Significantly, the COLPNCOL biphasic scaffold demonstrated two major morphological and biochemical characteristics which is representative of the tendon- fibrocartilage region of the native tendon-bone interface tissue: (1) A gradient increase in mineral deposits and a gradient decrease in the matrix alignment. (2) Elevated tenogenic expression with COL region and higher chondrogenic expression within PNCOL region.
This mechanically-conditioned PDGF/ASC-encapsulated COLPNCOL biphasic scaffold is expected to provide sustained cells and growth factor delivery to induce tenogenic differentiation and ECM secretion, and promote gradient matrix mineralization and collagen alignment of the de novo tissue, thereby aiding and accelerating the natural tendon-bone interface healing process for functional regeneration of the rotator cuff.
iv
To my friends and family, by blood and soul,
To them, who inspired me far beyond my goals,
To the magic of science & the mysteries it holds,
I dedicate this study, to each one and all...
Acknowledgements
I would like to express my sincere and warm gratitude to Dr. Eda Yildirim-Ayan for providing me with the opportunity and resources to pursue my passion in research. Her technical and editorial expertise along with her constant guidance and encouragement were instrumental in the successful completion of this study. I am truly fortunate to have had her as my advisor. Her mentorship has contributed immensely to my growth, both as an individual and a professional. I also would like to thank my committee members: Dr.
Beata-Lecka Czernik, Dr. Yakov Lapitsky, Dr. Ronald Fournier, and Dr. Arunan Nadarajah for their valuable inputs and feedback that improved the content of this dissertation.
A big thank you to all my previous and current Engineered Biosystems Lab members. In particular, Mostafa Elsaadany, for being a friend and a partner in crime during the last 5 years – right from setting up the lab, mentoring students, troubleshooting experiments, managing ‘crisis’ situations, attending conferences, going through the grind of I-Corps, to graduating together! Special thanks to Maggie Ditto – for the warm welcome I received when I first joined the lab; Callan Bialorucki – for those effective brainstorming sessions every time I hit a roadblock, and Andrew Trumbull – for being a good sounding board during the progress of my work.
Finally, my sincere thanks to Tammy Phares, for her unconditional help and support during the course of my research, and for making my 5-year stint as the Teaching Assistant of the
Bioprocessing Lab enjoyable and memorable. v
Table of Contents
Abstract ...... iii
Acknowledgements ...... v
Table of Contents ...... vi
List of Tables ...... xvi
List of Figures ...... xvii
List of Abbreviations ...... xxii
1 Background and Overview ...... 1
1.1 Rotator cuff tendon injuries ...... 1
1.2 Surgical repair of rotator cuff tendon ...... 2
1.3 Tendon anatomy and physiology ...... 3
1.4 Tendon healing process...... 5
1.5 Commercial solutions for rotator cuff repair ...... 7
1.6 Cells or growth factor delivery for tendon repair ...... 8
1.7 Tissue engineering approach for tendon repair ...... 9
1.8 State-of-the-art in tissue-engineered scaffolds for tendon repair ...... 11
1.8.1 Hydrogel-based scaffolds to augment tendon healing ...... 11
1.8.2 Synthetic and composite scaffolds for partial tendon repair ...... 13
vi
1.8.3 Mechanically-conditioned scaffolds for tendon reconstruction .... 16
1.8.4 Stratified scaffolds for tendon-bone interface repair...... 17
2 Objective and Thesis Outline ...... 19
2.1 Rationale ...... 19
2.2 Objective ...... 20
2.3 Thesis outline ...... 25
3 A Mechanical Loading Bioreactor to Apply Uniaxial Tensile Strains to Cell- encapsulated 3D Collagen Scaffolds ...... 28
3.1 Introduction ...... 28
3.2 Materials and Methods ...... 34
3.2.1 Design and fabrication of the uniaxial tensile strain bioreactor .... 34
3.2.1.1 Components of the uniaxial tensile strain bioreactor ...... 34
3.2.1.2 Loading chamber of the uniaxial tensile strain bioreactor .... 37
3.2.1.3 Material selection ...... 39
3.2.2 Experimental strain validation with collagen scaffolds ...... 40
3.2.3 Generation of Finite Element Model for the bioreactor ...... 42
3.2.4 Cell culture, scaffold synthesis, and mechanical loading
regimes…...... 45
vii
3.2.5 Cell viability and proliferation of mechanically-loaded
scaffolds…...... 47
3.2.6 Matrix organization of mechanically-loaded scaffolds ...... 48
3.2.7 Statistical analysis ...... 49
3.3 Results ...... 49
3.3.1 Operation and performance of the bioreactor ...... 49
3.3.1.1 Application of cyclic loads with the uniaxial tensile strain
bioreactor ...... 49
3.3.1.2 Experimental validation of the uniaxial tensile strain
bioreactor ...... 52
3.3.2 Characterization of bioreactor-induced strain profiles using
FEM…...... 54
3.3.2.1 Generation and validation of FEMs for the uniaxial tensile
strain bioreactor ...... 54
3.3.2.2 Bioreactor-induced longitudinal tensile strain profiles within
3D scaffolds ...... 57
3.3.2.3 Bioreactor-induced transverse compressive strains profile
within scaffolds...... 61
3.3.2.4 Bioreactor-induced creep strain profiles with 3D scaffolds . 62
viii
3.3.2.5 Effect of frequency and cycle number on the longitudinal
strain profile ...... 64
3.3.3 Biological characterization of mechanically-loaded scaffolds ...... 67
3.3.3.1 Viability of cells within mechanically-loaded 3D collagen
scaffolds...... 67
3.3.3.2 Cell proliferation within mechanically-loaded collagen
scaffolds...... 69
3.3.3.3 Matrix organization of mechanically-loaded 3D collagen
scaffolds...... 70
3.4 Discussion ...... 71
3.5 Conclusion ...... 80
4 Effect of Uniaxial Strains and Frequencies on the Matrix Organization and
Tenogenic Differentiation of Adipose-derived Stem Cells encapsulated within 3D
Collagen Scaffolds ...... 82
4.1 Introduction ...... 82
4.2 Materials and Methods ...... 89
4.2.1 Cell culture, scaffold synthesis and mechanical loading
regimes…… ...... 89
4.2.2 Viability of ASCs within 3D collagen scaffolds ...... 90
4.2.3 Proliferation of ASCs within 3D collagen scaffolds ...... 91
ix
4.2.4 Matrix directionality of ASC-encapsulated 3D collagen
scaffolds…...... 92
4.2.5 Gene expression analyses of ASCs encapsulated within
scaffolds…...... 93
4.2.6 Statistical analysis ...... 97
4.3 Results ...... 98
4.3.1 Matrix alignment of loaded ASCs-encapsulated scaffolds ...... 98
4.3.2 Viability and proliferation of ASCs within loaded 3D
scaffolds…… ...... 103
4.3.3 ECM gene expression of ASCs within loaded 3D scaffolds ...... 106
4.3.4 Tenogenic expression of ASCs within loaded 3D scaffolds ...... 112
4.3.5 Non-tenogenic lineage expression of ASCs within loaded
scaffolds...... 114
4.4 Discussion ...... 120
4.5 Conclusion ...... 129
5 Composite Collagen Scaffolds (PNCOL) Interspersed with Polycaprolactone
(PCL) Nanofibers for Controlled Growth Factor Delivery under Uniaxial Tensile
Mechanical Loading...... 132
5.1 Introduction ...... 132
5.2 Materials and Methods ...... 139
x
5.2.1 Synthesis of protein-immobilized composite collagen
scaffolds…… ...... 139
5.2.2 Protein retention capacity of PNCOL scaffolds ...... 142
5.2.3 Growth factor, cells, and culture media ...... 143
5.2.4 Bioactivity of GF-PNCOL scaffolds ...... 145
5.2.5 Growth factor release kinetics from GF-hb-PNCOL scaffolds ... 146
5.2.6 Long-term in vitro performance of GF-hb-PNCOL scaffolds .... 147
5.2.7 Protein release from PNCOL scaffolds under mechanical
loading…...... 149
5.2.8 Bioactivity of PDGF released from PDGF-hb-PNCOL ...... 150
5.2.9 Statistical analyses...... 152
5.3 Results ...... 153
5.3.1 Synthesis of PCL nanofibers-interspersed collagen scaffolds .... 153
5.3.2 Characterization of PNCOL scaffolds for protein delivery ...... 153
5.3.2.1 Protein retention capacity of PNCOL scaffolds ...... 153
5.3.2.2 Effect of PCL nanofibers on the protein bioactivity of within
PNCOL… ...... 155
5.3.3 Restoring and preserving protein activity within PNCOL ...... 157
5.3.3.1 Effect of heparin and BSA on protein bioactivity within
PNCOL scaffolds ...... 157
xi
5.3.3.2 Release kinetics of protein from GF-hb-PNCOL
scaffolds…… ...... 159
5.3.4 Long-term evaluation of GF-hb-PNCOL scaffolds ...... 160
5.3.4.1 Cell Proliferation within GF-hb-PNCOL scaffolds ...... 161
5.3.4.2 Bioactivity of GF-hb-PNCOL scaffolds ...... 162
5.3.5 Performance of GF-hb-PNCOL under uniaxial tensile loading .. 164
5.3.5.1 Protein release profile of mechanically-stimulated PNCOL
scaffolds...... 164
5.3.5.2 Bioactivity of PDGF released from PDGF-hb-PNCOL
scaffolds ……………………………………………………………..167
5.4 Discussion ...... 170
5.5 Conclusion ...... 178
6 PDGF/ASC-encapsulated COLPNCOL Biphasic Scaffolds with Gradient
Directionality, Mineralization, and Tenogenesis under Uniaxial Tensile Mechanical
Loading ...... 180
6.1 Introduction ...... 180
6.2 Materials and Methods ...... 186
6.2.1 Matrix stiffness of COL and PNCOL scaffolds ...... 186
6.2.2 Mineralization potential of COL and PNCOL scaffolds ...... 187
xii
6.2.3 Characterization of PDGF/ASC-encapsulated COLPNCOL
biphasic scaffold cultured under uniaxial tensile loading ...... 188
6.2.3.1 Synthesis and culture of biphasic scaffold ...... 188
6.2.3.2 Matrix morphology of biphasic scaffold...... 190
6.2.3.3 Gene expression profiles within biphasic scaffold ...... 191
6.2.3.4 Secreted extracellular matrix proteins from biphasic
scaffold… ...... 192
6.2.4 PDGF-induced migration and infiltration of fibrocartilage-like cells
into COLPNCOL biphasic scaffolds...... 193
6.2.5 Gradient mineralization within COLPNCOL biphasic scaffolds
cultured under uniaxial tensile loading ...... 195
6.2.5.1 ALP activity within COL and PNCOL regions of biphasic
scaffolds ……………………………………………………………..196
6.2.5.2 Mineralization within COL and PNCOL regions of biphasic
scaffolds……...... 197
6.2.6 Statistical analyses...... 198
6.3 Results ...... 198
6.3.1 Mechanical properties of Collagen and PNCOL scaffolds ...... 198
6.3.1.1 Material stiffness of COL and PNCOL scaffolds ...... 198
xiii
6.3.1.2 Mineralization potential of Collagen and PNCOL
scaffolds…...... 200
6.3.2 ASC differentiation and matrix alignment within mechanically-
conditioned COLPNCOL biphasic scaffolds ...... 202
6.3.2.1 Matrix morphology of COLPNCOL biphasic
scaffolds……...... 203
6.3.2.2 ECM gene expression profiles within COLPNCOL
biphasic scaffold ...... 208
6.3.2.3 Lineage-specific gene expression within COLPNCOL
biphasic scaffold ...... 209
6.3.2.4 Secreted extracellular matrix proteins from the biphasic
scaffold…… ...... 214
6.3.3 PDGF-induced migration and infiltration of fibrocartilage-like cells
into COLPNCOL biphasic scaffolds...... 215
6.3.4 Estimation of mineralization gradient within COLPNCOL
biphasic scaffolds cultured under uniaxial tensile loading ...... 218
6.3.4.1 ALP activity within COLPNCOL biphasic scaffolds ..... 218
6.3.4.2 Matrix mineralization within COLPNCOL biphasic
scaffolds ……………………………………………………………..220
6.4 Discussion ...... 225
xiv
6.5 Conclusion ...... 233
7 Future Work ...... 236
7.1 Recommendations for study described in Chapter 3…………………...232
7.2 Recommendations for study described in Chapter 4…………………...233
7.3 Recommendations for study described in Chapter 5…………………...234
7.4 Recommendations for study described in Chapter 6…………………...235
8 References ...... 243
xv
List of Tables
Table 1. Ogden model parameters used for the silicone loading chambers...... 43
Table 2. Viscoelastic Prony series parameters used for collagen scaffolds...... 44
Table 3. Quantification of viable cell number from live-dead images...... 68
Table 4. Forward and Reverse Primers used for Real-time PCR ...... 96
xvi
List of Figures
Figure 2-1. Schematic of the various tissue engineering components involved in each
Specific Aim of this thesis...... 23
Figure 3-1. Schematic of the uniaxial tensile strain bioreactor...... 35
Figure 3-2. Schematic of the silicone loading chamber: (A) Top view (B) Front view. .. 38
Figure 3-3. Experimental validation of the uniaxial tensile strain bioreactor. (A) Before loading (B) After loading...... 41
Figure 3-4. Generation of FEM for the uniaxial tensile strain bioreactor...... 44
Figure 3-5. Cyclic loading of 3D scaffolds using the uniaxial tensile strain bioreactor. .. 50
Figure 3-6. Performance of the uniaxial tensile strain bioreactor...... 53
Figure 3-7. Representative deformation contour from FEM...... 54
Figure 3-8. Finite Element Model validation for static loading...... 55
Figure 3-9. Finite Element Model Validation for Dynamic (Cyclic) Loading...... 56
Figure 3-10. Schematic of the regions in the collagen scaffold used to investigate the bioreactor-induced longitudinal tensile strain profile using FEM...... 57
Figure 3-11. Bioreactor-induced longitudinal tensile strain profiles across the length of
3D collagen scaffolds predicted using FEM...... 58
Figure 3-12. Bioreactor-induced longitudinal tensile strain profile across the width and thickness of 3D collagen scaffolds predicted using FEM...... 60
xvii
Figure 3-13. Bioreactor-induced tensile and compressive linear strains within 3D scaffolds predicted using FEM...... 62
Figure 3-14. Percentage equivalent creep strain experienced by the collagen scaffolds predicted by FEM...... 63
Figure 3-15. Bioreactor-induced strain profiles within 3D scaffolds at different loading frequencies predicted using FEM...... 64
Figure 3-16. Bioreactor-induced strain and stress profiles within 3D scaffolds during cyclic loading predicted using FEM...... 66
Figure 3-17. Effect of the uniaxial tensile strain bioreactor on cell viability within 3D scaffolds through experimental determination...... 68
Figure 3-18. Effect of bioreactor on cell proliferation within 3D scaffolds through experimental determination...... 69
Figure 3-19. Effect of bioreactor on collagen matrix organization within cell- encapsulated collagen scaffolds through experimental determination...... 70
Figure 3-20. Major results obtained in Chapter 3...... 80
Figure 4-1. Effect of uniaxial tensile loading at 0.1 Hz frequency on matrix organization of ASC-encapsulated 3D collagen scaffolds...... 99
Figure 4-2. Effect of uniaxial tensile loading at 1 Hz frequency on matrix organization of
ASC-encapsulated 3D collagen scaffolds...... 100
Figure 4-3. Effect of uniaxial tensile loading on matrix organization of ASC-encapsulated
3D collagen scaffolds...... 102
Figure 4-4. Effect of uniaxial tensile loading on ASCs viability within 3D scaffolds. .. 104
xviii
Figure 4-5. Effect of uniaxial tensile loading on ASC proliferation within 3D scaffolds.
...... 105
Figure 4-6. Effect of uniaxial tensile loading on Collagen expression of ASCs within 3D collagen scaffolds...... 107
Figure 4-7. Effect of uniaxial tensile loading on GAG expression of ASCs within 3D collagen scaffolds...... 110
Figure 4-8. Effect of uniaxial tensile loading on tenogenic gene expression of ASCs within 3D collagen scaffolds...... 113
Figure 4-9. Effect of uniaxial tensile loading on osteogenic gene expression of ASCs within 3D collagen scaffolds...... 116
Figure 4-10. Effect of uniaxial tensile loading on chondrogenic gene expression of ASCs within 3D collagen scaffolds...... 118
Figure 4-11. Effect of uniaxial tensile loading on myogenic gene expression of ASCs within 3D collagen scaffolds...... 119
Figure 4-12. Effect of uniaxial tensile loading on ASC differentiation within 3D scaffolds...... 128
Figure 4-13. Major results obtained in Chapter 4. The effect of uniaxial strains and frequencies on the matrix organization and tenogenic differentiation of adipose stem cells
(ASC) encapsulated within 3D collagen scaffolds...... 130
Figure 5-1. Synthesis of GF-hb-PNCOL composite scaffolds...... 139
Figure 5-2. Effect of PCL nanofiber concentrations in model protein retention within
PNCOL scaffolds...... 154
xix
Figure 5-3. Effect of PCL nanofiber concentrations on protein bioactivity within PNCOL.
...... 156
Figure 5-4. Restoring and preserving the protein bioactivity within PNCOL scaffolds. 158
Figure 5-5. Protein release kinetics of GF-hb-PNCOL scaffolds...... 159
Figure 5-6. Long-term evaluation of cell proliferation within GF-hb-PNCOL...... 161
Figure 5-7. Long-term evaluation of protein bioactivity within GF-hb-PNCOL...... 163
Figure 5-8. Protein release from PNCOL scaffold under uniaxial tensile loading...... 165
Figure 5-9. Bioactivity of PDGF released from hb-PNCOL scaffolds evaluated by scratch migration assay...... 168
Figure 5-10. Bioactivity of PDGF released from PNCOL scaffolds evaluated by cell proliferation assay...... 169
Figure 5-11. Major results obtained in Chapter 5...... 179
Figure 6-1. Schematic of mechanically-conditioned PDGF/ASC-encapsulated
COLPNCOL biphasic scaffolds...... 189
Figure 6-2. Schematic of the experimental design for PDGF-induced cell migration and infiltration within COLPNCOL biphasic scaffolds...... 194
Figure 6-3. Schematic of the experimental design to determine gradient matrix mineralization with COLPNCOL biphasic scaffolds...... 195
Figure 6-4. Effect of PCL nanofibers on material stiffness of PNCOL...... 199
Figure 6-5. Effect of PCL nanofibers on matrix mineralization of PNCOL...... 201
Figure 6-6. Morphology at the interface of mechanically-conditioned COLPNCOL biphasic scaffold...... 204
xx
Figure 6-7. Matrix alignment of mechanically-loaded PDGF/ASC-encapsulated
COLPNCOL biphasic scaffold...... 206
Figure 6-8. Amount of matrix directionality in mechanically-loaded PDGF/ASC- encapsulated COLPNCOL biphasic scaffold...... 208
Figure 6-9. ECM gene expression within mechanically-loaded PDGF/ASC-encapsulated
COLPNCOL biphasic scaffold...... 210
Figure 6-10. Musculoskeletal differentiation marker expression within mechanically- loaded PDGF/ASC-encapsulated COLPNCOL biphasic scaffold...... 211
Figure 6-11. Ratio of COL to PNCOL gene expression within COLPNCOL biphasic scaffolds...... 213
Figure 6-12. Secreted extracellular matrix proteins from COLPNCOL biphasic scaffold...... 215
Figure 6-13. PDGF-induced migration and infiltration of fibrocartilage-like cells into
COLPNCOL biphasic scaffolds...... 216
Figure 6-14. Alkaline Phosphatase activity within COLPNCOL biphasic scaffold. .. 219
Figure 6-15. Mapping of calcium deposits within COLPNCOL biphasic scaffolds. . 221
Figure 6-16. Spectral analysis for mineral deposits in COLPNCOL scaffolds...... 222
Figure 6-17. Gradient mineralization within COLPNCOL biphasic scaffold...... 224
Figure 6-18. Major results obtained from Chapter 6...... 234
xxi
List of Abbreviations
2D…………………...Two-dimensional 3D…………………...Three-dimensional
AC10………………..Cardiomyocytic cell line ACAN………………Aggrecan ALP…………………Alkaline Phosphatase ASC…………………Adipose-derived Stem Cells
BMP………………...Bone Morphogenetic Protein BMSC……………….Bone Marrow-derived Stem Cells BSA…………………Bovine Serum Albumin
C2C12……………….Myoblastic cell line COL…………………Collagen
DCN…………………Decorin DMEM………………Dulbecco’s Modified Eagle’s Medium
ECM…………………Extra Cellular Matrix EDS………………….Electron Dispersive X-ray Spectroscopy ELISA……………….Enzyme Linked Immuno Sorbent Assay
FBS………………….Fetal Bovine Serum FEA………………….Finite Element Analysis FEM…………………Finite Element Model FGF………………….Fibroblast Growth Factor
GAG…………………Glycosaminoglycan GAPDH……………...Glyceraldehyde 3-phosphate dehydrogenase xxii
GDF………………….Growth Differentiation Factor GF……………………Growth Factor (protein)
IGF…………………..Insulin Growth Factor
MC3T3-E1…………..Pre-osteoblastic cell line MEM………………...Minimum Essential Media MSC…………………Mesenchymal Stem Cells
OB6………………….Pre-osteoblastic cell line
PBS………………….Phosphate Saline Buffer PCL………………….Polycaprolactone PDGF………………..Platelet-Derived Growth Factor PEG………………….Poly-(ethylene glycol) PGA…………………Poly-(glycolic acid) PLA…………………Poly-(lactic acid) PLGA……………….Poly-(lactic-co-glycolic acid) PNCOL…………….. Polycaprolactone Nanofibers interspersed Collagen qPCR………………...Quantitative Polymerase Chain Reaction
SCX…………………Scleraxis SEM…………………Scanning Electron Microscope
TCN…………………Tenascin TNMD ………………Tenomodulin
VEGF……………….Vascular Endothelial Growth Factor
xxiii
Chapter 1
Background and Overview
1.1 Rotator cuff tendon injuries
More than 32 million people are affected by tendon and ligaments-related injuries, at an annual cost of approximately $30 billion [1, 2]. Injuries to tendons, either due to trauma or wear and tear, are highly common. It is seen that in the United States alone, nearly 30% of injury treatments in hospitals are due to soft tissue musculoskeletal injuries, and around
33,000 tendon repair procedures are carried out in the US annually [3]. The most commonly injured tendons include patellar tendon, Achilles tendon, and rotator cuff [4].
Rotator cuff injuries are highly prevalent in the aging population, with more than 30% of people above the age of 60 suffering from varying degrees of rotator cuff tear. The injury is characterized by pain, weakness, stiffness, and loss of mobility of the shoulder, and can result in significant disability in the individual’s day-to-day activities [5-7]. The pathophysiology of a rotator cuff injury is often atraumatic, caused due to a combination
1
of repetitive motion of the shoulder and age-related degeneration. It progresses from tendinosis, which is the inflammation of tendons, to partial tendon tear, and finally culminates in a full rupture of the tendon [8, 9]. Conservative treatments that include cortisone injection, physical therapy, and oral analgesics most often fall short because of the slow healing rate of the tendon tissue. Thus, surgical intervention has become the standard method of care [10].
1.2 Surgical repair of rotator cuff tendon
Around 600,000 surgical procedures are performed in the United States for rotator cuff repair in a year. The type of surgical treatment is dictated by the seriousness of the injury, ranging from partial repair to full tendon reconstruction [11, 12]. Direct suturing is employed when the injury is caused by a sharp rupture while arthroscopic surgery is performed for small tears. Partial or complete tear of collagen fibrils that cannot be treated by classic surgical techniques are rectified by tendon transplantation or grafting [13, 14].
Three types of grafts namely autografts, allografts, and xenografts are commonly used for tendon reconstruction. Autografts involve using the patient’s own tissue for transplant and hence do not have the risk of immune rejection or transmission of diseases. However, they are severely limited by availability of a tissue with comparable size, shape, and characteristics and require two surgical procedures. Disadvantage of allografts and
2
xenografts are that they are obtained from a donor, and hence have higher chances of tissue rejection and infection [13].
Despite advancements in surgical techniques and sutures, a reoccurrence of the injury is observed in 20-80% of the cases [11, 12]. Pathological examinations have revealed that inadequate healing of the rotator cuff and lack of matrix structural organization are the two major reasons for the re-ruptures; both of which can be associated with the distinct tissue properties and physiological environment of native tendons [15]. Hence, the next section gives an overview of the structure, function, and composition of tendons.
1.3 Tendon anatomy and physiology
Tendons are dense connective tissues that connect muscles to bones and thereby are found in body joints. The main role of tendons is to permit body locomotion by transmitting high tensile forces. They also allow connective flexibility to enhance the stability of joints and act as a temporary reservoir to store and recover energy. Healthy tendons are characterized by low cellularity and vasculature, slow metabolic rate, and high mechanical strength [16].
Tendons routinely experience cyclic uniaxial tensile strains in their in vivo environment
[17]. It has been established that presence of mechanical forces is vital for the homeostasis and regular function of tendon [18].
3
The tendon extracellular matrix (ECM) is predominantly is composed of collagens, glycosaminoglycans, proteoglycans, water, and cells. Out of these, collagen is the most abundant, accounting for 60% of the dry mass of tendon. Significantly, 95% of the collagen found in tendon is Collagen I, with the remaining 5% comprised of Collagen III and
Collagen V. The Collagen I fibers within tendons show a high degree of organization.
Cluster of collagen molecules form fibrils of 100-200 nm diameter, which assemble into fibers (50-100μm). The fibers aligned parallel to each other into the fascicles of a tendon unit are bundled in the hydrated proteoglycan rich matrix [19]. This hierarchical multi- stranded structure and anisotropic nature of the tendon ECM is responsible for their superior biomechanical properties and viscoelastic behavior. Crosslinking between fibrils results in higher mechanical strength than the individual units. The collagen fibers also have a characteristic crimp pattern that bestows a non-linear elastic property and low compressive stiffness to tendons (Awad 2012). Further, at the healthy tendon-bone insertion site, tendons demonstrate a specialized structural composition characterized by a gradient of mineralized matrix and a reverse gradient of aligned collagen fibers that enables tendons to achieve a strong attachment to the bone [20, 21].
Elongated, spindle-shaped fibroblasts called tenocytes are the primary cells of tendon.
They are found between the densely packed collagen fibers. The main role of tenocytes is to secrete and remodel the tendon ECM with the help of a myriad of growth factors, most prominently bone-morphogenetic proteins (BMP)-12 and BMP-14, platelet-derived growth factor (PDGF), and insulin growth factor-1 (IGF-1) [4]. In addition to chemical 4
cues, mechanical cues also play a significant role in stimulating tenocyte maturation and matrix remodeling that contribute to the tendon functional properties [17].
The other cells in the tendon tissue include round cells within the endotenon that separate each layer of the hierarchical structure, including the fibrils, fiber, and fascicles. Epitenon comprised of fibroblast-like cells cover the entire tendon unit. Both epitenon and endotenon layers contain collagen III protein. The paratenon layer between the tendon and its sheath is highly vascularized and provide nutrition to the tenocytes [17].
1.4 Tendon healing process
On subject to injuries, tendon healing process follows three stages that include the inflammatory, proliferation, and remodeling phases. The healing process is regulated by a large number of growth factors including PDGF, fibroblast growth factor (FGF), IGF, and transforming growth factor-beta (TGF-β) [22]. The inflammatory phase is initiated by the immediate response of the tendon tissue upon an injury. In the first 24 hours, inflammatory cells such as neutrophils and macrophages along with erythrocytes and platelets are recruited to the injury site by secretion of chemotactic factors such as platelet derived growth factor (PDGF). These cells migrate from epitenon, endotenon, and the tendon sheath and undergo phagocytosis to eliminate the foreign material, as well as release growth factors like TGF-β to induce tenocytes to produce and deposit collagen [23]. The
5
proliferation phase also called the reparative phase begins five days after the inflammatory phase and lasts for around 1 to 2 months. Here, more undifferentiated mesenchymal stem cells are recruited to the site of injury and differentiated into tenocytes in presence of growth factors such as BMP-12, BMP-14, and PDGF. They begin laying down collagen, proteoglycans, and other tendon ECM components around the wound site. Collagen III is deposited in random orientation during this phase. Vascular endothelial growth factor
(VEGF) induces formation of blood vessels to provide nutrition to the proliferating cells
[23]. When sufficient ECM components have deposited, the remodeling phase starts.
During remodeling phase, the previously laid Collagen III matrix get slowly replaced by highly directional Collagen I fibers. The number of tendon fibroblasts at the injury site reduces, leading to decrease in the synthesis of collagen. The tissue starts remodeling, and the collagen deposited assemble to fibrous aligned tissue. Due to the high density of collagen, vascular degeneration occurs, and the tissue metabolic rate reduces. The majority part of the remodeling phase lasts for about six months, but the tissue continues remodeling for many years. The timeline of the healing response depends on the severity of the injury and the nature of treatment [23].
The native healing process is limited in its capacity to regenerate the tendon tissue which is highly complex in structure, organization and function. This can be attributed to four main reasons: (1) the scarcity of resident tissue cells required to produce sufficient ECM,
(2) lack of sustained growth factor cues to accelerate the ECM remodeling process, (3) poor compaction and directionality of newly deposited collagen fibers, and (4) loss of 6
spatial mineral gradient at the tendon-bone insertion site [24-27]. A combination of these aspects results in inadequate Collagen I deposition and remodeling, a higher level of randomly oriented thin Collagen III fibers, and mismatch of mechanical properties at the tendon-to-bone interface, which further lead to scar tissue formation susceptible to re-tears
[10, 22].
1.5 Commercial solutions for rotator cuff repair
In an attempt to enhance the tendon healing process and to address the aforementioned issues, many different strategies involving biologics have been created. In last decade, two- dimensional (2D) acellular collagen-based patches have successfully cleared clinical trials for tendon healing and are commercially available to incorporate in surgical procedures
[28]. The most popular patches including Restore® (Depuy, Warsaw, IN), Zimmer
Collagen RepairTM (Warsaw, IN), GraftJacket® (Wright Medical Technology, Arlington
TN) show an increase in tissue ingrowth and thickening as they get absorbed into the repair site with time [10, 29, 30]. Thus, these patches mainly used in surgical procedures as reinforcement devices for mechanical stability [31, 32]. However, they are ineffective in addressing the primary concern of tendon healing with no significant differences seen in the tissue regeneration and functional rotator cuff restoration [33-35]. Rotation Medical’s
Rotator Cuff System (Plymouth, MN) have recently introduced a bio-inductive implant to
7
promote rapid healing of injured rotator cuff tendons, but its performance in a clinical setting is still unknown [36].
1.6 Cells or growth factor delivery for tendon repair
Apart from commercial solutions, a vast number of academic researchers have focused on developing improved strategies for tendon healing. Two such explored avenues are growth factor delivery and stem cells injection at the repair site to enhance ECM synthesis at the repair site. Studies show that in vivo administration of growth factors such as PDGF and
FGF can elicit positive response from cells in terms of increased proliferation and ECM synthesis of tenocytes. However, due to the limited number of cells available in the native tendon tissue at the repair site, none of them could lead to an increase in strength of the healing tissue [37-40]. On the other hand, cell-based therapies, involving mesenchymal stem cells (MSCs) injection into the repair site show an acceleration in the healing process at early stages with increased collagen alignment and stiffness. But due to lack of sustained cell delivery, no significant improvement is observed in the functional performance of tendons [41-43]. Also, injecting uncommitted stem cells capable of differentiating into multiple tissue lineages apart from tendons have the potential risk of undesirable effects such as ectopic bone formation and tumor-like tissue growth [44].
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1.7 Tissue engineering approach for tendon repair
Limitations in injured rotator cuff healing and regeneration through introducing collagen- based patches or cells or growth factors call the necessity for a combinatorial approach in which cell expression and function can be directed using biochemical and biomechanical cues delivered through a biomaterial carrier with appropriate stiffness and mechanical environment. Tissue-engineering based strategies have thus emerged to be promising candidates to augment and accelerate tendon repair and healing and comprise of four main components: biomaterial, cells, growth factors, and mechanical stimuli.
Currently used biomaterials for scaffolds in tendon tissue engineering can be classified into three groups including natural, synthetic, and hybrid materials. Natural materials such as collagen, chondroitin sulfate, and chitosan have been widely used in tendon tissue engineering for their excellent biofunctionality and biocompatibility. However, they exhibit poor mechanical strength, limited processability, and batch-to-batch variation since they are derived from natural sources. On the other hand, synthetic materials such as polycaprolactone (PCL) and poly-lactic-co-glycolic acid (PLGA) are biocompatible, biodegradable, possess good mechanical properties and processability, but are limited by their poor cell adhesion due to their hydrophobicity and acidic degradation products.
Surface hydrophobicity of synthetic polymer scaffolds is often overcome by surface modification strategies such as plasma surface treatment, RGD modification or coating with fibronectin in order to enhance cell adhesion. The third class, hybrid materials employ
9
a combination of natural and synthetic polymers and is recognized to be the best approach for creating a suitable scaffold for tendon tissue engineering [16, 22].
Cell lines used in tendon tissue engineering are majorly mesenchymal stem cells (MSC) that can be differentiated into tendon cells. They are widely used because they can be easily obtained from the bone marrow or adipose tissue. Tenocytes, the primary cells that comprise tendons are another choice of cells for incorporation into scaffold. However, tenocytes are very low in abundance within the native tissue. Another type of cell line employed is the dermal fibroblasts, again because of their easy accessibility and their potential to differentiate into tendon fibroblasts [16, 20, 44].
The inclusion of growth factors in scaffolds can achieve various favorable responses, such as cell proliferation, differentiation, migration, and recruitment of intrinsic cells. Some growth factors that have been incorporated in tissue engineering strategies are basic fibroblast growth factor (bFGF) and growth differentiation factor-5 (GDF-5) that enhances proliferation of tenocytes, PDGF that increases granulocytes recruitment and VEGF that promotes vascularization of the tissue. Additional growth factors that might be employed are TGF-β, insulin-like growth factor IGF-1 and cartilage derived morphogenetic protein
(CDMP) [16, 26].
Also, the use of mechanical stimulation to stretch cellular scaffolds in order to mimic the native tendon environment has been gaining popularity for tendon tissue engineering strategies. Since the tendon tissue is under uniaxial tensile forces within the body,
10
mechanical loading bioreactors that can apply cyclic uniaxial tensile strains to tissue- engineered scaffolds is of great interest to researchers. Many studies are focused on investigating the effect of strain, loading frequency, and duration on the regeneration and alignment of tendons scaffolds [18, 20]. Further, there are evidences that healing tendons show a positive response to mechanical loading [45].
1.8 State-of-the-art in tissue-engineered scaffolds for tendon
repair
The current efforts in developing tissue-engineered tendon scaffolds can be broadly classified into four categories based on its target application, which include tendon healing, partial tendon repair, whole tendon reconstruction, and recreating the tendon-bone interface tissue.
1.8.1 Hydrogel-based scaffolds to augment tendon healing
The first category of tissue-engineered tendon scaffolds designed to enhance healing of tendons at the repair site are usually composed of soft biomaterials or hydrogels that are predominantly biologic-based molecules such as collagen, fibril, hyaluronic acid, elastin, and gelatin. Due to their bioactivity and biocompatibility, they are excellent biomaterials 11
to incorporate cells or growth factor for delivering at the tendon repair site [23, 46-48].
Their weakness in stiffness and encapsulated protein retention have been overcome enhanced by using crosslinking agents and chemical treatment in order to increase their protein retention properties [46, 49, 50].
In a study conducted by Oka et al., gelatin hydrogels were chemically crosslinked with glutaraldehyde and then conjugated with simvastatin, a pro-angiogenesis drug to stimulate in vivo tendon-bone healing. Angiogenesis was induced in the short-term study, but no change in the tissue mechanical properties was observed in the long-term [51]. Tokunaga et al. used commercially available gelatin hydrogel sheets (MedGEL) and impregnated them with PDGF to use it as a delivery system in a rat tendon repair model. PDGF delivery lasted over a 2-week period where the cells showed increased proliferation, along with collagen orientation and higher material stiffness, thus providing evidence of enhanced tendon healing [52].
Some researchers have also explored the use of tendon-derived ECM to synthesize hydrogels to enhance the healing of tendon injuries. For instance, Farnebo et al. extracted the ECM solution from lyophilized and enzymatically-digested decellularized tendons to make tendon hydrogels. The hydrogels were found to possess appropriate porosity for cell invasion, good structural alignment, and supported proliferation of ASCs within its matrix and were envisioned as an injectable scaffold to stimulate tendon healing at the repair site
[53]. Crowe et al. observed that platelet-rich plasma could enhance the migration of ASCs
12
within the tendon hydrogels and had the ability to augment tendon healing [54].
Chattopadhyay et al. assessed the performance of dermal fibroblasts and ASCs within tendon hydrogels in terms of their in vivo proliferation and tenogenic differentiation. They concluded that delivery of dermal fibroblasts through the tendon hydrogels yielded better proliferation and enhanced tenogenesis when compared to ASCs [55].
Thus, the current hydrogel-based scaffolds mainly target to enhance the healing rate of the in vivo tendon defects and are not designed to provide significant mechanical support at the repair site.
1.8.2 Synthetic and composite scaffolds for partial tendon repair
The second category of tissue-engineered tendon scaffolds for partial repair of tendons are mostly composed of synthetic polymer-based biomaterials such as polylactic acid (PLA), polyglycolic acid (PGA), and polycaprolactone (PCL). They have higher mechanical strength compared to biologic scaffolds, so they are commonly used to make tissue- engineered tendon grafts implanted at the site of repair in order to provide mechanical stability [23, 56, 57]. However, their biodegradation products are often acid-based or esters that can be toxic to the native cell environment [58].
James et al. adopted a simple approach to develop a fibrous scaffold for tendon tissue engineering by using electrospun PLGA nanofibers. They confirmed the differentiation
13
potential of scaffold by seeding adipose stem cells on it and adding BMP-14 to the media to differentiate them into tendon fibroblasts [59]. Sahoo et al. looked to design a fiber- based scaffold constituting both microfibers and nanofibers with incorporation of cells and growth factor, to form a bio-functional environment with suitable mechanical properties and slow degradation rate. Knitted silk microfibers were combined with bFGF-coated electrospun PLGA nanofibers to form the composite scaffold. Bone marrow stromal cells
(BMSC) seeded in it showed enhanced attachment, proliferation and were also stimulated to differentiate into tenocytes, thus proving both inductive and conductive potential of the scaffold [60].
Though these scaffolds provide good mechanical stability, they are limited in their protein bioactivity due to the harsh environment during scaffold manufacturing. Also, the structure and biocompatibility of these scaffolds allows cells to be seeded only as a 2D layer on the scaffold surface [23]. Hence, synthetic biomaterials are often used in combination with natural biomaterials to encapsulate cells within a 3D structure or to prolong the bioactivity of growth factors.
Manning et al. developed a composite scaffold by alternating layers of hydrogel based delivery system and electrospun PLGA nanofibers, in order to create a scaffold for tendon repair with good mechanical and surgical handling properties. This composite scaffold was capable of delivering adipose-derived stem cells and PDGF in a controlled fashion. PDGF was retained substantially in the heparin-based delivery system for a two weeks; while
14
PLGA nanofiber backbone provided good mechanical integrity to the scaffold [27].
Vaquette et al. encapsulated alginate hydrogel in PLGA knitted scaffold to design a composite scaffold for tendons that had lower density and macroscopic interconnected pores, with good mechanical strength and slow degradation rate. As a part of this study, rabbits implanted with autologous BMSC seeded composite scaffold showed tendon regeneration with enhanced elastic modulus after 13 weeks [61].
Interestingly, collagen too has been a popular choice of biomaterial to engineer grafts that can restore tendon defects. Methods have been devised to dramatically improve the mechanical strength of collagen in order to design a successful scaffold. Kew et al. enhanced the mechanical properties of collagen-poly ethylene glycol composite by crosslinking it with a carbodiimide crosslinker and performed structural characterization
[62]. Shepherd et al. created a collagen-chondroitin sulfate composite and reinforced it using carbodiimide crosslinking, and obtained much higher values of tensile strength from the modified fibers [63].
Thus, synthetic and composite scaffolds are able to mechanically support the repair site along with some evidences of increased ECM synthesis and material stiffness of the healing tendons.
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1.8.3 Mechanically-conditioned scaffolds for tendon reconstruction
The third category of scaffolds are functional tissue-engineered tendon grafts developed and cultured under uniaxial tensile strains using mechanical loading bioreactor systems for full tendon reconstructions [5, 18, 64-66]. Mechanical conditioning of the scaffolds prior to implantation at the injury site is revealed to elicit enhanced cellular function and material stiffness due to simulation of the in vivo physiological environment of tendons in in vitro culture settings. The study conducted by Webb et al. employed polyurethane constructs encapsulated with fibroblasts that were subjected to uniaxial cyclic stretching at 10% strain with 0.25 Hz frequency for 8 hours/day over 7 days. They observed significant increases in the expression of tendon ECM markers as well as the elastic modulus of the mechanically-stimulated scaffolds [67]. Butler et al. synthesized MSC-encapsulated collagen sponge scaffolds and subjected them to 2.4% strain and 0.033 Hz frequency for
14 days. They also obtained increases in tendon-specific ECM marker, Collagen I, and the material stiffness of the scaffold [68]. The work by Chen et al. involved cyclic stretching of silk-collagen sponge scaffolds seeded with embryonic stem cells at 10% strain with 1
Hz frequency for 14 days. Elevated expressions of Collagen I and III genes were reported along with morphological changes with higher cell alignment and thickness of collagen fibrils, implying increase in material stiffness properties [2].
One other popular approach has been utilizing decellularized tendon grafts to create mechanically-conditioned tissue-engineered scaffolds. One such study was conducted by
16
Woon et al., who seeded dermal fibroblasts within acellular flexor tendon scaffolds and applied mechanical loading under various regimes using the LigaGen system (Tissue
Growth Technologies) [66]. Another study by Angelidis et al. involved mechanical conditioning of decellularized flexor tendons seeded with either ASCs or fibroblasts [64].
Both studies demonstrated that the ultimate tensile strength and elastic modulus of the mechanically-stimulated scaffolds were significantly higher compared to the non-loaded control samples.
Thus, mechanical loading of tissue-engineered scaffolds helps recreating the native tendon environment and induces increased ECM secretion and material stiffness of the scaffolds, thereby having the potential to provide good mechanical stability for full tissue reconstructions. However, these scaffolds are not incorporated with growth factors since the structure of their matrix, and the dynamic culture conditions makes it challenging to employ them as growth factor delivery systems.
1.8.4 Stratified scaffolds for tendon-bone interface repair
The fourth category of tissue-engineered tendon scaffolds are composed of stratified biomaterials designed to specifically target the tendon-bone insertion point and regenerate the gradient structural and mechanical properties of the interface tissue. These biomaterials have gradient mechanical properties to simulate the tendon-bone interface, in which there is a transition from a soft tissue to a hard tissue [20, 69]. One such scaffold was developed by Li et al., who applied a gradient coating of calcium phosphate on electrospun PLGA 17
and PCL fibers. The mineral gradient within the bone-tendon interface was able to influence gradient stiffness of the scaffold that in turn modulated the osteoblast activity depending on its spatial location within the scaffold [70]. Harley et al. developed an interface scaffold using collagen, GAG, and calcium phosphate with controllable pore size to mimic the osteochondral surface. Further, the scaffolds were crosslinked to enhance their mechanical properties, and the increased matrix stiffness influenced the morphology of the cells [71, 72]. These studies were followed by the development of biphasic and multi- phasic scaffolds, with the aim of creating a smooth transition from soft to hard tissue at the tendon-bone interface, and have shown promising in vitro results [73-75]. However, their performance in vivo is still unexplored.
In sum, despite the various different approaches for tendon repair via tissue engineering, a successful scaffold in terms of both biological and mechanical properties is yet to be achieved. The complexity in the structure, composition, and function of the tendon tissue make it very challenging for one single scaffold to address all essential aspects. Even though the scaffolds show promising indications in vitro, the biggest challenge remains the translation of the results in vitro to success in vivo. Hence, developing a scaffold that can meet the demands of the corresponding tendon injury has been the main focus of the current era in tendon tissue engineering.
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Chapter 2
Objective and Thesis Outline
2.1 Rationale
Although there are individual studies that highlight the importance of mechanical loading in the functional conditioning of tendon fibroblasts and MSCs within biomaterials [76-79], to the best of our knowledge, there is no study that employs mechanical loading-based approach which further incorporates a growth factor and cell-encapsulated tendon-to-bone scaffold to enhance healing of partial rotator cuff tears.
In this dissertation, we synthesized an innovative tendon-bone interfacial hydrogel scaffold cultured under physiologically relevant mechanical loading conditions for the augmentation of rotator cuff tears healing. This biphasic composite scaffold is predominantly composed of Collagen I hydrogel encapsulated with growth factor PDGF and adipose-derived stem cells (ASC). The scaffold possesses excellent biological properties to allow cell encapsulation, proliferation, differentiation, and migration along
19
with sufficient structural integrity to modulate controlled delivery of growth factors; and spatial material stiffness to promote in vivo gradient mineralization and matrix compaction.
Significantly, the cells and matrix of the scaffold are mechanically-conditioned by applying physiologically relevant uniaxial strains using our unique custom-built uniaxial tensile bioreactor to mimic native tendon environment before implantation into the repair site. We hypothesize that the scaffold, albeit having poor mechanical properties compared to native tendons, would be able to provide sustained cells and growth factor delivery to induce tenogenic differentiation and ECM secretion, and organize the scaffold matrix to promote gradient matrix mineralization and collagen alignment of the de novo tissue, thereby aiding and accelerating the natural tendon healing process for functional regeneration of the rotator cuff.
2.2 Objective
The objective of this dissertation was to create a mechanically-conditioned biphasic composite collagen scaffold to enhance rotator cuff regeneration with (1) controlled delivery of adipose-derived stem cells and platelet-derived growth factor to augment and accelerate tendon healing, and (2) spatial material stiffness to promote gradient mineralization and matrix directionality at the tendon-bone insertion site. The different components that constitute the interfacial scaffold were evaluated by a series of in vitro
20
studies conducted in mechanical loading conditions to characterize the cell and matrix response to biomechanical cues, growth factor release and bioactivity, and the collagen organization and mineralization potential of the gradient matrix. Our goal is to provide a viable solution to increase the healing and regeneration of tendon for rotator cuff injuries.
To meet this objective, the specific aims and the respective hypothesis of this dissertation were:
Specific Aim 1: Design and characterize a mechanical loading bioreactor to apply uniaxial tensile strains to cell-encapsulated 3D collagen scaffolds
Hypothesis 1: Eliminating grips to secure the 3D collagen scaffolds would enhance the region of homogenous strain profile and minimize scaffold disintegration during mechanical stimulation
Specific Aim 2: Investigate the effect of uniaxial strains and frequencies on the matrix organization and tenogenic differentiation of adipose stem cells (ASC) encapsulated within the 3D collagen scaffolds
Hypothesis 2: ASCs are capable of differentiating into tenocytes solely due to mechanical cues. Mechanical loading can induce directionality of the collagen matrix which would influence the orientation of cells within the scaffold.
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Specific Aim 3: Synthesize and evaluate composite collagen scaffolds (PNCOL) interspersed with polycaprolactone (PCL) nanofibers for controlled delivery of growth factors under uniaxial tensile loading
Hypothesis 3: The nanofibrous structure of PCL would provide higher surface area for growth factor immobilization and minimize burst release. Incorporation of biomolecules such as heparin or/and BSA would help long-term preservation of the protein activity.
Specific Aim 4: Engineer the PDGF/ASC-encapsulated COLPNCOL biphasic scaffold for tendon-bone interface and determine its potential for ASC tenogenic differentiation and gradient mineralization under uniaxial tensile loading
Hypothesis 4: Incorporation of PCL nanofibers into collagen matrix would improve the material stiffness of PNCOL, and that the increased matrix stiffness would correlate to higher cell-induced mineral deposition. The combination of half COL and half PNCOL would provide a spatial variation in the stiffness of the scaffold matrix that mimics the tendon-bone interface.
Figure 2-1 demonstrates the schematic representations of the tissue engineered scaffolds and components involved in each aforementioned specific aim in this study.
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Figure 2-1. Schematic of the various tissue engineering components involved in each Specific Aim of this thesis. (1) Design of a mechanical loading bioreactor to apply strains to 3D collagen scaffolds. (2) Employing the bioreactor to study ASCs response to uniaxial tensile loading regimes within 3D collagen scaffolds. (3) Incorporating PCL nanofibers into collagen 3D matrix for controlled release of bioactive PDGF. (4) Synthesis of COLPNCOL biphasic scaffolds encapsulated with ASCs and PDGF to augment healing of tendon-bone interface.
In brief, we first designed a mechanical loading bioreactor that can apply homogenous uniaxial strains to cell-encapsulated 3D collagen scaffolds. It was characterized for its
23
strain profiles along its length at various applied loads, loading frequencies, and cycle number.
This bioreactor was then used to mechanically stimulate ASCs encapsulated within collagen scaffolds at a different physiological strain and frequency regimes. The loaded samples were evaluated based on the cell viability and proliferation, differentiation, and matrix alignment in order to identify the appropriate strain and frequency to mechanically enhance matrix production and tenogenesis of ASCs.
We next improved the poor protein retention properties of collagen and made it suitable for growth factor incorporation with sustained release and activity that can further enhance
ASC proliferation and ECM secretion under mechanical loading conditions. This was achieved by creating a composite scaffold (PNCOL) made by interspersing PCL nanofibers into collagen solution to provide mechanical integrity and increased surface area for growth factor immobilization. Heparin and bovine serum albumin (BSA) biomolecules were incorporated into PNCOL for long-term preservation of the bioactivity of the growth factor, after which PDGF release and its activity on tendon fibroblasts-like cells was investigated under mechanical stimulation.
With the mechanical loading regime for tenogenic differentiation of ASCs determined, and
PNCOL characterized for PDGF release and activity, we then created the biphasic
COLPNCOL scaffold having gradient material stiffness, encapsulated with ASCs and
PDGF. The synergistic effect of PDGF, mechanical loading, and matrix stiffness was
24
investigated in terms of ASC tenogenic differentiation and gradient mineralization of the matrix to evaluate the in vitro performance of the biphasic composite scaffold for tendon- bone interface.
2.3 Thesis outline
Chapter 3 is a description of the introduction, methods, results, discussion, and conclusion related to Specific Aim 1. Section 3.1 introduces the current mechanical loading bioreactors developed for applying uniaxial strains to 3D tissue-engineered scaffolds.
Section 3.2 describes the design of the custom-built mechanical loading bioreactor that can apply homogenous uniaxial strains to cell-encapsulated 3D collagen scaffolds, and characterization methods employed to investigate its strain profiles and biological performance. Section 3.3 demonstrates the FEM-predicted bioreactor-induced strain and stress profiles experienced by the 3D collagen scaffolds, and the cell viability and matrix alignment observed within the scaffolds. Section 3.4 elaborates on the performance and advantages of the bioreactor in comparison to the current available systems. Section 3.5 covers the major conclusions obtained from Chapter 3 and how that would be employed in the next chapter.
Chapter 4 is a description of the introduction, methods, results, discussion, and conclusion related to Specific Aim 2. Section 4.1 gives an overview of the cells for tendon tissue
25
engineering strategies, and the role of chemical and mechanical stimulation in inducing cell differentiation to tendon lineage. Section 4.2 describes the mechanical loading regimes chosen to apply uniaxial tensile strains on ASC-encapsulated 3D collagen scaffolds, and the various biological characterization assays performed. Section 4.3 displays the results obtained in terms of matrix alignment, ASC viability, proliferation, and differentiation in response to different mechanical loading regimes. Section 4.4 discusses the effect of different strain and frequencies on cell behavior and scaffold structure and identifies the appropriate uniaxial tensile strain and loading frequency to induce pure tenogenic differentiation of ASCs. Section 4.5 outlines the major conclusions obtained from Chapter
4 and how that would be used in the subsequent chapter.
Chapter 5 is a description of the introduction, methods, results, discussion, and conclusion related to Specific Aim 3. Section 5.1 gives a background on the importance of growth factors during tendon healing, the various tissue-engineering based growth factor delivery systems explored, and the current challenges faced. Section 5.2 describes the synthesis of the composite collagen scaffold PNCOL and elaborates the characterization methods used to evaluate its protein retention capacity and bioactivity. Section 5.3 presents the protein retention profiles under static and uniaxial tensile loading conditions, short-term and long- term protein bioactivity within the scaffold, and bioactivity of the release media. Section
5.4 speculates on the possible mechanisms underlying the performance of the composite collagen scaffold exhibiting controlled release of bioactive proteins. Section 5.5
26
summarizes the major conclusions obtained from Chapter 5 and how that would be utilized in the subsequent chapter.
Chapter 6 is a description of the introduction, methods, results, discussion, and conclusion related to Specific Aim 4. Section 6.1 reviews the structure and composition of the tendon- bone interface and the various tissue engineering strategies developed to simulate the complexity of the interface tissue. Section 6.2 defines the methodology employed to synthesize and characterize the COLPNCOL biphasic scaffolds in terms of ASC gene expression, protein secretion, matrix directionality, cell infiltration, and matrix mineralization. Section 6.3 demonstrates the key differences between ASC response within
COL and PNCOL in terms of matrix structure, quantified mineral deposits, and gene expression. Section 6.4 evaluates the performance of the biphasic scaffold by taking the native tendon-bone interface tissue as the reference. Section 6.5 describes the major conclusions obtained from Chapter 6.
Chapter 7 elaborates the scope of the dissertation with recommended future work for each study presented in this dissertation.
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Chapter 3
A Mechanical Loading Bioreactor to Apply Uniaxial Tensile Strains to Cell-encapsulated 3D Collagen Scaffolds
3.1 Introduction
Tendons function in a very dynamic in vivo environment where they produce different body movements by transmitting high uniaxial tensile forces from muscle to bone along with providing stability to joints [16]. Their ability to withstand large mechanical forces on subject to loading comes from the high degree of organization of the tendon extracellular matrix predominantly composed of parallel bundles of Collagen I fibers [4]. Given its functional role, tendons inherently have the ability to undergo mechanical adaptation in response to uniaxial tensile stretching. The dynamic mechanical environment of tendons plays a huge part in the tissue maturation and function effectuated by inducing specific biological responses in residing tendon fibroblasts, which can further translate into changes in the composition and structural properties of the tissue [17]. 28
It is well-established that physical activity increases the tissue stiffness, tensile strength, and cross-sectional area of tendons [80, 81]. Conversely, there is also evidence that immobilization of tendon results in inferior tensile modulus [82, 83]. This is further confirmed by in vitro studies where in a static culture, the tendon fibroblasts lose their elongated morphology and functional responses [84]. It is has been recognized that since an appropriate mechanical stimulus is critical for tendon homeostasis, providing suitable mechanical cues is very important for the success of tendon tissue engineering. Thus, uniaxial tensile strain bioreactor systems that can simulate the tensile microenvironment of in vivo tendons hold great promise for generating mechanically-conditioned tissue- engineered cellular scaffolds that can induce cell proliferation, differentiation, and matrix production to enhance the healing and regeneration of tendons [18].
Several mechanical loading bioreactors have been designed and employed in the recent years to apply uniaxial tensile strains for tissue engineering applications. They can be broadly classified into two categories based on the mechanical integrity of the scaffolds desired to be loaded, which determines the actuation system and the gripping mechanism employed in the bioreactor.
The first group, comprising of bioreactors used for loading decellularized, ex vivo tendons, and synthetic tendon scaffolds. The commercial Bose® ElectroForce® BioDynamic®
29
system and the LigaGen system [64, 66] along with a few custom-built bioreactors [2, 5,
18, 85, 86] are representative bioreactors from this group. The loaded scaffolds from this group tend to be mechanically robust and easy to handle due to which they are usually mounted directly to the fixture of the bioreactor to subject them to cyclic tensile loading.
For instance, Angelidis et al. used the commercially-available Ligagen L30-4c (DynaGen systems, Tissue Growth Technologies) to apply cyclic uniaxial tensile strains to decellularized tendon constructs. The bioreactor system accommodates 4 tendons per run, and the tendons were clamped at the ends for load application. The authors observed slipping of the constructs from the clamps due to the vertical set up of the bioreactor which led to loss of samples, which was tried to overcome using glue and sandpaper at the edges of the constructs [64]. In the study by Wang et al., the bioreactor was designed to accommodate two ways of fixing the samples; clamps for synthetic scaffolds, and hooks for tendon tissues that are prone to slippage. Further, the bioreactor is set up on a horizontal platform, minimizing the effect of gravity that can cause slippage of the constructs from the fixture [65]. Juncosa-Melvin et al. used silicone dishes as the loading chamber having pins protruding from the base. Two 4-mm diameter holes were punched through either ends of cell-seeded collagen sponge constructs in order to secure them to the silicone dish. They were stimulated using a pneumatic computer-controlled mechanical loading system (Steri-
Cult Model 3033, Forma Scientific). Significantly, the pins restraining the collagen sponge constructs contributed to a highly uneven strain profile within the construct [87]. Thus, though direct gripping using tissue clamps, hooks, or pins, are widely used and are
30
relatively successful in mounting robust scaffolds and decellularized tendons to the mechanical loading systems. However, these tools lead to construct damage, slippage and produce non-uniform strain contours within the construct.
The second group consists of bioreactors used for loading 3D hydrogel-based cellular scaffolds. The commercially available bioreactors for 3D cellular scaffolds include the popular Flexcell® Tissue Train® Culture system, STREX Cell Stretching system, and
CellScale MechanoCulture system [88-90]. These bioreactors that load soft biomaterials, predominantly collagen, require innovative techniques to mount the scaffold without compromising its structural integrity [89]. This is commonly achieved by the use of tabs, nylon mesh or foam anchors onto which the scaffold solution is poured onto and allowed to polymerize [88, 90, 91]. Also, they are usually driven with a pneumatic non-contact actuation system that deforms the membrane onto which the cell-embedded collagen scaffolds are anchored at the ends [89, 92, 93]. Flexcell®, in particular, is currently used in numerous studies for mechanical loading of 3D collagen scaffolds due to its robust set up and has reproducible and well-characterized strain profiles [92, 94]. The Tissue Train® system has a trough for the gel, eliminating the need for sample transfer, it uses a nylon mesh to grip the ends of the loaded samples for effective strain transfer from the stretchable membrane to collagen scaffolds. This, however, compromises the region of the homogenous strain profile experienced by the scaffolds due to the stress shielding effect that happens due to the difference in the mechanical properties of the loaded gels and the
31
nylon mesh. Further, the Tissue Train® system utilizes a vacuum-based loading apparatus which requires very complicated sealing and necessitates a routine maintenance scheme to make sure the apparatus is very sealed, and the right amount of the applied pressure is transferred to the membrane that stretches the samples [88, 92]. Among the custom-made bioreactors, the bioreactor designed by Zimmermann et al. has an innovative way of applying loads without using clamps to grip the construct. The authors use pins inserted into the loop of ring-shaped collagen scaffolds in order to produce stretch during loading
[95]. However, the contact region of the pin with the collagen rings during the stretch is still likely to cause stress concentration and subject the cells in that region to much higher strain magnitudes when compared to the rest of the construct.
The above studies demonstrate that both clamps and mesh anchor along with their respective actuation system lead to non-homogenous strain profiles within the cellular scaffold due to stress concentration at the gripping regions. Thereby, in these mechanical loading bioreactors, the strains experienced by the cells inside the scaffold vary significantly based on their spatial location resulting in non-homogenous strain profile over the major part of the construct [18, 89]. This usually results in a narrow field of homogenous strain near the center of the scaffold, with wide variations in strain magnitudes near the ends of the scaffold [18, 79, 90]. On the other hand, non-tethered constructs have very low attachment to the stretching membrane which leads to inadequate strain application to the cell-encapsulated scaffold. The current mechanical loading bioreactors
32
are also too complicated in design, operation, and scaffold handling which may risk the structural integrity of the construct [88-90]. Thus, there is an unmet demand for uniaxial tensile strain bioreactors that can maximize the uniform strain region within the cell- encapsulated 3D scaffolds with the minimal risk of scaffold damage during mechanical loading.
The objective in this study was to design, fabricate, and characterize a mechanical loading bioreactor that can apply homogenous uniaxial tensile strains to 3D cellular collagen scaffolds for tendon tissue engineering applications. Collagen I, having excellent biological properties and being the predominant constituent of the tendon ECM is our preferred choice of biomaterial for devising the tissue-engineered scaffold for tendon regeneration.
Our research hypothesis was that eliminating grips to secure the 3D collagen scaffolds would enhance the region of homogenous strain profile and minimize scaffold disintegration during mechanical stimulation.
It is expected that mimicking the native tendon environment by applying physiologically- relevant mechanical loading conditions using the bioreactor would play a major role in accelerating and enhancing the cellular function and matrix organization of tissue- engineered collagen scaffolds when implanted at the repair site.
To this end, the uniaxial tensile strain bioreactor was designed with two key features: (1) a loading cum culture chamber that can effectively stretch the collagen scaffold without
33
direct gripping thereby enhancing the field of uniform strain within the scaffold, and (2) an easy-to-use controller-actuator system that can produce precise and reproducible strain values at physiological loading frequencies to mimic the in vivo dynamic environment in which tendon fibroblasts exist and function. Further, the bioreactor is simple to operate and maintain, and compact enough to fit into the shelf of a standard cell culture incubator. The design specifications of the loading chamber also minimizes handling procedures, scaffold disintegration, and risk of contamination.
A detailed study using Finite Element Analysis was conducted to predict the bioreactor- induced strain and stress profiles experienced by the 3D collagen scaffolds during static and cyclic loading. Next, a set of biological characterization studies were performed to provide basic evidence that cells are capable of survive, proliferate and influence matrix organization within the mechanically-stimulated collagen scaffolds.
3.2 Materials and Methods
3.2.1 Design and fabrication of the uniaxial tensile strain bioreactor
3.2.1.1 Components of the uniaxial tensile strain bioreactor
The schematic of the uniaxial tensile strain bioreactor (Patent Application Number
PCT/US17/20706) built for mechanically stimulation of 3D cell-encapsulated collagen scaffolds is displayed in Figure 3-1.
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Figure 3-1. Schematic of the uniaxial tensile strain bioreactor. (1) Silicone loading chambers containing the 3D collagen scaffolds (2) Fixed plate (3) Moving plate (4) Guiding sleeves for supporting plates (5) Connecting rods to transmit motion to the moving plate (6) Bearing supports to hold the ball-screw mechanism (7) Ball screw mechanism to produce precise linear motion (8) Coupling connecting the screw and the motor shaft (9) Motor support (10) 2-phase high torque stepper motor connected to programmable controller to produce controlled stretch [96].
The mechanical parts of the bioreactor were designed using SolidWorks 3D CAD design software (SolidWorks, MA), and manufactured and assembled at a high-precision local machine shop (OBARS Machine and Tools Company, OH) outside the University of
Toledo Main Campus. The loading chambers made out of silicone are supported by a polycarbonate base consisting of fixed and moving plates. These plates have aluminum 35
pins fitted on one of their ends that insert into the pinholes of the loading chamber in order to secure it to the bioreactor. The fixed plate helps supporting the weight of the loading chamber containing the tissue engineered scaffold and cell culture media. The moving plate is primarily responsible for transmitting motion provided by the driving mechanism to the loading chambers. A 2 mm gap exists between the fixed and the moving plate to prevent them from knocking against each other during cyclic motion. The plates are supported by polycarbonate guiding sleeves on either side. The sleeves allow translation motion of the moving plate, while the fixed plate is held in place by screwing it into the guiding sleeves.
The inner surfaces of both guiding sleeves are given a smooth finish to reduce friction and abrasion while the moving plate is in motion.
Two connecting rods are screwed into the free end of the moving plate to transfer the motion from the driving mechanism to the loading chambers. The driving mechanism of this bioreactor is a high precision ball screw assembly (Thomson Linear Motion Systems,
VA). The rotational motion of the ball screw is efficiently converted to translational motion of the ball nut. The connecting rods attached to the ball nut, in turn, produce motion of the moving plate and thus stretch the cell encapsulated 3D collagen scaffolds within the loading chambers. A two-phase high torque stepper motor coupled with a driver-controller
(Lin Engineering, CA) drives the motion of the ball screw assembly. The entire bioreactor assembly is secured on a polycarbonate base, with a polycarbonate lid in order to cover the samples cultured in the loading chambers during the operation of the bioreactor.
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3.2.1.2 Loading chamber of the uniaxial tensile strain bioreactor
The unique part of this uniaxial tensile strain bioreactor are the stretchable silicone loading chambers with a specific design and geometry. The key design consideration of the loading chamber was to maximize the strain transfer to the collagen scaffolds by minimizing its sliding with respect to the loading chamber during mechanical stimulation without the use of direct gripping methods such as clamps, hooks, or pins. The main features of the silicone loading chamber is indicated in Figure 3-2.
The cell-encapsulated 3D collagen scaffold is added within the groove of the silicone loading chamber that consists of two linear regions connected by semi-circular strips on either side. The groove holds the scaffold securely from both sides while the semi-circular sections provide mechanical support and stability to the scaffold during tensile loading.
The island in the middle separates the two linear strips and ensures that the scaffolds get stretched along with the loading chamber in the direction of load. The groove and island are enclosed within a well to hold media and other culture nutrients required for the survival, proliferation, and differentiation of cells encapsulated within the scaffolds. The well is flanked by flaps having pinholes on each side through which aluminum pins from the fixed and moving plates of the bioreactor is inserted in order to producing stretching of the silicone loading chambers. A mold corresponding to the geometry of the chamber was made to fabricate the silicone loading chambers. Dragon Skin® 10 High Performance
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Silicone rubber components A and B were mixed in a ratio of 1:1, poured into the mold and polymerized overnight to obtain the stretchable loading chambers.
Figure 3-2. Schematic of the silicone loading chamber: (A) Top view (B) Front view. (1) Groove into which cellular collagen scaffolds are polymerized comprises of linear strips (shaded) with dimensions of 25×4×2 mm (L×W×T) indicating the effective region for characterization and (2) semi-circular strips at either ends linking the two linear strips of the scaffold. (3) Island at the center provides support to the scaffolds during loading. (4) Well to hold sufficient media for cell survival, proliferation, and differentiation. (5) Pin holes through which the supporting base plates are inserted in order to apply loading [96].
This design of the loading chamber eliminates the necessity for grips to hold the sample and minimize the risk of scaffold disintegration. Further, since the loading chamber is also the culture chamber for the samples, there are no procedures required to transfer the samples from culture conditions to loading fixtures, thus minimizing the handling labor,
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possible scaffold damage, and risk of contamination. The linear strip having the dimension of 25×4×2 mm (L×W×H) is considered as the region of interest of the scaffold that would be used for future characterization studies. The thickness and width of the scaffold is kept small enough to reduce the strain gradient along those respective axes, and thus enhance the uniaxial nature of the strain experienced along the axis of load application.
3.2.1.3 Material selection
Silicone polymer was chosen as the material for the loading chamber due to its various favorable characteristics. It is robust, bio-inert and biocompatible; can be sterilized easily by autoclaving, and is cost-effective. It is easy to use and provides flexibility in chamber design since it can be poured into a mold of desired shape and size and allowed to solidify by polymerization. The curing occurs at room temperature with no negligible shrinkage.
Significantly, Dragon Skin silicone is highly elastic and possesses high tensile strength, and as stated earlier, it is capable of sustaining 1000% elongation-to-break without any noticeable plasticity or damage in structural integrity.
The components of the uniaxial tensile strain bioreactor are manufactured out of polycarbonate, stainless steel, or aluminum (McMaster-Carr, IL). The materials are chosen keeping in consideration the humid, warm, and sterile environment in which the bioreactor would be during operation. Polycarbonate is used to manufacture the main framework of the bioreactor, owing to its biocompatibility and resistance to corrosion at high humidity and physiological temperature. Aluminum is used for the pins and connecting rods due to 39
its anti-corrosion properties and ease of machining, while stainless steel being robust, durable and anti-corrosive is chosen for parts subjected to repetitive motion, such as the ball screw assembly and motor coupling.
3.2.2 Experimental strain validation with collagen scaffolds
The uniaxial tensile strain bioreactor was validated experimentally to determine the efficacy of the device in transmitting the load to the silicone chamber and the tissue- engineered scaffold within the chamber. This was studied by measuring the initial and final displacement undergone by the linear region of the groove of the silicone chamber and the linear 3D collagen scaffold within the groove. The point of load application is defined as the center of the aluminum pins attached to the moving plate that are responsible for transmitting motion from the moving plate to the loading chambers. Acellular 3D scaffolds were prepared using 2.5 mg/ml Collagen Type-I solution (Corning Life Sciences, US) neutralized with chilled 1 N NaOH solution to pH 7~8 along with sterile water and phosphate buffer saline (PBS), according to the manufacturer’s instructions. The scaffold solution was added into the grooves of each silicone loading chamber and polymerized at
37 oC for 1 hour, after which PBS was added into the well of each loading chamber.
Figure 3-3 is the schematic of the embedded markers placed for displacement measurements. The embedded markers were placed at the ends and the middle in the linear region of the silicone chamber and collagen scaffold, corresponding to 0 mm, 12.5 mm, and 25 mm in length from the end nearest to the point of load application. The scaffolds 40
were subjected to loads of 1 N, 2 N, and 3 N to investigate the deformation undergone under static loading.
Figure 3-3. Experimental validation of the uniaxial tensile strain bioreactor. (A) Before loading (B) After loading. Schematic of the placement of embedded markers and deformation measurement through image-based analysis for experimental validation of the uniaxial tensile strain bioreactor [96].
Digital images of the silicone loading chambers and collagen scaffolds were captured instantaneously at initial and final positions for each applied load in triplicates. The images were analyzed using ImageJ (NIH, US) image processing software by measuring the initial and final positions of the embedded markers for each sample at each applied load.
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For evaluating the deformation under dynamic (cyclic) loading conditions, collagen scaffolds with the embedded markers were loaded at 0.1 Hz, 0.5 Hz, and 1 Hz loading frequencies at the applied load of 2 N. High resolution video clips were recorded to capture the first five loading, and unloading cycles underwent by the scaffolds. Using image-based analysis, the frame at which maximum stretching occurred during cycle 1 was stored as an image, and the displacement of the embedded markers was determined by comparing their initial and final positions. Four samples were used for each set of loading condition for both static and cyclic loading, and the data was further used to validate the subsequently generated static and cyclic Finite Element Model.
3.2.3 Generation of Finite Element Model for the bioreactor
A Finite Element Model (FEM) was generated to conduct detailed investigation of the strain and stress contours experienced by the 3D collagen scaffolds in response to both static and cyclic mechanical loading using the uniaxial tensile strain bioreactor.
Geometry and Meshing: The model assembly was meshed with C3D10I; which is a 10- node general purpose tetrahedron with enhanced surface stress formulation elements. Mesh sensitivity studies were performed to obtain the most computationally competent mesh that does not affect the obtained results.
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Material properties: The loading chamber made of silicone was modeled as a hyperelastic material according to Ogden model. Ogden material parameters were generated through performing uniaxial tensile testing of the silicone chamber and subsequent curve fitting of the data. A summary of Ogden material parameters obtained via nonlinear curve fitting and used in the FEM is given in
Table 1.
Table 1. Ogden model parameters used for the silicone loading chambers [96].
i 훼푖 휇푖 (MPa) 1 0.68 0.075 2 0.001 0.03 3 0.35 0.114
The moving and fixed plates made out of polycarbonate were modeled as an elastic material. Young’s modulus and Poisson’s ratio of 2.4 GPa and 0.35, respectively were obtained from the supplier’s material data sheet.
Scaffolds made of collagen being viscoelastic and a time-dependent material were modeled according to Prony series model. Prony series material parameters were obtained by the nonlinear curve fitting of the stress relaxation curve that was obtained by Pyrse et al. [97].
The experimental stress relaxation was fitted to Prony series, and a summary of the material parameters used in the FEM for the 3D collagen scaffold is given in Table 2 [98].
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Table 2. Viscoelastic Prony series parameters used for collagen scaffolds [96].
i 푔푖 휏푖 1 0.0009 0.08 2 0.4045 0.136 3 0.1289 84.49 4 0.0001 290.5 5 0.0151 350.1 6 0.1206 1145
Loading and boundary conditions: Two representative views of the loading chamber assembly with the surfaces used to apply the boundary conditions and loading are shown in Figure 3-4.
Figure 3-4. Generation of FEM for the uniaxial tensile strain bioreactor.
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Schematic for the boundary conditions applied in order to generate the FEM for characterizing the strain and stress profiles experienced by the 3D collagen scaffolds when subjected to mechanical loading using the uniaxial tensile strain bioreactor [96]. The bottom surfaces of the fixed and moving plates (1) were constrained in both X and Z axes. Surface (3) was constrained in the X axis. The load was applied on surface (2) via kinematic coupling with a reference point that is constrained in all degrees of freedom except the X direction. The load was applied as displacement controlled boundary condition on the reference point. Surface-to-surface interaction was applied to the interface of silicone loading chamber and the collagen scaffold and also between the silicone chamber and polycarbonate plates. For the interaction between silicone and polycarbonate, normal penalty hard contact and tangential static-kinetic exponential decay frictional contact were used. The static coefficient of friction was set to 1.0, and the kinetic coefficient of friction was 0.5, and the exponential decay coefficient was 0.2. For the silicone-collagen interactions, normal hard contact and tangential frictional exponential decay with the static and kinetic coefficients were set to 2.0 and 1.0, respectively were employed. The exponential decay coefficient was 1.0.
3.2.4 Cell culture, scaffold synthesis, and mechanical loading regimes
The effect of bioreactor on cell encapsulated within the 3D collagen scaffolds was next investigate by performing a series of biological characterization studies. Human cardiomyocytes AC10, mouse myoblasts C2C12, and mouse osteoblasts OB6 were the
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three model cell lines used to characterize the performance of the uniaxial tensile strain bioreactor in terms of viability, proliferation, and matrix alignment. AC10s were maintained in 1:1 ratio of Dulbecco’s Modified Eagle Medium (DMEM) and F-12
(HyClone, US) with 12.5% Fetal Bovine Serum (FBS, Gibco, US), DMEM/low glucose
(HyClone, US) supplemented with 10% FBS was used to culture C2C12s, while OB6 culture media comprised of α-Minimal Essential Medium (α-MEM, Life Technologies,
US) also supplemented with 10% FBS. Penicillin-Streptomycin solution (Life
Technologies, US) at a final concentration of 1% was added into each culture media.
Cellular 3D scaffolds were prepared by encapsulating each cell line within neutralized
3 mg/ml Collagen type-I (Corning, US) solution at 1x106 cells/ml seeding density. The scaffold solutions were added into the groove of the silicone loading chambers and polymerized for 1 hour at 37 oC. The respective culture media of each cell line was added to the polymerized scaffolds, and the samples were returned to the cell culture incubator for 48 hours. The scaffolds were then subjected to cyclic stretching using the uniaxial tensile strain bioreactor at 1 N applied load (equivalent to 2% linear strain) and loading frequency of 0.1 Hz for 1 hour/day over a 3-day period. The bioreactor was placed in the incubator during its operation, thus ensuring that the cells within the scaffold continue to remain in their preferred environment. Scaffolds subjected to no loading (non-loaded) were used as control samples. The scaffolds were harvested at the end of the experiment, and the linear regions of interest were excised from the scaffolds to characterize their cell viability, proliferation, and matrix alignment.
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3.2.5 Cell viability and proliferation of mechanically-loaded scaffolds
The viability of the cells within scaffolds after three days of mechanical loading using the uniaxial tensile strain bioreactor was determined using Live-Dead Assay kit (Life
Technologies, US) according to the manufacturer’s protocol. Working solutions of Calcein and Ethidium Homodimer-1 dyes at 1:2 ratio were prepared in PBS. The dyes were added to the samples, incubated at 37 oC for 30 minutes, and fixed with 4% paraformaldehyde
(Sigma-Aldrich, US) at room temperature for 20 minutes. The scaffolds were visualized under the confocal microscope (Leica Microsystems) at 490/525 nm excitation and
557/576 nm emission wavelengths to identify live and dead cells, respectively. Three images each were taken at regions corresponding to 0, 12.5, and 25 mm of the collagen scaffold as depicted in Figure 3-3. The percentage viable cells in each scaffold was quantified by counting the number of live and dead cells obtained in each image.
The total number of cells within the loaded and non-loaded scaffolds after three days of uniaxial tensile loading was indirectly determined through DNA quantification performed using Picogreen assay (Life Technologies, US). The scaffolds were harvested and snap- frozen in liquid nitrogen. The frozen scaffolds were mechanically disrupted by homogenization with a pestle in order to liberate the cells from the scaffold. The cells were then resuspended in DNA lysis buffer (50 mM Tris HCl, 1 mM CaCl2, 400 µg/ml proteinase K, pH = 8) and incubated at 55oC overnight in order to degrade cellular protein 47
and collagen. The DNA lysate was centrifuged, and the supernatant was diluted 1:10 in TE buffer. A 1:200 dilution of Picogreen dye was prepared and was mixed with the diluted
DNA lysate in 1:1 ratio in microwell plates. The dye-lysate reaction was incubated at room temperature for 5 minutes. The samples were measured for their fluorescence intensities at
480/520 nm excitation/emission wavelengths using a microplate fluorometer (Wallac
1420). The amount of DNA from cells encapsulated within 3D collagen scaffolds was determined using a standard curve generated with different amounts of calf thymus DNA in ng and their respective fluorescence readings.
3.2.6 Matrix organization of mechanically-loaded scaffolds
The changes in morphology of collagen matrix due to mechanical stimulation of cell- encapsulated 3D collagen scaffolds using the uniaxial tensile strain bioreactor was visualized using the Scanning Electron Microscope (SEM). The scaffolds were fixed overnight with 4% paraformaldehyde (Sigma-Aldrich, US). The samples were then sequentially dehydrated by incubating them for 15 minutes each in a series of 30%, 50%,
70%, and 100% ethanol/water gradients. This was followed by dehydration with hexamethyldisilazane/ethanol gradients (Sigma-Aldrich, US) also in series of 30%, 50%,
70%, and 100% with incubation times of 20 minutes each. The scaffolds were allowed to dry overnight, mounted, sputter coated with gold, and examined under SEM (FEI Quanta
3D FEG) to observe the structure of the matrix.
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3.2.7 Statistical analysis
Four samples (n=4) were used for experimental validation and biological characterization studies performed with the uniaxial tensile strain bioreactor. Student t-test with a confidence interval of p < 0.05 was used to determine the statistical significance between two groups. The data is presented as ± standard deviation.
3.3 Results
3.3.1 Operation and performance of the bioreactor
3.3.1.1 Application of cyclic loads with the uniaxial tensile strain bioreactor
The uniaxial tensile strain bioreactor is comprised of uniquely-designed silicone loading chambers that contain cell-encapsulated 3D collagen scaffolds driven by a precise and versatile actuation system consisting of a miniature ball screw assembly driven by a high torque stepper motor.
Figure 3-5A shows the overall schematic of the uniaxial tensile strain bioreactor for cyclic loading while Figure 3-5B and Figure 3-5C are detailed schematics of the uniaxial tensile strain bioreactor at rest and during cyclic loading visualized from the top and front views.
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Figure 3-5. Cyclic loading of 3D scaffolds using the uniaxial tensile strain bioreactor. (A) Schematic of the major components involved in the operation of the uniaxial tensile strain bioreactor. A versatile driver-controller system programmed through LabVIEW- based software produces the defined rotation of the high torque stepper motor. This rotates the ball-screw actuation system that translates into the linear movement of the moving plate, resulting in the stretching of cellular 3D collagen scaffolds within the silicone loading chamber. (B) Top view and (C) Front view of the uniaxial tensile strain bioreactor with the silicone loading chambers when at rest and during cyclic loading. The numbers represent the same mechanical components described in Figure 3-1 [96]. 50
The design of the loading chamber eliminates the necessity for direct gripping of the 3D collagen scaffolds, generally achieved by using clamps, hooks, or pins, thereby greatly minimizing stress concentration at its either ends and the risk of scaffold disintegration.
This is accomplished by the groove-and-island ‘loop’ configuration of the silicone loading chamber demonstrated in Figure 3-2. The samples are secured from either sides by the groove and further supported by the island to ensure stretching along with the chamber during uniaxial tensile loading.
LabVIEW-based programming software is used to give input commands to the controller, which drives the stepper motor to rotate at the desired speed and number of revolutions, along with alternating the direction of motion to provide cyclic loading. The motor shaft, in turn, transmits its rotation to the ball screw, which causes the ball nut to move linearly in forward and backward directions. The linear motion from the ball nut is transmitted to the moving plate through the connecting rods attached to the ball nut assembly. Silicone loading chambers inserted into the pins of the fixed and moving plates get stretched along with the translational movement of the moving plate.
The silicone loading chambers are secured to the moving and fixed plates of the bioreactor within a laminar flow biosafety cabinet, thus ensuring aseptic culture techniques throughout the process. The friction between the loading chambers and the plates underneath is eliminated by adding a few drops of sterile water as a lubricant. A transparent lid is used to cover the loading chambers during mechanical stimulation. The strain
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bioreactor along with its electronic components can be placed in a standard tissue culture incubator during operation, allowing long term culture in a temperature and pH-controlled environment.
The bioreactor is capable of loading four silicone loading chambers simultaneously in one loading regime, which equates to eight tissue engineered scaffolds in a single run. The high torque stepper motor (1.8o per step) in combination with its controller is capable of high- precision, smooth operation at a broad range of strain and frequencies that match the physiological and pathological loading regimes in in vivo musculoskeletal tissues, including tendons.
3.3.1.2 Experimental validation of the uniaxial tensile strain bioreactor
Experimental validation of the uniaxial tensile strain bioreactor was conducted to evaluate the efficiency of the loading chamber design in transferring the applied loads to the 3D collagen scaffolds without substantial attenuation of strain. Loads of 1 N, 2 N, and 3 N were applied to the silicone loading chamber. The deformation undergone by the embedded markers at the specified points of 0, 12.5, and 25 mm in the linear region of the collagen scaffold (solid line) and loading chamber (dotted line) was estimated through image-based analysis and is presented in Figure 3-6.
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Figure 3-6. Performance of the uniaxial tensile strain bioreactor. Experimentally-determined deformation undergone by the linear region of the silicone loading chamber groove and collagen scaffold at the ends (0 mm and 25 mm) and center (12.5 mm) at applied loads of 1 N, 2 N, and 3 N. No significant difference seen between the deformation values obtained for silicone and collagen indicates that the applied load is being transferred effectively to the 3D collagen scaffold [96].
The results indicate that there is no significant difference in the deformation experienced by the loading chamber versus the collagen scaffolds along its linear length. This demonstrates that even without the use of direct gripping methods such as anchors, clamps, or hooks, the applied load is effectively transferred from the loading chamber to the 3D collagen scaffolds. It also confirms that the groove-and-island design of the loading chamber is able to act as an indirect support for the scaffold to remain in place during mechanical loading and help preserve the structural integrity of the scaffold.
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3.3.2 Characterization of bioreactor-induced strain profiles using FEM
3.3.2.1 Generation and validation of FEMs for the uniaxial tensile strain bioreactor
A Finite Element Model was generated to investigate the strain and stress profiles experienced by the 3D collagen scaffolds during mechanical stimulation at static and cyclic loading using the uniaxial tensile strain bioreactor. A representative deformation contour of the collagen scaffold predicted by the model when subjected to a static load of 3N depicted in Figure 3-7.
Figure 3-7. Representative deformation contour from FEM. Top and front views of the 3D collagen scaffold within the silicone loading chamber at an applied load of 3N. The contour proved to be in accordance with the experimental deformations values obtained at the 3N applied load presented in Figure 3-6 [96].
The static model was then validated using the experimental deformation data for the linear region of the stretching chamber and collagen scaffold at applied loads of 1 N, 2 N, and 3
N and presented in Figure 3-8.
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Figure 3-8. Finite Element Model validation for static loading. Comparison between the deformations experienced by collagen scaffolds experimentally and numerically at various applied loads of 1 N, 2 N, and 3 N. The data obtained from the FEM was found to be within ±10% of the experimentally measured data [96].
FEMs simulating the dynamic (cyclic) loading of the 3D collagen scaffolds were validated by comparing with the experimental deformation data generated at loading frequencies of
0.1 Hz, 0.5 Hz, and 1 Hz at an applied load of 2 N. This data is displayed in Figure 3-9. 55
Figure 3-9. Finite Element Model Validation for Dynamic (Cyclic) Loading. Comparison between the deformations experienced by the collagen scaffolds determined experimentally and numerically through FEM for the applied load of 2 N at various applied frequencies of (A) 0.1 Hz (B) 0.5 Hz (C) 1 Hz. The data obtained from the FEM was found to be within ±10% of the experimentally measured data [96]. 56
The results shown in Figure 3-8 and Figure 3-9 demonstrate that the FEM-predicted values are within ±10% accuracy when compared to the experimentally measured data.
3.3.2.2 Bioreactor-induced longitudinal tensile strain profiles within 3D scaffolds
The validated FEMs (Figure 3-8) were then used to elucidate the strain contours experienced within the 3D collagen scaffolds at applied loads of 1 N, 2 N, and 3 N. The longitudinal tensile strains that act parallel to the applied load were determined along the length (a) to (b), width (e) to (f), and thickness (g) to (h) of the linear part of the scaffold as displayed in Figure 3-10.
Figure 3-10. Schematic of the regions in the collagen scaffold used to investigate the bioreactor-induced longitudinal tensile strain profile using FEM. Length is defined as (a to b), width as (e to f) and thickness as (g to h) across the collagen scaffold. (c) to (d) represents the homogenous tensile strain region. (i) depicts the linear part and (ii) corresponds to the circular part of the full collagen scaffold [96].
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First, the longitudinal tensile strain profiles experienced across the full length of the collagen construct including the linear region (i) and semicircular regions (ii) when subjected to applied loads of 1 N, 2 N, and 3 N were predicted through FEM to generate
Figure 3-11. The predicted longitudinal tensile strain across the linear region of the scaffold (i) is observed to be very homogenous, over a 15 mm length from (c) to (d).
Figure 3-11. Bioreactor-induced longitudinal tensile strain profiles across the length of 3D collagen scaffolds predicted using FEM. Longitudinal tensile strain profiles over the total length of the scaffold at applied loads of 1 N, 2 N, and 3 N. Regions corresponding to the (i) linear and (ii) semi-circular parts of the collagen scaffold indicated in the schematic above the x axis. The applied loads of 1 N, 2 N, and 3 N correspond to 2%, 4%, and 6% linear strains. The strain profile near the gripped regions of the 3D scaffold (ii) shows decreased strain magnitudes when compared to the linear region of the scaffold, unlike the abnormally high strains usually generated with traditional gripping of scaffolds [96].
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At applied loads of 1 N, 2 N, and 3 N, the longitudinal tensile strain within the linear region of collagen scaffold (i) has a relatively uniform value of at 2±0.11%, 4±0.19% and
6±0.24% over the 15 mm length (c to d), respectively. The magnitude of strains from (c) to (a) and (d) to (b) that represent the ends of the linear region are seen to increase from the homogenous strain value by 1.2%, 1.5%, and 1.8%, respectively. This can be attributed to the linear to semi-circular switch in the scaffold geometry. Interestingly, the strain values decrease dramatically in the semi-circular regions (ii) of the scaffold and reach as low as
1% strain as they approach the ends of the whole scaffold. These results indicate that the loading chamber design has remarkably reduced the strain magnitudes usually experienced at the ends of the scaffold due to direct gripping. Also, the chamber design is able to significantly decrease the variation in the strain profile across the length of the scaffold, thereby increasing the region of homogenous strain.
Figure 3-12 demonstrates the longitudinal strain profile across the width and thickness of the 3D collagen scaffold at the applied loads of 1 N, 2 N, and 3 N. It is observed that there is negligible variation in the longitudinal tensile strain contours across the width and thickness of the scaffold. In terms of strain magnitudes, the range of strains across both width and thickness of the scaffold are within ±0.2-0.3% of the uniform strain values of
2%, 4%, and 6% corresponding to applied loads of 1 N, 2 N, and 3 N, respectively. The drop of 0.5% in strain observed near (f) for the longitudinal tensile strains estimated across the width is likely due to the island that resists the load.
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Figure 3-12. Bioreactor-induced longitudinal tensile strain profile across the width and thickness of 3D collagen scaffolds predicted using FEM. (A) Longitudinal tensile strain profiles across the width and (B) thickness of the scaffold at applied loads of 1 N, 2 N, and 3 N. The combined results for Figure 3-11 and Figure 3-12 demonstrate that the strain experienced by the collagen scaffold is homogenous over the region of 15×4×2 mm of the scaffold that is around 60% of the effective region of characterization [96]. 60
Based on the results from Figure 3-11 and Figure 3-12, it is established that the region of
15×4×2 mm ranging from lines (c) to (d) achieves uniform longitudinal tensile strain profile, with 1 N, 2 N, and 3 N loads corresponding to uniform strain values of 2%, 4%, and 6%. Further, this homogenous strain region has a volume of 120 µl which is around
60% of the linear part of the 3D collagen scaffold. Thus, for all biological characterization studies, this region can be defined as the region of interest.
3.3.2.3 Bioreactor-induced transverse compressive strains profile within scaffolds
It is known that uniaxial loading, in addition to longitudinal tensile strains, also generates a transverse compressive strain due to Poisson’s effect that acts perpendicular to the direction of the applied load. Figure 3-13 displays the ratio of transverse compressive strain to longitudinal tensile strain experienced by the linear length of the collagen scaffold at a load of 2 N.
The transverse compressive strain is found to be approximately 33±0.4% of the total strain experienced by the scaffold in the region of uniform longitudinal tensile strain (c to d). For instance, the predicted transverse compressive strain is 2.23% at the center of the scaffold with a uniform tensile strain of 4%. The result signify that the transverse strains are significantly and consistently lower than the longitudinal strains along the length of the region of interest of the scaffold.
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Figure 3-13. Bioreactor-induced tensile and compressive linear strains within 3D scaffolds predicted using FEM. Comparison of the longitudinal tensile strain and transverse compressive strain experienced by the collagen scaffold across its length on subject to a uniaxial tensile load of 2 N. Transverse compressive strains account for only 33% of the total strain experienced by the scaffold. The homogenous tensile strain across the 15×4×2 mm region is predominantly ‘uniaxial’ in nature [96].
3.3.2.4 Bioreactor-induced creep strain profiles with 3D scaffolds
Collagen being a viscoelastic material is known to undergo creep phenomenon, which is characterized by a time-dependent increase in deformation at a constant applied load. Thus, the creep undergone by the 3D collagen scaffolds at an applied load of 2 N was predicted using FEM. The 2 N load was applied monotonically and then held for 500,000 seconds (~
6 days) to record the equivalent creep strain as shown in Figure 3-14. 62
Figure 3-14. Percentage equivalent creep strain experienced by the collagen scaffolds predicted by FEM. Creep data recorded at 0 mm, 12.5 mm, and 25 mm region of the loaded sample after applying 2 N load monotonically then holding the load for 500,000 seconds. Static creep strain is around 1000-fold lower than the longitudinal tensile strains experienced by the collagen scaffold and can be considered insignificant [96].
The equivalent creep strain for the period of six days resembles both the primary and secondary creep behaviors of viscoelastic materials. However, the creep strain is found to be almost three orders of magnitude lower than the uniform longitudinal tensile strain value obtained at an applied load of 2 N. Thus the creep strain experienced by the collagen scaffolds in static loading conditions can be considered negligible in its contribution to the overall strain experienced by the scaffold.
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Overall, based on the various FEM-predicted strain profiles, it can be concluded that the volume of 15×4×2 mm3 of the 3D collagen scaffold experiences uniform tensile strains that predominantly act in the longitudinal direction and hence can be considered uniaxial.
3.3.2.5 Effect of frequency and cycle number on the longitudinal strain profile
Since in vivo tendons undergo repetitive cyclic motion, we next examined the effect of different loading frequencies and number of cycles on the strain profile within the 3D collagen scaffolds through the validated FEM demonstrated in Figure 3-9.
Figure 3-15. Bioreactor-induced strain profiles within 3D scaffolds at different loading frequencies predicted using FEM. Strain distribution profiles generated by the bioreactor at loading frequencies of 0.1 Hz, 0.2 Hz, 0.6 Hz, 0.8 Hz and 1 Hz across the length of scaffold for an applied load of 2 N. No significant variation seen in the strain values with change in loading frequencies [96].
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Figure 3-15 shows the strain profile across the scaffold length at loading frequencies of
0.1 Hz, 0.2 Hz, 0.4 Hz, 0.6 Hz and 1 Hz for an applied load of 2 N. It can be concluded that there is negligible variation in the strain magnitudes across the length of the scaffold due to change in the frequency of mechanical loading, indicating that the bioreactor is stable over a wide range of cyclic loading regimes.
Further, the effect of cycle numbers on the stress and strain profiles was studied by simulating cyclic loading at 0.5 Hz for a total of 40 cycles as shown in Figure 3-16.
Figure 3-16A and Figure 3-16B represent the FEM-predicted longitudinal tensile strain profile and von Mises stress profile experienced by the 3D collagen constructs during cyclic loading at 0.1 Hz. The profiles at cycle number 1 and cycle number 40 are compared for each applied load of 1 N, 2 N, and 3 N. There are no significant differences observed in the trend and magnitudes of the strain profiles between cycle 1 and cycle 40 (Figure
3-16A). Interestingly, both strain and stress profiles at cycle 40, in fact, look more uniform when compared to cycle 1. Also, lower stress concentration is observed at the ends of the linear region of the scaffold at the higher cycle number (Figure 3-16B). This demonstrates that the uniaxial tensile strain bioreactor is not only capable of applying uniform strain to the linear region of the collagen scaffold, but is also able to consistently reproduce the homogeneity in strain and stress contours with progression in cycle numbers.
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Figure 3-16. Bioreactor-induced strain and stress profiles within 3D scaffolds during cyclic loading predicted using FEM. Comparison of profiles at Cycle 1 vs Cycle 40 during cyclic loading of collagen scaffolds at 0.5 Hz loading frequency in terms of longitudinal tensile strain, and (C) von Mises stress, across its length at applied loads of 1 N, 2 N, and 3 N. The strain and stress profiles smoothen out at the ends with progression in cycle numbers and the magnitude of both stresses and strains in the region of homogenous strain remain fairly constant [96]. 66
3.3.3 Biological characterization of mechanically-loaded scaffolds
The effect of bioreactor-induced strains on the cell viability, proliferation, and matrix alignment of 3D collagen scaffolds seeded within the silicone loading chambers was experimentally evaluated using biological assays. Cell lines belonging to various musculoskeletal lineages that are known to experience uniaxial tensile strains on a daily basis were used to perform this set of basic characterization studies. Cell-encapsulated collagen constructs were loaded at 2% strain and 0.1 Hz frequency for 1 hour/day over a
3-day period.
3.3.3.1 Viability of cells within mechanically-loaded 3D collagen scaffolds
Figure 3-17 shows representative images of live-dead assay performed for loaded and non- loaded scaffolds seeded with OB6, C2C12, and AC10 cells using confocal microscopy.
The cell viability was imaged at 3 regions corresponding to 0, 12.5, and 25 mm distance of the linear part of the scaffold for each group. The representative live-dead images are taken from the 0 mm region that also corresponds to the region that experiences the maximum linear strain within the scaffold. The images indicate that there is no visible difference in cell viability between the non-loaded and loaded scaffolds in all three sample sets.
This observation is supported by the quantified cell numbers provided in Table 3, confirming that the uniaxial tensile strain bioreactor does not induce cell cytotoxicity within the 3D collagen scaffolds.
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Figure 3-17. Effect of the uniaxial tensile strain bioreactor on cell viability within 3D scaffolds through experimental determination. Loaded (2% strain, 0.1 Hz, 1 hour/day) and non-loaded 3D collagen scaffolds encapsulated with OB6, C2C12, or AC10 cells were visualized under confocal microscope at day 3. Green color represents live cells while red indicates dead cells. Scale bar is 100µm. The results combined with the quantified cell number show in Table 3 indicate no significant effect on the cytotoxicity of the loaded cellular scaffolds compared to the control [96].
Table 3. Quantification of viable cell number from live-dead images.
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3.3.3.2 Cell proliferation within mechanically-loaded collagen scaffolds
The cell numbers within the loaded and non-loaded collagen scaffolds was indirectly quantified by determining the amount of DNA. This data is presented in Figure 3-18.
Figure 3-18. Effect of bioreactor on cell proliferation within 3D scaffolds through experimental determination. DNA quantification of loaded (2% strain, 0.1 Hz, 1 hour/day) and non-loaded 3D collagen scaffolds encapsulated with OB6, C2C12, or AC10 cells at day 3. The black dotted line represents the initial cell density within the scaffolds. The results show no significant differences in cell proliferation of loaded scaffolds compared to the control [96].
A 2.5-fold increase in the amount of DNA is observed in OB6s, 2-fold rise in C2C12s, and around a 4-fold increase in AC10s for both non-loaded and loaded samples with respect to the initial DNA amount of 4500 ng (p < 0.05). This demonstrates that not only are the
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loaded cells viable, but they are also metabolically active and proliferating within the mechanically stimulated scaffold.
3.3.3.3 Matrix organization of mechanically-loaded 3D collagen scaffolds
The matrix morphology and structure of the loaded and non-loaded cell-encapsulated collagen scaffolds visualized using SEM is depicted in Figure 3-19.
Figure 3-19. Effect of bioreactor on collagen matrix organization within cell- encapsulated collagen scaffolds through experimental determination. Scanning Electron Micrographs of the loaded (2% strain, 0.1 Hz, 1 hour/day) and non- loaded cell-encapsulated 3D collagen scaffolds at day 3. Scale bar is 100 µm. Matrix organization is visible in all loaded samples, with the orientation of the fibers being parallel to the axis of load application [96].
The loaded cell-encapsulated scaffolds exhibit an increased level of matrix organization of collagen fibers when compared to the random organization in non-loaded samples. 70
Significantly, the loaded samples show clear matrix orientation, with the directionality of the matrix being predominantly parallel to the axis of applied load in all the three groups.
Further, it is observed that the extent of matrix compaction and the structure among the loaded groups vary depending on the cell-line encapsulated within the collagen scaffolds.
For instance, the scaffolds encapsulated with OB6 show prominent matrix compaction with thick and distinct collagen fibers, while the scaffolds embedded with AC10 though show alignment, appear the have the collagen matrix fused together. The results establish that mechanical loading using the uniaxial strain bioreactor, in presence of cells is able to induce a definitive organization of collagen in the scaffold matrix.
3.4 Discussion
Uniaxial tensile strain bioreactors systems that can allow culture of cellular 3D scaffolds under mechanical loading conditions are considered to hold great promise in developing tissue engineering strategies for tendon healing and regeneration. Their ability to mimic the in vivo mechanical environment of tendons makes them a valuable tool to create mechanically-responsive tissue engineered scaffolds in terms of cell proliferation, gene expression, and protein synthesis [18]. However, to the best of our knowledge, there are no existent commercial or custom-built devices that can apply homogenous uniaxial strain to three-dimensional cell-encapsulated hydrogel-based scaffolds without using gripping
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apparatus that usually compromise the gels integrity during and after loading. Further, the stress concentration at the ends due to the gripping effect usually leads to uneven strain distribution profiles within the mechanically-stimulated scaffolds.
In this chapter, we have created a simple and cost-effective mechanical loading bioreactor to apply precise homogenous uniaxial tensile strains to cell-encapsulated 3D collagen scaffolds. In addition, it is straightforward in design and construction, easy to operate and maintain, and compact when compared to existing bioreactors for uniaxial tensile loading of 3D collagen scaffolds. The bioreactor can be operated at physiologically-relevant uniaxial strains (0-12%) and loading frequencies (0.01-1 Hz) that mimic the in vivo environment of tendons. It also can be used to mimic various other tissues such as skin
[99], ligament, bone (Verbruggen et al. 2012), nerve [100] and skeletal muscle (Fukunaga et al. 2001) that experience uniaxial strains on a daily basis (Trumbull et al. 2016).
Additionally, the system can be assembled and reproduced by a novice user without detailed instructions which maximize its usefulness and significantly reduce its cost of machining, assembly, and maintenance. The uniaxial tensile strain bioreactor is capable of loading four silicone loading chambers simultaneously, with two cell-encapsulated 3D scaffolds within each chamber. The loading chambers are mounted onto the bioreactor within a biosafety cabinet to ensure aseptic culture techniques throughout the experiment.
The silicone chambers are handled similarly to any regular cell culture well plate. The
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media change is simply carried out in the biosafety cabinet by lifting off the bioreactor lid, removing the spent media from the loading chamber and replacing with fresh media using a regular micropipette. The entire bioreactor assembly is compact enough to be placed on a shelf of a standard tissue culture incubator during operation, allowing culture in a pH- and temperature-controlled environment. The bioreactor can support both short and long- term cultures based on the purpose and objective of the experiment conducted.
The performance of the bioreactor in stretching the collagen scaffolds along with the silicone loading chamber was experimentally determined by tracking the deformation undergone by strategically-placed embedded markers through image-based analysis
(Figure 3-6). No significant difference was noticed between the deformation undergone by the linear region of the loading chambers and collagen scaffolds at any measured position. This established that even without the use of direct gripping methods, the magnitude of applied load is effectively getting transferred from the chamber to the scaffold. Thus, the ‘loop’ configuration of the silicone loading chamber provides an indirect support for the scaffold to move along with the chamber during mechanical loading along with preserving its structural integrity. Additionally, since the loading chamber also acts as the culture chamber for the scaffolds, no procedures are required to transfer the scaffolds from culture condition to the loading apparatus. This significantly minimizes the need handling labor, possible scaffold disintegration, and risk of contamination.
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Since the performance of a bioreactor is largely directed by its design considerations such as chamber design, clamping technique, driving mechanism, and dimensions of the scaffold, detailed characterization of its strain profile is imperative before its use for biological applications. Surprisingly, amongst the currently studies dealing with bioreactors for 3D cellular scaffolds in literature, very few articles exist that focus on investigating the performance of a bioreactor. Instead, most articles directly focus on the biological application of their bioreactor system [90]. In this study, to elucidate the strain and stress profiles generated during static and cyclic mechanical stimulation using our uniaxial tensile strain bioreactor, FE-based models were generated. Both static and cyclic models were validated with a ±10% accuracy when compared to experimentally-obtained deformation data prior to employing them for generating bioreactor-induced strain contours (Figure 3-8 and Figure 3-9).
The validated static FEM was first used to predict the longitudinal tensile strain profile along the total length of the scaffold that include both its linear and semi-circular regions at applied loads of 1 N, 2 N, and 3 N (Figure 3-11). First focusing on the semicircular regions, it was evident that the ‘loop’ configuration of the scaffold was able to significantly minimize the gripping effect at its ends and reduce the abnormal high strain experienced within that region. In fact, with this design configuration, the semi-circular regions
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experienced very low strain magnitudes when compared to the linear region of the scaffold, with the values going as low as even 1% when approaching the ends of the total length of the scaffold. This was a clear contrast to the abnormally high strain magnitudes generated in the gripping region in traditionally gripped scaffolds and is a unique feature of this custom-built uniaxial tensile strain bioreactor.
Further, in this ‘loop’ configuration, the maximum strain due to indirect “gripping” was observed at the transition from the semi-circular to linear geometry of the 3D collagen scaffold, indicated by the dotted lines in Figure 3-11. This maximum strain experienced, also, was higher by only around 1.5% from the uniform strain value on an average at the applied loads of 1 N, 2 N, and 3 N, which was still significantly lower than the strain magnitudes generally observed due to direct gripping of samples. For instance, it is well recognized that custom-built bioreactors that employ hooks or punch holes to grip collagen-based scaffolds experience a highly uneven strain distribution with major risk of scaffold disintegration [2, 87].
Coming to the longitudinal tensile strain profiles predicted in the linear region of the collagen scaffolds, the region between 5 mm and 20 mm along the scaffold length exhibited a highly homogenous strain profile (Figure 3-11). The applied loads of 1 N, 2 N, and 3 N produced tensile strains of 2%, 4%, and 6% within that specific region of the scaffold. 75
Further, the strain distribution across both width and thickness of the scaffold were found to be fairly similar to the aforementioned uniform strain values at their respective applied load (Figure 3-12). Thus, the region with dimensions of 15×4×2 mm and volume of 120
µl experienced uniform uniaxial tensile strains. This region, estimated to be around 60% of the linear part of the 3D collagen construct can thus be defined as the region of interest when using the bioreactor for biological applications.
The calculated homogenous strain region for the Flexcell® Uniflex® 2D system was a rectangular region of 140 mm2 at the center of the membrane. This was estimated to be only around 47% of the effective area of characterization [92]. Though the strain profile for the Tissue Train 3D Culture system is not described in published literature [88], it also employs a membrane in order to stretch the 3D constructs and hence is expected to be similar to the 2D system in terms of performance.
Thus, in this study, through the design configuration of the loading chamber, we have eliminated any kind of direct gripping effect induced by the bioreactor. This not only renders the scaffold to have an enlarged region of homogenous strain distribution of 60% but also ensures that the semi-circular regions of the scaffold (excluded from characterization studies) do not experience abnormally high strain values and influence the cell response within the adjacent linear part of the scaffold. This characteristic feature of 76
our system renders it unique amongst the current mechanical loading bioreactors as it provides a great tool for not only tissue engineering applications but also mechanobiological studies that requires consistency within the same sample and reproducibility of experiments.
Uniaxial tensile loading is known to produce transverse compressive strain that act perpendicular to the longitudinal tensile strain. Also, collagen being a viscoelastic material has the tendency to exhibit creep behavior. Predicting the transverse compressive strain and creep strain for the collagen scaffolds revealed that uniaxial longitudinal tensile strain was the dominant strain experienced by the 3D collagen scaffolds. While transverse compressive strains were found to be around 33% of the total strain underwent by the scaffolds (Figure 3-13), creep strain was found to be lower than over 1000-fold when compared to the longitudinal tensile strain and thus considered insignificant (Figure 3-14).
These findings establish that the region of characterization of 15x4x2 mm of the 3D collagen scaffold experiences homogenous tensile strains predominantly acting in the longitudinal direction and thus can be considered ‘uniaxial’ in nature.
Since in vivo tendon undergo repetitive cyclic motion, we next examined the effect of different loading frequencies and number of cycles on the strain profile within collagen scaffolds through the generated FEM. There was negligible variation in the strain 77
magnitudes across the length of the scaffold due to change in the frequency of mechanical stimulation (Figure 3-15). Further, the uniform strain region showed no significant differences with progression of cycles (Figure 3-16). Thus the uniaxial tensile strain bioreactor is not only capable of applying homogenous uniaxial tensile strains to the linear region of the collagen scaffold but is also able to consistently reproduce the uniformity in strain and stress profiles at a wide range of loading frequencies over extended periods of cyclic loading regimes.
Basic biological characterization studies were performed to understand the effect of the bioreactor-induced strains on cell viability and proliferation within 3D cell-encapsulated scaffolds. Since bone, skeletal, and cardiac muscles are some prominent tissues that experience routine uniaxial cyclic tensile strains in their native environment, the corresponding cell lines, namely osteoblasts (OB6), myoblasts (C2C12), and cardiomyocytes (AC10), were chosen for this study. No cell cytotoxicity was observed in any of the loaded groups when compared to the non-loaded samples (Figure 3-17) which was further confirmed from image-based quantification of the cell number (Table 3). The
DNA quantification data also established that cells within the loaded scaffolds are viable and proliferating similar to that in non-loaded scaffolds (Figure 3-18).
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Matrix alignment exhibits significant differences between loaded and non-loaded samples in each group set. While the non-loaded scaffolds display disorganized matrix, the loaded scaffolds exhibit an organized matrix with definitive orientation of collagen fibers along the axis of applied load (Figure 3-19). The collagen fibers directionality is attributed to both mechanical loading and cell-mediated matrix compaction. It is visually observed that the extent of matrix alignment varies from one loaded group to another even when stretched using the exact same loading parameters. This indicates that the type of cell line encapsulated within the scaffold has a definite influence in the reorganization of collagen fibers to the loading direction. This is in accordance with several earlier studies that have reported collagen fibers alignment and compaction due to mechanical forces as well as cell-mediated collagen gel-compaction during loading [101-103].
Interestingly, though the matrix shows prominent alignment (Figure 3-19), the cell morphology is mostly rounded (Figure 3-17). This might be due to a possible time lag between strain-induced matrix alignment and its influence on cell elongation and morphology. The period of 3 days thus would not have been sufficient for the cells show a visible response in their morphology with the change in their microenvironment.
Nevertheless, the biological data confirms the ability of the uniaxial tensile strain bioreactor to maintain cell viability, support cell proliferation, and direct matrix organization within 3D collagen scaffolds.
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3.5 Conclusion
In Chapter 3, we devised a mechanical loading bioreactor that can apply physiological range of uniaxial tensile strains (0-12%) and loading frequencies (0.01-1 Hz) to stimulate cell-encapsulated 3D collagen scaffolds for tendon tissue engineering applications. Figure
3-20 displays the overall conclusions and results obtained from Chapter 3.
Figure 3-20. Major results obtained in Chapter 3. A simple and cost-effective mechanical loading bioreactor to apply homogenous uniaxial tensile strains to cell-encapsulated 3D collagen scaffolds. 80
The unique design of the silicone loading chamber eliminates the necessity of direct gripping and ensures that the collagen scaffold experiences uniform uniaxial tensile strains across 60% of its length while preserving its structural integrity. Significantly, this homogenous strain profile is reproducible and consistent at different stretching frequencies and stable over extended periods of cyclic loading. Further, the bioreactor promotes cell viability, cell proliferation, and matrix organization within the loaded 3D collagen scaffolds. The bioreactor has the potential to be expanded to stretch biomaterials other than collagen and can be modified to load larger and longer scaffolds due to the excellent tensile strength of the loading chamber, the flexibility offered by the chamber design configuration, and the versatility of the actuation system.
This bioreactor would be used in Chapter 4 for applying physiologically relevant uniaxial strains to ASCs-encapsulated collagen scaffolds. The effect of uniaxial loading on the proliferation, matrix alignment, and tenogenic differentiation of ASCs would be investigated to identify the loading regime appropriate for conditioning these cells for tendon healing at the repair site. The tissue-engineered scaffolds cultured in this bioreactor system is envisioned to be a mechanically-conditioned cell-encapsulated biomaterial (with or without bioactive factors) that can be introduced to the site of injury to enhance the healing rate of tendon tissues. This mechanically-conditioned scaffold is expected to provide bioactive, cellular, and micro-environmental cues to the repair site for partial tissue reconstructions.
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Chapter 4
Effect of Uniaxial Strains and Frequencies on the Matrix Organization and Tenogenic Differentiation of Adipose-derived Stem Cells Encapsulated within 3D Collagen Scaffolds
4.1 Introduction
The resident cells of tendons called tenocytes or tendon fibroblasts are elongated, spindle- shaped cells that align themselves parallel to the collagen fibers and are responsible for the synthesis and remodeling of the tendon extracellular matrix [4]. However, owing to the hypocellular nature of the tissue, with the tenocytes constituting less than 5% of the total volume, native tendons have a limited capacity for self-healing when injured [104-106].
The healing process causes formation of scar tissue that is different in morphology, composition, and mechanical properties when compared to healthy tendons, which leads to inadequate tissue regeneration, weak matrix structure, and compromised function [10,
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22]. Therefore, cell-based therapies for tendon healing are paramount to augment the cell number at the repair site and aid the native healing process.
Mesenchymal stem cells (MSCs) that include bone-marrow derived stem cells (BMSC) and adipose-derived stem cells (ASCs) have emerged to be the popular cell choice for tendon repair strategies due to their proliferative capacity and ability to differentiate into tenocytes [87, 93, 107, 108]. However, using BMSCs comes with potential complications of triggering inflammatory reactions or ectopic bone formation which would be of concern for tendon repair [44, 109]. Nowadays, the use of ASCs for tendon tissue engineering is widely explored because of their numerous advantages over their bone marrow counterparts. ASCs are 10-100 times more abundant per unit volume of the adipose tissue when compared to BMSCs within bone marrow aspirates. Isolating ASCs from subcutaneous fat tissue is easier, less painful and has a lower risk of donor morbidity compared to the highly invasive technique to aspirate BMSCs from bone marrow.
Significantly, ASCs, being derived from adipose tissue, have very low susceptibility to ossification, which makes it an ideal cell source for tendon tissue engineering applications.
Further, ASCs are known to exhibit anti-inflammatory properties, which is a major positive especially for treatment strategies that target to accelerate and augment tendon healing
[110-113].
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Studies focused on injecting autologous stem cells into the in vivo site of tendon repair revealed that delivering MSCs alone was insufficient to improve the healing of rotator cuff tears. Though the cells were initially capable of accelerating the healing process of the repair, no significant differences in the tissue regeneration was observed at long-term evaluation [41-43, 114]. This suggested that a controlled and sustained delivery of cells using a biomaterial carrier with appropriate chemical and mechanical cues was essential for successful healing of tendons. In the recent years, promising results have been demonstrated by in vitro culture of MSCs in tissue-engineered scaffolds, in terms of tenogenic expression and cell proliferation [115-118]. For instance, Barber et al. seeded
MSCs on braided electrospun poly(L-lactic acid) and observed cell alignment, proliferation, and increase in Scleraxis gene (tenogenic marker) expression [119]. Sahoo et al. fabricated a nanofibrous polymer scaffold was fabricated using poly(lactic-co- glycolic acid) (PLGA) electrospun nanofibers and knitted PLGA scaffold. BMSCs were seeded onto the scaffolds with fibrin gel as the carrier. The cell-seeded scaffolds exhibited increased cell proliferation, higher expression of Collagen I and glycosaminoglycans
(GAGs), but no statistically significant differences in the mechanical properties [120]. This indicates that though cells introduced with a biomaterial carrier show enhanced cellular function, they might be inadequate to restore tissue in a tendon defect. Thus, along with the biomaterial carrier, MSCs require biochemical or/and biomechanical cues in order to accelerate their tenogenic differentiation and augment the healing of tendon injuries.
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Growth factors belonging to the bone morphogenetic proteins family (BMPs), specifically
BMP-12 and BMP-14 are known to induce in vitro as well as in vivo tenogenesis of MSCs
[17, 26, 111]. In the study conducted by Park et al., ASCs stimulated with various BMP-
14 (GDF-5) concentrations exhibited time-dependent increases in ASC proliferation, ECM markers, and tenogenic mRNA and protein expression. As a result, they concluded that a concentration of 100 ng/ml was appropriate to induce in vitro tenogenic differentiation of
ASCs [121]. Subsequently, Shen et al. demonstrated the ability of BMP-12 at a concentration of 1 µg/ml to induce elevated tendon-related gene and protein expression within monolayer ASCs. Further, they found BMP-12 to be the more potent tenogenic growth factor when directly compared with the effect of BMP-14 in stimulating tenogenesis of ASCs [111].
Even in the presence of chemical factors, mechanical cues are considered to be very important for the maturation and function of the differentiating tenocytes. Studies suggest that tenocytes can exhibit gradual loss of tendon-specific markers in absence of tensile loading [84]. There are also evidences of uniaxial strains alone inducing increased proliferation, collagen secretion, expression of tenogenic markers, and decrease matrix degradation markers in tenocytes [122-127]. Significantly, varying the loading parameters such as the mechanical strain, frequency, and duration of mechanical stimuli has the capability to influence the type of cellular response [128-130]. Also, mechanical
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stimulation is strongly suggested to guide the fiber organization within the scaffold, which in turn can direct cell orientation and improve the matrix stiffness [66, 87, 131]. Thus, providing mechanical cues to the MSCs-seeded scaffolds is a critical component for the success of tendon healing strategies.
Previous studies have demonstrated that uniaxial tensile loading at various strains and cyclic frequencies stimulates tenogenic responses in MSCs and BMSCs in both 2D and 3D cultures [130, 132, 133]. Juncosa et al. cultured MSCs within collagen sponge scaffolds and loaded at 2.4% peak strain for 2 weeks once every 5 minutes for 8 hours/day. Increases in Collagen, I and Collagen III expression, were obtained along with 2.5-fold rise in linear stiffness and 4-fold rise increase in linear modulus compared to non-stimulated scaffolds
[87]. Scott et al. compared various loading modalities to determine their effect on the induction of tenogenic genes in MSCs encapsulated with 3D collagen constructs. They demonstrated that cyclic loading at 5% strain for 2 hours a day was more effective in inducing tenogenesis in comparison to static loading. Further, cyclic loading at the higher strain magnitude of 10% and cycle repetitions of 1000 resulted in increased Scleraxis expression of MSCs [134]. In studies involving BMSCs, the most prominent one was
Morita et al. who conducted an in-depth investigation with BMSCs cultured on 2D elastic surfaces using 5%, 10%, and 15% elongations at 1 Hz loading frequency for 1 or 2 days to determine the loading regime that directs BMSC tenogenesis. BMSCs stretched at 10% at
1 Hz were found to show the highest tenogenic response with increased expression of
Collagen I, Collagen III, Tenascin, and Scleraxis [130, 135]. 86
Interestingly, studies published in literature involving mechanical stimulation of ASCs are more focused on myogenic [78, 136] or osteogenic [137, 138] differentiation lineages.
There are only a few studies that have attempted to evaluate the effect of uniaxial tensile stimulation on the morphological or tenogenic response of ASCs. Rabbani et al. investigated the role of cyclic tensile loading on the organization of ASCs cultured on silicone strips. They concluded that cyclic loading produced a significant increase in both
ASC elongation and orientation [139]. In the study by Raghavan et al., ASCs seeded on silicone membranes exhibited higher proliferation, increased collagen production, and aligned morphology perpendicular to the axis of loading [140]. Yang et al. showed that
ASCs-encapsulated within collagen constructs in presence of tendon-derived ECM subjected to uniaxial tensile static loading undergo tenogenic differentiation, along with reduced matrix metalloproteases activity, and enhanced mechanical strength [141].
However, there is no comprehensive study yet that evaluates the effect of mechanical loading alone on tenogenic response of ASCs within the 3D environment.
Hence, in this section, we aimed to investigate the effect on the morphological and biochemical maturation of ASCs encapsulated within the 3D collagen scaffolds in response to different physiological uniaxial mechanical strains and loading frequencies.
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The objective of this study was to identify the uniaxial tensile loading strain and frequency appropriate for matrix alignment and in vitro differentiation of ASCs into pure tenogenic lineage within 3D collagen scaffolds for tendon tissue engineering applications.
Our research hypothesis was that ASCs are capable of differentiating into tenocytes solely due to mechanical cues and that mechanical loading can induce directionality of the collagen matrix which would influence the orientation of cells within the scaffold.
This is pursued with the vision that a tailored mechanical loading regime can direct ASCs lineage commitment towards tenocytes, which can both accelerate the functional response of these cells at the repair site as well as limit non-specific cell differentiation leading to potential complications such as ectopic bone formation. Further, mechanical loading would induce matrix directionality of the collagen scaffold which would influence the orientation of cells within the scaffold as well as enhance the alignment of newly synthesized collagen fibers at the in vivo repair site and improve the mechanical properties of the regenerating tissue.
The custom-made uniaxial tensile strain bioreactor introduced in Chapter 3 was utilized to apply cyclic loading at 2%, 4%, or 6% linear strains, and 0.1 Hz or 1 Hz loading frequencies to the ASC-encapsulated 3D collagen scaffolds for 2 hours/day over a seven-day period.
Loaded and non-loaded scaffolds were first evaluated in terms of matrix organization,
ASCs viability, morphology, and proliferation of ASCs within the 3D collagen scaffolds.
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Next, an in-depth gene expression analysis was conducted to estimate the mRNA levels of different musculoskeletal differentiation markers. In specific, the expression levels of ECM genes (Collagens and GAGs), tenogenic, osteogenic, chondrogenic, and myogenic genes were quantified. The data obtained from morphological and gene expression analyses was used to identify the most appropriate linear strain magnitude and cycling frequency that is capable of inducing tenogenic differentiation of ASCs cultured within 3D collagen scaffolds.
4.2 Materials and Methods
4.2.1 Cell culture, scaffold synthesis and mechanical loading regimes
The commercially available human adipose-derived stem cells kit (ThermoFisher
Scientific, US) was used to investigate the effect of different loading regimes on ASCs- encapsulated in 3D collagen scaffolds. As per manufacturer’s instructions, ASCs were cultured in MesenPRO RS™ basal media with MesenPRO RS™ growth supplement
(ThermoFisher Scientific, US) along with 200 mM Glutamine (Sigma-Aldrich, US). ASCs at passage 4 and 80% confluency were used for all experiments.
Cell-encapsulated 3D collagen scaffolds were prepared by adding 3mg/ml Collagen Type-
I solution (Corning, US), 10X PBS and chilled 1 N NaOH to neutralize the pH to 7-8 as per the manufacturer’s instruction. ASCs cell suspension was mixed into the neutralized
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collagen solution at a density of 750,000 cells/ml. The ASC-collagen solution was added into the grooves of the silicone loading chambers of the uniaxial tensile strain bioreactor and allowed to polymerize at 37 oC for 1 hour. Each loading chamber was filled with 1ml of aforementioned ASC media along with 1% Penicillin-Streptomycin solution (Gibco,
US) and cultured in the cell culture incubator for 48 hours.
The ASCs-encapsulated collagen scaffolds were then mechanically stimulated using the uniaxial tensile strain bioreactor at 2%, 4%, or 6% uniform linear strains, and 0.1 Hz or
1 Hz loading frequencies for 2 hours per day over a period of 7 days. Non-loaded scaffolds
(0% strain) were used as control samples. The media in the loading chambers were replenished every 2 days during the culture period. The samples were harvested by excising the 15 mm region of interest in the linear part of the scaffold and processed for various biological characterization studies including cell viability, cell proliferation, matrix alignment, and gene expression analysis.
4.2.2 Viability of ASCs within 3D collagen scaffolds
The effect of the different uniaxial tensile loading regimes on the viability of ASCs encapsulated within the 3D collagen scaffolds was examined by using the Live-Dead assay kit (Life Technologies, US) after 7 days of mechanical stimulation. The samples loaded at
0% (non-loaded), 2%, 4%, or 6% strains at 0.1 Hz or 1 Hz frequency were incubated in
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Calcein (green) and Ethidium Homodimer-1 (red) dyes mixed at 1:1 ratio at 37 oC for 30 minutes according to the manufacturer’s protocol. The samples were fixed with 4% paraformaldehyde (Sigma-Aldrich, US) for 1 hour at room temperature and visualized under confocal microscope (Leica Microsystems) at excitation/emission wavelengths of
490/525 nm and 557/576 nm. The live cells are labeled green while the dead cells are stained red. Three images were taken each at the ends and the middle of the linear scaffold for each group.
4.2.3 Proliferation of ASCs within 3D collagen scaffolds
The effect of the different uniaxial tensile loading culture conditions on ASC proliferation was evaluated by estimating the total cell number within the scaffold through PicoGreen dsDNA quantification assay (ThermoFisher, US). The samples loaded at 0%, 2%, 4%, or
6% strain at 0.1 Hz or 1 Hz loading frequency were snap-frozen in liquid nitrogen and subsequently homogenized using a pestle in order to liberate cells from the collagen scaffold. ASCs were then lysed using DNA lysis buffer (50 mM Tris HCl, 1 mM CaCl2,
400 µg/ml proteinase K, pH = 8) and incubated at 55oC overnight to allow digestion of membrane proteins and collagen. The lysed samples were centrifuged, and the supernatant was diluted 1:10 in a microplate. PicoGreen dye at a 1:200 working concentration was mixed with the diluted DNA lysate at 1:1 ratio and incubated for 5 minutes at room temperature. The fluorescence intensities of the samples were read at 480/520 nm 91
excitation/emission wavelengths using a fluorometer (Perkin Elmer Wallac 1420). The standard curve was generated by isolated DNA from different amounts of ASCs and plotting it against the corresponding fluorescence values obtained.
4.2.4 Matrix directionality of ASC-encapsulated 3D collagen scaffolds
The effect of uniaxial tensile loading in presence of ASCs on the matrix alignment of the
3D collagen scaffolds was visualized using SEM followed by image-based analysis of the samples. Collagen scaffolds loaded at 0%, 2%, 4%, or 6% strain and 0.1 Hz or 1 Hz frequency were fixed with 4% paraformaldehyde overnight and were rinsed twice with
PBS. Next, they were incubated with 1% Osmium Tetroxide for 30 minutes and washed twice with water. Then the samples were dehydrated by incubating for 15 minutes each in a series of ethanol-water gradient, followed by 10-minute incubations in a series of hexamethyldisilazane/ ethanol gradients (Sigma-Aldrich, US), both ranging from 30% to
100%. The samples were air-dried overnight and sputter coated with gold for 40 seconds before examining the morphology and structure of the matrix with SEM (FEI Quanta 3D
FEG). Four images each were taken at three different magnifications for each group. The lowest magnification images were also used to estimate the amount of matrix organization exhibited by the collagen scaffolds. This was quantified by obtaining directionality histograms using the Directionality plugin, Fiji/ImageJ (NIH, US) [142, 143].
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4.2.5 Gene expression analyses of ASCs encapsulated within scaffolds
The ability of different uniaxial tensile loading regimes in directing lineage-specific differentiation of ASCs encapsulated within 3D collagen scaffolds was investigated by conducting in-depth gene expression analyses. The expression levels of extracellular matrix (ECM), tenogenic, osteogenic, chondrogenic, and myogenic markers within the scaffolds loaded at 0%, 2%, 4%, or 6% strain and 0.1 Hz or 1 Hz frequency were determined using quantitative real-time polymerase chain reaction (qPCR). In addition to the non-loaded (0%) and mechanically stimulated groups, ASC-encapsulated collagen scaffolds incubated in media containing 1000ng/ml of bone morphogenetic protein-12
(BMP-12) was used as the positive control. Previous studies have demonstrated the ability of BMP-12 to induce tenogenic differentiation of ASCs [111, 144]. Hence, this group was included to provide a reference for direct comparison of the role of mechanical stimulation versus chemical stimulation in eliciting a tenogenic differentiation response of ASCs in their 3D environment.
The gene expression analyses was performed in three steps: RNA extraction from ASCs encapsulated within collagen scaffolds, cDNA synthesis from the extracted RNA, and qPCR with the synthesized cDNA to quantify the changes in gene expression.
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RNA extraction: All procedures were conducted in RNase-free environment, the area was wiped down with 0.1% Diethyl pyrocarbonate (DEPC), and RNase-free filter tips were used for all pipetting. The freshly harvested scaffold were crushed with a pestle in order to liberate the cells from the scaffold. The samples were incubated in TRIzol reagent
(ThermoFisher Scientific, US) for 15 minutes, mixed well and stored at -80 oC until further analysis. The samples were next thawed on ice, and the manufacturer’s protocol was followed for RNA extraction. Briefly, chloroform (Sigma-Aldrich, US) was added into each of the tube containing the cells in TRIzol and mixed vigorously for 30 seconds. The samples were allowed to sit for 5 minutes and then centrifuged at 4 oC to separate the organic and aqueous phases. The aqueous phase containing RNA was carefully transferred into a fresh tube. The organic phase and interphase containing DNA and protein, respectively, were discarded. Isopropanol (Sigma-Aldrich, US) was added into the aqueous phase and mixed gently for 30 seconds. The samples were incubated at room temperature for 10 minutes and then centrifuged at 4 oC to pellet down the RNA. The supernatant was discarded, and the RNA pellet was washed with 75% ethanol (Sigma-Aldrich, US), centrifuged and the supernatant was discarded. The pellet was air-dried and resuspended in RNase-free water. The isolated RNA was stored at -80 oC until further use. cDNA synthesis: The Omniscript cDNA synthesis kit (QIAGEN, US) was used to reverse transcribe the isolated RNA into cDNA according to the manufacturer’s instructions. The
RNA yield was estimated to be between 0.1 - 0.2 µg/µl. All RNA samples were heated at
65 oC for 10 minutes. The cDNA synthesis reaction was set up by adding the kit reagents 94
that include 10X reaction buffer, dNTPs, oligo-dT primer, RNase inhibitor, Reverse transcriptase along with 6 µl of the isolated RNA. The reaction mix was incubated at 37 oC for 1 hour, centrifuged and stored at -20 oC until further use.
Quantitative PCR: SYBR green master mix (Life Technologies, US) was used to perform all qPCR. The reaction was set up by adding sterile water, 5 µM forward and reverse primers of the gene of interest, 1.5 µl of cDNA and 2X SYBR green master mix. The samples were subjected to thermocycling using the iCycler iQ detection system (Biorad) to obtain the amplification and melt curves for each primer set. Thermocycling conditions were 10 minutes at 95 °C followed by 40 cycles at 95 °C for 15 seconds and 56 °C for 60 seconds.
Various primers associated with ECM genes and musculoskeletal lineage markers were used to investigate the gene expression profiles of non-stimulated, mechanically-stimulated and chemically-stimulated ASCs encapsulated within 3D collagen scaffolds.
The ECM markers evaluated are collagens, specifically Collagen I (COL-I), Collagen III
(COL-III), and glycosaminoglycans (GAGs) that include Decorin (DCN), and Aggrecan
(ACAN). Tenascin (TCN), Scleraxis (SCX) and Tenomodulin (TNMD) are the three tenogenic markers investigated. Other musculoskeletal lineage genes studied are osteogenic markers Runt-related transcription factor 2 (RUNX2) and Alkaline Phosphatase
(ALP); chondrogenic markers Collagen II (COL-II) and SOX9; and myogenic markers
Myogenic Differentiation antigen (MyoD) and Myogenin (MYOG). Glyceraldehyde-3-
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phosphate dehydrogenase (GAPDH) was used as the reference gene to normalize the Ct values of each gene of interest. The list of primers used for each gene is given in Table 4 along with their melt temperatures and corresponding references from where they were obtained.
All primers were purchased from Integrated DNA Technologies (IDT, US). The primer efficiency was found to be more than 95%, and thus ΔΔCt method was used to analyze all qPCR results. The fold difference in gene expression exhibited by the mechanically and chemically-stimulated groups was calculated in comparison to the non-loaded samples.
Table 4. Forward and Reverse Primers used for Real-time PCR.
Gene Forward primer Reverse primer Tm Ref GAPDH 5’ AGAAGGCTGGGGCTGATTTG 3’ 5’ AGGGCCCATCCACAGTCTTC 3’ 56oC [145] COL I 5’ GGCTCCTGCTCCTCTTAGCG 3’ 5’ CATGGTACCTGAGGCCGTTC 3’ 56oC [141] COL III 5’ CAGCGGTTCTCCAGGCAAGG 3’ 5’ CTCCAGTGATCCCAGCAATCC 3’ 56oC [141] DCN 5’ CGCCTCATCTGAGGGAGCTT 3’ 5’ TACTGGACCGGGTTGCTGAA 3’ 56oC [141] ACAN 5’ CACTGTTACCGCCACTTCCC 3’ 5’ ACCAGCGGAAGTCCCCTTCG 3’ 56oC [146] TCN 5’ GGTGGATGGATTGTGTTCCTGAGA 3’ 5’ CTGTGTCCTTGTCAAAGGTGGAGA 3’ 56oC [141] SCX 5’ ACACCCAGCCCAAACAGA 3’ 5’ GCGGTCCTTGCTCAACTTTC 3’ 54oC [141] TNMD 5’ CCATGCTGGATGAGAGAGGT 3’ 5’ CTCGTCCTCCTTGGTAGCAG 3’ 54oC [146] RUNX2 5’ CAACCACAGAACCACAAGTGC 3’ 5’ TGTTTGATGCCATAGTCCCTCC 3’ 54oC [141] ALP 5’ GATCTTCTTTCTCCTTTGCCTGG 3’ 5’ TGTTTGCAGTGGTGGTTCTGGCA 3’ 54oC [147] COL II 5’ GGCAATAGCAGGTTCACGTACA 3’ 5’ CGATAACAGTCTTGCCCCACTT 3’ 54oC [148] SOX9 5’ CACACAGCTCACTCGACCTTG 3’ 5’ TTCGGTTATTTTTAGGATCATCTCG 3’ 51oC [148] MyoD 5’ GCAGGTGTAACCGTAACC 3’ 5’ ACGTACAAATTCCCTGTAGC 3’ 51oC [149] MYOG 5’ GCCACAGATGCCACTACTTC 3’ 5’ CAACTTCAGCACAGGAGACC 3’ 54oC [149]
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4.2.6 Statistical analysis
Four samples (n = 4) were used for ASC viability, ASC proliferation, and matrix alignment studies. Eight samples (n = 8) were used for gene expression studies to obtained statistically discernable results. Statistical significance was determined using one-way ANOVA along with Fisher LSD post-hoc analysis (IBM SPSS Statistics software). The data is presented as the average of the samples, and the error bars represent ± standard deviation of the average. * represents significant fold increase in loaded samples with respect to 0% non- loaded group. * denotes p < 0.05, ** depicts p < 0.01, *** indicates to p < 0.001. † corresponds to the statistical difference between 2% and 4% groups loaded at the same magnitude of frequency while ‡ is the significant difference with respect to 6% group loaded at the same loading frequency, both with a 95% confidence interval (p < 0.05). # indicates a statistical difference between 0.1 Hz and 1 Hz group at the percentage strain with p < 0.05. § depicts significant difference with respect to non-loaded samples chemically treated with BMP-12 with a confidence interval of 95% (p < 0.05).
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4.3 Results
4.3.1 Matrix alignment of loaded ASCs-encapsulated scaffolds
The changes in matrix morphology and structure of ASC-encapsulated 3D collagen scaffolds subjected to uniaxial tensile loading at different strains and frequencies regimes was examined through SEM. The representative SEM images of scaffolds loaded at 0%
(non-loaded), 2%, 4%, or 6% strain at 0.1 Hz or 1 Hz frequency are displayed in Figure
4-1A and Figure 4-2A. The SEM images of 0% strain (non-loaded) scaffolds used as control samples in both Figure 4-1A and Figure 4-2A exhibit random distribution of collagen fibers with no particular orientation. When loaded at both 0.1 Hz and 1 Hz frequency, the scaffolds subjected to any of the three applied strains of 2%, 4%, or 6% exhibit definitive change in the organization of the matrix and morphology of the collagen fibers.
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Figure 4-1. Effect of uniaxial tensile loading at 0.1 Hz frequency on matrix organization of ASC-encapsulated 3D collagen scaffolds. (A) SEM images, and (B) Directionality histograms of ASC-seeded collagen scaffolds subjected to 7 days of uniaxial loading at 0%, 2%, 4%, or 6% strains at 0.1 Hz frequency for 2 hours/day. Scale bar in the image represents 100 µm. Sharper and broader peak in the histogram demonstrates higher amount of directionality of the fibers. The matrix orientation is parallel to the axis of applied load. 99
Figure 4-2. Effect of uniaxial tensile loading at 1 Hz frequency on matrix organization of ASC-encapsulated 3D collagen scaffolds. (A) SEM images, and (B) Directionality histograms of ASC-seeded collagen scaffolds subjected to 7 days of uniaxial loading at 0%, 2%, 4% and 6% strains at 1 Hz frequency for 2 hours/day. Scale bar in the image represents 100 µm. Sharper and higher peak in the histogram demonstrates higher amount of directionality of the fibers. The matrix orientation is parallel to the axis of applied load. 100
Comparing at the same magnitude of strain for each set, the scaffolds loaded at 0.1 Hz appear to have thinner compacted fibers, predominantly parallel to each other, while the loaded samples at 1 Hz display thicker bundles of compacted fibers that overlap one other.
To quantify the degree of matrix orientation of each sample, directionality histograms were plotted using Fiji-ImageJ Directionality plugin. The representative histograms for scaffolds loaded at 0% (non-loaded), 2%, 4%, and 6% strains at 0.1 Hz and 1 Hz frequencies are shown in Figure 4-1B and Figure 4-2B, respectively. The amount of directionality of the collagen fibers in the histograms is directly correlated to the combination of the height and the width of the peak obtained.
From Figure 4-1 and Figure 4-2, the images for non-loaded sample (0% strain) show no clear peak in the histogram implying random distribution of collagen fibers, with an amount of directionality as 0.015. Significantly, the loaded samples exhibit higher directionality, with magnitudes of 0.03, 0.035, and 0.04 at 0.1 Hz, and 0.03, 0.04, and 0.05 at 1 Hz, when loaded at strains of 2%, 4%, and 6%, respectively. Further, it is apparent that the peak in each loaded sample occurs at a 0o angle, which indicates that the fiber orientation in the scaffold matrix is along the direction of the applied uniaxial tensile load.
To determine whether the changes in amounts of matrix directionality between various loaded groups are statistically significant, histograms obtained from images from each loaded and non-loaded group were analyzed and consolidated into one figure for direct comparison (Figure 4-3).
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Figure 4-3. Effect of uniaxial tensile loading on matrix organization of ASC- encapsulated 3D collagen scaffolds. Quantified directionality of ASC-seeded collagen constructs subjected to 7 days of uniaxial loading at 0%, 2%, 4%, or 6% strains at 0.1 Hz or 1 Hz frequency for 2 hours/day from directionality histograms of SEM images obtained using ImageJ analysis. * represents statistical difference from the adjacent strain group. # denotes statistical significance between same strain groups at different frequencies. Directionality of collagen fibers increase with increase in magnitude of strain but largely remains unaffected by change in frequency (except for 6% group).
Figure 4-3 demonstrates increase in the degree of alignment of the scaffold matrix with increase in applied uniaxial tensile strain magnitudes. Further, it is found that the amount of directionality at each strain is statistically higher from the adjacent lower strain group at the same loading frequency (p < 0.05). On the other hand, no significant difference is observed in the alignment with between 0.1 Hz and 1 Hz loading frequencies at the applied strain of 2% and 4%. Interestingly, at the higher strain magnitude of 6%, which corresponds to the pathophysiological loading regime of in vivo tendon, a significant difference in
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matrix directionality is seen with change in frequency, with scaffolds loaded at 1 Hz having more alignment than 0.1 Hz (p < 0.05).
These results suggest that mechanical loading of ASC-encapsulated collagen scaffolds induces directionality of the collagen matrix parallel to the axis of load application. Further, the degree of directionality induced has a direct correlation with the magnitude of applied strain but is independent of the frequency of cyclic loading specifically in the physiological range of strain values.
4.3.2 Viability and proliferation of ASCs within loaded 3D scaffolds
Representative images of ASCs encapsulated within 3D collagen scaffolds stained with the live-dead assay dyes and visualized using a confocal microscope is presented in Figure 4-4.
The images indicate no significant cell death in the non-loaded as well as any of the loaded samples, demonstrating that the different mechanical loading regimes employed do not cause cytotoxicity of ASCs. Though the number of dead cells are negligible in the 6% at 1
Hz group after 7 days of loading, onset of membrane blebbing is observed in some cells, indicating that the high strain and high frequency loading condition could adversely affect
ASC viability over longer days of loading.
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Figure 4-4. Effect of uniaxial tensile loading on ASCs viability within 3D scaffolds. Confocal images of ASC-seeded collagen constructs subjected to 7 days of uniaxial loading at 0%, 2%, 4% and 6% strains and 0.1 Hz and 1 Hz frequencies for 2 hours/day. Green represents live cells, and dead cells are stained red. Scale bar represents 100 µm. Cells are viable within the loaded collagen scaffolds. 104
Focusing on the morphology of ASCs subjected to mechanical loading, while most groups have rounded cells that appear similar to the non-loaded (0% strain) group, a prominent change is exhibited by the sample loaded at 2% strain and 0.1 Hz frequency (Figure 4-4).
Remarkably, the cells within the scaffolds loaded at 2% strain and 0.1 Hz appear elongated, with emerging cytoplasmic extensions that is characteristic of tendon cells and look to be in the process of orienting themselves within the matrix.
Further, ASC proliferation within the scaffold was estimated by quantifying the amount of
DNA within each sample and correlating it with cell number as depicted in Figure 4-5.
Figure 4-5. Effect of uniaxial tensile loading on ASC proliferation within 3D scaffolds. Cell number within scaffolds determined using DNA quantification of ASC-encapsulated scaffolds loaded for 7 days at 0%, 2%, 4%, or 6% strains and 0.1 Hz or 1 Hz frequencies for 2 hours/day. Red dotted line indicates the initial cell number in each sample. * represents statistical difference from the other groups. Cells subjected to mechanical stimulation remain viable but show limited proliferation.
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The results demonstrate that there is no decrease in cell number over 7 days of scaffold culture due to mechanical loading with all groups being higher or similar compared to the initial seeding density of 200,000 cells represented by the red dotted line (Figure 4-5).
Interestingly, there is no correlation in cell proliferation with respect both increases in strains and frequencies of the loading regimes. The loaded samples have cell numbers similar to the initial encapsulation density, with the numbers ranging between 200,000 and
250,000 cells. The non-loaded (0% strain) group, on the other hand, shows a 1.5-fold increase in cell proliferation within the scaffold (p < 0.05) when compared to the loaded groups. This indicates that though the cells in the loaded samples remain viable, they undergo limited proliferation in these mechanically-stimulated culture conditions.
Nevertheless, the results confirm that physiological loading regimes does not adversely affect the cell viability, and can induce changes in the cells morphology and orientation at specific combinations of strain and frequency.
4.3.3 ECM gene expression of ASCs within loaded 3D scaffolds
The effect of uniaxial tensile loading at 2%, 4%, or 6% strains and 0.1 Hz or 1 Hz frequency on ASC expression of ECM genes was investigated by determining the fold-increases in the levels of Collagen (Collagen I and Collagen III), and GAGs (Decorin and Aggrecan) when compared to the non-loaded (0% strain) group. Also, non-loaded samples chemically
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stimulated with a known tenogenic differentiation factor BMP-12 were used as positive controls to evaluate the performance of mechanically-stimulated samples in comparison to the chemically-induced group.
Figure 4-6. Effect of uniaxial tensile loading on Collagen expression of ASCs within 3D collagen scaffolds. Gene expression profiles of ASCs encapsulated within collagen scaffolds subjected to BMP-12 treatment or uniaxial loading at 0%, 2%, 4%, and 6% strains at 0.1 Hz and 1 Hz. The graphs depict fold changes in ECM genes Collagen I and Collagen III. * indicates significant fold increase with respect to the 0% samples. * indicates p < 0.05, ** denotes p < 0.01, *** corresponds to p < 0.001. † represents significant difference between 2% and 4% groups while ‡ is the statistical difference with respect to 6% group, both with a 95% confidence interval. # represents a significant difference between 0.1 Hz and 1 Hz group at the same magnitude of strain with p < 0.05. § depicts significant difference with respect to non-loaded samples chemically stimulated with BMP-12 with p < 0.05. The mechanically-stimulated samples display increased level of Collagen I and III compared to the non-loaded scaffolds. 107
Figure 4-6 demonstrates significant increases in both Collagen I and Collagen III expression in mechanically loaded samples at the aforementioned applied strains and frequencies when compared to the 0% strain group. The chemically-stimulated positive control group (BMP-12) is observed to have a 5-fold increase in Collagen III expression
(p < 0.05), but does not exhibit any significant increase in Collagen I expression. On the other hand, the ASC samples subjected to uniaxial tensile strains of 2%, 4%, and 6% at loading frequencies of 0.1 Hz and 1 Hz display statistically significant increases in both
Collagen I and III gene expression, except for samples loaded at 6% strain and 1 Hz. ASCs stimulated at the lower frequency of 0.1 Hz, and low and moderate strains of 2% and 4% exhibit 3-fold and 2-fold rise in Collagen I (p < 0.01), and 4-fold and 12-fold increase in
Collagen III expression (p < 0.05), respectively. At the loading frequency of 1 Hz, Collagen
I and III levels are seen to be 5-fold and 20-fold higher at 2% strain (p < 0.001) and 2-fold and 10-fold more at 4% strain (p < 0.01) when compared to the 0% (non-loaded) samples.
Observing the results for 0.1 Hz and 1 Hz, the trend indicates that among 2% strained samples, the higher frequency of 1 Hz is able to stimulate more collagen production than
0.1 Hz. However, statistical analysis between 0.1 Hz and 1 Hz groups strained at 2% reveals a significant increase only in case of Collagen III expression (p < 0.05). Samples strained at 4% show little variation in the expression profiles of the Collagen I and Collagen
III between the loading frequencies of 0.1 Hz and 1 Hz. The group loaded at 6% strain exhibits an increase in only Collagen III expression, which usually is stimulated in excess during over-loading of tendons.
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Significantly, between the mechanically-loaded and the BMP-12 treated groups, it is evident that uniaxial tensile loading, in general, is able to induce elevated expression levels of Collagen I in ASCs, in addition to Collagen III that is expressed also in the chemically- treated ASC-encapsulated scaffolds.
Thus, looking at the overall collagen data (Figure 4-6), it is evident that with respect to the non-loaded (0% strain) group, samples stimulated with 2% and 4% at both 0.1 Hz and 1
Hz, and the group loaded at 6% at 0.1 Hz show significant increases in both Collagen I and
Collagen III gene expression levels.
Figure 4-7 depicts the fold differences in GAGs (Decorin and Aggrecan) expression exhibited by ASCs-encapsulated collagen scaffolds loaded at the different strains and frequencies along with samples treated with BMP-12 after 7 days. It is observed that GAGs are predominantly expressed in samples stimulated at 2% strain at both 0.1 Hz and 1 Hz frequency. ASC-encapsulated scaffolds treated with BMP-12 do not show a change in the expression of Decorin but exhibit a 5-fold increase in Aggrecan expression (p < 0.05).
Samples subjected to 2% at 0.1 Hz result in a 2-fold increase in Decorin and Aggrecan in comparison to the control (p < 0.05), while 2% at 1 Hz group exhibits 8-10 times increase in Decorin (p < 0.001) and Aggrecan expression (p < 0.01) in ASCs, when compared to the non-loaded scaffolds. No significant rise in GAG expression is seen in the rest of the mechanically loaded regimes except for a 3-fold increase in Aggrecan at the loading condition of 4% and 0.1 Hz (p < 0.01).
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Figure 4-7. Effect of uniaxial tensile loading on GAG expression of ASCs within 3D collagen scaffolds. Gene expression profiles of ASCs encapsulated within collagen scaffolds subjected to BMP-12 treatment or uniaxial loading at 0%, 2%, 4%, and 6% strains at 0.1 Hz and 1 Hz. The graphs depict fold changes in GAGs: Decorin and Aggrecan. * indicates significant fold increase with respect to the 0% samples. * indicates p < 0.05, ** denotes p < 0.01, *** corresponds to p < 0.001. † represents significant difference between 2% and 4% groups while ‡ is the statistical difference with respect to 6% group, both with a 95% confidence interval. # represents a significant difference between 0.1 Hz and 1 Hz group at the same magnitude of strain with p < 0.05. § depicts significant difference with respect to non- loaded samples chemically stimulated with BMP-12 with p < 0.05. The mechanically- stimulated samples display increased levels of GAG at 2% strain at 0.1 Hz and 1 Hz, and 4% strain at 1 Hz when compared to the non-loaded scaffolds.
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Comparing the results for 0.1 Hz and 1 Hz for GAGs from Figure 4-7, the trend is again similar to that observed with Collagen, where among the 2% strained samples, the higher frequency of 1 Hz stimulates more ECM production than 0.1 Hz. However, they were not found to be statistically significant (p > 0.05). Samples strained at both 4%, and 6% show little variation in the expression profiles of both Decorin and Aggrecan between 0.1 Hz and
1 Hz loading frequencies.
In comparison to the BMP-12 stimulated group, all of the mechanically-loaded samples exhibit significantly elevated levels of Decorin which is absent in the chemically-induced group. On the other hand, Aggrecan that has higher expression in BMP-12 treated samples is observed only in 2% strain groups loaded at both 0.1 Hz and 1 Hz amongst the groups subjected to mechanical loading.
Thus, looking at the overall GAGs expression data (Figure 4-7) it can be inferred that with respect to the non-loaded (0% strain) group, the samples loaded at 2% and 4% and both
0.1 Hz and 1 Hz are the groups that show significant increases in most of the ECM markers.
To sum up, from the ECM results displayed in Figure 4-6 and Figure 4-7 , it can be concluded that (1) mechanical loading at specific loading regimes effect an increase in the expression of ECM genes of ASCs encapsulated in 3D collagen scaffolds, (2) there are clear differences in ASC response in terms of ECM stimulation with both varying strains and frequencies with respect to both non-loaded and BMP-12 treated samples, and (3) significant increases in both tendon-specific collagens and GAGs are seen for groups
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strained with 2% at both 0.1 Hz and 1 Hz (4) significant increases in collagen genes are seen for groups strained with 4% strain at both 0.1 Hz and 1 Hz frequency.
4.3.4 Tenogenic expression of ASCs within loaded 3D scaffolds
To investigate the potential of ASCs to commit to a tenogenic lineage in response to mechanical loading, the expression level of tenogenic markers Tenascin-C, Scleraxis, and
Tenomodulin was quantified for samples stimulated at 0% (non-loaded), 2%, 4%, and 6% strains at 0.1 Hz and 1 Hz loading frequencies. Samples treated with BMP-12 was again used as the positive control. The results obtained are presented in Figure 4-8.
Among the mechanically-stimulated groups, it is observed that 2% strain groups at both
0.1 Hz and 1 Hz display increases in tendon-related gene expression in ASCs. The 2% at
0.1 Hz group shows 4-fold increases of tenascin-C (p < 0.01) and Scleraxis (p < 0.05), and
8-fold rise in Tenomodulin (p < 0.05) while at 1 Hz Tenascin and Tenomodulin increase by 6-fold (p < 0.05) with Scleraxis rising as high as 15-fold (p < 0.01) in comparison to the
0% (non-loaded) samples. ASCs-encapsulated scaffolds stimulated with BMP-12 exhibit increased tenogenic response as conforming to the results observed in previous studies
[111], with 10-fold rise in tenascin and 2-fold increases in Scleraxis and Tenomodulin.
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Figure 4-8. Effect of uniaxial tensile loading on tenogenic gene expression of ASCs within 3D collagen scaffolds. Gene expression profiles of ASCs encapsulated within collagen scaffolds subjected to BMP-12 treatment or uniaxial loading at 0% (non-loaded), 2%, 4%, and 6% strains at 0.1 Hz and 1 Hz loading frequencies. The graph depicts fold changes in various tenogenic markers: Tenascin-C, Scleraxis, and Tenomodulin. * indicates significant fold increase with respect to the 0% samples. * indicates p < 0.05, ** denotes p < 0.01, *** corresponds to p < 0.001. † represents significant difference between 2% and 4% groups while ‡ is the statistical difference with respect to 6% group, both with a 95% confidence interval. # represents a significant difference between 0.1 Hz and 1 Hz group at the same magnitude of strain with p < 0.05. § depicts significant difference with respect to non-loaded samples chemically stimulated with BMP-12 with p < 0.05. Samples stimulated with 2% strain at both 0.1 Hz, and 1 Hz show increased level of all three tenogenic markers, along with 4% at 1 Hz loading condition.
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Similar to the trend seen with the ECM gene expression, though 2% loaded samples at 1
Hz appear to have higher tenogenic gene expression compared to 0.1 Hz loaded samples, only Scleraxis is statistically different between 1 Hz and 0.1 Hz at 2% loading regime (p <
0.05) (Figure 4-8). Also showing higher ASC tenogenesis are samples subjected to 4% strain at 1 Hz, with 15-fold, 9-fold, and 6-fold increases in Tenascin, Scleraxis, and
Tenomodulin, respectively. On the other hand, samples loaded at 6% strain at both 0.1 Hz and 1 Hz frequencies, and 4% at 0.1 Hz do not show any marked increases in tenogenic genes.
These results imply that the mechanical loading regimes of 2% strain at both 0.1 Hz and 1
Hz, and 4% strain at 1 Hz that show significant increases tenogenic markers (Figure 4-8) along with enhanced ECM gene expression demonstrated in Figure 4-6, and Figure 4-7 are capable of stimulating ASC differentiation into tenogenic lineage.
4.3.5 Non-tenogenic lineage expression of ASCs within loaded scaffolds
ASCs being mesenchymal stem cells have the potential to differentiate into various musculoskeletal lineages including bone, cartilage, and skeletal muscles in response to mechanical loading [150]. Thus, in order to identify the appropriate uniaxial tensile strain and loading frequency for tenogenic differentiation, it is important to check for other lineage markers to identify any cross-differentiation or synergistic expression of
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differentiation markers of multiple musculoskeletal tissue lineages. To this end, we investigated the levels of osteogenic, chondrogenic, and myogenic expression in ASCs- encapsulated collagen scaffolds subjected to uniaxial tensile loading at the specified strains and frequencies. The results are displayed in Figure 4-9, Figure 4-10, and Figure 4-11.
Osteogenic lineage
Figure 4-9 displays the profile of osteogenic genes RUNX2 and ALP expressed in ASCs encapsulated within collagen scaffolds when stimulated with BMP-12 or subjected to uniaxial tensile loading at 0%, 2%, 4%, and 6% strains at 0.1 Hz and 1 Hz frequency for 7 days.
Among the mechanically stimulated samples, 4% strain at 1 Hz frequency is the only uniaxial tensile loading regime that exhibits increases in osteogenic gene expression, with
4-fold increases in both osteogenic genes RUNX2 and ALP when compared to the 0%
(non-loaded) group. Samples loaded at 6% strain at 0.1 Hz (p < 0.01) and 1 Hz (p < 0.05) display an increased expression in only RUNX2, while increased ALP expression is observed in samples stimulated at 4% strain at 0.1 Hz (p < 0.05). Samples stimulated with the growth factor BMP-12 do not show any change in the level of osteogenic markers.
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Figure 4-9. Effect of uniaxial tensile loading on osteogenic gene expression of ASCs within 3D collagen scaffolds. Gene expression profiles of ASCs encapsulated within collagen scaffolds subjected to BMP-12 treatment or uniaxial loading at 0% (non-loaded), 2%, 4%, and 6% strains at 0.1 Hz and 1 Hz loading frequencies. The graphs depict fold changes in osteogenic markers: RUNX2 and ALP. * indicates significant fold increase with respect to the 0% samples. * indicates p < 0.05, ** denotes p < 0.01, *** corresponds to p < 0.001. † represents significant difference between 2% and 4% groups while ‡ is the statistical difference with respect to 6% group (p < 0.05). # represents a significant difference between 0.1 Hz and 1 Hz group at the same magnitude of strain with p < 0.05. § depicts significant difference with respect to non-loaded samples chemically stimulated with BMP-12 with p < 0.05. Samples stimulated with 4% strain at 1 Hz show increased level of both osteogenic markers.
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Chondrogenic lineage
Differentiation response of ASCs towards the chondrogenic lineage was studied by quantifying the expression of Collagen II and SOX9 genes in ASCs-encapsulated 3D collagen scaffolds subjected to uniaxial tensile loading at 0%, 2%, 4%, and 6% strains at
0.1 Hz and 1 Hz frequency as depicted in Figure 4-10. The results clearly demonstrate that
ASCs seeded within the collagen scaffolds undergo a chondrogenic response only when stimulated at 2% strain at 1 Hz frequency, with over 10-fold increases in both Collagen II
(p < 0.01) and SOX9 expression (p < 0.001) when compared to the 0% (non-loaded) samples. Rest of the groups, including the samples treated with BMP-12, do not exhibit any increase in chondrogenic markers.
Myogenic lineage
Finally, the myogenic lineage commitment potential of ASCs in response to uniaxial tensile loading at the specified strains and frequencies was evaluated by quantifying the expression of MyoD and Myogenin genes. Figure 4-11 reveals that there are no significant changes observed in the levels of myogenic markers in any of the groups, and the loading parameters of 2%, 4%, and 6% at 0.1 Hz or 1 Hz frequency were not able to elicit myogenic response from ASCs after 7 days in culture within 3D collagen scaffolds.
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Figure 4-10. Effect of uniaxial tensile loading on chondrogenic gene expression of ASCs within 3D collagen scaffolds. Gene expression profiles of ASCs encapsulated within collagen scaffolds subjected to BMP-12 treatment or uniaxial loading at 0% (non-loaded), 2%, 4%, and 6% strains at 0.1 Hz and 1 Hz loading frequencies. The graphs depict fold changes in various chondrogenic markers: Collagen II and SOX9. * indicates significant fold increase with respect to the 0% samples. * indicates p < 0.05, ** denotes p < 0.01, *** corresponds to p < 0.001. † represents significant difference between 2% and 4% groups while ‡ is the statistical difference with respect to 6% group, both with a 95% confidence interval. # represents a significant difference between 0.1 Hz and 1 Hz group at the same magnitude of strain with p < 0.05. § depicts significant difference with respect to non-loaded samples chemically stimulated with BMP-12 with p < 0.05. Samples stimulated with 2% strain at 1 Hz show increased level of both chondrogenic markers.
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Figure 4-11. Effect of uniaxial tensile loading on myogenic gene expression of ASCs within 3D collagen scaffolds. Gene expression profiles of ASCs encapsulated within collagen scaffolds subjected to BMP-12 treatment or uniaxial loading at 0% (non-loaded), 2%, 4%, and 6% strains at 0.1 Hz and 1 Hz loading frequencies. The graphs depict fold changes in various myogenic markers: MyoD and Myogenin. * indicates significant fold increase with respect to the 0% samples. * indicates p < 0.05, ** denotes p < 0.01, *** corresponds to p < 0.001. † represents significant difference between 2% and 4% groups while ‡ is the statistical difference with respect to 6% group, both with a 95% confidence interval. # represents a significant difference between 0.1 Hz and 1 Hz group at the same magnitude of strain with p < 0.05. § depicts significant difference with respect to non-loaded samples chemically stimulated with BMP-12 with p < 0.05. No significant increase of myogenic markers in ASCs was observed in any of the mechanically stimulated groups.
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Thus, from the gene expression results obtained from Figure 4-6 through Figure 4-11, it can be concluded that the loading regime of 2% strain at 0.1 Hz frequency is the only condition that produces a pure tenogenic response from ASCs embedded within 3D collagen scaffolds. The groups of 2% and 4% strains at 1 Hz, though show increased levels of tenogenic markers, also display elevated expression of chondrogenic and osteogenic markers, respectively. This demonstrates that the magnitude of strain and frequency employed during uniaxial tensile loading dictates whether ASCs undergo solo tenogenic response or synergistic differentiation into more than one musculoskeletal lineages.
4.4 Discussion
ASCs have been gaining popularity over BMSCs for tendon tissue engineering strategies in recent years due to their relative abundance, ease of isolation, and anti-inflammatory properties [110, 113]. Previous studies have demonstrated the ability of ASCs to differentiate into tenocytes with the help of chemical factors such as BMP-12, BMP-13, or
BMP-14 [111, 121, 144]. Also, it has been observed that ASCs have the ability to respond to mechanical stimuli by undergoing changes in their morphology and biochemical expression [139-141]. However, to the best of our knowledge, there is no systematic study that investigates the effect of different strain magnitudes and loading frequencies on the tenogenic response of ASCs when subjected to mechanical stimulation.
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Hence, in this study, we aimed to identify the magnitude of strain and loading frequency appropriate for mechanically-stimulated tenogenic differentiation of ASCs cultured in a
3D microenvironment in order to aid the generation of effective and physiological relevant tissue-engineered tendon scaffolds. Uniaxial tensile force govern the dynamic in vivo environment of tendons and hence was considered to be the most relevant type of mechanical loading to stimulate ASCs towards tenogenic differentiation [17]. Collagen, being the major constituent of the tendon ECM was the preferred biomaterial to encapsulate
ASCs in order to elucidate its behavior in a 3D environment. The in vivo tendon physiology was taken as the reference to decide the mechanical loading regimes that would be applied on ASCs encapsulated-collagen scaffolds. Three strain values of 2%, 4%, and 6% that correspond to normal physiological loading (low), intense physiological loading (medium), and pathophysiological loading (high), respectively of tendons in their native environment were chosen for the load application [17]. Further, 0.1 Hz (low) and 1 Hz (high) that fall within the physiological cyclic loading rates corresponding to gentle and rapid stretching of tendons during body movement were selected as the loading frequencies [151]. Thus,
ASCs-encapsulated 3D collagen scaffolds were subjected to cyclic uniaxial tensile strains of 2%, 4%, or 6% in magnitude, loaded at 0.1 Hz or 1 Hz frequencies for 2 hours/day over a period of 7 days and were characterized in terms of their viability, morphology, proliferation, differentiation and matrix organization.
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The orientation of collagen in the scaffold matrix of loaded and non-loaded samples was visualized using SEM (Figure 4-1 and Figure 4-2). It was observed that all of the loaded samples show distinct organization in the structure of their matrix when compared to the random distribution of collagen fibers in the non-loaded samples. This matrix alignment could be attributed to a combinatorial effect of mechanical loading and cell-mediated compaction of collagen fibers, as demonstrated in other studies [102, 103]. Further, the orientation of the collagen matrix was consistently parallel to the axis of load application in all the loaded groups. Quantifying the amount of matrix directionality using ImageJ, it was established that the loaded samples at 2%, 4%, and 6% strains showed statistically significant increases in their matrix alignment in comparison to non-loaded samples at both
0.1 Hz and 1 Hz loading frequencies Figure 4-3. Significantly, there was a distinct correlation between the magnitude of applied strain and the amount of directionality exhibited by the collagen matrix. The applied strain magnitudes of 2%, 4%, and 6% induced 2-fold, 3-fold, and 4-fold increases, respectively, in the degree of matrix compaction (Figure 4-3). Interestingly, no such correlation in matrix organization was apparent with change in loading frequencies between samples loaded at 0.1 Hz and 1 Hz.
Although the directionality was found to be significantly higher in scaffolds loaded at 6% strain at 1 Hz versus at 6% strain at 0.1 Hz, on re-examining the samples it was attributed mainly due to a very small region in samples that showed dense matrix organization, while for the substantial part, the directionality histograms was found to be similar to the ones obtained with 6% strain at 0.1 Hz.
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The ASCs encapsulated within collagen scaffolds were found to be predominantly viable after 7 days of applying various mechanical loading regimes as shown by the representative live-dead assay images in Figure 4-4. The lack of apparent dead cells (red) indicated that the applied loading conditions did not induce cytotoxicity of ASCs. Also, looking at the morphological appearance of ASCs, it was evident that the samples loaded at 2% strain at
0.1 Hz frequency were distinctly elongated when compared to the rest of the groups including the non-loaded samples, which were round in shape. Further, the indirect quantification of total cell number within the scaffold through PicoGreen dsDNA assay
Figure 4-5 demonstrated no decrease in the ASC number from the initial seeding density of 200,000 cells per region of interest within scaffold and thus supported the visually- estimated inference from live-dead assay. However, ASCs in all the loaded groups, though viable, showed limited proliferation when compared to the non-loaded scaffolds, which underwent a 1.5 fold increase in proliferation over 7 days of culture. This was in accordance with previously published literature who reported limited or inhibited proliferation exhibited by loaded ASCs and tendon fibroblasts that however was accompanied with significant increases in ECM gene expression and protein synthesis [127, 152]. Hence, the limited proliferation of ASCs observed in this study on application of uniaxial tensile loading could be an indicator for the onset of cell differentiation.
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Detailed gene expression analyses of the samples loaded at 0% (non-loaded), 2%, 4%, and
6% strain at 0.1 Hz and 1 Hz frequencies were performed using real-time qPCR to quantify the level of ECM genes and differentiation markers belonging to various musculoskeletal lineages. Non-loaded samples treated with BMP-12, an established tenogenic differentiation growth factor [111], were used as the positive control to compare the differences in gene expression of chemically and mechanically stimulated ASCs within 3D collagen scaffolds. The fold changes in ECM genes displayed in Figure 4-6 indicated that
Collagens, especially Collagen III, which is secreted in the early stages of ECM synthesis
[153, 154], exhibited significantly elevated expressions ranging from 5 to 25-fold in the loaded ASCs-encapsulated samples, as well as the BMP-12 stimulated group when compared to non-loaded scaffolds (p < 0.05). While all the uniaxial tensile loading regimes with the exception of 6% at 1 Hz induced 3- to 5-fold rise in Collagen I expression, interestingly, BMP-12 was unable to elicit higher Collagen I levels Figure 4-6. This result conformed to previous study that had stimulated ASCs with BMP-12 and did not observe a rise in Collagen I expression [111]. This suggested that mechanical stimulation was more effective when compared to chemical treatment in its ability to direct the expression of the mature Collagen I fibrils by ASCs. The formation of Collagen I fibrils was possibly triggered by the combination of cell and mechanical loading-mediated matrix organization.
GAGs, on the other hand, that are known to play a role in regulating the alignment and orientation of collagen fibers [153, 155], did not exhibit a global increase in expression with uniaxial tensile loading. In fact, the only groups that displayed predominantly
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significant increases in both Decorin and Aggrecan are the ones loaded at 2% strain at 0.1
Hz (3-fold) and 1 Hz frequencies (9-fold) (Figure 4-7). The combined results for Collagens and GAGs identified three specific uniaxial tensile loading regimes that exhibited statistically significant rises in the ECM markers of ASCs encapsulated within 3D collagen scaffolds: 2% strain at 0.1 Hz, 2% strain at 1 Hz, and 4% strain at 1 Hz (p < 0.05).
Next, we evaluated the expression levels of tenogenic markers, namely Tenascin, Scleraxis, and Tenomodulin in ASCs in response to the different uniaxial tensile loading regimes
(Figure 4-8). Tenascin, a protein expressed during tendon development; Scleraxis, a transcription factor detected in tendon precursor cells; and Tenomodulin, a regulator of cell differentiation and collagen maturation, are found to play critical roles in tenogenic differentiation of MCSs [153, 156-158]. In accordance to the findings in previous studies, non-loaded samples treated with BMP-12 showed significant increases in all three tenogenic markers by atleast 4-fold when compared to non-loaded control group (p < 0.05).
Amongst the mechanically-stimulated samples, the only three groups that exhibited significant increases in tendon-related genes ranging between 5- to 12-fold were 2% strain at 0.1 Hz, 2% strain at 1 Hz, and 4% strain at 1 Hz (Figure 4-8). Remarkably, the three loaded groups identified with elevated tenogenic expression coincided with the groups that demonstrated increased ECM gene expression in Figure 4-6 and Figure 4-7. This suggests that ECM synthesis along with collagen matrix organization influences the differentiation
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of ASCs into the tendon lineage. Thus, the three groups of 2% strain at 0.1 Hz, 2% strain at 1 Hz, and 4% strain at 1 Hz were selected as the uniaxial tensile loading regimes capable of differentiating ASCs encapsulated within 3D collagen scaffolds into tenocytes.
ASCs being mesenchymal stem cells have the potential to differentiate into various musculoskeletal lineages including bone, cartilage, and skeletal muscles in response to mechanical loading [113]. Earlier studies have observed elevation of osteogenic markers along with tenogenic expression in tendon-derived stem cells stimulated with mechanical loading [159]. This is undesirable for tendon tissue engineering strategies due to the risk of in vivo ectopic bone formation when implanted into the repair site [144]. Thus, in order to identify the appropriate uniaxial tensile strain and loading frequency for pure tenogenic differentiation, it was important to check for other lineage markers to identify any cross- differentiation into multiple musculoskeletal tissue lineages. To this end, we evaluated the expression levels of osteogenic, chondrogenic, and myogenic markers in ASCs encapsulated within 3D collagen scaffolds subjected to the aforementioned uniaxial tensile loading regimes. None of the BMP-12 treated samples displayed elevated expression of non-tenogenic markers (Figure 4-9, Figure 4-10, and Figure 4-11), indicating that BMP-
12 used in its dosage of 1000ng/ml over 7 days of culture directed solo tenogenic differentiation of ASCs within collagen scaffolds. Remarkably, the loading regimes of 2% at 1 Hz frequency, along with increased expression of tenogenic genes, exhibited elevated levels of chondrogenic (12-fold) markers (p < 0.05) (Figure 4-10). On the other hand, the uniaxial tensile loading modality of 4% strain at 1 Hz frequency resulted in elevated 126
expression of osteogenic markers (5-fold) along with higher tenogenic expression (p <
0.05) (Figure 4-9). A previous study investigating the effect of mechanical loading on
MSC had observed that the loading frequency of 1 Hz was able to stimulate osteogenic differentiation in MSCs, so that could explain our findings [138]. Interestingly, no loading regime was capable of inducing myogenic gene expression in ASCs embedded within 3D collagen scaffolds. Prior studies have shown that uniaxial tensile stimulation is capable of differentiating ASCs to myogenic lineage in monolayer culture [136]. In our case, the presence of 3D microenviroment composed of a predominantly collagen-based matrix and its stiffness could likely have influenced the ASC-matrix interaction and contributed to the lack of myogenic differentiation. Significantly, the samples loaded at 2% strain and 0.1 Hz frequency having higher expression of tenogenic markers did not display any of the non- tenogenic musculoskeletal differentiation genes.
For direct comparison, the gene expression of all the markers assayed for each loading regime that showed significant increases in tenogenic markers compared to control (non- loaded) group, namely 2% at 0.1 Hz, 2% at 1 Hz, and 4% at 1 Hz were each consolidated into one graph (Figure 4-12). Figure 4-12 clearly demonstrates that 2% at 0.1 Hz is the loading regime that is able to stimulate ASCs to undergo pure tenogenic differentiation, with no significant increases in both osteogenic and chondrogenic marker expression.
Hence, 2% uniaxial tensile strain at 0.1 Hz loading frequency, without the use of any chemical factors, was able to induce pure tenogenic differentiation of ASCs encapsulated within 3D collagen scaffolds. 127
Figure 4-12. Effect of uniaxial tensile loading on ASC differentiation within 3D scaffolds. Gene expression profile of ECM, Tenogenic, Osteogenic, Chondrogenic, and Myogenic markers mapped for samples loaded at 2% strain at 0.1 Hz, 2% strain at 1 Hz, and 4% strain at 1 Hz. Genes highlighted in yellow indicate statistically higher expressions when compared to non-loaded samples (p < 0.05). 128
Published studies that have investigated the effect of mechanical loading on MSCs and
BMSCs have often identified the 1 Hz frequency to be suitable for tenogenic differentiation
[130, 132, 135]. Even our results display equal if not higher ECM and tenogenic gene expression at 1 Hz when compared to 0.1 Hz (Figure 4-6, Figure 4-7, and Figure 4-8)
However, the risk of cross-differentiation into other musculoskeletal lineages seems to be significantly enhanced with the use of higher loading frequency of 1 Hz, and hence makes
0.1 Hz as the preferred choice of cycling frequency for tendon tissue engineering applications.
4.5 Conclusion
In Chapter 4, we applied different loading regimes including low, medium and high physiological strains and low and high physiological loading frequencies to ASCs-seeded within 3D collagen scaffolds. We further evaluated the cell morphology, viability and proliferation, matrix alignment, and differentiation of ASCs. Figure 4-13 displays the overall conclusion and results obtained from Chapter 4.
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Figure 4-13. Major results obtained in Chapter 4. The effect of uniaxial strains and frequencies on the matrix organization and tenogenic differentiation of adipose stem cells (ASC) encapsulated within 3D collagen scaffolds.
The combined results of the ASC-collagen scaffolds subjected to mechanical stimulation at 2% strain and 0.1 Hz frequency indicate key features: (1) there a definitive change in the
ASC morphology with the rounded cells resembling more like tendon fibroblasts, with their elongated shape and the cytoplasmic extensions, (2) the scaffold matrix shows distinct organization with the directionality of collagen fibers being parallel to the axis of load application, (3) the gene expression data demonstrates significant increases in ECM and
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tendon-related genes, and (4) no cross-differentiation potential of ASCs to osteogenic, chondrogenic or myogenic lineage is observed thus giving rise to pure tenogenic differentiation. Hence, we have identified 2% strain and 0.1 Hz frequency to be the appropriate uniaxial tensile strain and loading frequency to mechanically-induce pure tenogenic differentiation of ASCs encapsulated in a 3D environment.
Interestingly, this corresponds to the low strain-low frequency loading regime usually employed during physical therapy prescribed during the rehabilitation phase after tendon repairs [151]. Thus, mechanical conditioning of ASCs at 2% strain and 0.1 Hz frequency within the collagen scaffolds that enhances ECM production and stimulate tenogenesis has the potential to accelerate tendon healing when implanted at the repair site with the rotator cuff under controlled mobilization.
Now that the mechanical loading bioreactor is validated (Chapter 3) and the desired loading regime for mechanical conditioning of ASCs within collagen scaffolds for tenogenic differentiation has been identified (Chapter 4), in Chapter 5 we aim to improve the poor protein retention properties of collagen hydrogel. This is important in order to be able to incorporate growth factors within the scaffold with sustained release and bioactivity under mechanical loading conditions that could further stimulate ASC proliferation and ECM secretion at the tendon repair site.
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Chapter 5
Composite Collagen Scaffolds (PNCOL) Interspersed with Polycaprolactone (PCL) Nanofibers for Controlled Growth Factor Delivery under Uniaxial Tensile Mechanical Loading
5.1 Introduction
Healing of injured tendons occurs through three main stages: inflammatory, reparative, and remodeling phases. This highly complex process is orchestrated by a large number of growth factors with diverse cellular and molecular effects [22, 153]. The reparative and early remodeling phases characterized by increased cell proliferation, and abundant collagens and GAGs secretion is initiated within 48 hours after injury and lasting around three months. The growth factors involved during this phase play a major role in regulating the critical events of cell migration, proliferation, ECM secretion, and differentiation that helps the injured tendon to synthesize and remodel its matrix and to achieve its native 132
structural and mechanical properties [144]. The reparative stage is thus usually the most targeted phase for strategies involving exogenous introduction of growth factors into the repair site to aid the native tendon healing process [26].
Platelet-derived growth factor (PDGF) is one such healing promotive factor secreted during the reparative phase that is identified to be important for tendon healing, with its receptor expressed up to 6 months after the onset of injury [37, 160]. PDGF is a known chemoattractant, cell proliferative agent that induces expression of other growth factors including the highly anabolic insulin growth factor-1 (IGF-1) during the early repair phase
[153, 161]. Further, there is ample evidence demonstrating the efficacy of PDGF the in vivo tendon repair site. However, these studies also highlight that improved healing with
PDGF is largely dependent on the timing, dosage, and delivery vehicle used [37, 162-164].
The mode of delivery for any growth factor is critical because of their short biological half- lives (30-90min) that causes rapid degradation when administered intravenously [165,
166]. Their rapid clearance necessitates supraphysiological doses and multiple injections of these growth factors [167-169]. Using larger than normal doses of growth factor can, however, have undesirable effects such as tumor formation, cytotoxicity, inflammation, and uncontrolled ECM synthesis at non-target sites [170-172]. Many strategies have been employed in the recent years to ensure controlled delivery of growth factors along with 133
preserving its bioactivity. Gene therapy approaches that involve transducing cells with the growth factor coding sequence through viral vectors are utilized as a carrier, with the cells over-expressing and delivery the growth factor at the repair site [173, 174]. Although this technique ensures controlled and sustained release of the growth factor, using genetically- manipulated cells through introduction of virus raises concerns of a tumorigenic response in the host tissue [26]. Hence, tissue-engineered scaffolds built with synthetic biomaterials or hydrogels have been deployed as growth factor delivery vehicles to augment tendon healing. Depending on the structural and mechanical properties of the scaffold, different techniques are employed to encapsulate the growth factor within the matrix.
Tissue-engineered constructs composed of synthetic or fibrous biomaterials are mechanically robust but lack the ability to accommodate bioactive proteins within their matrix, due to which growth factors are usually incorporated by physical encapsulation methods. [60, 162, 175]. For instance, Kovacevic et al. implanted PDGF-soaked collagen sponge scaffolds into the rat rotator cuff repair site. Though they observed initial increases in cell proliferation and vascularization, PDGF was not able to enhance the maturity of collagen fibers or increase fibrocartilage formation [162]. Cheng et al. fabricated composite fibers composed of electrospun collagen containing PDGF-releasing nanoparticles. The delivery system significantly enhanced the in vitro tenogenic differentiation of ASCs in short-term culture [175]. Another nanofiber-based delivery approach by Sahoo et al.
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involved electrospinning PLGA polymeric solution blended with the growth factor to obtain FGF-releasing nanofibrous scaffolds that retained their bioactivity until 7 days of culture [60]. Increased proliferation, tenogenic expression, and collagen secretion was elicited from BMSCs seeded onto the scaffold surface. These results demonstrated the efficacy of growth factor introduction in promoting cell proliferation and tenogenesis on a short-term basis. However, the delivery systems synthesized by blending, coating, or soaking the growth factor in the scaffold matrix do not exhibit long-term retention of the physically encapsulated proteins within the scaffold due to which the cellular response to response to the growth factor is transitional.
Tissue-engineered scaffolds composed of soft biomaterials such as hydrogels have poor mechanical properties to physically retain any biologics within their matrix. Direct encapsulation of growth factors into such scaffolds typically lead to rapid burst release during the initial phase, followed by passive diffusion release of the remaining protein in the matrix [176]. Hence, to improve the protein retention capacity of hydrogels, they are subjected to chemical modifications such as polymer crosslinking to increase matrix density, covalent-binding through peptide groups to immobilize the growth factor, and micro/nanoparticle carrier systems to retard the protein release rate [54, 176-179]. In the study by Chen et al., BMP-2 was bound to poly (ethylene glycol) (PEG) hydrogels using a chemical crosslinker. The polymer solution was injected into a rabbit rotator cuff injury
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model, and after 8 weeks, increased fibrocartilage and bone layer formation was observed
[178]. A carrier system developed by Peret et al. consisted of bFGF-containing microparticles within alginate hydrogels. Controlled release of the growth factor was observed with increased cell proliferation [180]. Zisch et al. designed a bioresponsive system consisting of VEGF immobilized onto RGD adhesion peptides-conjugated PEG hydrogels in order to deliver the growth factor based on cell demand. Active delivery of
VEGF was confirmed and subcutaneous introduction of the hydrogel in rats induced tissue vascularization [181].
These delivery systems thus exhibited acceptable in vitro protein release profiles with increases in in vivo cell proliferation and function at the repair site. Despite promising results, two major limitations noticed with such delivery techniques are high burst release of the growth factor and decreased protein bioactivity due to the harsh chemical environment experienced during hydrogel crosslinking and scaffold fabrication [182, 183].
Significantly, most of the growth factor delivery systems currently developed have been designed and evaluated under static conditions. However, since tendons exist in highly dynamic environment, mechanical loading is expected to play a major role in the protein delivery profiles of the tissue-engineered scaffolds [176, 184]. Hence, it is critical to elucidate the effect of mechanical stimulation on controlled delivery of growth factor in 136
order to develop a physiologically relevant growth factor delivery system [184]. Lee et al. previously demonstrated the influence of mechanical loading on VEGF release from polymeric hydrogels and its positive effect on angiogenesis [185]. But, to the best of our knowledge, there is no tissue-engineered scaffold developed for growth factor delivery to augment tendon that has been characterized under physiologically relevant mechanical loading conditions.
The objective of this chapter was to synthesize a composite collagen scaffold (PNCOL) by the inclusion of polycaprolactone (PCL) nanofibers into the collagen matrix for controlled delivery of bioactive growth factor under mechanical loading environment. Since collagen in its hydrogel form was limited in its capacity to retain bioactive molecules within its matrix and is associated with large burst release, we aimed to reinforce the collagen scaffold by incorporating nanofibers of the synthetic biomaterial polycaprolactone (PCL).
Our hypothesis was that the nanofibrous structure of PCL would provide higher surface area for growth factor immobilization and minimize burst release, and incorporation of biomolecules such as heparin or/and BSA would help in the long-term preservation of protein activity. Further, evaluation of the composite scaffold under uniaxial tensile loading conditions was hypothesized to be a better representation of the in vivo tendon environment when implanted at the repair site.
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This study was pursued with the vision that along with mechanically-stimulated cells within an organized scaffold matrix, controlled long-term delivery of a growth factor that can enhance the proliferation and ECM synthesis of the differentiating ASCs as well as the resident tenocytes would help accelerating the healing process at the repair site.
To this end, the composite scaffold PNCOL was synthesized by interspersing homogenized electrospun PCL nanofibers into collagen matrix. The biological properties of PNCOL incorporated with different concentrations of PCL nanofibers were characterized in terms of protein release and bioactivity to identify the most viable concentration of PCL within
PNCOL for controlled delivery of bioactive growth factors. Next, the performance of heparin and bovine serum albumin (BSA) in restoring and preserving the growth factor activity was evaluated for a long-term period within cellular PNCOL scaffolds. After identifying the appropriate components of the PNCOL scaffold for growth factor delivery, the scaffolds were mechanically loaded using the uniaxial tensile strain bioreactor described in Chapter 3 with the mechanical loading regime identified in Chapter 4 to simulate the in vivo tendon environment at the repair site. Finally, the bioactivity of the released PDGF was assessed in terms of its effect on migration and proliferation of tendon fibroblast-like cells.
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5.2 Materials and Methods
5.2.1 Synthesis of protein-immobilized composite collagen scaffolds
Figure 5-1. Synthesis of GF-hb-PNCOL composite scaffolds. (A) PCL dissolved in an organic solvent mixture subjected to electrospinning to obtain PCL nanofibers. (B) Finely chopped PCL nanofibers undergo oxygen-based plasma treatment to introduce functional groups on its surface. (C) Growth factor incubated with heparin (h) and BSA (b) at 1:40:2000 ratio to preserve and protect its activity. The GF-hb complex added to the plasma-treated PCL nanofibers to allow immobilization of the growth factor via the functional groups onto the PCL surface. (D) Growth factor immobilized onto PCL nanofibers incorporated in neutralized cell-encapsulated Collagen Type-I solution and polymerized to obtain GF-hb-PNCOL scaffolds.
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The growth factor-immobilized polycaprolactone nanofibers interspersed within the collagen scaffolds (GF-hb-PNCOL) were synthesized in a three-step sequential process.
The schematic of the experimental procedure involved in the synthesis of GF-hb-PNCOL scaffolds is depicted in Figure 5-1.
Electrospinning of PCL: Polycaprolactone (PCL) is an FDA-approved synthetic polymer than has been widely used as a biomaterial for tissue engineering applications and can be processed by various techniques [186]. Electrospinning, a method to produce nanofibers from polymeric solution through the use of electric force, is one of the technique routinely employed with PCL-based scaffold manufacturing process
PCL nanofibers were obtained by electrospinning using our established protocol developed in our lab [186, 187]. Briefly, a 16% (w/v) solution of PCL (Mw = 45,000, Sigma-Aldrich) dissolved in an organic mixture of chloroform/methanol (Sigma-Aldrich, US) at 3:1 ratio was delivered through a syringe pump with 20 gauge needle at a feed rate of 8000 µl/hr with 20 KV voltage supply. The fiber mat collected on an aluminum foil at the collecting plate were dried for 72 hours, peeled off from the foil and chopped into tiny fragments using a high speed homogenizer (Ultra Turrax).
Growth factor immobilization on PCL nanofibers: The homogenized nanofibrous mats were subjected to oxygen-based plasma-treatment (Harrick Plasma) for 3 minutes at medium power to reduce its hydrophobicity by introducing oxygen functional groups onto the surface of PCL [188]. This also allows PCL to act as ideal surface for proteins such as
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growth factors to bind via the functional groups. Prior to immobilizing growth factors onto the surface of PCL nanofibers, the growth factor was blended with heparin ‘h’ (Sigma-
Aldrich, US) and Bovine Serum Albumin (BSA) ‘b’ (Sigma-Aldrich, US) at a mass ratio of 1:40:2000 for 15 minutes each at room temperature. Heparin and BSA were used to preserve and protect the long-term bioactivity of the growth factor. The GF-heparin-BSA complex (GF-hb) was then incubated with PCL nanofibers for 20 minutes at room temperature to allow the growth factor to bind to the surface of PCL.
Incorporation of GF-immobilized PCL nanofibers into collagen solution: Collagen Type-
I solution (Corning, US) solution at pH~3-4 was diluted to 2.5 mg/ml and neutralized with cold 1 N NaOH along with sterile water and PBS in accordance with the manufacturer’s protocol. For cellular scaffolds, sterile water was replaced with cells resuspended in their respective culture media. The GF-hb bound PCL nanofibers were incorporated into the collagen solution with or without cells at 0% (pure collagen), 1%, 3%, and 6% (w/v) concentration, and polymerized at 37 oC for 1 hour to synthesize GF-hb-PNCOL scaffolds.
Different concentrations of PCL nanofibers were incorporated inside the collagen scaffold to identify the appropriate amount of PCL that can exhibit controlled release of bioactive proteins.
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5.2.2 Protein retention capacity of PNCOL scaffolds
The effect of PCL nanofiber concentration on the short- and long-term protein retention capacity of PNCOL scaffolds was investigated by quantifying the release of a model protein lysozyme over 4 hours and over 21 days, respectively. The time period of 4 hours was used to evaluate the burst release profile exhibited by the scaffolds, while the 21-day culture would correspond to the diffusion release phase of the encapsulated protein.
Lysozyme (MW 14,500, Sigma-Aldrich, US) was chosen as the model protein because it resembles many growth factors including BMP-2 and PDGF in terms of its physicochemical properties and size, with an isoelectric point (pI 9–10), and molar extinction coefficient around 38,000 cm−1 M−1 [189].
Lysozyme was covalently labeled with Alexa Fluor 350 dye (Molecular Probes) according to the manufacturer’s protocol in order to detect and quantify release of lysozyme from
PNCOL scaffolds using a microplate fluorometer. Briefly, the chemical reaction between the succinimidyl ester groups of the dye with the primary amines of lysozyme was carried out in 0.1M sodium bicarbonate solution, pH 8.2, for 1 hour at room temperature to form stable dye protein conjugates. The labeled lysozyme was then separated from the unreacted free dye by size exclusion chromatography using Sephadex G-25 resin column (Sigma-
Aldrich), equilibrated in a buffer containing 0.1M potassium phosphate, 1.5M NaCl, with
2 mM sodium azide, pH 7.2. A dye-to-protein molar ratio of 0.785 was determined by spectrophotometric analysis (SOFTmax Pro) at 280 nm and 346 nm wavelengths, using
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the extinction coefficient value of lysozyme and Alexa Fluor 350 provided in the product datasheet.
The Alexa Fluor 350 tagged lysozyme with a final concentration of 50 μg/ml was incubated with PCL nanofibers and incorporated into the collagen solution at 0%, 1%, 3%, and 6%
(w/v) PCL nanofiber concentrations to obtain PNCOL scaffolds. The release kinetics of lysozyme from the PNCOL scaffolds was determined by aspirating the supernatant (PBS) from each sample completely and replacing it with equivalent volume of fresh PBS. To evaluate the initial burst release, samples were collected at 1, 2, 3, and 4 hours. Diffusion release was investigated by collecting samples at 1, 2, 4, 6, 10, 14, 17, and 21 days. The collected samples were assayed for fluorescence intensities at 346/442 nm excitation/emission wavelengths using a microplate fluorometer (Perkin Elmer Wallac
1420). A standard curve was generated by plotting the fluorescence readings against different known concentrations of the dye-labeled lysozyme in order to quantify the amount of lysozyme released at each time point.
5.2.3 Growth factor, cells, and culture media
The short- and long-term bioactivity of GF-PNCOL scaffolds was evaluated to identify the optimal concentration of PCL nanofibers within the GF-PNCOL scaffold and then characterize the performance of GF-hb-PNCOL scaffolds. For this purpose, Bone
Morphogenetic Protein-2 (BMP-2) was selected as the model growth factor, and pre- osteoblastic cells MC3T3-E1 were chosen as the model cell line. 143
BMP-2 was chosen for its similarities in structure and activity with other heparin-binding growth actors including PDGF [27], and because its activity could be tested on MC3T3-
E1 cells by using well-established osteoblastic differentiation kits such as the ALP assay
(Sigma-Aldrich) with good reproducibility and sensitivity.
MC3T3-E1 cells (ATCC, CRL-2593; US) were cultured in complete media consisting of
Alpha-Minimum Essential Medium (α-MEM) (Life Technologies, US) supplemented with
10% FBS (Gibco, US) and 1% Penicillin-Streptomycin (Life Technologies, US). To incorporate cells within PNCOL scaffolds, MC3T3-E1 between passages 3-5 at a seeding density of 1x106 cells/ml were mixed into neutralized collagen solution before the addition of PCL nanofibers. Next, 200ng/ml of BMP-2 (Peprotech, US) incubated with PCL nanofibers for 20 minutes were together incorporated into the cell-encapsulated collagen scaffold solution and polymerized to obtain the GF-PNCOL scaffolds.
The scaffolds were maintained in complete media for short-term evaluation of BMP-2 bioactivity. For long-term in vitro study, the scaffolds were incubated within osteogenic media comprising of osteogenic factors 10mM ß-glycerophosphate (Sigma-Aldrich, US) and 50μg/ml ascorbic acid (Sigma-Aldrich, US) added into the complete media. All short- term characterization studies were conducted for 10 days while scaffolds for long-term studies were cultured over a 21-day period, with the media being replaced every 3-4 days.
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5.2.4 Bioactivity of GF-PNCOL scaffolds
To understand the effect of PCL nanofiber concentration on protein activity, model protein
BMP-2 was incorporated within 0%, 1%, 3%, and 6% (w/v) PNCOL scaffolds. Upon identifying the appropriate PCL concentration within PNCOL scaffolds, the effect of heparin and BSA on preserving and prolonging the bioactivity of the growth factor was studied. To evaluate the solo and combined effects of heparin and BSA on the activity of the growth factor, four groups were used i) GF-PNCOL (control), ii) GF-h-PNCOL, iii)
GF-b-PNCOL, and iv) GF-hb-PNCOL. In these groups, ‘GF’ symbolizes BMP-2, ‘h’ refers to heparin, ‘b’ denotes BSA, and PNCOL specifically refers to 3% (w/v) PNCOL. GF-h was incorporated in 1:40 mass ratio, while GF-b was blended in 1:2000 mass ratio before encapsulating in the PNCOL scaffolds.
In both cases, the alkaline phosphatase activity of pre-osteoblasts MC3T3-E1 cells was measured after 10 days of culture in complete media. The ability of BMP-2 to increase the cellular ALP synthesis within the PNCOL scaffolds was measured to estimate its bioactivity. The ALP activity of MC3T3-E1 cells encapsulated in the respective scaffolds was estimated using our established protocol [187] after ten days of culture in complete media.
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5.2.5 Growth factor release kinetics from GF-hb-PNCOL scaffolds
Upon identifying the best configuration to preserve growth factor activity within PNCOL scaffolds, the release profile of the growth factor was measured over 21 days to elucidate the effect of heparin and BSA on the release kinetics of PNCOL scaffolds. This was investigated by again using BMP-2 as the model protein and quantifying its release by performing BMP-2 Quantikine enzyme-linked immunosorbent assay (ELISA) (R&D systems). GF-hb-PNCOL scaffolds and GF-PNCOL (control) scaffolds were synthesized and incubated in PBS such that 40 ng of the growth factor was loaded in each scaffold. The release profile of the growth factor from the scaffold was determined by collecting the supernatant from each sample and replacing it with equivalent volume of fresh PBS at 1,
2, 4, 6, 10, 14, 17, and 21 days. The BMP-2 concentrations in the aspirated samples were determined by following the manufacturer’s instructions. Briefly, the specified volume of assay diluent was added into the wells of the plate provided with the kit, followed by addition of the release media from the samples. The plate was sealed and incubated on a shaker at room temperature for 2 hours. The supernatant was removed, and each well was rinsed four times with wash buffer. Next, the BMP-2-specific antibody conjugate was added into each well; plate was sealed and incubated again for 2 hours on a shaker at room temperature. The supernatant was discarded, and the plate was rinsed 4 times with wash buffer. Finally, the substrate solution was added into each well and incubated at room temperature for 30 minutes in dark in order to initiate the colorimetric reaction. The
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reaction was terminated by adding stop solution, and the absorbance was measured at
450nm with a wavelength correction of 570 nm. The amount of BMP-2 in the samples was determined by comparing the obtained values with the standard curve generated by assaying known concentrations of BMP-2 standard provided along with the kit.
5.2.6 Long-term in vitro performance of GF-hb-PNCOL scaffolds
To evaluate the long-term in vitro performance of the cells and growth factor encapsulated within the GF-hb-PNCOL scaffolds, 3% (w/v) PCL scaffolds encapsulated with 200 ng/ml
BMP-2 mixed with heparin and BSA in 1:40:2000 ratio were maintained in osteogenic media for 21 days. PNCOL scaffolds without GF was used as a control group. The scaffolds were evaluated in terms of cell proliferation and ALP activity at days 0, 7, 14, and 21.
Cell proliferation: Cell proliferation within GF-hb-PNCOL and PNCOL (control) scaffolds was assessed using the alamarBlue® cell viability assay kit (Life Technologies,
US). The alamarBlue reagent at 20% (v/v) concentration in complete media was added to each sample in volumes of 0.5ml and incubated at 37 oC for 2 hours. The media from each sample was aspirated, and the fluorescence intensities were measured using a microplate fluorometer (Perkin Elmer Wallac 1420) at 565/585 nm excitation/emission wavelengths.
A standard curve generated with varying number of cells and their corresponding fluorescence values in order to determine the total cell number within each scaffold.
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Growth factor bioactivity: The cellular alkaline phosphatase (ALP) activity within GF-hb-
PNCOL and PNCOL (control) scaffolds was estimated by quantifying the amount of product p-nitrophenol (pNP) produced from p-nitrophenyl phosphate (pNPP) substrate
(Sigma-Aldrich, US) in presence of the ALP enzyme. Scaffolds were frozen in liquid nitrogen and mechanically disrupted using a pestle in order to liberate the cells. The liberated cells were then resuspended in an alkaline lysis buffer (10mM Tris-HCl, 2mM
MgCl2, 0.1% Triton X-100, pH 8), and lysed with a homogenizer (Ultra Turrax). The samples were centrifuged, and the supernatant and the pNPP substrate was mixed in a microwell plate at 1:1 ratio and incubated for 30 minutes at 37 oC. The colorimetric reaction was stopped using 3N NaOH, and the samples were measured at 405nm using a microplate spectrophotometer (SpectraMax). A standard curve obtained with dilutions of calf alkaline phosphatase enzyme was used to quantify the total ALP which was then normalized with respect to the total protein content in each respective sample.
The total protein content was quantified using Coomassie Bradford protein assay kit
(Thermo Scientific, US). Briefly, 300µl of the Coomassie Blue Bradford reagent was mixed with 15µl of the samples lysate obtained from ALP assay. The reaction was incubated for 10 minutes, and then the absorbance vales were measured at 595nm wavelength in a microplate spectrophotometer (SpectraMax). A standard curve was generated by measuring the absorbance obtained for different concentrations of BSA in order to quantify the amount of protein within each scaffold.
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5.2.7 Protein release from PNCOL scaffolds under mechanical loading
We established that PNCOL with 3% (w/v) PCL nanofiber concentration in collagen solution is able to both retain protein in its matrix and exhibit preserved and sustained bioactivity over long-term in vitro culture. Next, we investigated the protein release profile from PNCOL in dynamic culture conditions under uniaxial tensile mechanical loading. The goal of this study was to understand whether uniaxial mechanical loading affect the protein release rate from PNCOL scaffold at the bust release phase and at the diffusion phase.
Based on our previous studies, diffusion phase of protein release profile can be considered as a phase immediately after the burst release of the protein from the PNCOL scaffold, which is 3 days after the PNCOL synthesis. The uniaxial tensile loading regime of 2% linear strain at 0.1 Hz frequency for 2 hours/day identified in the Chapter 4 was the chosen mechanical loading condition to conduct this study. Three groups were used for the experiment: (i) Non-loaded PNCOL scaffolds as control samples, (ii) PNCOL scaffolds mechanically loaded starting from day 0 (D0-loaded) after scaffold synthesis, to obtain the release profile due to uniaxial tensile loading from the burst release phase as well as the diffusion phase, (iii) PNCOL scaffolds mechanically loaded starting from day 3 (D3- loaded) after scaffold synthesis, to obtain the release profile due to uniaxial tensile loading from only the diffusion release phase. The procedure employed was similar to the protein retention study conducted in static culture conditions with PNCOL scaffolds having different concentration of PCL. Briefly, PNCOL scaffolds from all three groups were
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encapsulated with 50 µg/ml of Alexa Fluor 350-tagged lysozyme. The scaffolds were polymerized in the silicone loading chambers of the uniaxial tensile strain bioreactor.
Samples from each group were cultured in PBS the aforementioned culture conditions, respectively. At 0, 1, 2, 3, 6, 8, 10, 12, and 14 days, the supernatant was collected, and the chambers were replenished with an equivalent amount of fresh PBS. The fluorescence intensities of the collected samples due to the released lysozyme were measured at 346 nm excitation and 442 nm emission wavelengths using a microplate fluorometer (Perkin Elmer
Wallac 1420). The previously generated standard curve was used to determine the percentage protein release from PNCOL scaffolds under the two mechanically-loaded regimes in comparison to non-loaded scaffolds.
5.2.8 Bioactivity of PDGF released from PDGF-hb-PNCOL
After conducting detailed optimization studies to preserve, enhance, and sustain the bioactivity of GF-hb-PNCOL scaffolds using BMP-2 as the model growth factor, the components identified that exhibit the desired protein retention and bioactivity were used to encapsulate our growth factor of interest, PDGF, within PNCOL scaffolds. The experimental procedure shown in Figure 2-1 was employed to fabricate the PDGF-hb-
PNCOL scaffolds. PDGF was incorporated within PNCOL scaffolds at a final concentration of 1 µg/ml. To confirm that the PDGF in presence of the heparin-BSA complex is able to preserve its bioactivity, the release media from PDGF-hb-PNCOL 150
scaffolds was collected at day 4. This release media estimated to have around 100 ng/ml
PDGF was added onto monolayer MC3T3-E1 cells (experimental group) and evaluated for its ability to induce cell migration and proliferation. PNCOL scaffolds without PDGF (0 ng/ml) were used as negative control samples. Media spiked with 100 ng/ml PDGF was used as positive control. MC3T3-E1 cells were chosen to evaluate the PDGF bioactivity since they are similar to tenocytes in terms of their response to PDGF. It is well-established that MC3T3-E1 cells, like tenocytes, respond to PDGF by undergoing migration accompanied with increased proliferation [190]. Serum-free complete media was used for these experiments in order to limit the rate of proliferation via serum deprivation.
Migration assay: MC3T3-E1 cells were seeded at 80,000 cells/well in a 48-well plate such to achieve a confluency of >90%. Once the cells attached, a scratch was created through the middle of each well using a pipette tip. The media was replaced in order to discard the floating cell debris caused by the removal of cells from the scratched area. The initial positions of the scratched region was marked in each well by identifying it through an optical microscope. The release media from day 4, and the media from the negative and positive control samples were then added to four wells each, and the effect of the media on cell migration was monitored at regular time intervals. During an 18-hour period, bright field images of the scratched region in each well was captured every 2 hours to track any changes in the thickness of the scratch or position of the cells. The reduction or disappearance of the scratched region can be attribute to cells migrating into that area.
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Proliferation assay: MC3T3-E1 cells were seeded at 50,000 cells/well in 48-well plate such that the confluency after cell attachment is around 20-30%. The cell proliferation at day 1 was assayed using alamar Blue cell viability kit (Life Technologies, US). A 10% alamar Blue solution was prepared in complete media with no serum and 0.5ml was added in each well and incubated for 2 hours. The media was collected and evaluated for its fluorescence intensities at 565 nm excitation and 585 nm emission wavelengths using a microplate fluorometer (Perkin Elmer Wallac 1420). A standard curve was generated using different cell concentrations and their corresponding fluorescence intensities to quantify the number of viable cells within the scaffold.
5.2.9 Statistical analyses
Four samples (n=4) were used in each group for all assays. Statistical analysis was conducted using Student’s t-test and One-way ANOVA (IBM SPSS Statistics software).
All values are reported as the average, and the error bars correspond to ± the standard deviation of the mean. All statistical difference was determined with a 95% confidence interval (p < 0.05).
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5.3 Results
5.3.1 Synthesis of PCL nanofibers-interspersed collagen scaffolds
The composite PNCOL scaffold engineered to improve the poor protein retention properties of collagen was fabricated by incorporating PCL nanofibers within the collagen matrix. PCL nanofibers were manufactured by electrospinning technique and homogenized. Based on the established O2 plasma functionalization protocols [188, 191,
192] oxygen-containing functional groups were introduced on PCL surface to decrease its hydrophobicity and enhance PCL nanofiber’s cell and protein binding capacity. The functionalized PCL nanofibers at different (w/v) concentrations were admixed with the growth factor and incorporated homogenously within the neutralized collagen solution containing cells and polymerized to obtain PNCOL scaffolds. The effect of different PCL nanofiber concentrations within PNCOL scaffolds was evaluated in terms of their protein retention capacity and bioactivity.
5.3.2 Characterization of PNCOL scaffolds for protein delivery
5.3.2.1 Protein retention capacity of PNCOL scaffolds
To investigate whether incorporation of PCL nanofibers within collagen matrix has a role in preventing protein burst release and increasing the protein retention, short-term (4- hours) and long-term (21-days) protein retention studies were conducted. Model protein 153
lysozyme was encapsulated in PNCOL scaffolds incorporated with 0% (pure collagen),
1%, 3%, and 6% (w/v) PCL nanofibers to evaluate their protein retention properties. Figure
5-2A presents the short-term protein retention profile and Figure 5-2B displays the long- term protein retention profile exhibited by the PNCOL scaffolds. Figure 5-2A demonstrates that 6% (w/v) PNCOL has the highest short-term protein retention rate compared to the scaffolds with lower concentrations of PCL nanofibers within them. During 4 hours,
60±2.18% of initially loaded protein is released for 0% (w/v) PNCOL scaffold (collagen), while this percentage is dramatically reduced to 20±1.28% for 6% (w/v) PNCOL.
Figure 5-2. Effect of PCL nanofiber concentrations in model protein retention within PNCOL scaffolds. (A) Lysozyme retention profile within PNCOL incorporated with 0, 1, 3, and 6% (w/v) PCL nanofibers during time = 0 to 4 h corresponding to the burst release phase and (B) over 21 days corresponding to the diffusion release phase. Increase in PCL concentration within collagen matrix increases the protein retention properties of the scaffold. [193]
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The long-term protein release data from PNCOL scaffolds displayed in Figure 5-2B further confirms that increased PCL nanofibers within the collagen enhances the protein retention capacity of scaffold. Over 21 days, 3% (w/v) PNCOL and 6% (w/v) PNCOL retains 62 ±
0.79% and 78 ± 0.98% of the protein within the scaffold respectively, while all of the protein within 0% PNCOL is leached out within 2 days. These results establish that incorporation of PCL nanofibers inside collagen scaffolds reduces protein burst release and increases long-term protein retention, with the retention increasing with increased PCL concentrations.
5.3.2.2 Effect of PCL nanofibers on the protein bioactivity of within PNCOL
To understand the effect of PCL nanofiber concentration on protein activity, model protein
BMP-2 was incorporated within 0%, 1%, 3%, and 6% (w/v) PNCOL scaffolds and alkaline phosphatase activity of pre-osteoblasts MC3T3-E1 cells was measured after 10 days of culture in basal media.
Figure 5-3 demonstrates that the ALP activity, which is a direct measure of the protein activity within the scaffold, reduces with the incorporation of PCL nanofibers. Though
PCL nanofibers within the collagen matrix result in higher protein retention capacity, it has an adverse effect on the protein bioactivity within the scaffold.
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Figure 5-3. Effect of PCL nanofiber concentrations on protein bioactivity within PNCOL. Bioactivity of BMP-2 within the scaffolds measured in terms of ALP activity of MC3T3- E1 cells encapsulated within PNCOL after ten days of culture shown in the column graph. The black line graph with red bullets indicates the percentage of protein retained in the respective PNCOL scaffolds as indicated by the secondary axis. Together, incorporation of PCL nanofibers shows an increase in protein retention but a decrease in protein activity. (*) indicates significant difference compared to the adjacent group p < 0.05. [193]
It is observed that 1% and 3% (w/v) PNCOL that have 38 0.48% and 62 0.58% protein retention, respectively, demonstrate 2-fold loss of growth factor bioactivity (p < 0.05) when compared to 0% PNCOL (pure collagen) scaffolds. 6% (w/v) PNCOL which is able to retain 78 0.48% of the protein has over 5-fold loss of bioactivity (p < 0.05). Thus, a reverse correlation between the scaffold’s protein retention capacity and protein bioactivity is observed with increase in PCL nanofibers concentration within the collagen matrix. 156
Based on above findings, 3% (w/v) PNCOL scaffold is found to possess the favorable balance between protein retention and biological properties desired for the tendon scaffold, with an ability to retain 62% of protein within the scaffold over 21 days with only 50% loss in bioactivity. Thus, 3% (w/v) PNCOL scaffold is chosen for all subsequent studies and henceforth would be simply referred to as PNCOL.
5.3.3 Restoring and preserving protein activity within PNCOL
5.3.3.1 Effect of heparin and BSA on protein bioactivity within PNCOL scaffolds
In order to restore the two-fold loss in growth factor bioactivity within PNCOL scaffolds and further preserve it for long-term applications, two biomolecules heparin and bovine serum albumin (BSA) were investigated.
To identify the solo and combined effects of heparin and BSA on the protein activity, BMP-
2 was again used as the model protein. The bioactivity estimated through quantification of the ALP synthesized by model cell line MC3T3-E1 encapsulated within the various
PNCOL scaffolds after ten days in culture. BMP-2 (GF) and heparin (h) were incorporated at a mass ratio of 1:40, and BMP-2 and BSA (b) were encapsulated at a mass ratio of 1:2000 within PNCOL scaffold. Figure 5-4 shows the changes in ALP activity with PNCOL scaffolds with heparin and/or BSA. Figure 5-4 demonstrates that the growth factor (GF) activity is protected best in the presence of both heparin and BSA, with over a 2-fold increase in ALP activity compared to GF-PNCOL (control group) (p < 0.05). GF-h- 157
PNCOL indicates that the solo effect of heparin in bioactivity preservation is not statistically different from control group.
Figure 5-4. Restoring and preserving the protein bioactivity within PNCOL scaffolds. Solo and combined effect of heparin (h) and BSA (b) on restoring the bioactivity model growth factor BMP-2 (GF) within PNCOL scaffolds evaluated by cellular ALP activity after ten days in culture. BMP-2 in presence of both heparin and BSA shows highest ALP activity within scaffold. [193]
BSA plays a dominant role in preserving the bioactivity of protein encapsulated within
PNCOL in comparison to heparin, with a 1.5-fold higher ALP activity with respect to the control group (p < 0.05), with no statistically significant difference seen with heparin alone.
However, for maximum protein bioactivity preservation, BSA and heparin need to be utilized together. Thus, with the introduction of BSA and heparin, the 2-fold loss in the protein activity within PNCOL scaffold was restored.
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5.3.3.2 Release kinetics of protein from GF-hb-PNCOL scaffolds
Since inclusion of heparin and BSA was demonstrated to be important in order to restore and sustain the protein bioactivity activity within PNCOL scaffolds, next we determined whether the presence of heparin and BSA affects the release kinetics of the growth factor.
This was investigated by obtaining the kinetic release profile of model protein BMP-2 from
GF-hb-PNCOL scaffolds quantified using ELISA as shown in Figure 5-5.
Figure 5-5. Protein release kinetics of GF-hb-PNCOL scaffolds. Amount of model growth factor BMP-2 (GF) released from GF-hb-PNCOL scaffolds over 21 days estimated through ELISA. GF-PNCOL samples used as control. The protein release profile obtained from the lysozyme study shown in Figure 1 is presented here for reference. The protein release profile from GF-hb-PNCOL similar to that observed with the lysozyme release profile. [193]
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GF-PNCOL scaffold without heparin and BSA were used as control samples. Also, the protein retention profile obtained using the model protein lysozyme for 3% (w/v) PNCOL was taken as a reference for the release profiles obtained from ELISA. The data presented in Figure 5-5 demonstrates that protein release profile from GF-hb-PNCOL was similar to that obtained from the lysozyme-based release study, with around 35% burst release in the first 48 hours followed by 5% release over the next 19 days. This establishes that GF-hb-
PNCOL scaffolds are able to exhibit controlled and sustained protein release over 21 days similar to the trend observed in Figure 5-2B.
The release profile of control GF-PNCOL samples without heparin and BSA, though show a 5% lower release in the burst phase when compared to GF-hb-PNCOL samples over the first 48 hours, exhibit very similar diffusion release profiles over the 21-day period. There is no statistical difference in the diffusion release properties of GF from GF-hb-PNCOL and GF-PNCOL due to the addition of heparin and BSA biomolecules into GF-PNCOL scaffold. This confirms that inclusion of heparin and BSA does not interfere with the long- term protein release kinetics of GF-hb-PNCOL scaffolds.
5.3.4 Long-term evaluation of GF-hb-PNCOL scaffolds
The long-term performance of GF-hb-PNCOL scaffolds was evaluated for 21 days through investigating the proliferation and ALP activity of model MC3T3-E1 cells encapsulated within GF-hb-PNCOL. BMP-2 was again used as the model growth factor. PNCOL 160
without growth factor was used as the control group. Both group of scaffolds were cultured in osteogenic media for 21 days.
5.3.4.1 Cell proliferation within GF-hb-PNCOL scaffolds
Cell proliferation of MC3T3-E1 within GF-hb-PNCOL and PNCOL were measured for 21 days using alamar Blue assay and displayed in Figure 5-6.
Figure 5-6. Long-term evaluation of cell proliferation within GF-hb-PNCOL. Cell proliferation within GF-hb-PNCOL and PNCOL scaffolds cultured in osteogenic media from day 0 to day 21 evaluated using alamar Blue assay. (*) indicates significant difference in comparison to PNCOL (control group) at the same time point. Cell proliferation shows no significant difference between the groups with and without the protein, confirming that GF-hb-PNCOL does not adversely affect the viability of cells. [193] 161
The cell proliferation data demonstrates that there is a sharp increase in cell numbers in the first 7 days and the cells continue to proliferate until day 14 within both GF-hb-PNCOL and control scaffolds. At day 14 and day 21, there is no statistical difference between the cell numbers obtained within GF-hb-PNCOL and PNCOL scaffolds (p>0.05). A dramatic decrease in the rate of proliferation is seen after day 14 within both scaffold groups due to possibly onset of terminal differentiation that would be confirmed with ALP activity obtained over 21 days. The data indicates that GF-hb-PNCOL and PNCOL scaffolds allow proliferation of MC3T3-E1 cells within their scaffold matrix.
5.3.4.2 Bioactivity of GF-hb-PNCOL scaffolds
The ALP expression profile over 21 days within GF-hb-PNCOL and PNCOL scaffolds is shown in Figure 5-7. It is observed that the ALP activity increased sharply throughout the
21 days within GF-hb-PNCOL, while there is no statistically different increase in ALP activity after day 14 in case of the control PNCOL scaffolds. Furthermore, at day 14 and day 21, cells within GF-hb-PNCOL had higher ALP activity compared to those within the control group (p < 0.05). Thus, the higher ALP activity exhibited by cells within GF-hb-
PNCOL scaffold proves that the growth factor activity within the scaffold is not only restored but also preserved and sustained over 21 days with the help of heparin and BSA,
Thus, PNCOL scaffolds having superior protein retention properties compared to collagen were synthesized. The bioactivity of the growth factor encapsulated within the scaffold was
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restored with the help of heparin and BSA and was proved to be active even during long- term cultures.
Figure 5-7. Long-term evaluation of protein bioactivity within GF-hb-PNCOL. BMP-2 bioactivity within GF-hb-PNCOL and PNCOL scaffolds cultured in osteogenic media from day 0 to day 21. (*) indicates significant difference in comparison to PNCOL (control group) at the same time point. (#) indicates significant differences as compared to earlier time points of the same group (p < 0.05). ALP activity exhibits significant increases at days 14 and 21 as compared to PNCOL scaffolds (control). [193]
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5.3.5 Performance of GF-hb-PNCOL under uniaxial tensile loading
After identifying the appropriate components for PNCOL for growth factor delivery, we investigated the performance of PNCOL scaffolds in presence of uniaxial tensile mechanical loading. The below set of results displaying the protein release kinetics and bioactivity of the released PDGF from GF-hb-PNCOL scaffolds were investigated under the uniaxial tensile loading regime of 2% strain and 0.1 Hz frequency for 2 hours/day identified in Chapter 4.
5.3.5.1 Protein release profile of mechanically-stimulated PNCOL scaffolds
The protein release kinetics was studied using model protein lysozyme was encapsulated in PNCOL scaffolds and subjected to aforementioned uniaxial tensile loading regime. In order to study the effect of uniaxial tensile loading on the burst and diffusion release phases individually, one set of scaffolds were loaded starting from day 0 (D0), coinciding with the burst release phase. Another set of scaffolds were loaded starting from day 3 (D3), coinciding with the diffusion release phase. Non-loaded scaffolds were used as the control group. The protein release profile obtained for the three sample sets over 15 days is displayed in Figure 5-8.
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Figure 5-8. Protein release from PNCOL scaffold under uniaxial tensile loading. Lysozyme release profile from PNCOL loaded at 2% strain and 0.1 Hz frequency for 2 hours/day over the 15-day period. Samples loaded from day 0 onwards to obtain the release profile due to uniaxial tensile loading during burst phase followed by the diffusion phase. Samples loaded from day 3 to generate the release profile due to uniaxial tensile loading during the diffusion phase alone. Non-loaded samples used as control. * indicates statistical significance between the loaded and non-loaded groups (p < 0.05), ** represents p < 0.01 and *** corresponds to p < 0.001. Uniaxial tensile loading during diffusion release alone induces controlled release of protein from PNCOL with around 0.5% of the encapsulated protein released per day.
Figure 5-8 demonstrates that the PNCOL scaffolds in presence of uniaxial tensile loading are still able to exhibit controlled and sustained protein release over the 15-day culture period. Though differences are observed in the protein release profiles between the loaded and non-loaded groups, significantly, there is no drastic leaching of the encapsulated 165
protein from PNCOL, indicating that the integrity of the scaffold matrix is not compromised due to mechanical loading.
The samples subjected to loading from D3 predictably have similar burst release profiles
(until 48 hours). During the diffusion release phase, the mechanically-loaded PNCOL samples exhibit increased slope of the diffusion curve, implying a higher rate of protein release when compared to the non-loaded PNCOL samples. The cumulative amount of protein released becomes statistically different between D3-loaded and non-loaded samples from day 10 onwards (p < 0.05). Further, it is observed that around 6.5% protein diffuses out of the scaffold over 12 days from the D3-loaded samples, compared to only
2.8% from the non-loaded scaffolds over the same time period.
The scaffolds loaded from D0 during the burst release phase display significantly higher protein burst release from the scaffold when compared to non-loaded scaffolds (p < 0.05).
The D0-loaded samples have 30 ± 2.2% of protein released from the PNCOL matrix, compared to just 19 ± 2.4% released from non-loaded scaffolds. Interestingly, the slope of the release curve exhibited by the D0-loaded scaffolds during the diffusion release phase is similar to that obtained for D3-loaded scaffolds, indicating that the release induced by mechanical loading in the burst phase does not influence the release during the diffusion phase. Thus, the results depicted in Figure 5-8 establish that mechanically-loaded PNCOL scaffolds are capable of controlled and sustained release of growth factor with an average protein release rate of 0.5% per day.
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5.3.5.2 Bioactivity of PDGF released from PDGF-hb-PNCOL scaffolds
Next, we evaluated whether the combination of heparin and BSA is able to preserve the bioactivity of the PDGF released from hb-PNCOL scaffolds, which would be important in vivo to stimulate the migration and proliferation of resident tenocytes at the repair site.
Model MC3T3-E1 cells that are known to migrate in presence of PDGF were used to conduct the migration and proliferation studies. Based on the protein release profile, the concentration of PDGF in the release media to be used on monolayer cells was estimated to be around 100 ng/ml. Hence, 100 ng/ml of fresh PDGF was used as the positive control, with no PDGF (0 ng/ml) as the negative control group.
Migration: The effect of the release media from PDGF-hb-PNCOL scaffolds on the migration of MC3T3-E1 cells is displayed in Figure 5-9. Figure 5-9 demonstrates that the release media obtained from mechanically-stimulated PDGF-hb-PNCOL scaffolds after 4 days of culture is able to induce migration of cells within 18 hours, indicating that the
PDGF has retained its bioactivity. Negligible cell migration is induced in cells cultured in media containing no PDGF (0 ng/ml), with the scratched area indicated by the red parallel lines having little or no cells. The positive control samples incubated in 100 ng/ml PDGF media exhibit significant cell migration by 6 hours, with around 30% of the scratched region covered by cells, which increases to approximately 70% at 16 hours. Cells maintained in the release media from PDGF-hb-PNCOL scaffolds show an onset of
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migration at 6 hours, with a few cells invading the scratched region. By 18 hours, the entire scratch region is fully covered with cells, reaching almost a 100% coverage of the scratched area. This demonstrates that PDGF released from the scaffold is bioactive and, like the positive control, is able to elicit a migratory response from the MC3T3-E1 cells.
Figure 5-9. Bioactivity of PDGF released from hb-PNCOL scaffolds evaluated by scratch migration assay. Release media collected after four days from mechanically-stimulated PNCOL scaffolds encapsulated with PDGF and added onto monolayer MC3T3-E1 cells. Migration of scratched cells evaluated at 0, 6, and 18 hours. Red parallel lines indicate scratch region. Scale bar is 50 µm. Release media from scaffold induces cell migration. 168
Cell proliferation: The effect of release media from PDGF-hb-PNCOL scaffolds on the proliferation of monolayer MC3T3-E1 cells after one day of culture was quantified by alamar Blue assay as shown in Figure 5-10.
Figure 5-10. Bioactivity of PDGF released from PNCOL scaffolds evaluated by cell proliferation assay. Release media collected after four days from mechanically-stimulated PNCOL scaffolds encapsulated with PDGF and added onto MC3T3 cells. Proliferative capacity of cells evaluated at day 1 using alamar Blue assay. * represents statistical difference from media containing no PDGF (p < 0.001). # denotes significant difference from samples cultured in PDGF-hb-PNCOL release media (p < 0.05). PDGF released from PNCOL scaffolds shows a 2-fold increase in proliferation compared to media without PDGF and is bioactive.
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It is observed that the release media is able to induce almost a 2-fold increase in the proliferation of MC3T3-E1 cells within a day of culture compared to media containing no
PDGF (0 ng/ml) (p < 0.001). Though positive media containing 100 ng/ml of PDGF shows statistically higher amount of proliferation than the release media (p < 0.05), the differences in the results can be attributed to not knowing the exact PDGF concentration in the release media.
Thus, Figure 5-9 and Figure 5-10 clearly establish that the growth factor within PDGF- hb-PNCOL is bioactive and, when secreted into the media, is able to stimulate migratory and proliferative responses from tendon fibroblasts-like cells.
5.4 Discussion
Collagen in its hydrogel form is very popular for tissue engineering applications as along with its high biocompatibility, bioactivity, and inherent biochemical properties; it is the primary component of the tendon ECM [17, 194]. Collagen-based hydrogel scaffolds are, however, limited by their inferior structural properties that renders them incapable of acting as a reservoir for growth factors in their native state. Hence, different techniques have been employed to chemically or physically immobilize the growth factor within the collagen matrices [176]. However, despite advancements in the field of growth factor delivery, two major limitations continue to persist: (1) high initial burst release of the growth factor; and
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(2) loss of protein bioactivity due to the chemical treatment undergone during the immobilization process [183]. Thus, there is a great need to engineer hydrogel-based biomaterials with proper biological, mechanical, and structural properties to ensure controlled release of growth factors without loss of protein bioactivity to accelerate tendon healing at the repair site. Ideally, the biomaterial should be capable of acting as a growth factor reservoir and exhibit controlled and sustained release at the implanted site in order to stimulate proliferation, migration, and infiltration of resident tissue cells during the healing phase. Further, the effect of mechanical loading should be considered to engineer physiologically relevant delivery systems.
In this study, we employed a mechanistic approach in synthesizing a growth factor (GF)- encapsulated composite collagen scaffold that is capable of increased protein retention with no significant loss in protein bioactivity and promotes the cell viability, proliferation, and function within its matrix. To achieve this goal, we created PCL-interspersed collagen scaffolds (PNCOL), which combined the superior mechanical properties of electrospun
PCL nanofibers with the outstanding biological properties of collagen to yield a structurally stable composite scaffold that can host both cells and growth factor within its 3D matrix.
The growth factor was immobilized to the surface of PCL nanofibers via plasma-induced functional groups before adding the fibers into the collagen matrix (Figure 5-1). PNCOL incorporated with 0%, 1%, 3%, and 6% (w/v) PCL nanofibers were evaluated in terms of
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their protein retention capacity and bioactivity to identify the appropriate PCL nanofiber concentration within the composite collagen scaffold.
The addition of PCL nanofibers into the collagen matrix demonstrated a remarkable effect in the protein retention capacity within the scaffold, with increased PCL nanofiber concentration exhibiting lower initial burst release and long-term retention of the protein
(Figure 5-2). Significantly, 3% (w/v) PNCOL and 6% (w/v) PNCOL retained 620.79% and 780.98 % of the protein within the scaffold, respectively over 21 days of culture. Pure collagen scaffolds, on the other hand, released the entire amount of encapsulated protein within 2 days in culture. The superior protein retention capacity of PNCOL scaffolds can be attributed to the nanofibrous internal structure of the scaffold due to electrospun PCL nanofibers combined with the functional groups present on the surface of PCL nanofibers.
Secondly, the oxygen-rich functional groups introduced on the PCL surface through plasma treatment [188, 191, 192] facilitates the binding of protein and PCL nanofibers, which is instrumental in preventing the initial burst release (Figure 5-2A) retaining the protein within the scaffold matrix over long-term cultures (Figure 5-2B). The carboxyl and hydroxyl functional groups being electron donors increase the negative charge on the
PCL surface [195]. Growth factors, on the other hand, are positively charged proteins with a free-end amine group (-NH2) in aqueous solutions [196, 197]. Thus, a strong electrostatic attraction between positively charged GF and negatively charged PCL nanofiber is achieved with the binding between (-NH2) of the growth factor and the oxygen-based
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functional groups on plasma-treated PCL nanofibers. Hence, incorporating of GF- immobilized PCL nanofibers within the collagen matrix shows ability to retain the protein within the composite PNCOL scaffold for the duration required corresponding to the healing phase of tendon (~ 2 months) [23].
Though increased PCL nanofiber concentration within the PNCOL scaffold exhibited higher protein retention, a reverse correlation was observed with respect to the bioactivity of GF encapsulated within PNCOL. Significant loss of protein activity was observed at higher concentrations of PCL within the scaffold, with 2-fold and 5-fold decreases in activity for 3% (w/v) and 6% (w/v) PNCOL scaffolds, respectively (Figure 5-3). This finding could be associated with the increased shear forces experienced by the GF- immobilized PCL nanofibers during PNCOL synthesis. With higher PCL concentrations, the collagen-PCL nanofiber scaffold solution was subjected to vigorous pipetting in order to ensure homogenous mixing of PCL nanofibers within collagen. This shear force applied on the immobilized protein could have possibly disrupted the protein structure that led to a reduction in its bioactivity.
Based on above findings (Figure 5-2 and Figure 5-3), 3% (w/v) PNCOL scaffold, with a
62% retention of protein within the scaffold over 21 days and only a 2-fold loss in bioactivity was identified as the appropriate PCL concentration desired for the tendon
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scaffold. Thus, PNCOL scaffolds, in all subsequent studies, were incorporated with 3%
(w/v) PCL nanofibers.
The 2-fold loss in the bioactivity of the GF within PNCOL incorporated with 3% (w/v)
PCL nanofibers was restored by introducing heparin and bovine serum albumin (BSA) biomolecules around the protein. On evaluating the solo and combinatorial effect of heparin and BSA on preservation of protein bioactivity, the results established that GF-hb-
PNCOL scaffolds, consisting of both heparin and BSA together, significantly minimized the loss of protein bioactivity (Figure 5-4). This finding can be linked to the unique properties of heparin and BSA, which play different roles in preserving the GF bioactivity.
BSA is widely used as a protective shell for growth factors to preserve their bioactivity in harsh physical and chemical environments [198, 199]. In our study, BSA is likely to provide a layer around the protein and shield it against possible residual solvents left on the PCL nanofibers [199, 200]. BSA also could protect BMP-2 from possible structural damage due to shear forces exerted on protein during incorporating PCL nanofibers within the collagen through pipetting by providing a cushion-like environment. Heparin, on the other hand, is known to bind to GFs possessing the heparin-binding site directly through electrostatic interactions between N-/O-sulfated residues of heparin and the lysine/arginine residues of the GF. This electrostatic interaction helps in retaining protein bioactivity by
(i) prolonging its half-life as much as 20-fold in vitro [201, 202], and (ii) decreasing GF localization to other possible binding sites [201, 203].
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Further, it was determined that the inclusion of heparin and BSA within the scaffold matrix did not significantly alter the protein release kinetics from GF-hb-PNCOL scaffolds. The release profiles estimated using ELISA demonstrate similar burst and diffusion release profiles for GF-hb-PNCOL scaffolds when compared to the lysozyme-based release profile
(Figure 5-5). The release profile from GF-PNCOL scaffolds does show a lower burst release when compared to GF-hb-PNCOL scaffolds. This is most likely due to the decrease in the accessibility to the binding sites present on the PCL nanofibers due to the protective shell provided by BSA, thus a decrease in the initial number of GF molecules immobilized onto the PCL nanofibers. Significantly, no differences in the rate of protein release from
PNCOL scaffolds with and without heparin and BSA is observed during diffusion phase, with the similar slopes obtained beyond 2 days of culture (Figure 5-5). This confirms that the binding of GF to the PCL surface via oxygen-rich functional groups is not compromised due to heparin and BSA. Thus, we have used heparin and BSA together to create GF-hb-
PNCOL scaffolds with high protein retention and preserved bioactivity.
The long-term in vitro performance of cell-encapsulated GF-hb-PNCOL scaffolds was evaluated by measuring the cell proliferation and protein bioactivity within the scaffolds at
Days 0, 7, 14, and 21. PNCOL (without GF) was used as the control group. Cell proliferation was evident within GF-hb-PNCOL and PNCOL scaffolds with a steady increase in cell number in first 14 days. The decline in the rate of proliferation after 14 days
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can be attributed to the onset of terminal cell differentiation (Figure 5-6). Initiation of differentiation was confirmed with ALP activity. ALP being an early differentiation marker for MC3T3-E1 cells, its activity starts to pick up once proliferation reduces and differentiation starts [204]. Significantly, the ALP activity data (Figure 5-7) demonstrated that cells within GF-hb-PNCOL scaffolds exhibited higher ALP activity in comparison to the control PNCOL scaffolds. This result proves that bioactivity of the protein within the scaffold is preserved over 21 days with the help of heparin and BSA. BSA is likely to have provided a cushion-like environment to protect the protein during scaffold synthesis, while heparin would have prevented premature degradation of the GF, thus prolonging its activity during long-term culture.
Next, the performance of GF-hb-PNCOL scaffolds were evaluated under uniaxial tensile loading conditions of 2% strain and 0.1 Hz for 2 hours/day, thereby simulating the native tendon environment at the repair site where the rotator cuff would be under controlled mobilization during physical rehabilitation. The scaffolds under uniaxial tensile loading are still able to exhibit controlled and sustained protein release over the culture duration, with no dramatic leaching of the protein observed due to mechanical stimulation. The release kinetics demonstrate a steeper slope for D0-loaded samples and D3-loaded samples during the diffusion release phase when compared to non-loaded scaffolds (Figure 5-8).
This is most likely due to the loading-induced widening of the pores within the scaffold matrix combined with the increased pressure on the scaffolds, which results in
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mechanoresponsive release of growth factor bound to PCL nanofibers along with passive diffusion normally observed in non-loaded conditions [176, 185]. This mechanoresponsive diffusion rate in the present system is estimated to be 0.5% per day from the loaded scaffolds, compared to the passive diffusion rate of 0.2% per day exhibited by non-loaded scaffolds. Hence, if 1 µg of growth factor is encapsulated within the scaffold and subjected to uniaxial tensile loading from day 3, it would be expected to release 5 ng/day into the culture media. After the initial burst release of 200 ng, the remaining growth factor within the scaffold could last for at least 3-4 months if the release profile obtained is extrapolated.
Thus, these mechanically-conditioned scaffolds are expected to exhibit controlled and sustained release of growth factor at physiological levels of dosage at the implantation site for tendon repairs.
Further, the bioactivity of PDGF released from PDGF-hb-PNCOL scaffolds was assessed by evaluating the ability of the release media to stimulate proliferation and migration of tenocyte-like cells. The monolayer scratch assay demonstrated almost a complete coverage of the scratch area with 18 hours of incubation in the release media, indicating that the released PDGF is bioactive (Figure 5-9). This was further confirmed by the statistically significant increase in cell proliferation in presence of release media after one day of culture
(Figure 5-10). The results together established that the combination of heparin and BSA is able to preserve the bioactivity of PDGF within PNCOL scaffolds. Thus, the PDGF-hb-
PNCOL scaffolds are capable of controlled release of bioactive PDGF over a long-term
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culture. Since PDGF is known to induce proliferation and ECM synthesis in in vivo tendon fibroblasts [40, 205] the delivery of bioactive PDGF through PDGF-hb-PNCOL scaffolds would be critical to augment and accelerate tendon healing at the repair site.
5.5 Conclusion
In Chapter 5, we have synthesized a composite scaffold PNCOL by interspersing homogenized PCL nanofibers within collagen matrix that exhibits high protein retention properties along with long-term preservation of its bioactivity making the scaffold suitable for growth factor delivery for tendon repair applications. The key results obtained from this chapter are highlighted in Figure 5-11. The higher surface area of PNCOL and functionalized groups on the PCL nanofibers contributed to minimized burst release and superior long-term protein retention capacity when compared to pure collagen. PCL nanofibers at a concentration of 3% (w/v) within the collagen matrix demonstrated desirable protein retention and bioactivity was identified. Though there was a 2-fold reduction in the protein bioactivity within PNCOL, it was restored using a mixture of heparin and BSA solutions. The preservation of the protein bioactivity within the scaffold was confirmed over 21 days of in vitro culture rendering it suitable for long-term studies.
Further, the GF-hb-PNCOL scaffolds exhibited controlled protein release profiles even under uniaxial tensile loading conditions, with a protein release rate of 0.5% per day. The
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released PDGF from the composite scaffolds retained its bioactivity and were able to induce migration and proliferation of tenocyte-like cells.
Figure 5-11. Major results obtained in Chapter 5. Composite collagen scaffolds (PNCOL) interspersed with polycaprolactone (PCL) nanofibers for controlled delivery of bioactive growth factors under mechanical loading conditions
PNCOL will be evaluated for its material properties in the Chapter 6 to create the biphasic composite scaffold encapsulated with ASCs and PDGF, and subjected to 2% at 0.1 Hz mechanical loading regime identified in the previous study and investigate its performance.
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Chapter 6
PDGF/ASC-encapsulated COLPNCOL Biphasic Scaffolds with Gradient Directionality, Mineralization, and Tenogenesis under Uniaxial Tensile Mechanical Loading
6.1 Introduction
The highly challenging nature of repairing tendon injuries is due to its role of connecting a soft tissue (muscle) to a hard tissue (bone) [69]. The area of the tendon insertion point in bone, called enthesis, has unique tissue organization and mechanical properties that enables tendon to effectively transfer mechanical forces from soft tissues to bone [206]. The defining characteristic of the interfacial tissue matrix is the gradual increase in mineral content and a gradual decrease in collagen alignment moving from tendon to bone. This helps to form a strong and tough attachment at the tendon-bone insertion point [207]. Most rotator cuff repairs post-surgery are seen to fail at the tendon-bone interface due to a mismatch in material properties attributed to scar tissue formation and the loss of mineral
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gradient during the healing process [208, 209]. Thus, understanding the physiology of the structural gradient at the enthesis is critical in order to devise scaffolds that can minimize risks of tendon re-rupture and increase the success rate of rotator cuff surgeries.
Majority of tendon-bone entheses are fibrocartilaginous, which can be divided into four zones: (1) tendon, (2) non-mineralized fibrocartilage, (3) mineralized fibrocartilage, and
(4) bone; each distinct in composition yet continuous in structure. It is well-recognized that the structural complexity of the tendon-to-bone attachment site during its formation is modulated by the synergistic effect of tendon-related growth factors, cells, and mechanical cues [21]. While the tendon is comprised of elongated tenocytes residing in a matrix of highly aligned collagen fibers, the fibrocartilage zone of the enthesis consists of fibrochondrocytes that are rounded in appearance, produce glycosaminoglycans (GAGs), have less matrix directionality when compared to tendons, and can secrete calcified deposits to mineralize the matrix [20, 21, 208-210]. Studies have also demonstrated the importance of mechanical loading on enthesis growth and maturation. In absence of mechanical loading, a reduction in mineral deposits, fiber organization, and matrix stiffness were observed [211-213]. However, the mechanisms underlying the structural and cellular gradient achieved in the native tendon-bone interface and the molecular mechanisms involved in enthesis healing are poorly understood. The exact role of the different growth factors and cells contributing to the unique mechanical gradient of the tissue are still being
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explored [21]. Hence, the current tissue engineering efforts for tendon-to-bone healing are mainly focused on recreating the structural gradation of the enthesis with the thought that promoting the gradual mineralization from tendon to bone would improve the tissue transition from soft to hard and minimize risk of re-tears.
In the past decade, researchers have designed some innovative stratified scaffolds using soft and hard biomaterials having different layers with gradual increase in mechanical properties similar to the enthesis tissue [70, 73-75, 214, 215]. These studies have also incorporated cells such as fibroblasts or ASCs, and/or growth factors such as BMP-2,
PDGF or TGF-β within the stratified scaffold for enhancing healing of tendon-bone interfaces. One such study was conducted by Spalazzi et al., who designed a continuous triphasic scaffold with phase I composed of aligned PLGA nanofibers seeded with fibroblasts (soft tissue), phase II made of PLGA microspheres seeded with chondroblasts
(fibrocartilage tissue), and phase III formed of PLGA/glass microspheres seeded with osteoblasts (hard tissue). It was observed that the increasing gradient of matrix stiffness of this triphasic scaffold influenced the ECM synthesis of the seeded cells. Collagen I, specific to the soft and hard tissues found in phase I and phase III, while Collagen II, specific to fibrocartilage secreted only in phase II [75]. Soo Kim et al. went a step further and made a scaffold with 4 phases, to mimic all four zones of the enthesis region: phase I (tendon) made of collagen, phase II (non-mineralized fibrocartilage) made of collagen crosslinked
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with chondroitin sulfate, phase III (mineralized fibrocartilage) composed of collagen and lower concentration of hyaluronic acid, and finally phase IV (bone) composed of collagen and higher concentration of hyaluronic acid. They observed that the increased material stiffness resulted in decreased elongation capacity of the four-phase scaffold [74]. Liu et al. created a PLGA nanofiber scaffold with an increasing mineral gradient from tendon to bone seeded with ASCs and demonstrated that the increased mineral gradient induced higher osteogenic differentiation of ASCs [214]. Similar technique was used by Eriksen et al. who incorporated an increasing gradient of tricalcium nanoparticles into electrospun
PCL nanofibers [73]. They too observed that MC3T3-E1 cells showed higher osteogenesis with increase in calcium content within the gradient scaffold.
Promising results have been obtained with in vitro evaluation of these scaffolds, with evidences that gradient material stiffness directs gradient mineralization, increased material stiffness leads to decreased scaffold elongation capacity, and GAGs secretion can be induced in phases experiencing mechanoactive compression. However, most of these scaffolds have not yet been evaluated in vivo, with the exception of Lipner et al., who created a PLGA stratified scaffold with BMP-2 and ASCs that was implanted into the rat rotator cuff after seeing positive results in in vitro culture. Surprisingly it was observed in that study that the scaffold did not help enhancing the in vivo healing process. Instead, it contributed to additional bone loss, inflammatory reactions and scar tissue formation [215].
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This emphasizes the challenge faced with devising tissue-engineered scaffolds for tendon- bone interfaces, due to the complexity of enthesis tissue compounded by lack of sufficient information about the biological mechanisms involved in enthesis healing. Also, the effect of mechanical loading on the performance of the various multi-phasic scaffold strategies still remain unexplored.
The objective of this chapter was to adopt a straightforward approach in simulating the tissue near the tendon insertion point by engineering a mechanically-conditioned
PDGF/ASC-encapsulated COLPNCOL biphasic scaffold, with COL mimicking the tendon region and PNCOL representative of the fibrocartilage region of the enthesis tissue.
Our hypothesis was that the PNCOL composite scaffold synthesized to improve the protein retention properties of collagen also increased the mechanical properties of collagen and that the increased matrix stiffness would correlate to higher cell-induced mineral deposition. Thus the combination of half COL and half PNCOL would provide a spatial variation in the stiffness of the scaffold matrix that mimics the tendon-bone interface.
PNCOL due to the nanofiber inclusions would be more resistant to matrix re-organization and should result in a matrix with low directionality when compared to highly compacted collagen fibers, thus simulating the fibrocartilage environment.
This aim was pursued with the vision that the resident fibrochondrocytes of the enthesis tissue would be recruited to the scaffold site by PDGF-induced chemotaxis along with
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ASCs within PNCOL are expected to mineralize the matrix in a gradient fashion, increasing from COL PNCOL that could increase the mechanical strength of the regenerating tendon at the tendon-bone insertion point. This would be one of the early steps towards mimicking a very complex tissue that is governed by a synergistic effect of different growth factors, cells, and mechanical loading.
To achieve this objective, the material properties and their effect on cell-induced mineralization for individual PNCOL and collagen scaffolds were characterized. The
COLPNCOL biphasic scaffold possessing gradient material stiffness was engineered, encapsulated with PDGF and ASCs, and subjected to uniaxial tensile loading. PDGF was incorporated into PNCOL scaffolds employing the technique devised in Chapter 5. The scaffolds were mechanically-conditioned using the custom-built uniaxial bioreactor developed in Chapter 3 at the mechanical loading regime of 2% strain and 0.1 Hz frequency for 2 hours/day identified in Chapter 4 for a total of 10 days. The synergistic effect of
PDGF and uniaxial tensile loading on ASCs differentiation response within COL and
PNCOL regions of the biphasic scaffold was evaluated. Also, the biphasic scaffolds were assessed for their ability to promote gradient increase in mineralization and gradient decrease in directionality of the matrix under uniaxial tensile mechanical loading conditions.
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6.2 Materials and Methods
6.2.1 Matrix stiffness of COL and PNCOL scaffolds
The material stiffness properties of COL and PNCOL scaffolds were characterized by estimating the elastic and viscous properties of the scaffolds. This was done by performing rheological analysis (n = 6) using a parallel plate rheometer (Rheometric Scientific).
Acellular COL scaffolds were prepared by neutralizing 3mg/ml Collagen Type-I (Corning,
US) with chilled 1 N NaOH along with PBS and sterile water. To obtain PNCOL scaffolds,
3% (w/v) PCL nanofibers were added into the neutralized collagen solution. Samples from both groups were polymerized at 37 oC in 35 mm Petri dishes to obtain cylindrical samples of 2 mm height and incubated in PBS. The samples were placed between the parallel plates of the rheometer made of nonporous metal. After lowering the top plate to a gap distance of 0.5 mm, frequency sweep test from 1 to 100 rad/sec was conducted at 20% strain under dynamic conditions. The data collected was analyzed using RDA software (TA instruments) to provide the storage modulus (G′) and loss modulus (G”) of the respective scaffolds as a function of frequency, at a constant temperature. Storage modulus corresponds to the elastic properties of the material while loss modulus is indicative of the viscous properties of the material. The combination of G’ and G” demonstrate the viscoelastic properties of COL and PNCOL scaffolds that is a direct estimate of their matrix stiffness.
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6.2.2 Mineralization potential of COL and PNCOL scaffolds
The ability of COL and PNCOL scaffolds to promote cell-induced matrix mineralization within the scaffolds was investigated using MC3T3-E1 cells capable of secreting calcium deposits in presence of osteogenic factors. MC3T3-E1 cells (ATCC, CRL-2593; US) were maintained in complete media consisting of Alpha-Minimum Essential Medium (α-MEM)
(Life Technologies, US) along with 10% FBS (Gibco, US) and 1% Penicillin-Streptomycin
(Life Technologies, US). COL scaffolds were synthesized by neutralizing 3mg/ml
Collagen Type-I (Corning, US) with cold 1 N NaOH along with PBS and MC3T3-E1 cell suspension at a seeding density of 1×106 cells/ml. To prepare PNCOL scaffolds, 3% (w/v)
PCL nanofibers were added into the neutralized MC3T3-E1 cell-encapsulated collagen solution. The cell-encapsulated COL and PNCOL scaffolds were polymerized at 37 oC and cultured for 21 days in osteogenic media comprising of 10 mM ß-glycerophosphate
(Sigma-Aldrich, US) and 50 μg/ml ascorbic acid (Sigma-Aldrich, US) added into complete media. The media was replaced every 3-4 days.
Alizarin red assay (Millipore, US) was used to quantify the amount of mineral deposits within the scaffold matrix at day 7 and 21 of culture. The samples were snap-frozen in liquid nitrogen and mechanically disrupted with a pestle in order to liberate the cells from the scaffolds. They were incubated in Alizarin red S solution for 20 minutes to allow the dye to bind to the calcium deposits. The samples were then centrifuged, the supernatant was discarded, and the pellet was resuspended in PBS. This was repeated 6X times in
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order to wash the scaffold and get rid of the unbound dye. Acetic acid at a concentration of
10% was added to each sample and incubated for 30 minutes to dissolve the bound dye.
The samples were centrifuged to precipitate the scaffold debris. Equal volumes of the supernatant from each tube were transferred to fresh microfuge tubes and heated at 85 oC for 10 minutes to dissolve the calcium deposits. The solution was cooled on ice for 2 min and was neutralized with 10% ammonium hydroxide to bring the pH up to 4.3. The neutralized solution was read at 405 nm wavelength using a microplate spectrophotometer
(SpectraMax). A standard curve was generated with different concentrations of Alizarin red solution and their corresponding absorbance value in order to quantify the amount of alizarin red dye bound to each sample.
6.2.3 Characterization of PDGF/ASC-encapsulated COLPNCOL
biphasic scaffold cultured under uniaxial tensile loading
6.2.3.1 Synthesis and culture of biphasic scaffold
The biphasic scaffold with varying material stiffness was engineered by employing COL and PNCOL biomaterials, with COL representing the soft tendon tissue, while PNCOL mimicking the fibrocartilage region of the enthesis tissue. The schematic of the components within the biphasic scaffold is displayed in Figure 6-1.
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Figure 6-1. Schematic of mechanically-conditioned PDGF/ASC-encapsulated COLPNCOL biphasic scaffolds. The COLPNCOL scaffold was encapsulated with ASCs at 1.5×106 cells/ml density, with 600 ng/ml of PDGF within PNCOL scaffolds. The samples were loaded using mechanical loading regime of 2% strain and 0.1 Hz frequency for 2 hours/day for 10 days.
Human ASCs (ThermoFisher Scientific, US) were cultured in MesenPRO RS™ basal media with MesenPRO RS™ growth supplement (ThermoFisher Scientific, US) and
200mM Glutamine (Sigma-Aldrich, US). COL scaffold solution was prepared with 3 mg/ml Collagen Type-I (Corning, US) encapsulated with ASCs at 750,000 cells/ml density. For preparation of PNCOL scaffold solution, PCL nanofibers at 3% (w/v) were incubated with 600ng/ml PDGF blended with heparin and BSA at 1:40:2000 ratio and incorporated into the neutralized 3mg/ml Collagen Type-I solution encapsulated with
ASCs at 750,000 cells/ml density. The scaffold solutions were deposited into the groove of the silicone loading chambers designed for the uniaxial tensile strain bioreactor such that COL fills one half of the groove and PNCOL occupies the other half. The PDGF/ASC- 189
encapsulated COLPNCOL scaffolds were polymerized at 37 oC for 1 hour to obtain a biphasic scaffold with seamless transition from COL to PNCOL regions. The scaffolds were cultured in ASC culture media supplemented with 1% Pencillin-Streptomycin solution (Gibco, US). After 48 hours, the biphasic scaffolds were subjected to uniaxial tensile loading at 2% strain and 0.1 Hz frequency for 2 hours/day over a 10-day period using the uniaxial tensile strain bioreactor, with the media was replaced every 2-3 days.
Non-loaded ASC-encapsulated COLPNCOL scaffolds were used as control samples.
The synergistic effect of uniaxial tensile loading, matrix stiffness, and PDGF on the matrix morphology and differentiation response of ASCs was investigated after the specified culture duration.
6.2.3.2 Matrix morphology of biphasic scaffold
The differences in the matrix morphology of COL and PNCOL regions of the loaded and non-loaded PDGF/ASC-encapsulated COLPNCOL biphasic scaffolds was examined using SEM. The biphasic scaffolds were fixed in 4% paraformaldehyde (Sigma-Aldrich,
US) overnight and subjected to sequential dehydration with 15-minute incubations in series of ethanol/water and hexamethyldisilazane/ethanol gradients with concentrations ranging from 30% to 100%. The samples were air-dried, sputter-coated with gold, and viewed under SEM (FEI Quanta 3D FEG) to observe the morphology and structure of the matrix within the COL and PNCOL region as well as their interface. The images were used to
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obtain directionality histograms using the Directionality plugin of Fiji/ImageJ (NIH, US)
[142, 143] in order to quantify the amount of directionality in each sample.
6.2.3.3 Gene expression profiles within biphasic scaffold
The differentiation response of ASCs in presence of PDGF within COL and PNCOL regions of loaded and non-loaded COLPNCOL biphasic scaffolds was investigated by looking at their gene expression profiles using qPCR. COL and PNCOL regions of the biphasic scaffold were excised and processed separately. RNA was extracted from the samples using TRIzol reagent (ThermoFisher Scientific, US) following the manufacturer’s instructions. The isolated RNA was transcribed into cDNA using the Omniscript cDNA synthesis kit (QIAGEN, US). qPCR was performed using SYBR Green Select Master mix
(Life Technologies, US) to detect the gene expression of extracellular matrix markers
Collagen I (COL-I), Collagen III (COL III), Decorin (DCN), and Aggrecan (ACAN); tenogenic markers Tenascin (TCN) and Scleraxis (SCX), osteogenic markers Alkaline
Phosphatase (ALP); and chondrogenic marker SOX9 within COL and PNCOL regions.
Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was the housekeeping gene used to normalize the expression of each gene of interest. The iCycler iQ detection system
(Biorad) was used to run all qPCR analysis with thermocycling carried out for 40 cycles.
Data analysis was performed using ΔΔCt method. The fold differences in gene expression for COL and PNCOL regions of loaded samples were estimated in comparison to the COL and PNCOL regions of the non-loaded control samples, respectively.
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6.2.3.4 Secreted extracellular matrix proteins from biphasic scaffold
The amount of extracellular matrix proteins secreted by the PDGF/ASC-encapsulated
COLPNCOL biphasic scaffold was quantified by analyzing the cumulative culture media for collagen and GAG expression after ten days of culture.
Quantification of secreted collagen was performed using SircolTM Collagen assay
(Biocolors, UK) according to the manufacturer’s protocol. The culture media from each sample was mixed with 10X amount of Sircol dye reagent and incubated on a shaker at room temperature for 30 minutes to allow the dye to bind to soluble collagen. The samples were centrifuged, and the supernatant was discarded. The pellet was rinsed with acid-salt wash buffer to eliminate any unbound dye and then resuspended in alkali reagent. The samples were read at 555 nm in the microplate spectrophotometer (SpectraMax). The amount of collagen was estimated by using a standard curve generated by assaying known collagen amounts and determining their respective absorbance readings.
Quantification of secreted GAGs was conducted using Dimethylmethylene (DMMB) Blue
Assay kit (Amsbio, UK) by following the manufacturer’s protocol. The culture media was mixed with 2X DMMB dye reagent and incubated on a shaker at room temperature for 30 minutes to allow the dye to bind to sulfated GAGs. The samples were centrifuged, and the supernatant was discarded. The pellet was rinsed with an acid-salt wash solution in order to remove any unbound dye from the pellet. The samples were then resuspended in 10% sodium dodecyl sulfate (SDS) and assayed at 656 nm wavelength using the microplate
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spectrophotometer (SpectraMax). A standard curve was generated by determining the absorbance of known amounts of chondroitin sulfate in order to estimate the amount of
GAG in the culture media obtained from each sample.
The amount of collagen and GAGs determined from the assay were both normalized with the total protein present in the respective culture media. The total protein was estimated by reading the absorbance of the culture media at 280 nm and equating it with a standard curve obtained by using different concentrations of BSA in fresh culture media.
6.2.4 PDGF-induced migration and infiltration of fibrocartilage-like
cells into COLPNCOL biphasic scaffolds
The ability of PDGF to induce migration and subsequent invasion of fibrocartilage-like cells into PDGF-encapsulated PNCOL scaffold was evaluated by conducting an invasion study using biphasic scaffold made of PNCOL in one half and Collagen in the other half.
The schematic of the experimental design is given in Figure 6-2. PNCOL region of the scaffold was encapsulated with 1 µg/ml PDGF along with heparin and BSA in the pre- determined ratio to obtain PDGF-hb-PNCOL. MC3T3-E1 cells that are known to undergo chemotaxis in response to PDGF were encapsulated in Collagen region of the scaffold at
750,000 cells/ml seeding density to obtain MC3T3-Collagen scaffold. The scaffolds were
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cultured under mechanical stimulation at 2% strain and 0.1 Hz frequency for 2 hours/ day over a 7-day period.
Figure 6-2. Schematic of the experimental design for PDGF-induced cell migration and infiltration within COLPNCOL biphasic scaffolds. Biphasic scaffold composed of Acellular PDGF-hb-PNCOL and MC3T3-encapsulated COL loaded for 7 days at 2% strain and 0.1 Hz frequency for 2 hours/day. Scaffolds with Acellular hb-PNCOL with MC3T3-COL used as control samples. Cell migration from COL to PNCOL assayed through DNA quantification of COL and PNCOL regions.
Control biphasic scaffolds consisted of Collagen encapsulated with MC3T3-E1 cells, but in absence of PDGF within PNCOL. Thus, the initial cell density within Collagen region of the scaffold was to be 750,000 cells/ml, while PNCOL region has no cells. After seven days of loading, the PNCOL and Collagen regions of the scaffolds we carefully excised and separated. The total number of cells in each region was indirectly determined through
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PicoGreen dsDNA quantification assay (Life Technologies, US) following the manufacturer’s protocol in order to determine whether PDGF was able to induce MC3T3-
E1 cell invasion into PNCOL.
6.2.5 Gradient mineralization within COLPNCOL biphasic scaffolds
cultured under uniaxial tensile loading
The schematic of the experimental design employed to study gradient mineralization within
COLPNCOL scaffolds is displayed in Figure 6-3.
Figure 6-3. Schematic of the experimental design to determine gradient matrix mineralization with COLPNCOL biphasic scaffolds. MC3T3-E1-encapsulated COLPNCOL biphasic scaffolds cultured in osteogenic (differentiation) media for 21 days. Biphasic scaffolds cultured in complete (growth) media used as control samples. All scaffolds loaded at 2% strain and 0.1 Hz frequency for 2 hours/day.
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COL and PNCOL scaffolds solutions were encapsulated with a total of 1×106 cells/ml
MC3T3-E1 cells and deposited into the groove of the loading chamber. MC3T3-E1 cells are chosen to mimic fibrocartilage cells that are capable of mineralization of the matrix.
The polymerized COLPNCOL biphasic scaffolds were cultured in osteogenic media for
21 days under mechanical stimulation at 2% strain and 0.1 Hz frequency for 2 hours/day.
Biphasic scaffolds cultured in complete media without osteogenic factors were used as control samples. At the end of 21 days, the samples were harvested to visualize and quantify the matrix mineralization potential of MC3T3-E1 cells within COL and PNCOL regions of the biphasic scaffold.
6.2.5.1 ALP activity within COL and PNCOL regions of biphasic scaffolds
ALP activity in MC3T3-E1 cells acts as a precursor for onset of mineralization. Thus, measuring the level of ALP activity in the samples is an indirect indicator of the ability of
MC3T3-E1 cells within the COL and PNCOL regions of the biphasic scaffold to mineralize the matrix. ALP activity was measured using the Alkaline Phosphatase kit (Sigma-Aldrich,
US). COL and PNCOL regions of the biphasic scaffold were carefully excised and assayed separately. Frozen scaffolds were mechanically disrupted and resuspended in an alkaline lysis buffer (10 mM Tris-HCl, 2 mM MgCl2, 0.1% Triton X-100, pH 8), and lysed with a homogenizer (Ultra Turrax). The samples and the pNPP substrate provided in the kit was mixed at 1:1 ratio in a microwell plate, incubated at 37 oC for 30 minutes and the reaction was stopped using 3N NaOH. The product formed from each sample was measured at
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405 nm using a microplate spectrophotometer (SpectraMax). Total ALP was quantified using a standard curve obtained by determining the optical density readings of different dilutions of calf alkaline phosphatase enzyme. The total ALP was then normalized with the total protein content in each corresponding sample using Coomassie Bradford protein assay kit (Thermo Scientific, US) following the manufacturer’s protocol.
6.2.5.2 Mineralization within COL and PNCOL regions of biphasic scaffolds
Mineral mapping: The calcium deposits in the COL and PNCOL regions of the biphasic scaffold was visualized and mapped using Energy Dispersive X-ray Spectroscopy (EDS) through SEM. Samples were prepared for SEM using the protocol described in the previous section. Briefly, the COLPNCOL biphasic scaffolds were fixed, dehydrated, air-dried, sputter-coated, and analyzed through SEM-EDS (FEI Quanta 3D FEG) in the analytical mode with 20 KV excitation energy. The calcium deposits were mapped, and a line spectrum of the various elements present in the visualized region was obtained through the
EDS interface.
Alizarin red assay: The amount of mineralized deposits secreted by MC3T3-E1 cells within COL and PNCOL regions of the biphasic scaffold was estimated using Alizarin red assay (Millipore, US). COL and PNCOL regions of the biphasic scaffold were carefully excised, and the quantification of mineral deposits for each region was done individually following the protocol described in the previous section. After obtaining the spectrophotometric readings for the total bound alizarin red dye which is has a direct 197
correlation with amount of cell-secreted calcium deposits, they were normalized with the total protein content in the respective regions of the biphasic scaffold.
6.2.6 Statistical analyses
Four samples (n=4) were used in each group for all assays. Statistical analysis was conducted using Student’s t-test and One-way ANOVA (IBM SPSS Statistics software).
All values are reported as the average, and the error bars correspond to ± the standard deviation of the mean. All statistical difference was determined with a 95% confidence interval (p < 0.05).
6.3 Results
6.3.1 Mechanical properties of Collagen and PNCOL scaffolds
6.3.1.1 Material stiffness of COL and PNCOL scaffolds
The material properties of PNCOL and Collagen (COL) scaffolds were evaluated by performing rheological analysis using a parallel plate rheometer (Rheometric Scientific).
The data was recorded as storage modulus (G’) and loss modulus (G’’). The data obtained from rheology for COL and PNCOL as a function of frequency is displayed in Figure 6-4.
The storage modulus G’ describes the solid behavior of the material while the bulk modulus
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G” corresponds to the liquid properties of the material. Figure 6-4 demonstrates that the storage (elastic) modulus significantly increases with incorporation of PCL nanofibers within the scaffold. A 20-fold rise in G’, which is an estimate of the mechanical stiffness of a material, is observed for PNCOL when compared to pure collagen scaffolds and reached to around 2000 Pa.
Figure 6-4. Effect of PCL nanofibers on material stiffness of PNCOL. Rheological analysis for storage/elastic modulus (G′) and loss/viscous modulus (G″) as a function of frequency for PNCOL and COL (collagen) scaffolds. (*) indicates significant difference in comparison to COL at the same time point. The viscoelastic properties and matrix stiffness of collagen dramatically increase with the incorporation of PCL nanofibers in its matrix. [193]
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Figure 6-4 also indicates that there is a significant rise in the loss (viscoelastic) modulus with inclusion of PCL nanofibers within the scaffold. For collagen scaffolds, the viscoelastic modulus is around 10 Pa which indicates that they are much softer material when compared to PNCOL scaffolds, with a value of 1000 Pa. Thus, inclusion of PCL nanofibers into collagen matrix has helped in dramatically increasing the matrix stiffness of PNCOL when compared to pure collagen (COL) scaffolds.
6.3.1.2 Mineralization potential of Collagen and PNCOL scaffolds
The effect of matrix stiffness on the mineralization potential of COL and PNCOL scaffolds were studied by encapsulating model MC3T3-E1 cells within the scaffolds that can be induced to secrete mineral deposits on stimulation with osteogenic factors. The secreted minerals were quantified using Alizarin Red S solution that stains the calcium deposits as shown in Figure 6-5. No mineralization is seen at Day 7 during early stage of differentiation for either samples. Day 14 indicates the onset of calcification of the matrix within PNCOL, while COL scaffolds still do not exhibit any significant presence of mineralized deposits. Statistically higher values of calcified mineral deposits are found in
PNCOL scaffolds in comparison to COL at both Day 14 and Day 21, with a 2-fold difference at Day 14 and almost a 2-fold difference at Day 21 (p < 0.05). These results establish that the increased in stiffness of PNCOL scaffold due to incorporation of PCL nanofibers not only accelerate the onset of cell-induced mineralization but also stimulate
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the cells to secrete significantly higher amounts of mineral deposits when compared to
COL scaffolds possessing poor stiffness properties.
The material stiffness and matrix mineralization data together confirm that PNCOL possesses significantly higher mechanical stability and stiffness compared to COL, and varying the matrix stiffness of a scaffold can induce different levels of mineralization within the scaffold using the same cell type and culture conditions.
Figure 6-5. Effect of PCL nanofibers on matrix mineralization of PNCOL. Alizarin red quantification of mineralized deposits within PNCOL and COL scaffolds cultured in osteogenic media for 21 days. (*) indicates significant difference in comparison to COL at the same time point. PNCOL scaffold exhibits significant increases of mineralized deposits at day 21 in comparison to COL scaffold. [193]
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Based on these results, the COLPNCOL biphasic scaffold with spatially distributed matrix stiffness was engineered, with the goal to guide the maturation of the de novo tissue at the tendon-bone insertion point of the rotator cuff. COL was expected to mimic the soft tendon tissue while PNCOL was representative of the fibrocartilage region of the enthesis.
The performance of the COLPNCOL biphasic scaffold was evaluated through two broad characterization studies, namely (1) investigating the matrix directionality and tenogenic response of ASCs within the biphasic scaffold; and (2) examining the gradient mineralization potential of the biphasic scaffold.
6.3.2 ASC differentiation and matrix alignment within mechanically-
conditioned COLPNCOL biphasic scaffolds
The first study focused on investigating the synergistic effect of PDGF, matrix stiffness, and uniaxial tensile loading on the matrix alignment and tenogenic response of ASCs encapsulated within the COL and PNCOL regions of the PDGF/ASC-encapsulated
COLPNCOL biphasic scaffold. The samples were loaded using mechanical loading regime of 2% strain and 0.1 Hz frequency for 2 hours/day for 10 days. The COLPNCOL scaffold was encapsulated with ASCs at 1.5×106 cells/ml density, with 600 ng/ml of PDGF within PNCOL scaffolds. The matrix alignment, gene expression profiles, and secreted
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extracellular matrix protein were measured at the end of ten days of loading. Non-loaded
COLPNCOL scaffolds were used as control samples.
6.3.2.1 Matrix morphology of COLPNCOL biphasic scaffolds
The morphology of the biphasic scaffold was visualized using SEM to identify the response of COL and PNCOL regions of the matrix to uniaxial tensile loading in presence of ASCs.
Figure 6-6A and Figure 6-6B displays the structure of the loaded COLPNCOL biphasic scaffold at the interface of COL and PNCOL regions at low and high magnifications.
There are distinct differences in the matrix morphology observed between the two regions that make it very easy to identify the interface of COLPNCOL scaffold. COL scaffold appears to have high directionality in matrix with the fibers aligned parallel to each other.
Further, the scaffold looks to have undergone ASCs-mediated compaction producing thicker collagen fibers with decrease in the width of the COL region. The PNCOL region, on the other hand, exhibits no specific matrix organization, has thinner fibers when compared to COL scaffold, and retains its original structure.
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Figure 6-6. Morphology at the interface of mechanically-conditioned COLPNCOL biphasic scaffold. (A) Lower magnification, and (B) High magnification SEM micrographs of COLPNCOL scaffold interface at encapsulated with ASCs and PDGF, and mechanically stimulated at 2% strain, 0.1 Hz frequency for 2 hours/day over a period of 10 days. Scale bar represents 1mm. Clear distinction in morphology is visible between COL and PNCOL. 204
To investigate the morphological differences in the biphasic scaffold in detail, SEM images for COL and PNCOL regions were taken individually for both loaded and non-loaded scaffolds. The images were also used to estimate the amount of directionality of exhibited by the matrix in the respective regions. The SEM micrographs of the scaffolds and their respective histogram is presented in Figure 6-7. No specific organization is observed within the matrix of both COL and PNCOL regions of the non-loaded biphasic scaffold. The corresponding histograms support the observation with an amount of directionality of
0.015 in both cases, with no prominent height in the peak obtained. The matrix organization of loaded scaffolds, on the other hand, show remarkable differences between the COL and
PNCOL regions, similar to that seen in Figure 6-6 at the COLPNCOL interface. COL region of the scaffold is highly organized, with the matrix exhibiting aligned fibers parallel to the direction of applied load. This is confirmed by the corresponding histogram that depicts a clear peak with a magnitude of 0.03 in directionality.
Matrix compaction is evident within the COL region of loaded biphasic scaffold, due to which the collagen fibers obtained are visibly much thicker when compared to the non- loaded COL region. Loaded PNCOL region of the scaffold, on the other hand, exhibit a very low matrix organization despite the presence of uniaxial tensile loading. Though some thickening of fibers within the scaffold is observed, unlike the COL region, they do not orient themselves in any specific pattern.
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Figure 6-7. Matrix alignment of mechanically-loaded PDGF/ASC-encapsulated COLPNCOL biphasic scaffold. Representative SEM Micrographs and their respective directionality histograms of COL and PNCOL regions of the biphasic scaffold encapsulated with ASCs and PDGF and mechanically stimulated at 2% strain, 0.1 Hz frequency for 2 hours/day over a period of 10 days. Non-loaded COLPNCOL scaffolds used as control. Scale bar represent 100 µm. Loaded COL region appears to be more aligned with compacted fibers, with a 2-fold rise seen in the amount of directionality from the histograms. Loaded PNCOL region shows little directionality. 206
This lack of directionality within the loaded PNCOL region is further confirmed by the histogram with an amount of directionality of 0.017 which is very similar to the directionality of non-loaded PNCOL sample at 0.015 (Figure 6-7). The results demonstrate that uniaxial tensile loading in presence of ASCs elicits very different responses from COL and PNCOL scaffolds in terms of matrix structure and organization, with a highly-aligned
COL matrix while negligible directionality within PNCOL.
Further, the histograms obtained were consolidated into one graph to compare the directionality of COL and PNCOL regions of loaded and non-loaded scaffolds to determine whether the differences in matrix organization visualized are statistically relevant. This data is shown in Figure 6-8. It is evident from Figure 6-8 that the COL region of the loaded biphasic scaffolds has significantly higher matrix directionality, with at least at 2-fold increase the magnitude of directionality when compared to the non-loaded COL region (p
< 0.05). Significantly, within the loaded COLPNCOL biphasic scaffold, the COL region exhibits a 2-fold higher directionality when compared to the PNCOL (p < 0.05). On the other hand, no statistical difference between the matrix directionality of the loaded and non-loaded PNCOL regions is obtained.
These results confirm that uniaxial tensile loading in presence of ASCs plays a significant role in directing matrix alignment within COL scaffolds, while it does not significantly alter the structure of PNCOL scaffolds.
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Figure 6-8. Amount of matrix directionality in mechanically-loaded PDGF/ASC- encapsulated COLPNCOL biphasic scaffold. Quantified directionality of COL and PNCOL regions of the biphasic scaffold encapsulated with ASCs and PDGF and mechanically stimulated at 2% strain, 0.1 Hz frequency for 2 hours/day over a period of 10 days. Non-loaded COLPNCOL scaffolds used as control. Graph derived from consolidating the directionality histograms of SEM images obtained using ImageJ analysis shown in Figure 6-7. * represents statistical difference from the non- loaded group of the same region. # denotes statistical significance between COL and PNCOL at the same culture condition (p < 0.05). COL region exhibits significantly higher directionality of matrix when compared to PNCOL region of the loaded biphasic scaffold.
6.3.2.2 ECM gene expression profiles within COLPNCOL biphasic scaffold
The expression levels of ECM markers including Collagen I, Collagen III, and GAGs
(Decorin & Aggrecan) of ASCs within COL and PNCOL regions of loaded and non-loaded
PDGF/ASC encapsulated COLPNCOL biphasic scaffolds is shown in Figure 6-9.
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Loaded COL region of the scaffold shows significant increases in all four ECM markers, with a 4-fold and 8-fold rise in Collagen I and Collagen III expression, and 11-fold and 7- fold increase in Decorin and Aggrecan, respectively, when compared to COL region of the non-loaded scaffold (p < 0.05). The loaded PNCOL region exhibits higher expression levels of Collagen III, Decorin, and Aggrecan with increases of 4-fold, 11-fold, and 5-fold, respectively, compared to the non-loaded PNCOL region (p < 0.05). Significantly, within the loaded COLPNCOL biphasic scaffold, only COL region shows statistical increase in
Collagen I expression, while the expression in PNCOL is similar to the non-loaded samples
(p < 0.05).
6.3.2.3 Lineage-specific gene expression within COLPNCOL biphasic scaffold
The expression of tenogenic, osteogenic, and chondrogenic markers within COL and
PNCOL regions of loaded and non-loaded PDGF/ASC-encapsulated COLPNCOL biphasic scaffolds is displayed in Figure 6-10. Prominent increases in the expression of tenogenic markers Tenascin and Scleraxis is observed in COL region of the loaded scaffolds. There is an 8-fold rise in Tenascin and a 13-fold higher expression in Scleraxis when compared to the COL region of the non-loaded biphasic scaffolds (p < 0.05).
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Figure 6-9. ECM gene expression within mechanically-loaded PDGF/ASC- encapsulated COLPNCOL biphasic scaffold. Gene expression profiles of ECM markers Collagen I, Collagen III, and GAGs: Decorin and Aggrecan within COL and PNCOL regions of the biphasic scaffold encapsulated with ASCs and PDGF, and mechanically stimulated at 2% strain, 0.1 Hz frequency for 2 hours/day over a period of 10 days. Non-loaded COLPNCOL scaffolds used as control. * represents statistical difference between loaded and non-loaded groups for the same region. Both COL and PNCOL regions show significant increases of ECM markers. 210
Figure 6-10. Musculoskeletal differentiation marker expression within mechanically- loaded PDGF/ASC-encapsulated COLPNCOL biphasic scaffold. Gene expression profiles of Tenogenic markers: Tenascin (TCN) and Scleraxis (SCX), Osteogenic marker (ALP), and Chondrogenic marker (SOX9) within COL and PNCOL regions of the biphasic scaffold encapsulated with ASCs and PDGF, and mechanically stimulated at 2% strain, 0.1 Hz frequency for 2 hours/day over a period of 10 days. Non- loaded COLPNCOL scaffolds used as control. * represents statistical difference between loaded and non-loaded groups for the same region (p < 0.05). # represents significant difference between COL and PNCOL at the same culture condition (p < 0.05). COL region shows significant increases of tenogenic markers. 211
Interestingly, no significant differences in the expression of tenogenic markers is observed between the PNCOL region of loaded and non-loaded biphasic scaffolds (Figure 6-10).
This implies that ASCs within COL region of the biphasic scaffold, along with increased expression of Collagens and GAGs, are undergoing tenogenic differentiation. ASCs within
PNCOL scaffold though have enhanced Collagen III and GAGs expression, do not exhibit any potential of going down the tenogenic lineage.
Looking at the osteogenic marker ALP, no statistically different expression is obtained for the loaded and non-loaded COLPNCOL scaffold, indicating no osteogenic differentiation potential in either COL or PNCOL regions of the scaffold. Interestingly, the chondrogenic marker SOX9 shows a significant increase in expression within the PNCOL region of the loaded biphasic scaffold, with around 3.5-fold rise when compared to PNCOL region of the non-loaded scaffold (p < 0.05). No statistical difference is obtained in the
SOX9 expression between COL region of the loaded and non-loaded biphasic scaffolds.
This indicates that ASCs within the PNCOL region of the loaded scaffold have the potential to differentiate into chondrocytes.
Further, comparison of the different gene expression ratio of COL to PNCOL regions of the loaded and non-loaded scaffolds is given in Figure 6-11.
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Figure 6-11. Ratio of COL to PNCOL gene expression within COLPNCOL biphasic scaffolds. COL to PNCOL gene expression ratio of Collagens (Collagen I and III), GAGs (Decorin and Aggrecan), Tenogenic markers (Tenascin and Scleraxis), Chondrogenic marker (SOX9), and Osteogenic marker (ALP) within the biphasic scaffold encapsulated with ASCs and PDGF, and mechanically stimulated at 2% strain, 0.1 Hz frequency for 2 hours/day over a period of 10 days. Non-loaded COLPNCOL scaffolds used as control. * represents statistical difference between loaded and non-loaded groups for the same region (p < 0.05). Mechanically loaded COL region shows significant increases in collagen and tenogenic gene expression compared to PNCOL region.
It is observed that there is 2-fold increase in Collagen gene (Collagen I and Collagen III) expression within COL versus PNCOL regions between loaded and non-loaded scaffolds
(p < 0.05). Interestingly, the GAG (Decorin and Aggrecan) expression in COL versus
PNCOL regions is similar between loaded and non-loaded COLPNCOL scaffolds. There is a dramatic 6-fold rise in the expression of tenogenic markers (Tenascin and Scleraxis)
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within the loaded COL versus PNCOL regions when compared to non-loaded scaffolds (p
< 0.05). No significant differences are seen in the expression levels of osteogenic markers
ALP and chondrogenic marker SOX-9 between COL to PNCOL ratio for loaded and non- loaded biphasic scaffolds. However, a decreasing trend is observed in chondrogenic expression of COL to PNCOL ratio of the loaded scaffolds when compared to non-loaded scaffolds, indicating a potential of ASCs to undergo chondrogenesis within the loaded
PNCOL region.
6.3.2.4 Secreted extracellular matrix proteins from the biphasic scaffold
The amount of Collagens and GAGs secreted by the loaded and non-loaded PDGF/ASC- encapsulated COLPNCOL scaffolds after 10 days of culture was estimated by analyzing the culture media through colorimetric assays and normalized with the total protein content as depicted in Figure 6-12. Two-fold rises in both GAG and Collagen secretion is observed from the loaded COLPNCOL scaffolds when compared to non-loaded samples (p <
0.05). This demonstrates that the increase in gene expression of ECM markers of ASCs encapsulated within COLPNCOL biphasic scaffolds obtained through qPCR is translated to increase in protein expression.
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Figure 6-12. Secreted extracellular matrix proteins from COLPNCOL biphasic scaffold. Amount of GAG and Collagen secreted by the ASCs encapsulated within COLPNCOL scaffold mechanically stimulated at 2% strain, 0.1 Hz frequency for 2 hours/day over a period of 10 days estimated through quantitative DMMB and Sircol dye assays, respectively. Non-loaded COLPNCOL scaffolds used as control. * indicates significant difference between non-loaded and loaded groups of the same assay (p < 0.05). There is a 2-fold rise in secreted GAG and Collagen from the loaded COLPNCOL biphasic scaffold compared to non-loaded control.
6.3.3 PDGF-induced migration and infiltration of fibrocartilage-like
cells into COLPNCOL biphasic scaffolds
We checked the potential of PDGF within PDGF-hb-PNCOL scaffolds to act as a chemoattractant in order to recruit native fibrocartilage cells to migrate and infiltrate the
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PNCOL scaffold when implanted at the site of injury. This was conducted by performing a cell invasion assay with a biphasic scaffold consisting of the acellular PDGF-hb-PNCOL construct in one half and MC3T3-E1-encapsulated Collagen constructs in the other half.
The migration and subsequent infiltration of cells from collagen to PNCOL was evaluated after 7 days of uniaxial tensile loading through DNA quantification within PNCOL and
Collagen scaffolds as displayed in
Figure 6-13.
Figure 6-13. PDGF-induced migration and infiltration of fibrocartilage-like cells into COLPNCOL biphasic scaffolds. Biphasic scaffolds composed of acellular PDGF-hb-PNCOL on one side and MC3T3-E1- encapsulated Collagen on the other side loaded for 7 days at 2% strain and 0.1 Hz frequency for 2 hours/day. Scaffolds with acellular PNCOL with MC3T3-Collagen used as control samples. Cell migration from Collagen to PNCOL assayed through DNA quantification of (A) PNCOL and (B) Collagen regions. Red dotted line depicts the initial seeding density
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within the respective region of the scaffold. * represents a statistical difference from the other group (p < 0.05). PDGF exhibits a 2.5-fold increase in cell migration from Collagen to PNCOL region and is able to act as a chemoattractant for fibrocartilage-like cells.
Figure 6-13A demonstrates the number of cells within PDGF-hb-PNCOL (1 µg/ml PDGF) and PNCOL (control) scaffolds (0 µg/ml PDGF) after 7 days of culture with MC3T3-E1-
Collagen scaffolds. PNCOL scaffolds that were acellular on Day 0 show significant number of cells in their matrix in both PDGF-hb-PNCOL and PNCOL groups, with
100,000 cells within PDGF-hb-PNCOL and 40,000 cells within PNCOL. This indicates that PNCOL scaffold has the appropriate porosity to allow cell infiltration and subsequent cell proliferation. Significantly, the number of cells migrated into PDGF-hb-PNCOL is statistically higher by 2.5-fold when compared to cells within the PNCOL (p < 0.05).
This is further confirmed by
Figure 6-13B which depicts the number of cells present in the Collagen region of the respective PDGF-hb-PNCOL and PNCOL control scaffolds after 7 days of culture. There is a significantly lower number of cells in the Collagen region of scaffolds having GF-hb-
PNCOL in its adjacent region in comparison to Collagen scaffolds fused with PNCOL control scaffolds. This suggests that migration of cells from Collagen to PDGF-hb-PNCOL is likely to have contributed to the reduced number of cells in the Collagen region.
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6.3.4 Estimation of mineralization gradient within COLPNCOL
biphasic scaffolds cultured under uniaxial tensile loading
The ability of fibrocartilage-like cells to migrate and infiltrate PDGF-encapsulated PNCOL scaffolds was demonstrated in Chapter 5. Now, we examine the ability the infiltrated cells to undergo gradient mineralization within COLPNCOL scaffolds due to its gradient material stiffness properties. This was performed by encapsulating model MC3T3-E1 cells into the COLPNCOL biphasic scaffold and cultured in osteogenic (differentiation) media under uniaxial tensile loading conditions of 2% strain and 0.1 Hz frequency for 2 hours/day. Biphasic scaffolds cultured in complete (growth) media were used as control samples. The COL and PNCOL regions of the biphasic scaffolds were individually assessed for their ALP activity and mineral content at the end of 21 days.
6.3.4.1 ALP activity within COLPNCOL biphasic scaffolds
ALP, which is a precursor to matrix mineralization, was measured in order to evaluate the mineralization potential exhibited by MC3T3-E1 cells within COL and PNCOL regions of the biphasic scaffolds and displayed in Figure 6-14. The results shown in Figure 6-14 demonstrate that PNCOL is able to elicit a higher cellular ALP activity when compared to
COL scaffolds within scaffolds cultured in the differentiation media under mechanical loading conditions. Both COL and PNCOL regions of the biphasic scaffold show statistically significant increases in the ALP activity after 21 days of culture in
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differentiation media (p < 0.05). However, while COL region shows a 2-fold increase in
ALP activity with respect to the control samples, a 6-fold increase obtained within PNCOL scaffolds in comparison to the control samples cultured in growth media (p < 0.05). Further, the ALP activity within COLPNCOL biphasic scaffolds cultured in differentiation media, the PNCOL region has 2-fold higher ALP activity when compared to COL region, establishing that the difference in material stiffness is directing the cellular synthesis of
ALP (p < 0.05).
Figure 6-14. Alkaline Phosphatase activity within COLPNCOL biphasic scaffold. ALP activity of MC3T3-E1 cells within COL and PNCOL regions of COLPNCOL scaffolds loaded at 2% strain and 0.1 Hz frequency for 2 hours/day for 21 days in osteogenic (differentiation) media or complete (growth) media. # indicates statistical difference between COL and PNCOL (p < 0.05). * depicts significant difference between samples cultured in growth and differentiation media. COLPNCOL scaffolds exhibit 2- fold higher ALP activity in PNCOL compared to COL.
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6.3.4.2 Matrix mineralization within COLPNCOL biphasic scaffolds
Mineral mapping within COLPNCOL biphasic scaffolds
The calcium deposits secreted by the MC3T3-E1 cells within COL and PNCOL regions of the biphasic scaffold cultured in the differentiation media for 21 days were visualized using
SEM-EDS (Figure 6-15).
Figure 6-15A displays the SEM images and the corresponding calcium maps for COL and
PNCOL regions of biphasic scaffolds cultured in growth media (control) and differentiation media. Similar to that observed in Figure 6-7, the effect of uniaxial tensile loading in the matrix directionality is pronounced in the COL region of the biphasic scaffolds cultured in both growth and differentiation media, while PNCOL does not exhibit organization of the matrix. The calcium maps indicate that the amount of calcium is highest in PNCOL region of the scaffold cultured in the differentiation media.
The amount of calcium deposits are further quantified by performing spectral analysis and quantifying the percentage calcium obtained from the element spectra as displayed in
Figure 6-16. Figure 6-16A and Figure 6-16B depicts the spectral analysis of COLPNCOL biphasic scaffolds cultured in growth media and differentiation media, respectively, and correspond to the SEM images shown in Figure 6-15.
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Figure 6-15. Mapping of calcium deposits within COLPNCOL biphasic scaffolds. SEM micrographs and EDS distribution maps of calcium deposits of MC3T3 cells encapsulated within COLPNCOL scaffolds cultured in complete (growth) media and osteogenic (differentiation) media loaded at 2% strain and 0.1 Hz frequency for 2 hours/day for culture period. Scale bar represents 200 µm. Higher amount of calcium deposits visualized for PNCOL region of scaffolds cultured in differentiation media compared to COL region of the biphasic scaffold. 221
Figure 6-16. Spectral analysis for mineral deposits in COLPNCOL scaffolds. Element spectrum of MC3T3-E1-encapsulated COLPNCOL scaffolds loaded at 2% strain and 0.1 Hz frequency for 2 hours/day cultured in (A) complete (growth) media, and (B) osteogenic (differentiation) media. Red arrow indicates the Calcium peak in the spectra. (C) Quantification of percentage calcium from the spectral analysis. * depicts significant difference between samples cultured in growth and differentiation media. PNCOL exhibits 2-fold higher amount of calcium when compared to COL region of the biphasic scaffold. 222
The spectral analyses indicates the presence of calcium peaks in COL and PNCOL cultured in both growth and differentiation media. However, the peak appears to be the highest for the cells within the PNCOL region of the biphasic scaffold cultured in differentiation media with a count of 30000, compared a count of around 15000 for the COL region of the same scaffold. This conforms to the calcium map obtained in Figure 6-15. Further, the spectral analyses for three images for each region of the sample were consolidated into one graph for direct comparison to quantify the percentage calcium present within each sample as shown in Figure 6-16C. The results indicate that there is a 2-fold rise in the calcium amount within PNCOL region in comparison to COL region of biphasic scaffolds cultured in differentiation media (p < 0.05).
Quantification of mineral deposits within COLPNCOL biphasic scaffolds
In order to quantify the total amount of minerals secreted by MC3T3-E1 cells within each region of the biphasic scaffold cultured in growth media and differentiation media, alizarin red assay was performed. Figure 6-17 displays the quantified matrix mineralization of COL and PNCOL regions of the biphasic scaffold after 21 days of culture. COL region of the biphasic scaffold does not show any statistical differences in mineralization produced due to culture in differentiation media when compared to growth media. Significantly, PNCOL region of the scaffold exhibits a 2-fold increase in the amount of matrix mineralization compared to its COL counterpart when cultured in differentiation media (p < 0.05). This
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result is in accordance with the spectral analysis and mineral mapping data presented above and thus confirms that PNCOL is able to mineralize the matrix twice as much as COL.
Figure 6-17. Gradient mineralization within COLPNCOL biphasic scaffold. Alizarin red quantification of mineral deposits of MC3T3-E1 cells encapsulated within COLPNCOL scaffolds loaded at 2% strain and 0.1 Hz frequency for 2 hours/day for 21 days in osteogenic (differentiation) media or complete (growth) media. # indicates statistical difference between COL and PNCOL (p < 0.05). * depicts significant difference between samples cultured in growth and differentiation media. COLPNCOL scaffold exhibits gradient mineralization with PNCOL having 2 fold higher ability to mineralize the matrix compared to COL region.
Thus, the gradient material stiffness employed in the biphasic scaffold is able to direct gradient matrix mineralization of COLPNCOL scaffolds with fibrocartilage-like cells capable of infiltrating and mineralizing the scaffold matrix.
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6.4 Discussion
The gradient in composition, tissue organization, and matrix stiffness of the enthesis region mediates the strong attachment of tendon-to-bone at its insertion point. Recreating the complexity of the enthesis region is highly challenging since it is governed by a variety of growth factors, cell types, and mechanical stimuli. Though many innovative multiphasic scaffolds have been design in the past decade, most of them have not achieved success when implanted into the site of repair. In this chapter, we synthesized PDGF-encapsulated
COLPNCOL biphasic scaffolds to mimic two zones of the enthesis tissue, namely tendon, and fibrocartilage and evaluated the effect of matrix stiffness gradient on the mineralization, directionality and ASC differentiation under physiologically relevant uniaxial tensile loading conditions.
Initially, the material stiffness properties of COL and PNCOL scaffolds were evaluated through rheological analysis. The combined data obtained for elastic and viscous modulus established that PNCOL had remarkably higher mechanical strength and hence material stiffness when compared to COL scaffolds (Figure 6-4). Significantly, this difference in material stiffness between COL and PNCOL resulted in distinct differences in the capacity of fibrocartilage-like MC3T3-E1 cells to mineralize the respective scaffold matrix. A 20- fold increase in the elastic modulus and a 100-fold rise in the viscous modulus of PNCOL led to a 2-fold rise in the amount of mineral deposits within its scaffold matrix in comparison to the mechanically inferior COL scaffolds. This is in accordance with studies
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that have observed increased osteoblastic cell differentiation with increase in the material stiffness of the scaffold [74, 186].
This led to the fabrication of COLPNCOL biphasic scaffolds encapsulated with ASCs and PDGF, with COL representing the tendon region and PNCOL corresponding to the fibrocartilaginous region of the tendon-bone enthesis. Clear differences in the matrix organization and ASC gene expression profiles were observed within COL and PNCOL regions of the biphasic scaffolds subjected to uniaxial tensile loading at 2% strain and
0.1 Hz frequency for 2 hours/day.
Morphological evaluation of the mechanically-loaded biphasic scaffolds revealed remarkably differences in the morphology within COL and PNCOL regions. Specifically, the interface between COL and PNCOL regions demonstrated a compositionally distinct yet structurally continuous transition within the biphasic scaffold Figure 6-6. The fibers within COL region exhibited high degree of matrix organization along with ASC-mediated matrix compaction resulting in parallel alignment of the thickened collagen fibers in the direction of applied load Figure 6-7. In contrast, PNCOL region of the mechanically- loaded biphasic scaffolds largely composed of collagen microfibers and PCL nanofibers prominently displayed random and disorganized matrix (Figure 6-7). The non-loaded biphasic scaffolds, on the other hand, indicated no visible structural differences between 226
COL and PNCOL regions. The visual observations were supported by the directionality analysis that revealed a 2-fold higher amount of directionality within COL in comparison to PNCOL with loaded biphasic scaffolds as well as non-loaded COLPNCOL scaffolds
(p < 0.05) (Figure 6-8).
This established that the matrix stiffness properties of the biomaterial used within the scaffold influenced the extent of matrix organization induced by uniaxial tensile loading.
PNCOL, having a 20-fold higher elastic modulus than COL (Figure 6-4) was able to resist undergoing significant re-organization of its matrix even with the presence of ASCs, thus resulting in a randomly aligned fibers. On the other hand, the low elastic modulus of COL made them very responsive to mechanical loads, which resulted in parallel alignment of the fibers to the applied load of 2% strain and 0.1 Hz. Thus, by varying the material stiffness properties within the COLPNCOL biphasic scaffold, the amount of matrix directionality effected by cyclic loading within COL and PNCOL regions was modulated. The
COLPNCOL biphasic scaffold exhibited a high to low gradient of matrix directionality when transitioning from COL to PNCOL regions within the scaffold. The native enthesis tissue is known to have highly aligned tendon tissue that transitions to criss-cross fibers within the non-mineralized fibrocartilage region, finally giving rise to randomly distributed fibers within the mineralized fibrocartilage region [208]. Thus, the biphasic scaffold in this
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study is able to broadly mimic the structural organization of the in vivo tendon-bone insertion site.
The differentiation response of ASCs within COL and PNCOL regions of PDGF- encapsulated COLPNCOL biphasic scaffolds to uniaxial tensile loading were evaluated by estimating the fold-increases in the ECM, tenogenic, osteogenic, and chondrogenic markers with respect to non-loaded biphasic scaffolds. Three major differences were identified in the gene expression profiles between COL and PNCOL regions of the loaded biphasic scaffolds. First, a significant increase in Collagen I expression (4-fold) was obtained only in the COL region of the biphasic scaffold, with PNCOL region exhibiting no statistically different Collagen I levels (p < 0.05) (Figure 6-9). Second, a remarkable rise in tenogenic markers Tenascin and Scleraxis was obtained only within the COL region, with no significant increases of either genes observed within PNCOL region (p < 0.05)
(Figure 6-10). Third, while COL region displayed no substantial increase in the chondrogenic marker SOX9, a 3.5-fold increase is observed by the ASCs within the
PNCOL region of the loaded biphasic scaffolds (Figure 6-10). It is understood that the fibrocartilage region shows lower expression of Collagen I [21] which is in accordance with our findings, where PNCOL does not show significant rise in Collagen I. Neither COL nor PNCOL display any elevated levels of osteogenic gene ALP when compared to non- loaded scaffolds (Figure 6-10). Interestingly, significant increases in GAG markers
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Decorin and Aggrecan were obtained for both COL and PNCOL regions, with similar fold- increases within each region of the loaded scaffolds in comparison to non-loaded biphasic scaffolds (p < 0.05) (Figure 6-9). This observation is similar to studies where GAGs have been predominantly upregulated in both tenogenesis and chondrogenesis at its early stage.
In such cases, an increase in Collagen II along with GAGs was considered as a definitive indication of fibrocartilage tissue formation [21].
Thus, the gene expression data demonstrate the synergistic effect of uniaxial tensile loading, PDGF and matrix stiffness in dictating the differentiation lineage of ASCs encapsulated within the COL and PNCOL regions of the biphasic scaffolds. It is observed that the loaded COL region that experienced higher strains due to its elasticity (Figure 6-4) along with a highly aligned matrix (Figure 6-7) was able to direct ASCs to synthesize
Collagen I and III, and induce tenogenic differentiation, along with elevated GAG levels
(Figure 6-9 and Figure 6-10). This was very representative to the characteristics of a tendon scaffold. In contrast, PNCOL that was resistant to the applied uniaxial tensile load and exhibited disorganized fiber distribution within its matrix (Figure 6-7) was unable to stimulate elevated expressions of both Collagen I and tenogenic markers. Instead, PNCOL, along with higher expression of GAGs showed an onset of chondrogenesis (Figure 6-11).
These results matched well to the expected composition within a fibrocartilaginous tissue
[21]. It can hence be said that there is a decreasing gradient of Scleraxis and an increasing
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gradient of SOX9 within COLPNCOL biphasic scaffolds. Significantly, recent studies investigating the molecular mechanisms within the enthesis region have identified
Scleraxis and SOX9 as the critical genes driving tenogenesis of the tendon phase, and chondrogenesis of the fibrocartilage phase, respectively [209, 216]. Thus, the statistically significant increases of Scleraxis in COL region and SOX9 in PNCOL (Figure 6-10) indicates that ASCs directed by the mechanical properties of the biomaterial they are encapsulated within can be differentiated into both tenocytes and chondrocytes under the same uniaxial tensile loading modality.
Quantification of secreted proteins from the loaded COLPNCOL biphasic scaffolds demonstrated 2-fold rises in both collagen proteins and GAGs in comparison to non-loaded biphasic scaffolds (Figure 6-12). This implied that ASCs within COLPNCOL scaffolds are able to translate the fold-rises in the obtained ECM gene expressions into increased
ECM synthesis. Earlier studies have established that increased synthesis of ECM leads to increase in the overall mechanical properties of the scaffold. Thus, the increase in ECM secretion observed in this study could reduce the stress concentration between the interface region of between COL and PNCOL, thereby increasing the robustness of the biphasic scaffold [217].
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The ability of PDGF-encapsulated with COLPNCOL scaffolds to act as a chemoattractant is critical in order to induce resident fibrocartilage cells to migrate and infiltrate the COLPNCOL scaffold and subsequently mineralize the matrix when implanted at the repair site. Biphasic scaffolds composed of acellular PDGF-immobilized hb-PNCOL fused with MC3T3-E1-encapsulated COL displayed a 2.5-fold increase in cell infiltration from COL to PNCOL region in presence of PDGF (1 µg/ml) versus no PDGF
(0 µg/ml) control samples (p < 0.05). Figure 6-13A thus established two important features of PDGF-hb-PNCOL scaffolds: (1) PNCOL had appropriate porosity to allow infiltration of cells into its matrix, and (2) PDGF with PNCOL scaffolds induced fibrocartilage-like cells to migrate and subsequent infiltrate the PDGF-hb-PNCOL scaffold. Specifically, 15% of the cells encapsulated within COL scaffolds infiltrated into the PDGF-hb-PNCOL after
7 days of culture under uniaxial tensile loading conditions.
Interestingly, the number of cells within the COLPNCOL biphasic scaffold having no
PDGF (0 µg/ml) scaffolds were similar to those within PDGF-encapsulated (1 µg/ml) scaffolds (p < 0.05) (Figure 6-13B). Since we have already established in Chapter 5 that
PDGF is bioactivity within PDGF-hb-PNCOL scaffolds (Figure 5-10), the lack of proliferation, in this case, could be attributed to the dosage of PDGF with which the cells are stimulated. The monolayer proliferation study was conducted with 100 ng/ml PDGF in the release media, while PDGF within the scaffolds are loaded at a 10-fold higher
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concentration of 1 µg/ml. Higher concentrations of growth factor could have reduced the stimulatory effect on cell proliferation. Nevertheless, the COLPNCOL biphasic scaffold can be tailored to accommodate the appropriate amount of growth factor that positively stimulates the proliferation of cells both within the scaffold and outside the scaffold by conducting a detailed dosage-response study.
Finally, the ability of the infiltrated fibrocartilage-like cells to mineralize the mechanically- loaded COLPNCOL biphasic scaffolds in a gradient fashion was demonstrated using mineral mapping and quantification studies. The effect of gradient material stiffness in generating the gradient mineralization within the biphasic matrix was clearly evident from calcium mapping and element spectra that exhibited almost a 2-fold higher calcium levels within PNCOL when compared to COL scaffolds (p < 0.05) (Figure 6-15 and Figure
6-16). This was further confirmed by quantifying 2-fold increases in both the cellular ALP activity that is a precursor to mineralization and the total mineral deposits within PNCOL versus COL regions of the biphasic scaffold (Figure 6-14 and Figure 6-17). Thus,
COLPNCOL biphasic scaffold exhibits an increasing gradient of mineral deposits that correlates to its increasing gradient of matrix stiffness. This is very similar to that observed with in vivo enthesis tissues that have increasing mineral gradient from tendon to fibrocartilage to bone, thus creating a smooth mechanical stress distribution at the tendon- bone insertion point [218].
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6.5 Conclusion
In Chapter 4, we engineered a biphasic scaffold composed of COL and PNCOL phases encapsulated with ASCs and PDGF and evaluated its performance in terms of matrix directionality, matrix mineralization, and ASC differentiation under uniaxial tensile loading. Figure 6-18 summarizes the major highlights of this study.
The mechanically-conditioned PDGF/ASC-encapsulated COLPNCOL biphasic scaffolds exhibited distinct morphological, compositional, and biochemical differences between COL and PNCOL regions that are appropriate for a scaffold targeting the repair of tendon-bone interfaces.
The COL region of the loaded biphasic scaffold, due to its lower matrix stiffness demonstrate high matrix directionality and low matrix mineralization. ASCs within COL matrix have higher Collagen I, GAGs expression, and tenogenic markers, with no osteogenic and chondrogenic expression, thus exhibiting tendon-specific characteristics.
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Figure 6-18. Major results obtained from Chapter 6. Potential of PDGF/ASC-encapsulated COLPNCOL biphasic scaffolds for ASC tenogenic differentiation and gradient mineralization under uniaxial tensile loading conditions.
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PNCOL, on the other hand, having 20-fold higher elastic modulus is highly resistant to mechanical loading-induced structural changes, and thus has a predominantly random matrix organization. The higher matrix stiffness also contributes to a significantly higher ability for cell-induced matrix mineralization. The gene expression profiles, remarkably, do not show any elevation in tenogenic markers as well as collagen I expression. Instead, a significant increase in the expression of chondrogenic marker SOX9 is obtained along with increases in GAG expression, which indicates a potential for PNCOL to mimic the fibrocartilage region of the enthesis tissue.
The two phases of the mechanically-conditioned COLPNCOL biphasic scaffold thus exhibit some defining characteristics observed in the tendon-fibrocartilage region of the enthesis tissue at the rotator cuff insertion point. Further, PNCOL encapsulated with PDGF demonstrates the ability to chemo-attract cells and allow invasion into the scaffold matrix which is critical at the in vivo tendon repair site for resident fibrocartilage cells to infiltrate the scaffold and mineralize the matrix in a gradient fashion. The gradation in material stiffness within the biphasic scaffold results in gradient increase in mineralization and gradient decrease in matrix directionality which can smoothen the stress concentration at the tendon-bone interface.
Thus, the mechanically-conditioned PDGF/ASC encapsulated COLPNCOL biphasic scaffold has great potential to enhance healing and guide the maturation of the de novo tissue when implanted at the tendon-bone insertion point of the rotator cuff.
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Chapter 7
Future Work
In this dissertation, we engineered a mechanically-conditioned PDGF/ASC-encapsulated
COLPNCOL biphasic scaffold with gradient material stiffness properties to mimic the tendon-bone interface tissue. This biphasic scaffold exhibited controlled release of bioactive PDGF, gradient increase in matrix mineralization, gradient decrease in matrix directionality and expression of tenogenic markers within COL region and chondrogenic markers within the PNCOL region. The scaffold would next have to be evaluated in an animal injury model to assess its performance at the rotator cuff repair site.
The mechanically-conditioned biphasic scaffold is envisioned as a tissue engineering strategy to enhance tendon healing and guide the formation of the enthesis tissue at the rotator cuff repair site. The goal is to use the patient’s own ASCs obtained from liposuction and encapsulate them into the COLPNCOL biphasic scaffold along with PDGF. The scaffold would be mechanically-loaded using the rehabilitation loading regime of 2% strain and 0.1 Hz frequency to accelerate the cellular response. The mechanically-conditioned
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COLPNCOL biphasic scaffold would be arthroscopically implanted into the rotator cuff repair site and fixed on the area with the help of biodegradable sutures or staples. The scaffold could be delivered along with a commercially available acellular collagen patch.
The biphasic scaffold is expected to provide bioactive, cellular, and micro-environmental cues to the repair site and enhance cell proliferation and ECM secretion, while the acellular patch would provide mechanical stability to the repair site.
The below sections provide thoughts and recommendations on possible future work for each study presented in this dissertation.
7.1 Recommendations for study described in Chapter 3
“A mechanical loading bioreactor to apply homogenous uniaxial tensile strains to cell-encapsulated 3D collagen scaffolds.”
The uniaxial tensile strain bioreactor described in Chapter 3 is specifically targeted to mechanically condition collagen-based scaffolds for tendon repair strategies. However, the bioreactor presented in this study is a mere framework of a mechanical loading platform and has great potential to be modified to accommodate various other applications. This is possible mainly due to the excellent tensile strength of the loading chamber material, the
237
flexibility in the design configuration of the chamber, and performance of the driving mechanism. Some possible modifications include scaling up of the loading chamber to hold longer or larger scaffolds for repair of large tissue defects, employing stiffer biomaterials with superior mechanical properties, or designing different geometries of the silicone loading chamber to cater to various implant shapes. With these modifications, the uniaxial tensile strain bioreactor can be employed to create engineered scaffolds for a wide range of tissues, such as bone, ligament, skin, heart, muscle, and nerve that routinely experience uniaxial tensile strains in their in vivo environment.
Further, the potential use of this uniaxial tensile strain bioreactor does not restrict solely to tissue engineering strategies. This bioreactor can be also be applied to conduct mechanobiology and biochemical expression studies to elucidate cellular mechanotransduction signaling pathways within a 3D microenvironment. Another major application of this bioreactor is to create mechanically-conditioned human tissue substitutes to use as in vitro platform for animal-free efficacy and toxicity testing of drugs and cosmetics products.
7.2 Recommendations for study described in Chapter 4
“Effect of uniaxial tensile strains and frequencies on the matrix organization and tenogenic response of ASCs within 3D collagen scaffolds.” 238
The study described in Chapter 4 conducted a detailed investigation on the morphological changes and gene expression profiles exhibited by ASCs within 3D collagen scaffolds when subjected to a range of strains and frequencies of mechanical loading. This was performed to identify the magnitude of uniaxial tensile strain and loading frequency appropriate for pure tenogenic differentiation of ASCs. This study could be further expanded to investigating the protein expression profiles exhibited by ASCs through
Western analyses and immunohistochemistry by checking for ECM and tendon-related gene markers. Further, mechanobiological studies could be conducted to identify the signaling pathway and genes involved that induces ASC tenogenic differentiation in response to mechanical loading. Also, the mechanism behind the change in cell morphology and orientation could be investigated by checking for actin proteins and calcium ion signaling pathways.
7.3 Recommendations for study described in Chapter 5
“Composite collagen scaffolds (PNCOL) interspersed with polycaprolactone
(PCL) nanofibers for controlled delivery of growth factors under uniaxial tensile loading.”
The study in Chapter 5 described the synthesis of composite collagen scaffolds (PNCOL) with improved protein retention properties for controlled delivery of bioactive growth 239
factors under uniaxial tensile loading for tendon repairs. The versatility of PNCOL allows it to be used as a biomaterial for various other tissue engineering applications. Addition of functionalized PCL nanofibers into the collagen matrix was seen to increase the structural integrity of the scaffold and provide a larger surface area for protein immobilization. Thus, by varying the PCL concentration within the collagen matrix, the release profile and the duration of bioactivity preservation of the growth factor within PNCOL composite scaffolds can be tailored based on the target release kinetics required for a treatment strategy. Further, the versatility of PNCOL as a composite biomaterial allows it to be used as a dual-growth factor delivery system with sequential release of growth factors. The growth factor loaded directly into the collagen matrix would exhibit a faster release profile, while the growth factor bound to PCL would undergo a slow and controlled release.
Finally, PNCOL remains a hydrogel-based biomaterial despite incorporation of synthetic
PCL fibers. This allows the composite biomaterial PNCOL to be delivered as an injectable scaffold to fill the defect site, with controlled and sustained delivery of cells and growth factor for tissue regeneration.
240
7.4 Recommendations for study described in Chapter 6
“PDGF/ASC-encapsulated COLPNCOL biphasic scaffold with gradient matrix directionality, mineralization, and tenogenic differentiation under uniaxial tensile loading for tendon-bone interface.”
In the study described in Chapter 6, a straightforward biphasic scaffold was engineered with COL and PNCOL having gradient material stiffness properties. The mechanically- loaded COLPNCOL scaffolds encapsulated with ASCs and PDGF exhibited gradient decrease in matrix alignment and tenogenic differentiation, and gradient increase in matrix mineralization, thus representing the gradient structure and composition at the native tendon-bone interface. This is, however, the first step in mimicking a highly complex and evolved enthesis tissue at the tendon-to-bone insertion site. The gradation in mineral deposits and matrix directionality can be further improved by making a tri-phasic scaffold, with the third phase comprised of PNCOL with 1% (w/v) PCL added between COL and
PNCOL regions. This would provide a smoother transition in the material stiffness properties of the tri-phasic scaffold and reduce the stress concentration at the original COL-
PNCOL interface. The COLPNCOL biphasic scaffold could be further enhanced by incorporating fibrochondrocytes in the PNCOL region along with ASCs in the COL region, to accelerate fibrocartilage formation within the PNCOL region. Also, though PDGF has been employed in this study, other growth factors such as TGF-β or BMP-2 involved in the
241
maturation of the native tendon-bone interface can incorporated within the COLPNCOL scaffold matrix and evaluated for its performance. Finally, mechanical testing of the mechanically-conditioned biphasic scaffold should be performed to evaluate its ultimate tensile strength and matrix stiffness.
242
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