MESENCHYMAL STEM CELL MECHANOBIOLOGY AND …MESENCHYMAL STEM CELL MECHANOBIOLOGY AND TENDON...

MESENCHYMAL STEM CELL MECHANOBIOLOGY

AND TENDON REGENERATION

Daniel Warren Youngstrom

Dissertation submitted to the faculty of the

Virginia Polytechnic Institute and State University

in partial fulfillment of the requirements for

the degree of

Doctor of Philosophy

in

Biomedical and Veterinary Sciences

Jennifer G. Barrett, Chair

Linda A. Dahlgren

William R. Huckle

Willard H. Eyestone

20 March 2015

Blacksburg, Virginia, USA

Keywords: regenerative medicine, tissue engineering,

mesenchymal stem cells, mechanobiology, tendon

Copyright© 2015 Daniel Warren Youngstrom

unless otherwise stated

MESENCHYMAL STEM CELL MECHANOBIOLOGY

AND TENDON REGENERATION

Daniel Warren Youngstrom

ABSTRACT

Tendon function is essential for quality of life, yet the pathogenesis and healing of

tendinopathy remains poorly understood compared to other musculoskeletal disorders. The aim

of regenerative medicine is to replace traditional tissue and organ transplantation by harnessing

the developmental potential of stem cells to restore structure and function to damaged tissues.

The recently discovered interdependency of cell phenotype and biophysical environment has

created a paradigm shift in cell biology. This dissertation introduces a dynamic in vitro model for

tendon function, dysfunction and development, engineered to characterize the mechanobiological

relationships dictating stem cell fate decisions so that they may be therapeutically exploited for

tendon healing.

Cells respond to mechanical deformation via a complex set of behaviors involving

force-sensitive membrane receptor activity, changes in cytoskeletal contractility and

transcriptional regulation. Effective ex vivo model systems are needed to emulate the native

environment of a tissue and to translate cell-matrix forces with high fidelity. A naturally-derived

decellularized tendon scaffold (DTS) was invented to serve as a biomimetic tissue culture

platform, preserving the structure and function of native extracellular matrix. DTS in concert

with a newly designed dynamic mechanical strain system comprises a tendon bioreactor that is

able to emulate the three-dimensional topography, extracellular matrix proteins, and mechanical

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strain that cells would experience in vivo. Mesenchymal stem cells seeded on decellularized

tendon scaffolds subject to cyclic mechanical deformation developed strain-dependent alterations

in phenotype and measurably improved tissue mechanical properties. The relative tenogenic

efficacies of adult stem cells derived from bone marrow, adipose and tendon were then compared

in this system, revealing characteristics suggesting tendon-derived mesenchymal stem cells are

predisposed to differentiate toward tendon better than other cell sources in this model.

The results of the described experiments have demonstrated that adult mesenchymal

stem cells are responsive to mechanical stimulation and, while exhibiting heterogeneity based on

donor tissue, are broadly capable of tenocytic differentiation and tissue neogenesis in response to

specific ultrastructural and biomechanical cues. This knowledge of cellular mechanotransduction

has direct clinical implications for how we treat, rehabilitate and engineer tendon after injury.

iv

DEDICATION

To my love, Catherine E. M. Youngstrom, who shows me the beauty in life.

May our work help alleviate suffering.

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ACKNOWLEDGEMENTS

First and foremost I owe gratitude to Jennifer G. Barrett: a brilliant scientist, dedicated

mentor and friend. Your contagious work ethic and high expectations have inspired me to

achieve to the best of my ability. It was an honor growing with you as the first PhD student in

your lab. Thank you to Linda A. Dahlgren, William R. Huckle and Willard H. Eyestone for

providing valuable insight into my project as committee members, as well as for the stimulating

discussions within the Interdisciplinary Graduate Education Program in Regenerative Medicine.

Thank you to Roger J. Avery and Becky Jones in the Program in Biomedical and Veterinary

Sciences. Thank you to George J. Christ for serving as the external examiner for my defense.

I am appreciative to have worked alongside an incredible group of fellow PhD students at

the Equine Medical Center: Tracee L. Popielarczyk, Jade E. LaDow, Ibtesam Rajpar and Sophie

H. Bogers. I look forward to following the excellent work you all continue to do. A huge thank

you also goes out to the research and regenerative medicine laboratory staff for your technical

assistance and companionship: Elaine Meilahn, Laurie Fahl, Zijuan Ma, Sara Stapp, Andrew

Hogan, Hannah Dawson and Shay Strauss. I wish the best to the undergraduate students I was

fortunate to work with, especially Vidya Ganesh and Austin Dunn. Thank you to all of the

veterinary residents and hospital staff for joining me in this journey.

This work has granted me opportunities to work with talented collaborators. It was a

pleasure working on the first bioreactor study with Shannon L. Smith, Rod R. Jose, Jonathan A.

Kluge and David L. Kaplan at Tufts University. I must also thank those I worked with during my

Fulbright year in Latvia at the Cell Transplantation Center: Inese Čakstiņa, Mārtiņš Borodušķis,

Lita Grīne, Vika Telle and Ēriks Jakobsons. While our work is published elsewhere, you exposed

me to new techniques and ideas while enriching my time abroad.

I am indebted to those who served as mentors and friends during my undergraduate years

at the University of Michigan. I owe particular thanks to Remy L. Brim, Brandon H.

McNaughton, Kevin Hartman, Jeffrey R. Brender and Ayyalusamy Ramamoorthy, without

whom I may never have pursued a PhD. You initiated me into laboratory culture, served as

excellent role models, treated me as a peer and helped me realize my professional potential.

Above all, I must thank the dear friends and family members who provided unconditional

support during the highs and lows of the past five years. I could not have done this without you.

vi

ATTRIBUTIONS

This dissertation represents an original body of work by Daniel W. Youngstrom. The

roles of co-authors and collaborators in this research are described below.

Chapter 3 is based on a manuscript co-authored with Jennifer G. Barrett, Rod R. Jose and

David L. Kaplan: “Functional characterization of detergent-decellularized equine tendon

extracellular matrix for tissue engineering applications,” published in PLoS ONE.

Jennifer G. Barrett, DVM PhD is the Theodora Ayer Randolph Professor of Equine

Surgery at the Marion duPont Scott Equine Medical Center at Virginia Tech. She participated in

study design, data acquisition and analysis, critical revision, and manuscript approval.

Rod R. Jose, PhD was a graduate student in biomedical engineering at the Tissue

Engineering Resource Center at Tufts University at the time of publication. He assisted in data

acquisition and analysis of biomechanical data, critical revision, and approved the manuscript.

David L. Kaplan, PhD is the Stern Family Professor of Engineering, Chair of Biomedical

Engineering and Director of the Tissue Engineering Resource Center at Tufts University. He

assisted in data acquisition and analysis, critical revision, and approved the manuscript.

Chapter 5 is based on a manuscript co-authored with Ibtesam Rajpar, Jennifer G. Barrett,

and David L. Kaplan: “A bioreactor system for in vitro tendon differentiation and tendon tissue

engineering,” published in the Journal of Orthopaedic Research.

Ibtesam Rajpar, MS was a laboratory technician at the Marion duPont Scott Equine

Medical Center at Virginia Tech at the time of publication. She assisted in data acquisition and

analysis of qPCR data, critical revision, and approved the manuscript.

Jennifer G. Barrett, DVM PhD participated in study design, data acquisition and analysis,

critical revision, and manuscript approval.

vii

David L. Kaplan, PhD assisted in data acquisition and analysis, critical revision, and

approved the manuscript.

Chapter 6 is based on a manuscript co-authored with Jade E. LaDow and Jennifer G.

Barrett: “Relative tendon differentiation of bone marrow-, adipose- and tendon-derived stem

cells in a dynamic bioreactor,” which is currently under review for publication.

Jade E. LaDow, MS is graduate student in biomedical and veterinary sciences at the

Marion duPont Scott Equine Medical Center at Virginia Tech. She assisted in data acquisition

and analysis, critical revision, and approved the manuscript.

Jennifer G. Barrett, DVM PhD participated in study design, data acquisition and analysis,

critical revision, and manuscript approval.

viii

TABLE OF CONTENTS

ABSTRACT ii

DEDICATION iv

ACKNOWLEDGEMENTS v

ATTRIBUTIONS vi

TABLE OF CONTENTS viii

LIST OF FIGURES xiii

LIST OF TABLES xv

PREFACE xvi

OVERVIEW OF DISSERTATION xvii

CHAPTER 1: GENERAL INTRODUCTION

Tendon Structure, Function and Development 1

The Pathogenesis of Degenerative Tendinopathy 4

Mesenchymal Stem Cells and the Stem Cell Niche 8

Regenerative Medicine: Cell Therapy and Tissue Engineering 10

Mechanobiology: Differentiation and Phenotype 12

CHAPTER 2: MODEL SYSTEMS IN MUSCULOSKELETAL CELL BIOLOGY

Animal Models of Tendinopathy 17

Current Status of Biomimetic Scaffolds and Culture Systems 19

Review of Scaffold Systems 21

Review of Bioreactor Systems 26

Applications of Bioreactors 33

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CHAPTER 3: A DECELLULARIZED EQUINE FLEXOR TENDON SCAFFOLD

Abstract 35

Introduction 36

Materials and Methods 39

Experimental design 39

Scaffold preparation 40

Biochemical characterization 41

Microscopic evaluation 41

Scanning electron microscopy 42

Biomechanics 42

Biocompatibility 43

Cell culture 44

Statistical analysis 45

Results 45

2% SDS removes cellular debris from tendon explants 45

Tendon composition is not significantly altered by decellularization 47

Decellularized tendon maintains desired ultrastructure 48

Scaffolds are biocompatible and free of endotoxin 50

MSCs proliferate on and penetrate deep into DTS 51

Discussion 51

Acknowledgements 58

CHAPTER 4: DEVELOPMENT OF A TENDON BIOREACTOR

Hardware 59

x

Software 61

CHAPTER 5: MECHANICAL STRAIN MODULATES STEM CELL PHENOTYPE

Abstract 65

Introduction 66



Production of decellularized tendon scaffolds (DTS) 68

Derivation of primary mesenchymal stem cell (MSC) lines 69

Construct seeding and bioreactor culture 69

RNA isolation and gene expression analysis 71

Mechanical testing 72

Spectrophotometric biochemistry assays 73

Histology 73


Results 74

Cyclic strain promotes a tenocytic gene expression profile 74

3%-strained constructs mimic the mechanical properties of native FDST 75

MSCs integrate into DTS and modulate scaffold composition 77

Discussion 79

Acknowledgements 84

CHAPTER 6: COMPARISON OF TENOGENIC EFFICACY OF ADULT STEM CELLS

Abstract 85

Introduction 86

xi



Decellularized tendon scaffolds 89

Cell isolation and culture 90

Cell-laden bioreactor constructs 90

Cyclic strain bioreactor 91

Quantitation of gene expression 91

Microscopic imaging 93

Biochemical composition 93

Mechanical testing 93


Results 95

BM MSCs form fewer colonies than AD or TN counterparts 95

MSCs integrate into DTS during bioreactor culture 95

Gene expression profiles during tenogenesis differ by MSC source 96

Scaffold composition was not heavily influenced by cell type 98

Construct mechanical properties are altered by resident cells 99

Discussion 99

Acknowledgements 105

CHAPTER 7: CONCLUSIONS AND FUTURE RESEARCH

Summary of Results 106

Future Directions 108

REFERENCES 109

xii

APPENDIX: SPECIALIZED EXPERIMENTAL PROTOCOLS

Research Protocol: Decellularization of Equine Flexor Tendons 157

Technical notes 157

Materials 157

Methods 158

Research Protocol: RNA Isolation and Purification from Tendon 162

Technical notes 162

Materials 162

Methods 163

xiii

LIST OF FIGURES

Figure 1.1 - Overview of integrin signaling 14

Figure 3.1 - Calculus for biomechanical data 43

Figure 3.2 - Nuclear content following decellularization 46

Figure 3.3 - Image analysis of nucleic acid fluorescence 47

Figure 3.4 - Treatment alters tendon matrix composition 48

Figure 3.5 - Ultrastructural imaging 49

Figure 3.6 - Representative DTS stress/strain curve 51

Figure 3.7 - Biocompatibility of DTS to MSC culture 53

Figure 3.8 - Cellular integration into DTS 55

Figure 4.1 - Bioreactor wiring diagram 60

Figure 4.2 - Photograph of bioreactor clamps 61

Figure 4.3 - Bioreactor LabVIEW program 62

Figure 4.4 - Software front end 63

Figure 4.5 - Tensile testing program 64

Figure 5.1 - Photograph of bioreactor vessel 70

Figure 5.2 - Experimental timeline 71

Figure 5.3 - Strain amplitude alters gene expression 75

Figure 5.4 - Real-time strain monitoring 76

Figure 5.5 - Construct resilience during exercise 77

Figure 5.6 - 3% strain improves failure strength 78

Figure 5.7 - Biochemical content varies by amplitude 80

xiv

Figure 5.8 - MSC integration into DTS in bioreactor 83

Figure 6.1 - Bioreactor protocol and experimental timeline 92

Figure 6.2 - Proliferative capacities of MSCs derived from BM, AD and TN 95

Figure 6.3 - Histology and confocal microscopy 97

Figure 6.4 - Confocal cellular infiltration 97

Figure 6.5 - Seeding density and 3-D reconstruction 98

Figure 6.6 - Relative gene expression profiles 100

Figure 6.7 - Endpoint biochemical analysis and GAG release 101

Figure 6.8 - Daily secant moduli monitored via load cells 102

Figure 6.9 - Biomechanical characterization 103

Figure A.1 - Scaffold cutting template for bioreactor 160

Figure A.2 – Pictorial tendon decellularization protocol 161

xv

LIST OF TABLES

Table 2.1 - Tendon bioreactors by generation 29

Table 2.2 - Summary of generation-4 bioreactors based on cell-laden DTS 32

Table 3.1 - DTS tensile testing data 50

Table 5.1 - List of qPCR primers and probes in Chapter 5 72

Table 6.1 - Surface marker profiles of BM, AD and TN cells 88

Table 6.2 - List of qPCR primers and probes in Chapter 6 94

xvi

PREFACE

Manuscripts and figures reproduced from previously published materials and the roles of

those who are acknowledged in subsequent chapters are described below.

Chapter 1 contains a figure reproduction. Figure 1.1 was adapted with permission from

Macmillan Publishers Ltd: Nature Rev Mol Cell Bio (Wang, et al.), copyright 2009.

Chapter 3 is based on a published manuscript. Assistance with electron microscopy was

provided by the Morphology Laboratory at the Virginia-Maryland College of Veterinary

Medicine and the Nanoscale Characterization and Fabrication Lab at the Virginia Tech Institute

for Critical Technology and Applied Science. This manuscript is published and available open-

access as: Youngstrom, et al. (2013) PLoS ONE - DOI: 10.1371/journal.pone.0064151.

Chapter 4 contains original, unpublished information on bioreactor hardware and

software. The concept was loosely based on previously published information, which has been

cited. This system is patent pending (US patent #62050792). Co-inventors are Jennifer G.

Barrett, Daniel W. Youngstrom and David Simmons, who also fabricated the design at Virginia

Tech. This system was used to produce Figures 5.4-5.5, as well as all of the data in Chapter 6.

Chapter 5 is based on a published manuscript. Shannon L. Smith and Jonathan A. Kluge

provided equipment and training for use of the bioreactor at Tufts University. Laurie Fahl

assisted with cell culture and sample shipping at Virginia Tech. This manuscript is published as:

Youngstrom, et al. (2015) J Orthop Res - DOI: 10.1002/jor.22848.

Chapter 6 is based on a manuscript that is under peer review. Zijuan Ma conducted the

flow cytometry experiments included in Table 6.1. Kristopher E. Kubow generously provided

voluntary assistance with confocal microscopy at James Madison University.

Financial support was provided by the Virginia-Maryland College of Veterinary

Medicine, the Virginia Tech Institute for Critical Technology and Applied Science, the Morris

Animal Foundation, the Grayson-Jockey Club Research Foundation, and the Tufts University

Tissue Engineering Resource Center, sponsored by the National Institutes of Health. Funding

agencies had no role in study design or interpretation. No conflicts of interest are declared.

xvii

OVERVIEW OF DISSERTATION

Chapter 1 contains an outline of the clinical problem of tendinopathy and the new

treatment paradigm of stem cell-based regenerative medicine. We have a basic understanding of

how cells integrate environmental information to produce and repair injured tendon, but new

techniques are needed to translate this knowledge to the bedside.

Chapter 2 serves as a review of model systems in musculoskeletal cell biology, with an

emphasis on bioreactor systems incorporating mechanical stimulation. This demonstrates the

broad array of applications for bioreactors, including their use in cell therapy and tissue

engineering, putting context to the developments contained in this dissertation.

A novel detergent-based decellularization protocol is presented in Chapter 3, specifically

optimized for the equine flexor digitorum superficialis tendon (FDST) due to its size, mechanical

strength and availability. These decellularized tendon scaffolds (DTS) are designed to serve as

three-dimensional platforms for tendon tissue culture, and can withstand heavy physical

manipulation while translating mechanical forces to cells with high fidelity relative to what they

might experience in an animal. DTS is produced using 2% sodium dodecyl sulfate, followed by

treatment with trypsin, DNase-I and ethanol. DTS is cell-free and retains the ultrastructure,

mechanical properties and biochemical composition of native tendon. DTS is shown to be

hospitable to MSC culture.

Chapter 4 describes a tendon bioreactor system designed, fabricated and programmed at

Virginia Tech. Hardware, software and basic applications are outlined.

The merging of DTS and bioreactor is the subject of Chapter 5. Bone marrow-derived

MSCs were cultured on matched DTS and cultured for 11 days subject to increasing durations of

xviii

0%, 3% or 5% strain at 0.33Hz for up to one hour per day. Cells integrated into DTS over that

time and developed elongated, fibroblastic phenotypes in parallel with collagen fibers. The 3%-

strained group expressed the most scleraxis, collagen-I and decorin mRNA, representing the

most tenocytic phenotype. Strained constructs were mechanically robust relative to native FDST,

and the 3% group developed a significant increase in failure stress relative to DTS. The results of

this experiment led to the conclusion that MSCs are responsive to varying amplitudes of cyclic

strain, and provided a protocol for subsequent tissue culture studies.

Principles from Chapter 5 are then applied in Chapter 6 to elucidate basic epigenetic

differences between MSCs derived from bone marrow, adipose and tendon. All three cell types

began to acquire a tenocytic morphology and phenotype in response to culture in the bioreactor,

and improved construct mechanical properties relative to unseeded DTS. Tendon-derived cells

proliferated the fastest and expressed more scleraxis, collagen-I and cartilage oligomeric matrix

protein mRNA than MSCs derived from bone marrow or adipose. Although tendon-derived

MSCs are more difficult to obtain, their relative propensity to form tendon may be particularly

beneficial for cell therapy or tissue engineering applications using allogeneic cell lines.

Major conclusions, a general discussion, and recommendations for future work using this

tendon bioreactor system are included in the final chapter.

1

CHAPTER 1: GENERAL INTRODUCTION

TENDON STRUCTURE, FUNCTION AND DEVELOPMENT

Tendons connect muscle to bone, functioning in force translation and energy storage

during movement [1]. Up to 80% of the dry mass of tendon is fibrillar collagen, and the specific

mechanical properties of tendon are largely the result of type-I collagen organization within the

extracellular matrix (ECM) [2]. Three α-helical molecular [Gly-x-y]n collagen strands form the

triple helical foundation of tendon structure: these are quarter-staggered into banded microfibrils

that are hierarchically arranged into secondary and tertiary bundles [3, 4]. Collagen expression

within tendon is regulated by a number of molecules including the transcription factor scleraxis,

which will be discussed in greater detail elsewhere in this work [5, 6]. Procollagen molecules

then undergo post-translational modifications and their assembly is regulated by molecular

chaperones [7, 8]. Type-III collagen is an important minority component of tendon ECM, and its

elevated presence is associated with decreased fiber diameter [9] during development and repair

[10]. Type-V collagen is also present in the core of fibrils and contributes to the structural

arrangement of ECM more than its mechanical properties [11].

Partially a result of its characteristically low cellularity relative to other tissues, tendon

matrix was once thought to be inert of metabolic activity [12]. However, much like the classic

frameworks of bone remodeling (Wolff’s Law and Davis’ Law, respectively) [13], tendons

dynamically react to different loading conditions [14], with different tendons exhibiting

variations by function [15]. Tenocytes are the terminally differentiated cells resident to tendon,

and are generally responsible for maintaining ECM homeostasis. Tenocytes align along the

2

proximal-distal axis parallel to fiber direction, extending projections deep into their extracellular

environment and maintaining cell-cell connectivity through cadherin-11 junctions [16]. Matrix

metalloproteinases (MMPs) degrade ECM and include secreted gelatinases (MMP-2 and -9),

collagenases (-1 and -8), and stromelysins (-3, -10 and -11) among others [17, 18]. Dysregulation

of these enzymes is a trait of degeneration: MMP-2, -3, -14 and -19 are significantly upregulated

in human tendinopathy [19]. Healthy tenocytes deposit collagen to counteract this degradation

[20]. The metabolic activities of tendon cells differ from tendon to tendon, and it is uncertain if

this is a developmental trait or the result of adaptation to a specific mechanical environment [21].

Glycosaminoglycans (GAGs) are linear polysaccharide chains covalently bonded to

proteoglycan cores within the ECM [3]. GAGs are important extracellular regulators, assisting in

the lateral aggregation of type-I collagen [22] assisting in water homeostasis [23], altering tissue

biomechanics and resisting compression [24, 25]. GAGs also non-specifically bind growth

factors, giving the ECM additional regulatory properties [26]. GAG content must be carefully

maintained, as over- or under-production has negative effects. In addition to remodeling

collagen, tenocytes are responsible for managing proteoglycan turnover. In bovine deep flexor

tendon explants, large proteoglycans have half-lives of approximately two days [27]. This is

comparatively rapid for ECM proteins, particularly as carbon turnover virtually ceases in humans

after adolescence [28]. Changes in proteoglycan turnover likely come before detectable structural

alterations [29]. Increased GAG content has been found in diseased tendons [30], while GAG

digestion lowers viscoelasticity [31, 32]. GAG exposure in damaged tendon may also be an

irritant related to the pain of tendinopathy [33]. Thus GAGs are inexorably tied to tendon health.

There are many cell types to be found within tendon, resident to both the tendon mid-

substance and its associated vascularization and innervation. Three of these cell types are of

3

particular note in tendon: tenocytes, tendon stem/progenitor cells (TSPCs) (also known as

tendon-derived MSCs) and pericytes. TSPCs are rapidly-dividing, colony-forming multipotent

mesenchymal stem cells [34] that represent between 0.05% [35] and 4% [36] of the resident cell

population in tendon. As there are no definitive markers for mesenchymal stem cells, a

combination of phenotypic characteristics is usually referenced. For example, TSPCs are positive

for CD-44, CD-73, CD-90, CD-105, CD-146, Stro-1, Sca-1, nucleostemin, Oct-4 and SSEA-4,

while negative for CD-18, CD-31, CD-34, CD-45, CD-106, CD-117, CD-144 and Flk-1 [37].

Molecular markers for tenocytes include tenomodulin, scleraxis and thrombospondin 4 [38, 39].

Pericytes may contribute to primary mixed cell populations derived from tendon and have

significant phenotypic overlap with TSPCs, though they express 3G5, NG2, ALP, CD-146,

PDGF-, and -SMA [40]. All of these markers may drift over time with increasing numbers of

cell divisions during cell culture [41] and donor age [42]. Tenocytes, TSPCs and pericytes likely

all contribute to tendon damage propagation and healing.

The behavior of tendon and other materials is frequently represented in a stress/strain

curve, which plots elongation versus force per cross-sectional area [43]. The classical response

demonstrates an initial toe region, a linear-elastic region, a plastic deformation region, and a

point of failure. This model fits the deformation properties of ligament and tendon, and

parameters of this relationship change during injury and aging [44, 45], ultimately altering

cellular behavior [46]. As a complex viscoelastic biomaterial, tendons also exhibit force-

relaxation, creep, and hysteresis [47]. Hysteresis is the dependence of a tissue’s mechanical

properties on its deformation history, such as occurs under repetitive loading. The area of a

hysteresis loop represents energy loss in the form of heat during repetitive loading: greater

hysteresis means decreased efficiency of locomotion [48].

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There are three principal modes of tendon elongation. Under low strain, periodic bends,

called crimps [49], within collagen fibers straighten [50], distributing tension without resulting in

major structural changes [51]. This is the toe region of the stress/strain curve, an example of

which is later presented in Figure 3.6. Molecular extension and within-fiber sliding are the

fundamental mechanisms of linear-elastic deformation in tendon, giving rise to the distinctive

biomechanical properties of the tissue and acting on the cellular scale (15-400µm) [31, 52-54]. In

response to persistent strain, load relaxation occurs via between-fiber sliding, reducing protective

crimp which can result in damage [24, 55]. In response to high strain, tendons deform by

secondary-scale elongation: plastic sliding of antiparallel fascicles [51, 56]. Molecular damage

begins to accumulate when the capacities of these three modes are exceeded, and basic

biomechanical deficiencies precede major damage [55]. In acute circumstances of rapid

elongation, fibers may catastrophically tear [57].

THE PATHOGENESIS OF DEGENERATIVE TENDINOPATHY

Tendon disease is a significant public health problem for working animals and humans.

Damaged tendons exhibit ECM disorganization and loss of elasticity, hypercellularity, increased

vascularity, inflammation and pain [58]. 14% of Thoroughbred racehorses are retired due to

tendinopathies [59], with an incidence of approximately 1:1000 starts [60]. Spontaneous Achilles

tendon rupture occurs in humans at an incidence of 11.33:100,000 per year and is most common

among males in the third decade of life [61]. Despite the evolutionary importance of distance

running [62], 50% of modern runners suffer from exercise-induced musculoskeletal injuries

annually, with 25% injured at any given time [63]. Risk factors include heavy use during manual

5

labor or athletic activity, anatomical misalignments and comorbid health conditions [19, 64, 65].

Tendon injuries necessitate compensation of energy loss by associated muscles [66] and hinder

proper movement biomechanics organism-wide [67], leading to further musculoskeletal damage.

Even healed tendon lacks full function, and is prone to re-injury. For example, approximately 1:5

surgical rotator cuff tendon repairs fail within 6 months [68].

Tendinopathy, often considered a preventable exercise-induced injury [69], can be

viewed as a continuum encompassing acute and chronic forms of structural degeneration [70].

Until recently most tendon overuse injuries were categorized predominantly as inflammatory

disorders [71], hence the outdated terminology, “tendinitis” [72]. It is now understood that the

inflammation characteristic of tendon damage is just once factor in the disease process [73], and

the utility of anti-inflammatory treatment is still debated [74-76]. Despite the high morbidity of

tendon dysfunction and its debilitating effects on quality of life, relatively little is known about

its etiological mechanisms. The most prominent hypotheses involve the progressive

accumulation of structural microdamage, the buildup of high temperature during exercise, and

the interaction of these two factors with the pathological behavior of resident cells.

It is widely accepted that tendinopathies develop via the buildup of microscale ECM

defects [77] that are either too prevalent or too severe to be fixed by innate cellular repair

mechanisms [78]. Tendons maintain a characteristically low cell population, making the tissue

slow to adapt to damage or rapid changes in use. Excessive loading may contribute to

tendinopathy by activating prostaglandin E2-mediated pathways, reducing tendon stem cell

population size and encouraging differentiation of TSPCs into non-tenocyte lineages [79, 80].

Conversely, the loss of cellular mechanical stimulus resulting from localized breakages in ECM

induces a catabolic response that contributes to disease progression [81, 82]. Thus, once intrinsic

6

repair mechanisms are exhausted, there is a positive feedback loop whereby overuse induces

ECM damage that reduces tissue mechanical function and induces improper tendon cell function.

Hypoxia and angiogenesis also play a role in the progression of tendinopathy [78].

Hyperthermia can severely damage tissue [83]. Increased temperature occurs within

viscoelastic materials as energy is released during repeated loading, and hyperthermia is a likely

contributor to degenerative changes observed in tendinopathies. While the hyperthermic origin

theory has not been thoroughly studied, it is nevertheless a likely contributor to chronic

tendinopathies and will be discussed in detail here. Collagen unfolding is a temperature-

dependent irreversible rate process that is accelerated by heat, the result of the properties of a 66-

residue thermally labile domain near the N-terminus [84]. Conformational freedom within this

domain in response to heat and mechanical overload may make tendon ECM susceptible to

proteolysis and alter mechanical signals experienced by resident cells. Localized, transient

unfolding events are believed to commonly occur in collagenous tissues [85], though these

structural changes are typically rectified by intrinsic cellular repair mechanisms [86].

Calorimetric data suggests that the increased conformational freedom of heated collagen

alters the lattice structure but does not fully denature the matrix [57]. However, the synergy of

both heat and strain results in residual elongation that can be understood in a rheological model

of the tissue [87, 88]. While tendon stiffness has not been found to be significantly influenced by

temperature between the range of 20-41C [89], this is only one facet of a complex tissue

response. For example, mechanical deformation has steric influences on small-molecule

diffusion and the configurational entropy of collagen. Strain decreases the interaxial spacing

within collagen fibers, stabilizing them [90] but decreasing the space available for unbound

7

water molecules [91]. Alterations in water content may interfere with adequate heat dissipation

and exacerbate the resulting mechanical damage.

The thermodynamic properties of pathological tendon samples present alterations versus

their healthy counterparts. One study found that degenerate tendon samples melted at the same

temperature as matched control tissue (70°C), but a decrease in enthalpy and increased water

content was found within the lesion [92]. Even extreme transient temperatures in vivo [93, 94] do

not approach denaturation temperatures (65C) [95, 96], though above 45C there is a reduction

in elastic modulus and the first signs of irreversibility begin to emerge [97]. Since friction from

adjacent tissues increases hysteresis values recorded in live animals, localized hyperthermic

conditions approaching this transition point in theoretical models may tip over the threshold for

ECM damage in an animal [47]. The onset of mechanically-induced structural damage may

decrease collagen melting point by 3-5C [57], accelerating ECM deterioration.

In addition to its alteration of basic connective tissue biomechanics, prolonged exposure

to hyperthermic conditions is a known cause of cell death. In vivo temperature readings up to 43-

45C correlate with a threshold of tendon cell death in vitro [93, 98]. Although tendon cells are

more resistant to heat than cells derived from other tissues, there is still a steep drop in

survivability after only 10 minutes of exposure to 45-48C [99]. While temperatures in vivo are

likely insufficient to kill cells, hyperthermia may alter metabolism of ECM proteins. For

example, tendon cells heated to 40C for 30 minutes showed a three-fold increase in TNF-

mediated proMMP-9 synthesis, which returned to normal levels after cooling [18]. Another study

using a rodent overuse model documented upregulation of heat shock proteins and apoptotic

mediators in hyperthermic conditions [100]. In addition to heat, apoptosis can be initiated by

8

strain [101], and has been correlated with tendinopathy [102], under which circumstances there

is rapid cell death, accompanied by a slow subsequent die-off phase [103].

MESENCHYMAL STEM CELLS AND THE STEM CELL NICHE

Stem cells are present in nearly every adult tissue, including sources as diverse as liver

[104], synovium [105] and brain [106]. While the full diversity of cell lineages emerges before

birth, these tissue-resident stem cells participate in tissue renewal and are activated in response to

injury [107, 108]. Due to their tremendous importance to the adult organism, stem cells are

confined to specialized physiochemical milieus that regulate proliferative activity and preserve

characteristics of “stemness” [109]. As a result of this regulation, stem cells have a younger

“biological age” than their host tissues, representing an intermediate between embryonic and

adult stages [110]. Unfortunately, the population size of stem cells declines with age, an effect

correlated with degeneration of bone [111], tendon [112] and other tissues late in life.

Mesenchymal stem cells (MSCs) are multipotent stem cells within the mesenchymal

hierarchy [113]. First identified as the non-hematopoietic stem cells in the bone marrow stroma,

MSCs are capable of undergoing discrete hierarchical differentiation: from a naïve state through

committed progenitor cells to terminally-differentiated cells [114]. MSCs are present in all

musculoskeletal tissues [115] and have similar properties in different anatomical locations [116].

Fibroblasts and MSCs represent a large and diverse family of cells that are difficult to distinguish

phenotypically, with specializations lying on a continuum that is largely defined by extracellular

niche, historically leading to confusion in the literature [117]. Perivascular stem cells (pericytes)

are distributed throughout the body [118] and may or may not be distinct from tissue-specific

9

MSCs [119]. While their true definition remains nebulous [113], MSCs are minimally defined in

vitro by their self-renewal, plastic adherence (due to high expression of integrins and the

adhesive glycoproteins CD44H and CD54 [120]) and trilineage differentiation capacity (into

osteogenic, chondrogenic and adipogenic cells) [121]. MSCs by definition express CD105,

CD73 and CD90 while being negative for a panel of hematopoietic cell surface markers [122].

While MSC lines are not immortal, their very long culture lives can be extended with

overexpression of telomerase reverse transcriptase [123]. However, MSCs can be expanded a

billion-fold without manipulation and safely withstand cryopreservation [110]. Nevertheless,

early-passage cells are more desirable for therapeutic use due to senescence [124], reduced

immunomodulatory properties [125] and an increased likelihood of chromosomal aberrations

increasing risk of malignancy [126] occurring after extended laboratory culture.

The stem cell niche is a distinct environment that regulates stem cell activity in vivo

[127]. These anatomical locations have been described in detail for many highly proliferative

tissue types, including epithelial stem cell niches within small-intestinal crypts [128] and the

outer root sheath bulge in skin [129]. Tendon stem cells first began to be characterized in the

early 2000s following the discovery of a population of highly proliferative cell lines exhibiting

developmental plasticity [130] and universal mesenchymal stem cell features across species

barriers [131]. Unsurprisingly, ECM proteins such as biglycan and fibromodulin play an

important role in the stem cell niche [34]. Dysregulation of stem cells within tendon, in part due

to accumulating ECM damage , may result in altered differentiation and contribute to the disease

process [132]. Mimicking the stem cell niche in vitro is critical to prevent loss of phenotype and

to improve long-term maintenance of healthy tendon cell populations [133, 134].

10

REGENERATIVE MEDICINE: CELL THERAPY AND TISSUE ENGINEERING

There has long been a desire to fully regenerate damaged body parts: this scientific field

is called regenerative medicine. In evolutionary exchange for reduced incidence of neoplasia,

mammals undergo epigenetic lockdown of genetic developmental programs, thus preventing the

extreme regeneration seen in more basic vertebrates [135, 136]. However, the more naïve state of

adult MSCs can potentially be harnessed medicinally for tissue repair and the prevention of non-

functional fibrosis. Stem cells are largely responsible for the therapeutic efficacy of several

widespread 20th

century medical procedures including bone marrow transplantation and skin

grafting [137-140]. More recently, the developmental plasticity of adult cells has been exploited

to encourage direct and endogenous tissue healing [141]. MSCs readily differentiate into tissues

other than their source with minimal persuasion [142]. In addition to their ability to form

multiple tissue types [143], MSCs are anti-inflammatory and immunomodulatory [144]. MSCs

have been used in humans to treat a diverse spectrum of conditions including graft-versus-host

disease [145], myocardial infarction [146], and congenital skeletal defects [147].

The use of cell therapy to repair tendon [148] has been explored since stable multi-

lineage differentiation protocols were first established [143]. MSC therapy in the horse has a

track record over a decade long, having been used to treat tendon lesions, since 2003 [149]. The

most common site of tendon injury and therefore cell therapy is the flexor digitorum superficialis

tendon (FDST), commonly known as the superficial digital flexor tendon. Current data indicates

that MSCs are safe [150], integrate into the site of injury [151, 152], improve collagen

organization [153] and encourage a healthy gene expression profile including type-I collagen and

cartilage oligomeric protein [154, 155]. While many mechanistic questions remain, early data on

11

MSC transplantation supports its use to treat FDST injuries, particularly in the absence of

effective pharmacological therapies. Advances in veterinary cell therapy have provided evidence

of safety and efficacy [156] that are beginning to be translated to human medicine [157, 158].

There are other cell types competing with MSCs for the limelight in tendon cell therapy.

Embryonic stem cells (ESCs) are pluripotent cells derived from the blastocyst inner cell mass in

pre-implantation embryos. The enormous therapeutic potential of ESCs was recognized from the

time they were first isolated [159, 160], and while ESCs have been used to treat tendon lesions

with short-term safety [152, 161] ethical controversy [162, 163] and tumorigenic danger [164]

prevents their widespread use. Personalized ESCs can be derived from adult patients using

somatic cell nuclear transfer, also known as therapeutic cloning [165-167], but the efficiency of

this process is variable, influenced by the epigenetic state of the donor nucleus [168]. It is

possible to use xenogenic oocytes as recipient cells for this technique [169], though this does not

circumvent its technical difficulty or ethical criticism [170]. Induced pluripotent stem cells

(iPSCs) produced by overexpression of the Yamanaka factors Oct4, Sox2, Klf4 and c-Myc in

adult cells possess embryonic-like properties [171] and can be used to treat tendon lesions [172],

but still contain epigenetic memory of their tissue of origin [173]. In contrast to MSCs, iPSCs

differentiate less readily [174], are potentially immunogenic [175] and if not properly controlled

can aggressively form teratomas [176]. MSCs therefore represent a better balance of regenerative

utility, availability, ease of use and ethical justification for regenerative medicine at this time.

Tissue engineering emerged as a new interdisciplinary field in in the 1980’s, when

scientists, engineers and clinicians began to combine cells, scaffold material and environmental

stimuli with the aim of producing replacement tissue in vitro [177]. This approach has brought

tremendous advances in the reconstruction of simple tissues such as bladder [178] and trachea

12

[179]. This dissertation brings closer the goal of engineering replacement tendon with ideal

cellular, ultrastructural and biomechanical properties [180]. A thorough outline of tissue

engineering applications relevant to tendon and ligament are reviewed in the next chapter.

MECHANOBIOLOGY: DIFFERENTIATION AND PHENOTYPE

Cells sense mechanical stimulation and ECM damage, integrating environmental signals

to alter function (Figure 1.1) [181]. Tendon cells maintain a homeostatic cytoskeletal tension,

which forms a reference point for perception of mechanical signals transferred through the ECM

[182]. Tendon cells extend projections deep into this surrounding matrix [29] and the properties

of cells and ECM are dynamically interdependent [183]. Cell-ECM communication is mediated

by a number of mechanotransduction mechanisms. Five pathways specifically implicated in

tendon cell fate decisions are kinase-associated integrin pathways, direct cytoskeletal coupling,

force-sensitive ion exchange, growth factor signaling and primary ciliary transport. Scaffold

architecture, alignment and deformation influence these processes, linking extracellular stimuli

to changes in MSC morphology and differentiation [184].

Integrins are a class of transmembrane heterodimeric proteins that form complexes

linking the ECM to the cytoplasm. As such they facilitate a broad range of intracellular responses

to environmental mechanical signals, including strain. Integrins are concentrated in large

membrane-associated protein assemblies known as focal adhesions, and serve as major centers of

proprioception and signal integration [185]. The intracellular domain of the integrin -subunit

mechanically couples to a number of actin-binding proteins, including talin, -actinin and

filamin [186]. Vinculin is an essential intracellular focal adhesion regulator that bears force [187,

13

188]. Direct cytoskeletal coupling via vinculin and other integrin-associated structural proteins

results in stress waves that propagate mechanical stimuli to the nucleus on the millisecond scale.

This may alter gene expression through several mechanisms including chromatin recombination,

distortion of the nuclear matrix and modified nuclear protein translocation [189]. In contrast,

classical integrin signaling cascades are diffusion-limited and occur on the time scale of several

seconds. A central regulator of this method of transduction is the pseudokinase, integrin-linked

kinase (ILK). ILK, through its association with the accessory proteins PINCH and parvin [190,

191], passes integrin stimuli through signaling intermediates via phosphorylation of the

serine/threonine protein kinases, glycogen synthase kinase 3 (GSK-3) and protein kinase B

(PKB) [192]. These kinases regulate diverse cellular behaviors including multipotency, self-

renewal, differentiation and apoptosis [193, 194]. As more than 20 different molecules are

known to directly complex with intracellular integrin domains during cell-ECM adhesion [186],

integrins serve as central participants in many diverse mechanotransduction functions.

Receptor tyrosine kinases (RTKs), such as discoidin domain receptor 2 (DDR2) in

mesenchymal cells [195], bind growth and survival factors. RTKs activate Ras-family pathways

[196], attributed primarily to proliferation and differentiation functions. The wingless/integrase-1

(Wnt) is a highly evolutionarily conserved pathway that is also implicated in cell polarity, self-

renewal and differentiation [197]. Wnt ligands bind to the Frizzled family of G protein-coupled

receptors (GPCRs), and canonical β-catenin accumulation shares pathway overlap with integrins

[198]. The Wnt pathway is a common regulatory target implicated in most stem cell niche

environments [199]. If uninhibited by ILK, GSK-3 positively regulates expression of the cell

cycle protein cyclin D1 and the ECM turnover protease MMP-9 through the intermediates,

cAMP response element-binding protein (CREB) and activator protein-1 (AP-1) [192].

14

Figure 1.1 – Overview of integrin signaling. Integrin-mediated mechanobiological signals are

translated from ECM to the nucleus via diffusion and stress wave propagation.

Wang, N, JD Tytell and DE Ingber. (2009). Mechanotransduction at a distance: mechanically

coupling the extracellular matrix with the nucleus. Nature Rev Mol Cell Bio 10:75-82. Used

with permission of Nature Publishing Group, 2015.

c-Jun N-terminal kinase (JNK) 1 and 2 are constitutively expressed downstream mitogen-

activated kinases (MAPKs) that are responsive to stress. JNKs, which function downstream of

integrins, RTKs and GPCRs, control cell survival and programmed death depending on context

[200-202]. These kinases are increasingly phosphorylated in early tendinopathy and hypoxia

[203], are upregulated in tendon cells in response to cyclic strain in an amplitude- but not

frequency-dependent manner, and are mediated by calcium, a critical secondary messenger of

15

mechanotransduction [204]. Ca2+

ions may arrive as biproducts of another cascade or by calcium

ion influx through the cell membrane, since human tenocytes express both voltage-operated

calcium channels and the stretch-activated potassium channel, TREK-1 [205]. Thus,

depolarization or raw mechanical stimulation may alter MAPK activity by action of increased

intracellular cation concentrations. Calcium waves are further propagated through adjoining

tenocyte populations via connexin 32, increasing collagen expression [206].

Aside from the direct and indirect influence of mechanical stimulation on integrin-

associated signaling cascades and mechanosensitive ion channels, transforming growth factor

(TGF-), a cytokine implicated in cell cycle control, differentiation and matrix metalloproteinase

expression [207], plays a complicated and poorly-understand role in tendon development,

including by increasing scleraxis expression [208], likely through a Smad 2/3 pathway [209].

Smad pathways are particularly important in integrating pro- and anti-tenogenic stimuli in naïve

MSCs [210]. Scleraxis associates with tendon-specific element 2 of the procollagen Col1A1

promoter to induce collagen-I expression [5, 209], and is an early and sustained marker of tendon

tissue [211]. Scleraxis expression continues during tendon tissue homeostasis, and is further

upregulated in response to exercise adaptation [212]. TGF- also provokes activation of the Wnt

pathway by inactivating GSK-3 [213], and regulates tenocyte contraction [214].

Finally, each tenocyte also possesses a single cilium [215], oriented approximately

parallel to the ECM axis [216], providing yet another method of gathering mechanical

information. Cilia are narrow projections involved in cell cycle control and differentiation of

MSCs [217]. Cilia house an array of sensory elements including RTKs that interact with various

components of the ECM [218]. Aligned surface topography has been found to decrease

intracellular actin-myosin tension, increase cilia prevalence and length, and suppress canonical

16

Wnt signaling [219]. As non-canonical Wnt signaling serves as a switch toward differentiation,

subcellular compartmentalization of β-catenin within the ciliary body, sequestered by

intraflagellar transport protein 88 (IFT88), ties scaffold alignment with MSC differentiation

[220]. IFT88 thereby mediates early lineage commitment in MSCs, and its knockdown is

associated with decreased differentiation [221] and increased proliferation [222]. Ciliary length

is correlated with the establishment of baseline cytoskeletal tension in tendon cells [223, 224],

and deflect in response to strain [225]. The precise mechanisms by which primary cilia regulate

MSC and tendon cell phenotype will likely be fruitful topics of future investigation.

17

CHAPTER 2: MODEL SYSTEMS IN MUSCULOSKELETAL CELL BIOLOGY

ANIMAL MODELS OF TENDINOPATHY

An ideal animal model captures the full disease process of tendinopathy, including

clinical/imaging features, histopathological organization and loss of function with pain and gait

abnormalities [226]. Unfortunately, no single model will fit all injury conditions, with variables

including function and insertion type (tendon/ligament), lubrication and vascular supply

(intrasynovial/extrasynovial), and severity of the condition (partial/complete/critical) [227]. In

vivo models provide a more holistic picture than reductionist systems including the interaction

with surrounding tissues, but complicate basic mechanistic studies while requiring the sacrifice

of research animals [228]. Current animal models of tendinopathy are generally created using

one of three methods: 1) forced overuse, 2) surgical induction or 3) intratendinous injection.

Several models have been based on the etiological hypothesis that most tendon injuries

result from overuse. One hour of treadmill running per day on a 10° decline induces gross,

histological and biomechanical alterations consistent with supraspinatus tendinopathy in rats

[229]. This procedure results in downregulation of tenocytic genes and upregulation of

chondrocytic genes [230], which is reversible with two weeks of rest [231]. Soslowsky’s model

has led to a number of important findings including the involvement of early and sustained

angiogenesis and inflammation [232], nitric oxide [233], glutamate signaling [234], upregulated

heat shock proteins [100] and insulin-like growth factor 1 (IGF-1) [235] and GAG sequestration

of heparin affinity regulatory peptide (HARP) [236] in the early disease process. Uphill treadmill

running was similarly found to target the Achilles tendon [237]. A competing technique

18

involving cyclic loading of flexor muscles by direct electrode stimulation has been used with

some success in a rabbit model [77, 238-240]. This approach allows enhanced control of loading

in a single target but may not match the deformation patterns that would naturally occur.

Tendon lesions are commonly created surgically, either to model an acute condition [241,

242] or the combination of chronic and acute injury [243]. Window defects or full-thickness

tenotomies have been used to evaluate intra- and post-operative treatments including sutures

coated with transforming growth factor-β (TGF-β) [244], gene therapy with bone morphogenetic

proteins BMP-12 [245] and BMP-14 [246], the use of cell-laden hydrogels [148, 247-249] and

stem cells overexpressing scleraxis [250, 251], among many other strategies. Surgical tendon

lesions are a common testing ground for tissue engineering techniques that will be discussed in

greater detail elsewhere in this mini-review. Repaired laceration models have also been used to

examine the influence of frequently comorbid conditions in humans. For example, type-II

diabetes has significant negative effects on histological and biomechanical outcomes in the

healing rat supraspinatus tendon [252]. Sub-failure strain results in a different damage profile

and reparative response than laceration, and can also be induced surgically [253]. This technique

has revealed cycle-dependent changes in biomechanical properties [55] and immediate, strain-

dependent upregulation of interleukin-1 β (IL-1β) and matrix metalloproteinase 13 (MMP-13)

[254]. In situ freezing as also been used to model the “ideal” acellular autologous graft [255].

Agents that induce extracellular matrix (ECM) degeneration, inflammation or cell death

may be delivered via intratendinous injection. Bacterial collagenase has been used to model the

collagen degeneration characteristic of tendinopathy in several organisms including horses [256],

rats [257] and rabbits [258]. These injuries have then been used to track biomarkers of the

healing process [259] and characterize novel treatment methods such as growth factor [260] or

19

stem cell [153] supplementation. Corroborating overuse models, non-tenocytic phenotypes have

been observed in collagenase-induced lesions, including ectopic calcification and expression of

SOX9 as well as type-II and type-X collagen [261]. ECM damage can also be induced by

cytokines [262], prostaglandins [263], and fluoroquinolones [264].

Genetic engineering techniques have helped to elucidate developmental cues leading to

healthy tenogenesis [208, 265] and the interdependency of ECM molecules during fibril

assembly [266]. Genetic predisposition to tendon disorders has long been recognized [267], and

polymorphisms in tenascin-C [268] and type-V collagen [269] have been associated with

Achilles tendinopathy. In the future we are likely to see techniques such as conditional

knockouts and RNA interference used more frequently to model tendon pathologies in animals.

Beyond these techniques, tendinopathies naturally occur in some species, most notably

involving the superficial digital flexor tendon in horses [270] and the supraspinatus tendon in

dogs [271]. Patient to patient heterogeneity and an inability to control disease stage and

progression complicate the use of natural models [272]. However, veterinary patients may best

simulate the human condition, and additional animal lives need not be expended in the process.

CURRENT STATUS OF BIOMIMETIC SCAFFOLDS AND CULTURE SYSTEMS

The realm of soft tissue bioreactors has remained relatively unexplored. Even rarer are

systems incorporating naturally-derived scaffolds, despite the long history of tendon allografts in

clinical practice. A PubMed.gov literature search conducted on December 3, 2014 for

“(decellularized OR acellular) AND tendon” in all fields yielded 159 results with full texts

available in the English language. 6 of these articles were reviews. One hundred and six articles

20

were excluded as being unrelated, the most common reason being false hits on studies involving

acellular dermal matrix for tendon reconstructive surgery (51 articles). The remainders were

sorted as pertaining principally to either the development or application of cell-free tendon

scaffolds.

Twenty-six articles recovered in this search relate to applications of tendon scaffolds or

graft materials. These results are confounded with a boom of in vivo allograft studies in the

1980s utilizing pretreatment methods that are difficult to characterize into discrete categories.

Another 18 articles were specifically classified as methodological papers, meaning that

development of a novel cell-removal protocol was either the primary goal of the study or a

necessary prerequisite included in the methods section. The most common model organisms

among this group are rabbit (5 articles) and pig (4 articles), and the most common

decellularization methods are based on detergent treatment (12 articles) or freeze/thaw cycling (4

articles).

Three laboratories have experimented with homogenized tendon ECM as a supplement to

monolayer- or hydrogel-based cultivation methods. However, disrupting tendon ultrastructure in

this manner, while useful in its own right, precludes the use of the resulting biomaterial for

grafting: an important downstream application that is currently in demand.

In the context of tissue engineering, the term “bioreactor” describes any (typically in

vitro) culture system that not only sustains the life of cells/tissues, but enriches the environment

with dynamic stimuli designed to promote a particular phenotype. A PubMed.gov literature

search conducted on December 3, 2014 for “(tendon OR ligament) AND bioreactor” in all fields

yielded 92 results with full texts available in the English language. To put this result in

perspective, corresponding searches for cartilage and bone yielded 456 and 754 results,

21

respectively. Nineteen of the 92 articles were reviews or protocols, and the rest were excluded as

relating to ex vivo characterization (6 articles) or unrelated tissues (13 articles).

The three-dimensional tendon/ligament scaffolds identified in this search can be divided

into two groups: those that were subjected to mechanical stimulation and that attempt to mimic

the anisotropic alignment and mechanical properties of native tendon (26 articles), and those that

do not (28 articles). Among the former, the most common scaffold materials are synthetic

polyesters (emerging in 2004, 10 articles), silk fibroin (emerging in 2002, 5 articles), and

decellularized tendon (emerging in 2010, 4 articles). Constructs based on decellularized tendon

scaffolds were seeded with allogeneic tendon stem/progenitor cells (3 articles, rabbit or chicken)

or dermal fibroblasts (1 article, human). Whitlock, et al. 2013 is the only one of these studies to

examine outcomes outside of microscopy and tissue mechanical properties.

There are certainly many relevant articles that did not meet the search criteria due to

atypical descriptors or keywords used by the authors: these articles are still included in the

following mini-review to the best of the author’s ability. However, the above summary

represents an accurate cross-section of the research in tendon-derived scaffolds and their

application in bioreactor systems. It is clear that the relationship of tendon/ligament cell biology

to three-dimensional environment and mechanical stimulation remains critically understudied,

despite the prevalence and severity of tendinopathies. This remains a significant shortcoming in

hypothesis-driven research and development of cell- and tissue-based therapeutic modalities, and

holds back the potential for clinical translation.

22

REVIEW OF SCAFFOLD SYSTEMS

Tendon autotransplantation developed as a discipline in response to unprecedented

numbers of combat casualties in the wake of World War I [273], but it was not until the mid-

1950s that allografts [274] and artificial tendons [275] first entered trials. Tendon healing after

trauma is ordinarily accomplished by a combination of cells both intrinsic and extrinsic to the

tendon mid-substance [276], but it was uncertain how free tendon grafts integrated with the host,

and in turn, how that process might be improved. Tendon scaffold was first used for the purpose

of basic cell biological research in 1986, when rabbit quadriceps patellar tendon autografts were

flash-frozen in liquid nitrogen and used for anterior cruciate ligament (ACL) reconstruction in

order to demonstrate the donor origin of repopulating cells [277]. While not “decellularized”,

these constructs were devoid of live donor cells – allowing the first observations into the active

role of cells in tendon homeostasis. Infiltration by peripheral cells was found to be insufficient to

restore full biomechanical functionality. The failure strain of freeze-killed medial collateral

ligaments (MCLs) orthotopically transplanted in a rabbit model decreased by 25% versus their

fresh cell-containing counterparts nearly one year after the operation [278]. Cells quickly became

the focus of tendon reconstructive research after this discovery [279]. Achilles tendon prostheses

containing autologous MSCs dramatically enhanced gap defect healing in rabbits [247]. Since

that time, a tissue engineering approach combining cells and scaffolds has been widely explored

in effort to enhance tendon regeneration [280].

Decellularization protocols were invented in order to prevent the immunogenicity seen

following anterior cruciate ligament repairs with freeze-dried allograft of xenograft [281].

Though some aspects of collagen structure are not conserved, the primary agents responsible for

23

tendon graft rejection are donor cells [282]. Decellularization is necessary to promote an M2

decision in the “fight or fix [283]” host macrophage response [284]. The predominance of type-I

collagen in tendon ECM poses a particularly low immunological risk once cells are removed

[285]. Chloroform-methanol extraction was the first chemical decellularization technique to

achieve widespread use [286, 287]. Since that time, techniques for cell removal have been

incrementally improved, with increasing emphasis on tissue engineering.

Modern decellularization protocols most commonly apply detergents to solubilize cell

debris. Detergents can disrupt collagen banding and mechanical properties [288], so a balance

must be found in removing cells but preserving ECM. A limited number of systematic detergent

decellularization protocols exist in the literature, and all use surfactants such as sodium dodecyl

sulfate (SDS) and 4-octylphenol polyethoxylate (Triton X-100) or the organophosphorus solvent

tri(n-butyl)phosphate (TnBP). Jeffrey S. Cartmell and Michael G. Dunn compared the effects of

Triton X-100, TnBP and SDS at 0.5-2.0% concentration on rat tail tendons for 12-48 hours and

found that 1% SDS for 24 hours or 1% TnBP for 48 hours most effectively removed cells and

maintained histological and biomechanical features of normal tendon [289]. Terence Woods and

Paul F. Gratzer observed that, of two-step detergent protocols involving 48 hour incubation in

1% Triton X-100 followed by another 48 hours in 1% solutions of SDS, Triton X-100 or TnBP,

the Triton-SDS combination worked best for 300µm-thick canine Achilles tendon ribbons [290].

Interestingly, the same group discovered that a different combination (Triton-TnBP) was more

effective at porcine ACL decellularization [291]. Shuxing Xing and colleagues compared 24

hour incubations of 1% Triton X-100, 0.5% SDS, 1% TnBP, 1% Triton X-100 with 0.5% SDS,

1% TnBP with 0.5% SDS and 1% TnBP with 1% SDS to decellularize rabbit semitendinosus and

flexor digitorum tendons, and found 1% Triton X-100 with 0.5% SDS to be most effective in

24

removing cells without damaging mechanical strength [292]. Deeken, et al. compared 1% Triton

X-100, 1% Triton X-100 with 1% TnBP, 2% TnBP, 1% TnBP, 1% SDS and 2% SDS, and found

1% TnBP to be most effective at decellularizing porcine diaphragm tendon [293].

Our group was the first to use an equine tendon scaffold for tissue engineering

applications, as will be described in subsequent chapters. This model is advantageous due to the

size, availability, low vascularity and high mechanical strength of equine tendons relative to

other species. Briefly, we compared the effect of 1% TnBP, 1% SDS, 2% SDS, and 0.5% Triton

X-100 with 0.5% SDS with a detergent-free group on 400µm-thick equine flexor digitorum

superficialis tendon (FDST) ribbons, finding 2% SDS in combination with other methods

provided a nearly cell-free, biomechanically robust and biocompatible scaffold material [294].

Flexor tendon allografts needed for hand reconstruction typically fall within the range of 2-7cm

[295], and decellularized equine FDST ribbons may prove to be ideally suited for this

application. Burk, et al. later decellularized full-thickness equine FDST samples using 48 hours

of 1% Triton X-100 incubation in combination with freeze/thaw cycles [296]. On the human

side, Pridgen, et al. compared 1% Triton X-100, 1% TnBP, 1% SDS and 0.1% SDS, and 0.1%

SDS was sufficient to decellularize Achilles tendon [297]. Hammer, et al. decellularized human

iliotibial tract using 1% SDS, noting incomplete DNA removal (44.7% residual) and native

tensile properties, and incorrectly stating that theirs was the first study to compare matched

native and decelluarized tendon samples [298]. Other techniques such as freeze/thaw cycles

[299], hypotonic solutions [300], nucleases [301], oxidizing agents [302] and irradiation [303]

have also been used on tendon, alone or in combinations with detergents.

Human flexor digitorum profundus tendons, decellularized in SDS and implanted into

outbred rats, had decreased immunogenicity and improved mechanical properties versus non-

25

decellularized counterparts [304]. A similar result was seen in 2% SDS-decellularized rat

Achilles tendons [305]. Of the commercial tissue augmentation materials surgically used for

tendon reconstruction, those that are based on natural collagen matrices are stronger and better

retain sutures [306, 307]. However, the most common products, such as Graftjacket RTM

(Kinetic Concepts, Inc.), Allopatch HD (Musculoskeletal Transplant Foundation), and

TissueMend Soft Tissue Repair Matrix (Stryker) are (human, human and bovine, respectively)

dermal allografts and lack the standalone biomechanical properties of tendon. Full coverage of

non-tendinous ECM scaffolds is beyond the scope of this work, but some groups have

experimented with surface modification or reinforcement techniques to add mechanical strength

to ECM sheets [308-310], while others embrace the sacrificial nature of weak but rapidly

remodeled scaffolds as encouraging de novo tissue growth. As previously stated, tendon grafts

lacking live cells are in widespread use for ACL replacement, but do not fully restore native

function [311]. It can take three years or longer for allografts to reach peak integration, which

still remains incomplete and weak compared to healthy tissue [312]. While some groups are

experimenting with functionalization or composite techniques [313], another potential solution to

this problem is to provide a cell population that remodels and strengthens the scaffold over time.

Reseeded decellularized tendon scaffolds have demonstrated strong potential as graft

materials [314], but animal testing is required [315]. Multilayer composites of decellularized

canine infraspinatus tendons laden with rabbit bone marrow-derived MSCs remained vital, began

to express tenomodulin, and altered collagen and matrix metalloproteinase activity [316]. Rabbit

rotator cuffs repaired with matrix performed better after 8 weeks when seeded with tendon cells

[317]. Human flexor digitorum profundus tendons (FDPT) decellularized in 0.1% SDS and

peracetic acid then seeded with adipose-derived MSCs implanted subcutaneously in nude mice

26

remained healthy and viable for one month [318]. Reseeded Triton X-100-treated rat Achilles

tendons were stronger and better-organized after 24 in a surgical replacement model [319].

However, cell-laden tendon constructs cultured ex vivo without mechanical manipulation

actively degrade their scaffolds, necessitating culture techniques that fulfill the need for cells to

experience stimulation [320].

REVIEW OF BIOREACTOR SYSTEMS

In vitro cell culture was pioneered by Alexis Carrel at the turn of the 20th

century. By

nourishing tissue explants in plasma enriched with various animal-derived extracts, Carrel

extended the life expectancy and proliferative capacity of cells in culture from weeks to months

[321, 322]. Realizing that diffusion-limited nutrient flow hindered not only the size of tissue

explants but the vitality of transplant organs, Carrel, in collaboration with Charles A. Lingbergh,

co-invented “an apparatus for the culture of whole organs” [323] – the first perfusion bioreactor.

Musculoskeletal loading plays an important role in tissue homeostasis – a fact that gained

increasing attention as humans established technologies facilitating spaceflight. Astronauts

experience reversible bone demineralization [324], which is attributed to attenuated bone

formation but retained bone resorption while in orbit [325]. It was recognized that rat bone

marrow cells lose osteogenic potential when unloaded in vivo [326], but have improved

osteogenic potential when cultured on flexible-bottom culture dishes subject to 1Hz deformation

cycles [327]. As gravitational and locomotive forces have been essential evolutionary conditions,

many tissues experience similar phenomena. ECM forces and cell shape result in fate decisions

including differentiation and apoptosis [328], and transplanting cells can induce progenitor cell

27

differentiation into tissues other than their source of origin [329]. Modern musculoskeletal

bioreactors are direct decedents of these discoveries and attempt to manipulate these effects.

Deformation protocols are most efficacious when they resemble the in vivo environment [330],

as cells respond differently to deformations induced by compression versus tension [331].

Elastic deformation of tendon occurs at the 100µm level by straightening of fibrillar

crimp, and then at the 10-15nm level by molecular elongation of collagen helices [332].

Sensation of these cell-scale forces, such as through integrin-activated MAP kinase and NF-κB

signaling or by stress-responsive gene enhancers, is essential to proper tenocyte phenotype [333].

Stretch also alters the availability and rate of ECM binding domains involved in assembly and

degradation [334]. Tendons deprived of mechanical stimulation in vitro experience profound

negative alterations including changes in anatomical size, a near complete loss of biomechanical

function, hypercellularity and decreased ECM alignment [82]. This occurs even in tendons

frozen in situ [335]: a treatment which may in fact exacerbate ECM catabolism through

proteolytic enzyme release. Cells exhibit sensitivity to subtle changes in their mechanical

environment, with small deformations frequently leading to an anabolic and anti-inflammatory

response, and large deformations leading to inflammation and ECM damage [336]. Selectivity to

static versus dynamic forces of the same magnitude is also provided by differences in resistance

to conformational change among components of intracellular focal adhesion complexes [188].

While other types of bioreactors such as microcarriers [337, 338], flow perfusion and shear

systems [339-342] and hydrostatic bioreactors [343] exist, this mini-review focuses on systems

involving stretching or mechanical load that attempt to model natural deformation.

Albert J. Banes made pioneering advances in the musculoskeletal mechanobiology field

with his invention of the Flexcell bioreactor platform in 1985 [344]. This system uses flexible-

28

bottom circular culture wells deformed via vacuum to deliver controlled mechanical signals to

cells, allowing mechanistic in vitro studies of tendon and ligament signaling [345]. For example,

ligaments undergo atrophy when unloaded, as sensed via fibronectin-specific integrin α5β1 and

other mechanisms [346]. Cell stretching not only encourages tissue anabolism, but also results in

cell-mediated ECM recomposition. Primary human MCL cells increasingly express type-III

collagen under 7.5% but not 5% strain in a Flexcell bioreactor at 1Hz for 16-25 hours [347]. This

effect is not observed in cells derived from synovium [348]. Human tendon fibroblasts secrete

the growth factors TGF-β, bFGF and PDGF in response to stretch on the timescale of hours

[349]. Flexcell bioreactors can be programmed to conform to virtually any waveform desired,

but the resulting deformation is only precise in the center of each well [350], even after careful

calibration [351]. A slightly more uniform approach is to use a uniaxial stretch system to deform

rectangular silicon dishes using a stepper motor, the use of which demonstrated TGF-β1-

mediated expression of type-I and type-III collagen in human ACL fibroblasts [352].

Scaffold-free flexible plastic may be considered the first generation of tendon/ligament

bioreactors. While they represent a tremendous increase in complexity versus traditional

monolayer culture, they are far from modeling the in vivo environment (Table 2.1). Second

generation bioreactors are based on three-dimensional scaffolds, but do not mimic the alignment

or function of native tendon. As the principle component of tendons and ligaments, type-I

collagen is the most common base, but the lack of hierarchical organization makes them

structurally inferior for translational use [353]. Tendon cells do, however, interact with collagen

gels, resulting in changes in phenotype [354]. David L. Butler’s group has developed

functionalized type-I collagen-based sponge constructs for tendon repair that can withstand

forces experienced in rabbits [355-357]. Rabbit bone marrow-derived MSCs embedded in these

29

sponges established maximum stiffness when cultured at 2.4% strain at 1Hz for 50 minutes per

day [358]. Human tenocytes cultured in reconstituted rat tail collagen stretched at 5% strain at

1Hz for 48 hours resulted in differential expression of several proteases and matrix proteins, as

well as TGFβ activation [359]. Many natural but isotropic materials used for surgical

augmentation are occasionally used in bioreactors, including porcine small intestine submucosa

[360] and human umbilical veins [361], as well as designer scaffolds such as the woven

hyaluronic acid-based Hyalonect [362]. Biaxial deformation is possible in these bioreactors

[363], but multi-axial strain is not typically applicable to tendon physiology.

Table 2.1 – Tendon bioreactors by generation. Attributes of tendon tissue culture systems sorted.

Synthetic hydrogels are also commonly implemented, and their properties are tunable

[364]. A commercial human MSC line encapsulated in poly(ethylene glycol) (PEG)-based

scaffolds under 10% strain at 1Hz for alternative 3 hour on/off periods for 21 days resulted in

upregulation of tenocytic genes including collagens and tenascin-C [365]. Rabbit Achilles tendon

cells in porous poly(L-lactide-co-ε-caprolactone) (PLCL) scaffolds under 10% strain at 0.25Hz

for 400 minutes per day for 4 weeks demonstrated enhanced proliferation and type-I collagen

deposition [366]. 3-D culture is critical for proper morphology, but disordered hydrogels do not

deform via the same mechanisms of native ECM. Hydrogels contract and remodel, and may

begin to establish alignment, but do not resemble tendon [367, 368]. Cultured MSC sheets

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deposit ECM on plastic, which can also be used as a simple matrix. One study using such a

system discovered that both mechanical forces and scleraxis can independently induce

tenogenesis, but their influences are synergistic and most effective when combined [369]. This

strategy falls into the same technical class as hydrogels: useful but not biomimetic. Nevertheless,

fundamental signaling information, such as the nature of the refractory period following periods

of mechanical stimulation [370], are likely conserved. However, heterogeneity in the structural

properties of different systems complicates comparisons, such as cell-scale felt strain.

Third-generation bioreactors implement aligned scaffolds, natural or synthetic, and more

faithfully recapitulate tendon structure and alignment, but are not biomechanically functional as

standalone replacement tendons. Rat MCL cells aligned on collagen-coated

polydimethylsiloxane (PDMS) scaffolds demonstrated increased cell-cell calcium signaling

sensitivity versus non-aligned controls [371]. Another option is to use synthetic

nano/microfibers. These have better structural similarity to tendon, induce spindle morphology,

and when sized correctly provide structural cues that aid in tenogenesis [372]. Human ligament

fibroblasts cultured on electrospun polyurethane nanofibers increased ECM production in

response to 5% strain at 0.2Hz in a modified Flexcell bioreactor [373]. Cardwell, et al. cultured

C3H10T1/2 MSCs on electrospun poly(ester urethane) urea fibers, loaded daily by 4% strain at

0.5Hz for 30 minutes, witnessing alignment and a tenocytic gene expression profile [374]. Silk

scaffolds are widely used by David L. Kaplan and his collaborators as a platform for MSC

differentiation toward ligament [375]. Aligned, cross-linked collagen fibers most closely

approximate the structural and mechanical properties of native tendon. In a recent study by Qiu,

et al., human bone marrow-derived MSCs cultured under 10% strain at 1Hz for alternating 3

31

hour on/off cycles for 14 days proliferated more and expressed greater amounts of scleraxis,

tenascin-C and collagens than their static counterparts [376].

The most holistic tendon bioreactors use decellularized tendon matrix as a scaffold

subject to cyclic strain. These are the fourth generation tendon bioreactors, which not only

replicate the mechanical environment of native tendon, but also the complex ultrastructure,

composition and biomechanical properties. There are currently only four principle investigators

using this technique: James Chang of Stanford University, Mark E. Van Dyke of Wake Forest

University, Jennifer G. Barrett at Virginia Tech and Chunfeng Zhao at Mayo Clinic (Table 2.2).

Chang’s group released back-to-back Tissue Engineering: Part A papers in 2010, characterizing

the response of rabbit and human flexor tendon constructs cultured in Ligagen L30-series axial

bioreactors. P1 rabbit FDPT-derived cells cultured on decellularized FDPT increased construct

elastic modulus and ultimate tensile strength in response to cyclic 1.25N strains at 0.0167Hz for

alternating one-hour periods over 5 days [377]. In a similar study with matched human FDST

and EDST sets (P4) with 0.625N-2.5N strains over 3-8 days, the investigators observed time-

dependent improvements in biomechanical properties, but no differences between groups of

different load magnitudes [378]. Constructs then degraded after two days of disuse. Van Dyke’s

group released a 2013 paper using allogeneic chicken FDPT constructs (P4) exposed to 5% strain

at 1Hz for one hour per day for 7 days in the same Ligagen system. Cells preserved construct

mechanical properties versus unseeded controls, but no significant changes in mRNA profiles

were seen resulting from strain [379]. Our group used a custom bioreactor to characterize

amplitude-dependent gene expression profiles of P2 horse bone marrow MSCs, finding that 3%

strain at 0.33Hz for one hour per day increased ultimate tensile strength and induced high

expression of scleraxis, type-I collagen and proteoglycans [294]. Zhao’s group released the

32

results of a similar study after our 2015 Journal of Orthopaedic Research article was published,

reaching analogous conclusions in a canine Achilles tendon model [380]. Most recently, we

followed up with a comparison of the relative tenogenic efficacies of MSCs derived from bone

marrow, adipose and FDST, finding tendon-derived cells to have optimal differentiation and

remodeling performance [381]. The full results of our studies are contained in Chapters 4-5 of

this dissertation.

Table 2.2 – Summary of generation-4 bioreactors based on cell-laden DTS.

Earlier bioreactor designs have greatly expanded our knowledge of mechanotransduction.

However, fourth generation bioreactors possess even higher clinical relevance, including faithful

cell-scale force translation and scaffold material properties that match natural tissue. Within this

class of bioreactors, of all model organisms currently in use, equine tendon is ideal due to its

structural similarity to human tissue, its high performance characteristics and the relative size

and availability of donor material. Our bioreactor is based on relatively inexpensive modular

hardware, accommodates samples of any size simultaneously, gathers individual load data, can

33

be sterilized, and is scalable. We therefore believe that our model is ideal as a basic model of cell

biology, as a method for priming therapeutic cells and for cultivating replacement tissue.

APPLICATIONS OF BIOREACTORS

Bioreactors are used for 1) basic pathway studies, 2) growing replacement tissues, 3)

maintenance of organ vitality ex vivo and 4) priming therapeutic cells prior to cell

transplantation. The first two aims, which focus on cell phenotype and biomechanics,

respectively, have been covered in depth in the previous section. There is increasing demand for

techniques that maintain or promote a desired phenotype during in vitro cell culture [382].

Preconditioning cells prior to implantation [383] is not currently in widespread use for clinical

cell therapy, but this is a potentially fruitful future application for bioreactors. To accelerate

clinical use, fresh human bone marrow aspirate behaved similarly to subcultured bone marrow

MSCs on decellularized tendon scaffolds in a flow perfusion system, potentially providing a

shortcut when autologous cells are rapidly needed [384].

Bioreactors are often effectively used to culture direct tissue explants. The first of such

studies in tendon were conducted by Steven P. Arnoczky’s group, when Hannahin, et al. found

that canine FDPT maintained its mechanical properties ex vivo for 4 weeks while unloaded

controls degraded [385]. Michael Lavagnino, et al. later performed multiple sets of experiments

to determine that frequency and amplitude resulted in dose-dependent increases in MMP-1

expression in rat tail tendons [386]. This technique is still being used to elucidate pathways

involved in tendon adaptation to exercise, such as collagen and IL-6 expression [387], as well as

34

damage resulting from repetitive loading [388] and cellular maintenance of biomechanical

properties [389].

Living animals also provide valuable information for in vitro bioreactors. For example,

following a single loading episode in rat Achilles tendons, gene expression returns to baseline

after one day [390], but as little as 5 minutes of loading over 4 days is enough to improve

mechanical properties [391]. While the use of animals cannot be eliminated, bioreactors reduce

the necessity of laboratory animal experiments consistent with the 3Rs [392], namely by

reducing the numbers needed (by harvesting tissues at necropsy from unrelated studies for

bioreactor use) and replacing them with comparable methods (synthetic scaffolds). In addition to

tendon/ligament, cyclic-strain bioreactors are excellent platforms for cultivating muscle material,

such as for the repair of critical-size volumetric defects [393, 394] or even artificial meat [395].

Tendons are dynamic tissues, and tendon pathologies significantly reduce quality of life.

There are no effective pharmacological therapies currently available to treat tendinopathies, and

should tissue replacement be necessary, donor-matched prostheses are unavailable. Recent

developments in ex vivo modeling techniques have allowed us to dissect structure/function

relationships and elucidate cell-ECM interactions with unprecedented accuracy and

environmental control. Bioreactors are the best available tools for developing novel regenerative

treatments and cultivating functional replacement tissue. Future iterations of bioreactor

technology may be even better suited to these aims, and will likely be capable of replicating

multi-tissue or transitional structures by becoming increasingly complex.

35

CHAPTER 3: A DECELLULARIZED EQUINE FLEXOR TENDON SCAFFOLD

Published as:

Youngstrom DW, Barrett JG, Jose RR, Kaplan DL. Functional characterization of

detergent-decellularized equine tendon extracellular matrix for tissue engineering

applications. PLoS ONE (2013) DOI: 10.1371/journal.pone.0064151.

Reprinted under the PLoS ONE Creative Commons Attribution (CC BY) license.

ABSTRACT

Natural extracellular matrix provides a number of distinct advantages for engineering

replacement orthopedic tissue due to its intrinsic functional properties. The goal of this study was

to optimize a biologically derived scaffold for tendon tissue engineering using equine flexor

digitorum superficialis tendons. We investigated changes in scaffold composition and

ultrastructure in response to several mechanical, detergent and enzymatic decellularization

protocols using microscopic techniques and a panel of biochemical assays to evaluate total

protein, collagen, glycosaminoglycan, and deoxyribonucleic acid content. Biocompatibility was

also assessed with static mesenchymal stem cell (MSC) culture. Implementation of a

combination of freeze/thaw cycles, incubation in 2% sodium dodecyl sulfate (SDS),

trypsinization, treatment with DNase-I, and ethanol sterilization produced a non-cytotoxic

biomaterial free of appreciable residual cellular debris with no significant modification of

biomechanical properties. These decellularized tendon scaffolds (DTS) are suitable for complex

36

tissue engineering applications, as they provide a clean slate for cell culture while maintaining

native three-dimensional architecture.

INTRODUCTION

Extracellular matrix (ECM) has emerged as a fundamental tool for developing

regenerative tissue prostheses [396]. Simultaneously, bioengineered scaffolds (either natural or

synthetic) are critical to improving our understanding of the complex relationship between three-

dimensional topographical and biomechanical environments and stem cell growth and

differentiation [397, 398]. Regardless of the intended application, decellularization protocols are

designed to remove cells and debris while preserving the three-dimensional organization and

ultrastructure of the extracellular matrix. Biomaterials based on stem cells seeded on

decellularized tissue scaffolds are emerging as exciting options for clinical therapy, by obviating

the need for traditional organ transplant or autologous donation techniques which are associated

with significant morbidity [280, 399-401]. Moreover, the study of stem cell/matrix interactions

in a native scaffold environment furthers our understanding of basic cellular behavior and stem

cell differentiation.

Tendon is an important target for tissue engineering due to its frequent involvement in

musculoskeletal pathology and its comparatively simple organization. Tendon tissue repairs

slowly, has a poor functional endpoint after healing, and often suffers re-injury [402].

Furthermore, there is an unmet need for functional and readily integrated graft material for

human patients that suffer from traumatic tendon rupture or loss [403, 404]. The horse is an

excellent model for tendon research due to its pathophysiological similarities with human

37

degenerative orthopedic disease [215]. Additionally, there is a relatively large quantity of donor

tissue available compared with other model animals, as well as significant clinical demand.

Equine athletes routinely function close to the mechanical threshold for tendon damage [405] in

a mildly hyperthermic environment [99], resulting in cumulative cellular and extracellular

breakdown as well as changes in tissue biochemistry [406]. Tendinopathy results when this

deterioration exceeds the capacity for restorative remodeling [407, 408]. Since ECM components

are highly conserved, potential immunogenic reactivity is minimal [409]. Furthermore, due to the

high in vivo tensile load experienced by the equine flexor digitorum superficialis tendon (5%

average strain at a speed of 7m/s) [410], a maximal load on the order of 10kN [405], and the low

cellularity and vascularity of the tissue, it is a strong and homogeneous starting material as a

source of xenogeneic scaffold material.

It has long been recognized that cells demonstrate a complex array of behaviors in

response to culture on natural collagenous matrices [411]. This characteristic has been exploited

in biotechnology, as natural matrices inherently possess bioactive factors that induce host-

mediated healing [412, 413]. MSC differentiation pathways are dependent on the structural and

chemical composition of ECM [414], as well as its mechanical properties [415]. This is

particularly true for tendon progenitor cells, whose niche is dominated by extracellular proteins

[34]. Therefore, successful engineering of scaffolds for orthopedic research requires the correct

physical and chemical environment to induce de novo tissue generation. Preparation of these

tissues ideally removes cellular material, but leaves behind the critical fibrillar collagen

ultrastructure, as well as the majority of glycosaminoglycans (GAGs), to aid in tissue

regeneration.

38

Successful decellularization is a tissue-dependent procedure, and optimization of

protocols has until now been lacking for dense strong tendon such as equine flexor digitorum

superficialis. Nevertheless, testing protocols published for other fibrous tissues could translate

into an optimized protocol for this novel tissue. Low concentrations of tri(n-butyl)phosphate

(TnBP) and SDS have shown utility in a preliminary decellularization study of rat tail tendon

[289]. An analogous study of porcine diaphragm tendon revealed that TnBP caused adequate loss

of cellularity and preserved ECM architecture, while other commonly implemented detergents,

including Triton X-100, did not [293]. Conversely, Triton X-100, when used in combination with

other methods, reportedly aided in decellularization of flexor digitorum superficialis tendons in a

chicken model [416], and has also been used in combination with SDS [290, 417]. SDS has

effectively removed cellular debris from connective tissues when other methods have failed

[418], and has been used successfully in whole-organ [419, 420] and even multi-organ [421]

decellularization. Enzymatic decellularization without detergent exposure has also been

documented for certain tissues [301, 422, 423], and it is common practice to combine physical

agitation, chemical manipulation, and enzymatic digestion to achieve a finished product [409]. It

is apparent that decellularization procedures must be optimized by tissue type and donor species,

and tailored to the needs of the application.

Our aim was to test several different decellularization protocols and compare their ability

to decellularize equine flexor digitorum superficialis tendon while maintaining collagen

ultrastructure and minimizing loss of GAG content. Our hypothesis was that a treatment protocol

using SDS would remove the majority of cellular debris without reducing collagen content or

compromising structural organization of the DTS, preserving scaffold topography and

mechanical properties. We anticipated higher detergent concentrations would result in GAG loss,

39

so multiple concentrations were compared against each other and against other decellularization

procedures referenced in the literature. We additionally hypothesized that our DTS would be

compatible with allogeneic MSC culture, resulting in no significant loss of viability or

proliferative potential.

MATERIALS AND METHODS

Experimental design

Equine flexor digitorum superficialis tendons were aseptically harvested from the

forelimbs of eight castrated male sporting horses (mean age 12.8±1.4 years) euthanized for

conditions unrelated to musculoskeletal disease. All procedures were approved by the

Institutional Animal Care and Use Committee of Virginia Tech. Tendons were longitudinally

sectioned into 400µm-thick ribbons using a Padgett Model B electric dermatome (Integra

Lifesciences). Samples were sectioned into squares approximately 3cm2 in area and were

randomly assigned to treatment groups. With three replicates of each control and treated scaffold

per horse, a total of 144 tendon samples were required for initial characterization. Each outcome

variable was performed in triplicate on samples from each of the eight horses.

Two small representative sections were removed from every scaffold sample for

biochemical characterization. These sections were dehydrated overnight in an 80°C oven,

weighed, and placed into low binding affinity microcentrifuge tubes (Eppendorf). One portion

from each sample (4.7mg mean dry mass) was digested in 1mL of 1mg/mL papain [299] (Sigma)

for 36 hours in a heated water bath at 65°C, while the other (2.8mg mean dry mass) was digested

with 0.1mg/mL pepsin (Sigma) in 0.5M acetic acid for 48 hours at 4°C. All concentrations were

40

normalized to scaffold dry mass. The remaining portions of the original scaffold samples were

reserved for microscopy.

Following biochemical and microscopic characterization, the optimal protocol (2% SDS)

was selected for additional comparison versus the untreated tendon control, including analysis of

ultrastructure, biocompatibility and cellular integration. This was performed in triplicate on

material from four horses.

Scaffold preparation

Tendon samples were assigned to either the untreated control group and immediately

frozen at -80°C, or to a treatment group consisting of one of the following detergent types: (1)

phosphate buffered saline [PBS] (Lonza) control; (2) 1% tri(n-butyl)phosphate [TnBP] (Aldrich);

(3) 1% SDS (Sigma) plus 0.5% Triton X-100 (Sigma) [SDS/TX100]; (4) 1% SDS; (5) 2% SDS.

All detergents were buffered in 1M Tris-HCl (Fisher Scientific), pH 7.8. In a previous study, we

determined that no live cells and no residual mRNA were present in tendon samples after four

freeze/thaw cycles [299]. Following that procedure, samples were individually suspended in

2mL of their assigned detergent types in six-well plates in a refrigerated shaker at 4°C for 48

hours to allow adequate perfusion of tendon matrices. Tendon samples were then rinsed in PBS

six times to remove residual detergent [424] before incubation with 0.05% trypsin-EDTA

(Gibco) for 10 minutes and washed with water. Samples were then stored at 4°C for 24 hours

with the addition of 1µl/mL of a mammalian tissue protease inhibitor cocktail (Sigma).

Additional treatment steps included incubation in DNase-I (STEMCELL Technologies) for 30

minutes and 95% ethanol (Sigma) for two hours at 4°C, separated with and followed by three 10-

41

minute washes in water. Treatment steps were conducted in a gyratory shaker (New Brunswick

Scientific). Samples were divided for each assay as described above.

Biochemical characterization

DNA content, as a marker for cell debris, was quantitatively measured in the papain

scaffold digests following a single ethanol-based extraction technique using a fluorometric dye,

Quant-iT PicoGreen (Molecular Probes) in a ratio of 170µL working solution to 30µL

samples/standards in a 96-well plate.

Protein content in 1:10 dilution papain digests was measured via a standard Bradford

absorbance assay with Coomassie Brilliant Blue G-250 (Fisher Scientific). 50µL of each sample

was combined with 200µL reagent solution in a 96-well plate, referencing type-I rat tail collagen

(Gibco) as a standard.

Solubilized collagen following digestion was assessed in 100µL aliquots of acid/salt-

washed pepsin scaffold digests using a Sircol kit (Biocolor Ltd.), which is based on the specific

binding of Sirius red to the [Gly-x-y]n triple helix motif characteristic of collagen.

Sulfated GAG content in the papain digests was quantified through a spectrophotometric

assay based on 1,9-dimethylmethylene blue (Sigma) and compared to a standard curve of

chondroitin sulfate A from bovine trachea (Sigma), with a combination of 50µL

samples/standards and 200µL DMMB solution. Data was in all cases normalized to scaffold dry

weight, obtained prior to digestion.

Microscopic evaluation

Samples for microscopy underwent fixation in 4% paraformaldehyde and were submitted

for paraffin embedding, sectioning at 5µm, and routine histological staining (Histoserv, Inc.).

42

Longitudinal cross sections were stained with hematoxylin and eosin (H&E) or Masson’s

trichrome. Images were acquired using standard brightfield techniques on an Olympus IM

inverted microscope.

DNA was microscopically observed in tendon samples with ethidium homodimer-1

(EthD-1) (Invitrogen) [528EX, 617EM] under an AMG EVOSFL digital inverted fluorescence

microscope. Identical brightness and exposure settings were used for each image. A similar

procedure was replicated for cell culture analysis, with the addition of calcein, AM [494EX,

517EM] via an Invitrogen Live/Dead cytotoxicity assay. Cell viability over a period of four days

was assessed by a combination of live/dead fluorescence and the CellTiter 96 assay (Promega).

Scanning electron microscopy

Samples for SEM were dehydrated in a graded ethanol series (15%, 30%, 50%, 70%,

95%, and 100%), critical-point dried in CO2, and sputter coated with gold. Samples were

visualized in an FEI Quanta 600 FEG scanning electron microscope and representative images of

scaffold ultrastructure were acquired.

Biomechanics

Mechanical testing was conducted on duplicate samples from native tendon and DTS

from four horses, with a mean gauge length of 16.15±0.29mm, 5.79±0.14mm width, and

439±17µm thickness. Scaffolds underwent failure testing at a 1mm/s extension rate parallel to

the fiber orientation while submerged in a 37°C PBS bath on an Instron 3366 using a 100N load

cell and pneumatic clamps with 100-grit sandpaper. A tangent modulus of 0.003MPa was set for

0% strain following conversion from the raw load/extension relationship. Ultimate tensile stress

43

was measured empirically, and its corresponding strain was calculated using a fourth-order

polynomial fit to the stress/strain curve. Elastic modulus was reported at the maximum first

derivative of this relationship, and yield point was determined by the corresponding maximum of

the second derivative (Figure 3.1).

Figure 3.1 – Calculus for biomechanical data. Polynomial fits and the corresponding derivatives

were used to precisely identify points for elastic modulus and yield.

Biocompatibility

Tendon samples (0.4cm2) from the freeze/thaw group and from the 2% SDS experimental

group were dehydrated and sterilized in the graded ethanol series as above, rehydrated in sterile

PBS, and tested for biocompatibility with equine MSCs according to the procedures below. For

longer-term cell culture, 1x4.5cm DTS sections were prepared in accordance with the 2% SDS-

based decellularization procedure. Additionally, a section of each scaffold was soaked in 0.05%

Tween-20 (Fisher Scientific), and tested with a commercial limulus amoebocyte lysate-based

44

chromogenic endotoxin assay (GenScript). Values were compared to positive and negative

controls, and referenced to a broad-range standard curve of endotoxin.

Cell culture

Bone marrow aspirate was obtained from the sternum of a 3 year old male horse and

processed using routine stem cell separation procedures [299] that were approved by the

Institutional Animal Care and Use Committee. Bone marrow-derived MSCs were cultured in

low-glucose GlutaMAX DMEM with 110µg/mL sodium pyruvate (Gibco) supplemented with

10% MSC FBS (Gibco) and 100U/mL sodium penicillin, 100µg/mL streptomycin sulfate

(Sigma) at 37°C, 5% CO2, and 90% humidity. Cells were expanded and passaged twice at 80%

confluence, then 250,000-cell aliquots were directly seeded over the longitudinal surface of the

scaffolds in a 200µL meniscus of the same media. Samples were then incubated for 48 hours,

after which scaffolds were transferred to fresh containers. Adhesion efficiency was assessed by

performing a manual count of non-adherent cells in a hemocytometer following scaffold transfer

and trypsinization of the culture wells. A CellTiter 96 assay was performed for to quantify

viable cells after four days. A live/dead cell staining kit was also used to visualize scaffolds

using fluorescence microscopy on day four as described above.

Cellular integration within engineered tissue constructs was examined after an 11-day

culture period. An autologous MSC/DTS pair from a 9 year old female horse was cultured at low

density (20,000 cells/cm2) in the same media as previously described with the addition of

35.7µg/mL ascorbic acid and clamped on both ends. The sample was incubated for a seeding

period of 3 days followed by 8 days of immersion in media, with the solution changed every 3

45

days. At the experimental end point, the sample was collected and analyzed histologically to

assess cellularity deep in the scaffold.

Statistical analysis

Quantitative data is presented as mean ± standard error for all reported assays.

Differences across treatment means for each of the eight research horses were assessed using

one-way repeated measures multivariate analysis of variance (MANOVA). Data was grouped by

significance with p≤0.05 via alphabetical notation in applicable figures; data points that share a

letter are not statistically different from one another. Additionally, decellularized groups were

individually tested against the untreated controls for each of the biochemical, mechanical, and

cytocompatibility outcome variables with a one-way t-test. Statistical significance was declared

for those values that were found to meet the criteria of α≤0.01 and β≤0.02, noted with an

asterisk. All analyses were performed using the commercial software programs JMP 9 (SAS

Institute Inc.), Excel 12 (Microsoft), and Mathematica 8 (Wolfram).

RESULTS

2% SDS removes cellular debris from tendon explants

Histological sections stained with H&E are displayed alongside equivalent EthD-1

labeled scaffolds in the first figure. Untreated tendon histology shows tenocyte nuclei exhibiting

a characteristic elongated morphology, in parallel alignment with collagen fibrils in lacunae.

Nuclear remains of resident tenocytes are evident, as blue staining in the H&E sections, and as

red fluorescence following EthD-1 hybridization due to membrane disruption from repetitive

46

freeze/thaw cycles. A marked reduction in DNA content was demonstrated in all three SDS

experimental groups (Figure 3.2). Quantitative analysis of these data was performed using ImageJ

software (National Institutes of Health), and this difference was statistically significant (Figure

3.3). Most notably, the 2% SDS decellularized group retained only 0.11±0.05µg/mg residual

DNA compared with 0.67±0.12µg/mg in untreated tendon. Subjectively, no major alterations in

scaffold architecture were noted in histological samples.

Figure 3.2 – Nuclear Content following decellularization. Brightfield images of H&E-stained

histological cross-sections (5µm-thick) and fluorescence micrographs of scaffolds imaged

with EthD-1 (400µm-thick). Marked loss of DNA content is evident in SDS-decellularized

experimental groups, with 2% SDS resulting in the most dramatic decellularization. All

images are representative and were acquired from the same horse for ease of comparison.

47

Figure 3.3 – Image analysis of nucleic acid fluorescence. Quantitation of EthD-1-stained

decelluarized scaffolds was normalized to untreated tendon.

Tendon composition is not significantly altered by decellularization

Tendon samples were assessed for the following biochemical outcome variables: DNA,

protein, soluble collagen, and GAG levels (Figure 3.4). All detergent-treated groups exhibited a

statistically significant reduction in DNA versus untreated tendon. The 2% SDS treatment

reduced DNA content to 16.8±6.9% of untreated tendon, and this reduction was significant

(p=0.0081 with 0.0023 sphericity, power of >0.999). There was some protein loss in the SDS-

treated samples, reaching significance (p<0.001) in the 1% concentration group and approaching

significance (p=0.0052) in the 2% group. There was no significant difference between SDS

treatment groups (p=0.30). The majority of GAG content was preserved. The 2% SDS

decellularized tendon resulted in a small reduction in GAG content (2.83±1.64µg/mg decrease)

with p=0.042 and a power of 0.86.

48

Figure 3.4 – Treatment alters tendon matrix composition. Biochemical analysis of tendon

scaffolds, including DNA (A), total protein (B), soluble collagen (C), and GAG content (D).

Statistical significance between untreated control and treatment is annotated by use of an

asterisk. Statistical differences between treatments are indicated by different letters (repeated

measures MANOVA).

Decellularized tendon maintains desired ultrastructure

Because the 2% SDS treatment maintained collagen content and resulted in minimal

GAG loss and maximal DNA removal, this treatment was chosen for further characterization.

Untreated tendon (Figure 3.5A) underwent structural comparison with scaffolds decellularized

with 2% SDS. The characteristic deep blue coloration of collagen after staining with Masson’s

trichrome was apparent even in decellularized tendon, consistent with maintenance of collagen

content (Figure 3.5B). Samples were also evaluated via scanning electron microscopy (Figure

3.5C), revealing no microscale topographical differences in tendon structure aside from a

49

subjective increase in porosity as observed in the transverse plane. No statistically significant

alterations in tensile properties were observed between native equine tendon and 2% SDS

decellularized scaffolds. A summary of biomechanical outcome variables is included in Table 1,

with a representative stress/strain curve presented in Figure 3.6.

Figure 3.5 – Ultrastructural imaging. (A) Scanning electron micrograph of untreated tendon strip,

angled to show both longitudinal and transverse section architecture. (B) Histological

sections stained with Masson’s trichrome indicate maintenance of collagen content with a

slight increase in porosity. (C) Comparison of SEM images obtained from untreated and 2%

SDS decellularized tendon samples, shown at 100x and 5000x magnifications longitudinally,

and 5000x transversely. Collagen ultrastructure is not adversely altered by detergent

treatment.

50

Table 3.1 – DTS tensile testing data. No statistically significant alterations in scaffold mechanics

following decellularization were indicated. Data represents mean ± standard error values.

Scaffolds are biocompatible and free of endotoxin

Scaffolds decellularized with 2% SDS were compared to untreated tendon for assessment

of biocompatibility (Figure 3.7). Prior to testing, all samples were screened and tested negative

for endotoxin (data not shown). There was no significant difference in seeding efficiency

between decellularized tendon and untreated tendon scaffolds; 45.9±7.0% of plated cells adhered

to decellularized scaffolds compared with 38.9±3.3% for untreated tendon (p=0.77). Cell counts

on day four post-seeding indicated a global mean cell density of 420,000±17,600 cells per square

centimeter of scaffold with no statistical significance between untreated and 2% SDS treated

tendon. Fluorescence micrographs on day four using a live/dead cell staining kit demonstrated a

healthy population of MSCs with no detectable cell death (Figure 3.7C). The untreated tendon

had background staining of DNA, as expected. Subjectively, seeded cells exhibited an elongated

morphology in parallel with the aligned scaffolds.

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Figure 3.6 – Representative DTS stress/strain curve. Acquired using a materials testing device.

MSCs proliferate on and penetrate deep into DTS

Culturing autologous MSCs on DTS over 11 days demonstrated marked cellular

proliferation and integration, as visualized histologically (Figure 3.8). This strongly supports the

suitability for DTS as a platform for tissue engineering applications.

DISCUSSION

DTS provides a structurally similar physical environment to native tendon with minimal

residual cellular debris. Engineered scaffolds including DTS are geared to provide a tunable

microenvironment in which we may study fundamental development as well as cell-mediated

52

mechanisms of tissue regeneration. Additionally, advanced scaffolds such as DTS may prove

valuable in extending the phenotypic stability of tenocytes [134] and tendon stem cells [40],

which experience drift over extended culture periods [41]. Our novel protocol resulted in

formation of biocompatible, acellular constructs that will prove valuable in this pursuit. There

are several ECM-based scaffolds on the market for clinical use, but most have demonstrated

limited clinical efficacy as standalone products [401] and no tissue-specific commercial options

for tendon and ligament engineering or augmentation are available from animal sources [425].

Moving forward, optimization of tendon ECM decellularization focusing on structural stability

and accessibility to cell seeding are crucial to the development of useful reconstructive scaffold

materials [426]. Equine DTS represents an improvement to current competing technologies in

that regard, as it provides a unique combination of long graft length, mechanical strength, and

inexpensive small-scale production.

We evaluated the effect of several candidate detergents in combination with physical and

enzymatic cell disruption, followed by a series of wash steps and sterilization in ethanol, on

longitudinal tendon ribbons. These sections were acquired from the core regions of tendon

explants to take maximum advantage of intrinsic properties including comparatively low cell

density and vascularity as well as uniform ultrastructure and mechanics compared with other

regions such as epitenon [427-429]. As 300µm is reported as the minimum desirable tendon

scaffold thickness from a biomechanical standpoint [430], we selected a thickness of 400µm to

compensate for potential loss of surface ultrastructure following decellularization. This thickness

allows complex tissue construction techniques such as stacking [431], rolling [432], and forming

composites [433], yet is strong enough to facilitate early range of motion. And, it would allow

for repair of partial tears, and replacement of small tendons such as flexor tendons of the hand

53

via a rolling or stacking technique. Scaffolds underwent rigorous comparison with validated

methods for efficacy of removal of cellular debris, maintenance of ECM composition and

functional properties, and compatibility for allogeneic MSC culture. 2% SDS treatment, in

combination with trypsinization, treatment with DNase-I, and ethanol sterilization, resulted in

optimal characteristics.

Figure 3.7 – Biocompatibility of DTS to MSC culture. (A) Plating efficiency calculated as a ratio

of MSCs adherent to scaffolds following a 48-hour incubation period. (B) Total cell count at

96 hours obtained via MTS assay, indicating no significant reduction in proliferation or

metabolic activity on DTS. (C) Fluorescence micrographs portraying representative samples

of untreated and decellularized tendon, with live cells stained green (calcein) and dead cells

stained red (EthD-1).

54

DNA was selected as a sensitive indicator of cell debris due to its high stability, and

common use as a proxy for other cellular debris. While DNA is not cytotoxic, it is a useful index

that strongly correlates with adverse host immunogenicity [434]. EthD-1 was selected as a

marker for nuclear content following confirmation that no live cells or mRNA remained after

four freeze/thaw cycles [299]. The data indicates that 2% SDS removed a mean 83% of DNA

from the tendon scaffolds, and 1% SDS resulted in a 76% reduction. This represents an

improvement over other effective published decellularization protocols for tendon (67% [416]).

SDS in combination with Triton X-100 removed 72% of DNA, and TnBP was the least effective

detergent-based method with 71%. Detergent-free decellularization (PBS group) removed only

44% of nuclear content. These numbers correlate with fluorescence images of intact scaffolds.

However, it is interesting to note that residual TnBP may enhance EthD-1 fluorescence, as the

intensity displayed on the representative micrograph does not correspond to results obtained with

complimentary techniques, including H&E staining.

The Bradford assay demonstrated a mean 24% reduction of total protein, which may

correspond to loss of soluble intracellular proteins, chromosomal proteins such as histones,

organelle proteins such as lysosomal proteases and/or noncollagenous extracellular matrix

proteins. The statistical significance of the protein loss following 1% SDS decellularization most

likely corresponds to intracellular proteins or non-collagenous matrix proteins, as soluble

collagen content following DTS digestion was statistically unchanged with all experimental

groups failing to approach any of the statistical significance criteria.

It is interesting to note that our procedure leaves biologically relevant residual GAG

content, as experiments in porcine ACL demonstrated that SDS decellularization dramatically

reduced sulfated GAG content compared with untreated tissue [435]. Maintenance of GAG

55

glycosaminoglycan

Figure 3.8 – Cellular integration in DTS. H&E staining demonstrates infiltration of MSCs deep

into DTS following 11 days of static tissue culture.

content is desirable due to the role of the molecules in signal modulation, water composition, and

tissue biomechanics [436]. The DMMB assay demonstrated a 33% reduction in GAG content

which approached statistical significance (as it reached in other experimental groups), but

residual GAG content was much higher than anticipated. Statistical differences between the 1%

and 2% SDS-treated groups in protein and GAG outcomes may be attributed to the relatively

small sample size. A canine tendon decellularization protocol that did not use detergent has been

shown to maintain native proteoglycan content [301], but our data indicates that equine tendon

requires detergent decellularization, which may fortunately still be accomplished while

preserving the integrity of functional macromolecules. This highlights the necessity for tissue-

specific optimization of decellularization protocols.

Scanning electron microscopy allowed analysis of potential ultrastructural modifications

to decellularized scaffolds. Subjectively, SEM images did not demonstrate disruption of collagen

alignment or fiber thickness at the micrometer level, though a slight reduction in fibril density

was evident in the SDS-treated group that may prove beneficial for seeding scaffolds with cells.

This may correspond with the 18% increase in seeding efficacy noted in subsequent cell culture,

56

as the increase in porosity and surface area was hospitable toward MSCs. Histological sections

stained with Masson’s trichrome revealed similar staining of ECM, indicative of native collagen

composition, while also suggesting a subtle increase in porosity, supporting the electron

micrographs.

Biomechanical parameters of DTS did not significantly deviate from control tissue,

contrary to results reported in other decellularization protocols [297]. An insignificant extension

of the toe region and a decrease in yield strain was observed (p=0.16) with a slight increase in

elastic modulus (p=0.11), indicating trends that may emerge with a larger sample size.

Interestingly, these data suggest the potential for safe elongation protocols up to 7% strain before

plastic deformation is observed, with an ideal linear extension region centered on 4.1±0.2%

strain (data not shown). This characterization is necessary for experiments involving mechanical

manipulation of cell-seeded constructs.

While endotoxin contamination has not been associated with long-term derogative effects

in tissue graft integration or remodeling [437], testing is essential for evaluation of biomaterials

destined for bioreactor cell culture and medical device applications [438]. Our DTS contained no

detectable levels of endotoxin. In order to assess the initial feasibility of using DTS as a raw

material for tissue engineering, we seeded our scaffolds with MSCs for four days due to their

frequent use in cell-based orthopedic tissue engineering studies, and the possibility that

autologous MSCs could be used on DTS as a therapeutic graft material [439]. MSC culture was

successful in terms of plating efficiency, cell proliferation, and viability. As stated, DTS had a

higher seeding efficiency than identically sectioned control tissue, perhaps due to increased

porosity resulting from the decellularization process. For viability, no dead cells were seen

adherent to the scaffolds among 2% SDS DTS samples. A statistical comparison of cell viability

57

on DTS versus control tissue could not be performed due to the presence of high background

staining of endogenous dead cells in non-decellularized control tendon. The influence of long-

term culture on the growth and differentiation of MSC-seeded DTS constructs has only begun to

be explored, and will be the subject of further investigation. However, the results of an 11-day

study of autologous MSC/DTS constructs showed excellent cellular integration throughout the

depth of the scaffold and further supports its lack of cytotoxicity as a scaffold material.

Acellular ECM represents a versatile, biocompatible scaffold for three-dimensional cell

culture. This study validated a novel natural scaffold for tendon tissue engineering purposes.

Equine flexor digitorum superficialis tendon is well suited for use in tissue engineering

applications, bioreactor cell culture studies and graft material as a result of its robust mechanical

properties, homogeneous low cellularity, low vascularity, and long length and width. Our

protocol, based on 2% SDS detergent decellularization in conjunction with free/thaw lysis,

trypsinization, DNase-I digestion, and ethanol sterilization induces practical acellularity without

compromising functionality. The remaining extracellular matrix material maintains the

biochemical composition, ultrastructure, and mechanics of native tendon, yet has minimal

residual cellular debris. This provides a clean slate for subsequent cell culture, allowing

exploitation of the features intrinsic to physiological tendon matrix without concern over

functional or immunological interference from the original resident cells.

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ACKNOWLEDGEMENTS

The authors would like to acknowledge the ICTAS Nanoscale Characterization and

Fabrication Laboratory (NCFL) and the Morphology Laboratory at the Virginia-Maryland

Regional College of Veterinary Medicine for their technical assistance with this study.

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CHAPTER 4: DEVELOPMENT OF A TENDON BIOREACTOR

HARDWARE

The bioreactor system (patent pending: US #62050792) may be divided into four units:

the base computer, the control hardware (external to the incubator), the stage (inside the

incubator), and the vessels.

The base computer is a Lenovo ThinkPad Edge E545 powered by a 2.90GHz AMD A6

processing unit with 4.0GB RAM running 64-bit Windows 7 Professional SP1. This computer is

networked to the control hardware via a category 5 Ethernet cable.

The control hardware consists of an assembly of modular units based on a National

Instruments (NI) CompactRIO (cRIO) 9076 4-slot 400MHz controller and chassis. Data from

each load cell is transferred through an NI 9949 screw terminal block through an RJ50 cable to

an NI 9237 analog input module docked in the first slot of the cRIO. The second through fourth

slots contain NI 9512 stepper drive interfaces. These interfaces are connected to terminal blocks

using 15-pin D-SUB and MDR to 37-pin D-SUB Y-cables, which are wired to Hayden-Kerk

DCM8027 micro-stepping chopper drives. Power for the whole system sans the base computer is

provided by three 5A 25VDC NI PS-15 supplies, with everything mounted on a DIN rail. A

wiring diagram is portrayed in Figure 4.1 to illustrate connectivity of the hardware.

The stage contains the moving parts and sensors responsible for bioreactor function.

Three pairs of Honeywell Model 31 miniature load cells and custom Hayden-Kerk NEMA 11

captive linear actuators with 2” of captive travel and proximity sensors are mounted to an

aluminum stage using machined brackets. Opposing load cells and linear actuators are attached

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to sample vessels using set screws. Wires are routed through a hole in the back of a Forma

Scientific Steri-Cult 200 Model 3033 air jacketed incubator.

Figure 4.1 – Bioreactor wiring diagram. Connectivity of hardware is illustrated.

Bioreactor vessels consist of standard T-175 tissue culture flasks with holes drilled into

the sides to accommodate sample clamps. Polytetrafluoroethylene brackets stabilize clamps and

lock flasks into matching slots in the aluminum stage. Custom stainless steel sample clamps were

designed and fabricated for this project (Figure 4.2). Preload is set manually at the point where

scaffolds are taut but not tensioned beyond 0% strain. Media changes are accomplished using

standard cell culture technique through screw-caps on the flasks.

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Figure 4.2 – Photograph of bioreactor clamps. Samples is held within T-175 culture vessels.

SOFTWARE

Custom software for bioreactor culture and tensile testing was designed in LabVIEW

2013 SP1 with following modules: Control Design and Simulation, Mathscript RT, SoftMotion

and Real-Time. Software versions must match between the control unit and the base computer.

There are two versions of the program. One version accepts inputs for time, displacement

and period for each of the three stages independently. The actuators are set to cycle with linear

displacement based on these parameters. The LabVIEW program is disclosed in Figure 4.3. The

front end that is accessible to the user is shown in Figure 4.4. When the program starts, there is

an initialization period in which old program variables are cleared the load cells are zeroed based

on their starting state. Actuators are controlled using Straight-Line Move Express VI

functionality provided by the SoftMotion module, contained within a for-loop. Raw load cell

voltages are converted to force using calibration information provided by the manufacturer,

using Mathscript equations. An example of strain versus time for the 3% cyclic strain protocol is

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Figure 4.3 – Bioreactor LabVIEW program. Designed to operate three parallel reactor stages.

63

Figure 4.4 – Software front end. Custom program with inputs for time, displacement and period.

later shown in Figure 6.1 B. Load data is continuously acquired over this period. At the end of an

experiment, this data is exported as a spreadsheet for processing. Status bars indicate

experimental progress, and an emergency stop button is provided.

A modified version of the standard bioreactor program is used for tensile testing, with the

following alterations. Inputs are restricted to sample length only. 11 cycles of 3% strain at

0.33Hz are undergone as described previously. Following the conclusion of this priming period,

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samples are slowly stretched to 30% strain at 0.1% strain per second, which will result in failure

of 100% of standard-sized bioreactor samples based on decellularized tendon scaffolds (Chapter

3). Load data is then exported for manual processing. The failure program (reduced to a single

platform for clarity) is disclosed below in Figure 4.5.

Figure 4.5 – Tensile testing program. Designed for failure testing of reactor constructs.

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CHAPTER 5: MECHANICAL STRAIN MODULATES STEM CELL PHENOTYPE

Published as:

Youngstrom DW, Rajpar I, Kaplan DL, Barrett JG. A bioreactor system for in vitro

tendon differentiation and tendon tissue engineering. Journal of Orthopaedic Research

(2015) DOI: 10.1002/jor.22848.

This article is protected by copyright. It has been reprinted with permission from

John Wiley and Sons under license #3570981486762. All rights reserved.

ABSTRACT

There is significant clinical demand for functional tendon grafts in human and veterinary

medicine. Tissue engineering techniques combining cells, scaffolds and environmental stimuli

may circumvent the shortcomings of traditional transplantation processes. In this study, the

influence of cyclic mechanical stimulation on graft maturation and cellular phenotype was

assessed in an equine model. Decellularized tendon scaffolds from four equine sources were

seeded with syngeneic bone marrow-derived mesenchymal stem cells and subjected to 0%, 3%

or 5% strain at 0.33Hz for up to one hour daily for 11 days. Cells cultured at 3% strain integrated

deep into their scaffolds, altered extracellular matrix composition, adopted tendon-like gene

expression profiles, and increased construct elastic modulus and ultimate tensile strength to

native levels. This bioreactor protocol is therefore suitable for cultivating replacement tendon

material or as an in vitro model for studying differentiation of stem cells toward tendon.

66

INTRODUCTION

Tendon dysfunction occurs with high morbidity in both humans and animals,

compromising freedom of movement and quality of life. Tendons are predominantly composed

of hierarchically organized, aligned collagen fibrils [440], and the specialized structure of tendon

extracellular matrix (ECM) provides tensile strength while transferring mechanical stimuli to

resident cells [80, 441]. There is a reciprocal relationship between ECM properties and cellular

behavior, and success of in vitro cultivation of tendon is dependent on recapitulating the natural

environment of the tissue.

The horse is a model organism for studies of human tendon pathophysiology [442, 443].

Injury of the flexor digitorum superficialis tendon (FDST) is particularly common [405], and

significant research has been dedicated to addressing the poor intrinsic regenerative capacity of

this tissue [444]. Mesenchymal stem cell (MSC) implantation has been safely used in the

treatment of tendon degeneration, and there is some evidence that the multipotency and

immunomodulatory properties of MSCs may improve healing [150]. Seeding cells on scaffolds

influences cellular behavior [445] and supports endogenous repair [446], but the utility of current

commercial tendon augmentation products remains limited [447]. An equine decellularized

tendon scaffold (DTS) has been previously developed in our laboratory as a step toward graft

material production and as an in vitro model for tendon injury and repair [448].

According to a recent review, inadequate knowledge of the tendon progenitor cell niche

is a primary barrier inhibiting the development of effective cell-based therapies [449]. Proteins

isolated from tendon extracellular matrix alone enhance proliferation and tenogenesis in three-

dimensional culture [450], yet intact, naturally-derived scaffolds have the benefit of 1)

67

biochemical composition, 2) three-dimensional topography and 3) tissue-relevant mechanical

properties. DTS is suitable for MSC culture under static conditions, and it was hypothesized that

subjecting these constructs to mechanical stimulation would induce differentiation toward tendon

and produce viable regenerative graft materials. From previous bioreactor studies on tendon it is

evident that, while loading is required for maintenance of a differentiated tenocytic phenotype

[451] and tissue biomechanical properties [379], mechanically stimulated tendon constructs

exhibit sensitivity to the characteristics of mechanical stimuli. This cellular response is likely

tissue- and model-dependent, requiring optimization based on construct properties and

environmental conditions.

The aim of this experiment was to compare three deformation protocols on MSC-seeded

DTS by examining the influence of strain on MSC phenotype. Two dynamic strain regimens of

varying amplitude (3% and 5%) were selected based on their physiological relevance and

compared to static (0%) controls. The approximate biomechanical transition between the toe

region and the linear elastic region of deformation of the tendon stress-strain curve is 3% strain

[452], while 5% is a standard linear amplitude of normal usage conditions [410]. Despite the

seemingly minor differences between these two groups, it was hypothesized that the distinctive

biomaterial behaviors delineating the two deformation regions [31] would differentially translate

mechanical stimuli to resident cells. We hypothesized that both 3%- and 5%-strained constructs

would exhibit stronger evidence of tendon differentiation than static culture. Furthermore, we

anticipated the 3% strain protocol would effectively induce tendon differentiation in adult MSCs

and promote an anabolic response. Effects of the three strain protocols were evaluated via the

expression of tendon marker genes, biomechanical properties and production of ECM following

11 days of bioreactor culture.

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Experimental design

MSC-seeded DTS was divided into three groups by strain amplitude: referenced in the

text as the 0%, 3% and 5% experimental groups. Microscopy, composition and biomechanics

data references either initial DTS (iDTS), control DTS (cDTS), or both as negative controls.

iDTS is the freshly prepared scaffold material, subject to no further manipulation. cDTS was not

seeded with cells, but underwent identical incubation conditions to the 0% experimental group.

Adult tendon (FDST) was used as a control to compare bioreactor gene expression data with

mature whole tissue (n=4).

Production of decellularized tendon scaffolds (DTS)

DTS was produced using sterile technique in accordance with methods developed in our

laboratory [448]. FDSTs were surgically excised from the forelimbs of four adult sport horses

aged 4.5±1.7 years, euthanized as a result of unrelated conditions in accordance with the

Institutional Animal Care and Use Committee of Virginia Tech. Tendons from these horses were

longitudinally sectioned using an electric dermatome (Integra Lifesciences) into ribbons 400µm

in thickness, then divided into samples 10x45mm in surface area. Briefly, these samples were

decellularized by four freeze-thaw cycles, a 48-hour detergent infusion with 2% SDS (Sigma) in

1M Tris-HCl, pH 7.8 (Fisher Scientific) at 4°C, incubations in 0.05% trypsin-EDTA (Gibco),

10µg/mL DNase-I (STEMCELL Technologies) and 95% ethanol (Sigma), followed by repeated

washings in H2O in a gyratory shaker (New Brunswick Scientific). The resulting scaffolds were

69

frozen at -20°C prior to use. Reference FDST samples for RNA analysis were flash frozen using

liquid nitrogen and stored at -80°C prior to processing in the same manner as the DTS samples

(described below).

Derivation of primary mesenchymal stem cell (MSC) lines

MSCs were collected and assessed via routine processing techniques [299] using bone

marrow aspirate collected from the sternum of the same four donor horses as the DTS material.

Cells were cultured at 37°C, 5% CO2 and 90% humidity in standard MSC media: low-glucose

GlutaMAX DMEM with 110µg/mL sodium pyruvate (Gibco) plus 10% MSC FBS (Gibco) and

100U/mL sodium penicillin, 100µg/mL streptomycin sulfate (Sigma). Cells were expanded in

monolayer culture to 80% confluence and passaged twice. Flow cytometry was conducted on

these four separate cell lines with a BD FACSCalibur using monoclonal antibodies previously

validated in our laboratory (data not shown). The results demonstrated that approximately 90%

of cells were positive for CD-90 and CD-44, common MSC surface antigens, as well as Oct-4,

which is indicative of a naïve stemness found in both embryonic [453] and adult [454] stem cell

populations.

Construct seeding and bioreactor culture

Following MSC expansion, DTS was thawed and saturated in tendon cell culture media:

standard MSC media as previously described with the addition of 35.7µg/mL L-ascorbic acid

(Sigma). This medium was used for the remainder of the study. DTS samples were clamped into

the bioreactor vessels along their natural axis of alignment (Figure 5.1), obscuring 0.5cm on each

end. MSC suspensions were deposited via micropipette over syngeneic DTS at a density of

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20,000 cells/cm2, which equates to the approximate surface density of a 40% confluent

monolayer and had previously been validated [448]. Seeded DTS was subsequently placed in an

incubator for 72 hours to allow cells to adhere, with the vessels filled to their maximum media

volume of 6mL after the first 24 hours. Following this seeding period, bioreactor culture was

initiated, and half of the media was changed every 2-3 days.

Figure 5.1 – Photograph of bioreactor vessel. The bioreactor consists of interchangeable,

enclosed modular vessels. This picture portrays an MSC-laden DTS construct with 10x35mm

of exposed surface area immediately following seeding.

A custom bioreactor at the Tissue Engineering Resource Center was used in this study.

The device, described previously [350], incorporates self-contained tissue culture vessels that

allow samples to be individually clamped and mechanically manipulated. A LabVIEW program

71

(National Instruments) operates four stepper motors in parallel stages. MSC-seeded DTS

constructs were cultured in this bioreactor for a total of 11 days: three days without stimulation,

three days subject to displacement for 30 minutes per day, then five days at 60 minutes per day

(Figure 5.2). Samples underwent linear deformation at 0.33Hz according to their experimental

group. These parameters were selected due to their physiological relevance as well as reports that

as few as 5-7 days of bioreactor culture at 0.0167-0.5Hz are sufficient to observe improvements

in material properties in fibroblast-laden tendon/ligament constructs [378, 455]. All groups were

repeated in triplicate for a total of 36 vessels: four horses, three experimental groups, and three

replicates. After the final day, constructs were removed from their vessels, divided for assays and

either flash frozen in liquid nitrogen or immersed in a fixative for preservation prior to analysis.

Figure 5.2 – Experimental timeline. The duration each construct spent in the bioreactor per day

gradually increased from 0 to 30 to 60 minutes over the cultivation period.

RNA isolation and gene expression analysis

Half of each sample was used for gene expression analysis, and all samples within each

treatment group were pooled. The experiment was repeated in its entirety to produce a replicate.

Bioreactor constructs were transferred from storage at -80°C directly into a cryomill (SPEX

SamplePrep) and pulverized in liquid nitrogen. RNA was isolated from tissue homogenates using

an acid guanidinium thiocyanate extraction protocol in phenol-chloroform, followed by

precipitation in isopropanol. The resulting pellets were resuspended and the solutions

72

concentrated in RNeasy spin columns (Qiagen), quantitated with RiboGreen RNA reagent (Life

Technologies) and reverse-transcribed with a high-capacity cDNA kit (Life Technologies).

cDNA was pre-amplified using a validated commercial TaqMan kit (Life Technologies) prior to

reaction in a 7500 Real-Time PCR System (Applied Biosystems) using custom TaqMan probes

(Life Technologies) in duplicate. A list of primers, probes and abbreviations used are included in

Table 5.1. Reactions were quantified with the 2-∆∆Ct

method using GAPDH as a reference gene

and are reported by fold-change with respect FDST.

Table 5.1 – List of qPCR primers and probes in Chapter 5. Custom-designed equine qPCR

primers and probes were designed to target: glyceraldehyde 3-phosphate dehydrogenase

(GAPDH), scleraxis (SCX), type-I collagen (COL-I), type-III collagen (COL-III), decorin

(DCN) and biglycan (BGN).

Mechanical testing

Daily load data was collected for replicates run on the EMC bioreactor. A representative

sample was collected from one replicate of each FDST and experimental group to undergo

failure testing (average dimensions 12.4±1.1mm x 1.7±0.1mm). Following measurement with a

digital caliper, samples were elongated at 0.5% per second until failure using a custom materials

testing device controlled by National Instruments components, generating stress-strain curves.

73

Elastic modulus was computed as the slope of the total linear region of this relationship. Ultimate

tensile strength was calculated as the maximum force per unit area endured prior to failure.

Spectrophotometric biochemistry assays

Sulfated glycosaminoglycan (GAG) content was assayed using the 1,9-

dimethylmethylene blue (Sigma) technique, referencing chondroitin sulfate A (Sigma). This

procedure was conducted in aliquots obtained during media changes, as well as in each

bioreactor construct on day 11. Solid samples were solubilized by enzymatic digestion in papain

(Sigma). cDTS was also included in this analysis, in addition to the typical FDST and iDTS

controls, to isolate the influence of cells on GAG maintenance over time under experimental

conditions and in tendon cell culture media. DNA content was quantified in the same digest

solutions using a Quant-iT PicoGreen (Molecular Probes) assay to determine relative cell

number. Solubilized collagen was measured using a Sircol kit (Biocolor Ltd.) in acid/salt-washed

pepsin (Sigma) digests of solid samples following the conclusion of the experiment. Values are

reported with respect to dry weight, obtained by dehydration in an oven.

Histology

A portion of each experimental sample, as well as of each FDST and iDTS control, was

fixed in a freshly prepared solution of 4% paraformaldehyde (Sigma) and submitted for

commercial histological preparation (Histoserv, Inc.). Samples were embedded in paraffin,

longitudinally sectioned into 5µm slices, and stained with hematoxylin and eosin. Images were

acquired using an Olympus IM inverted microscope and a Moticam 10 CMOS camera.

74


Data are reported as mean ± standard error in all figures. One-way multivariate analyses

of variance (MANOVA) with a repeated measures designs followed by standard F-tests were

used to determine statistical significance of all data points except qPCR data (p≤0.05). A

standard one-way MANOVA was used for qPCR results. Results are annotated in figures

alphabetically. One-way Student’s t-tests were also used in biochemical and biomechanical

analysis to specifically test experimental groups to DTS controls. Points of significance (p≤0.05)

are demarcated with an asterisk in applicable figures. Computation was performed in JMP Pro 11

(SAS Institute Inc.), Prism (GraphPad Software Inc.) and Excel 14 (Microsoft).

RESULTS

Cyclic strain promotes a tenocytic gene expression profile

Gene expression data are shown in Figure 5.3. SCX expression more than doubled from

the 0% to the 3% experimental group to 70±15% of FDST, and this difference was significant

(p=0.024). SCX in the 5% experimental group fell between the 0% and 3% groups, and was not

statistically different from either group. COL-I expression was greatest in the 3% experimental

group, with message levels present at 2.09±0.96 times what is observed in FDST: statistically

greater than the 5% group (p=0.041). COL-III expression was dramatically upregulated in the

0% experimental group versus FDST (p=0.005). The ratio of relative COL-I to COL-III

expression was 1.75 in the 3% experimental group, whereas it was less than or equal to 0.14 in

the 0% and 5% experimental groups. DCN expression changed in response to bioreactor

protocol, with greatest DCN expression in the 3% experimental group, significantly higher than

75

in the 0% (p<0.001) or 5% (p=0.011) experimental groups. BGN was most expressed in the 3%

experimental group, but this difference was not significant.

Figure 5.3 – Strain amplitude alters gene expression. Messenger RNA expression profiles of the

tenocytic marker genes scleraxis (SCX), collagen types-I/III (COL-I and COL-III), decorin

(DCN) and biglycan (BGN) varied by bioreactor protocol – 3% strain induced a phenotype

correlated with tenocytic differentiation and development. Data is reported by fold-change

with respect to FDST. Data points that share a letter are not significantly different.

3%-strained constructs mimic the mechanical properties of native FDST

Peak strain gradually decreased over each exercise cycle, with a representative graph

shown in Figure 5.4A. Natural log transformations of this data were fit to linear regressions, an

example of which is shown in Figure 5.4B. Hysteresis loops are shown in Figure 5.4C. The

slopes of the natural log linear regressions were interpreted as a measure of scaffold resilience.

76

Constructs in the 5% strain group experienced greater hysteresis loss than constructs in the 3%

strain group, but this difference did not reach statistical significance.

Figure 5.4 – Real-time strain monitoring. (A) Plot of strain by cycle number for a representative

bioreactor sample. (B) Linear fit of natural log by cycle number. (C) Hysteresis loops.

Constructs in the 3% experimental group failed at a mean stress of 17.7±3.8MPa, which

is more than double that of iDTS (p=0.041) (Figure 5.6A). Constructs in the 0% and 5%

experimental groups failed at significantly lower stresses than those in the 3% experimental

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group (p=0.009 and p=0.043, respectively). Constructs in the 0% and 5% experimental groups

had significantly lower elastic moduli than FDST (p=0.034 and p=0.019, respectively), while

this difference only approached significance for iDTS (p=0.090) (Figure 5.6B). Relative to iDTS,

the 3% experimental group exhibited a 2.56-fold increase in elastic modulus to 119±44MPa, a

value within 25% of the elastic modulus of matched native tendons (98±25MPa), and without

statistical significance between the two.

Figure 5.5 – Construct resilience during exercise. Endpoint resilience of bioreactor constructs,

indicating greater decrease in peak strain in the 5% experimental group.

MSCs integrate into DTS and modulate scaffold composition

Decellularization eliminated 95% of DNA from FDST, to 0.03µg/mg in DTS (p<0.001).

All MSC-seeded bioreactor constructs had significantly more DNA than native tendon

(p<0.001), equating to 6.6±0.2 times the value of FDST (Figure 5.7A). There were no statistical

differences in DNA content between the 0%, 3% and 5% experimental groups. Soluble collagen

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production from the 3% experimental group after 11 days was 12.0±1.9µg/mg (Figure 5.7B).

There were no significant differences in soluble collagen between groups.

Endpoint GAG composition in the 3% experimental group increased by a factor of 2.14

relative to iDTS to 13.5±3.1µg/mg (p=0.050), while unseeded cDTS released 74% of GAG

content into the culture media (p=0.004) (Figure 5.7C). GAG release into culture media was

tracked cumulatively, and it was found that cDTS lost 10.1±2.6µg/mL of GAG in the first three

days (Figure 5.7D). In contrast, the mean GAG release of the 0%, 3% and 5% experimental

groups was 4.6±0.26µg/mL three days into the bioreactor culture period. There was no further

GAG release between days 8-11 in the 3% experimental group (p=0.106).

Histological examination confirmed the high cellularity of MSC-seeded constructs

relative to native FDST (Figure 5.8). Cells integrated at least 200µm deep into DTS by 11 days.

Cells also established an anisotropic phenotype, elongating parallel to the DTS collagen fibers.

Figure 5.6 – 3% strain improves failure strength. Construct elastic modulus and ultimate tensile

strength were increased to native physiological levels by bioreactor culture at 3% strain. Data

points that share a letter are not significantly different. Asterisks demarcate t-test significance

from iDTS.

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DISCUSSION

There is an immediate clinical demand for tendon graft material as well as a long-term

need to better understand the tendon differentiation process. The purpose of this study was to

establish a tissue culture technique for investigating tenogenesis and engineering tendon graft

material. A novel approach was taken to this end, applying biologically-inspired mechanical

stimulation protocols to natural decellularized scaffolds laden with matched cells ex vivo. Cyclic

linear strain of 3% at 0.33Hz for up to 60 minutes per day was found to promote tendon

differentiation in bone marrow-derived MSCs on syngeneic DTS.

Although there is no single marker gene of the tenocyte phenotype, SCX and COL-I are

highly expressed in tendon [39] and are responsive to exercise [212]. SCX is a basic-helix-loop-

helix transcription factor acting on the COL1A1 promoter [5, 209], the expression of which

characteristically demarcates tendon from surrounding tissues beginning during fetal

development [211, 456]. SCX is instrumental in the MSC to tendon progenitor transition [457],

as well as in adaptability to injury [212]. SCX was expressed in all constructs, suggesting that

DTS promotes tenogenesis even in the absence of mechanical stimulation, but was highest in the

3% strain group. COL-I represents 75% of the dry mass of tendon [406], while COL-III is more

highly expressed in degenerate tendon as well as in early tendon healing [259]. COL-I was most

highly expressed in 3%-strained constructs with COL-III expressed at normal levels: a result

concurrent with physiologically normal tendon anabolism. The ratio of COL-III to COL-I was

greater than 5.0 in the 0% and 5% experimental groups, indicating that under- or over-

stimulation results in atypical collagen gene expression.

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DCN and BGN are small leucine-rich proteoglycans (SLRPs). DCN is the most abundant

non-collagenous ECM protein in tendon [458], and among other functions modulates

fibrillogenesis [459]. DCN was most heavily upregulated in the 3% experimental group. The

results of this study are consistent with a previous observation that DCN expression is increased

following exercise [460]. BGN is similarly important for directing assembly of collagens [133],

yet it is also notable for its essential role in the maintenance of the putative tendon stem cell

niche [34]. BGN transcription was not statistically different from FDST in the 0% or 3% groups,

but was downregulated in the 5% group (p=0.027). DCN and BGN expression exhibit temporal

dependence during the tendon healing process, with BGN peaking earlier in development [461].

Figure 5.7 – Biomechanical content varies by amplitude. Endpoint scaffold content of DNA (A),

soluble collagen (B) and GAG (C) were quantified by spectrophotometric assays. Cumulative

GAG release into cell culture media was similarly assessed (D). Data points that share a letter

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are not significantly different as determined via one-way MANOVA. Asterisks demarcate t-

test significance from iDTS.

Therefore, it may be of interest in future studies to follow trends in SLRP expression over a

variety of time points.

Failure testing confirmed one of the most critical factors in tendon graft candidacy: that

bioreactor-cultured constructs were mechanically robust relative to FDST. The elastic modulus

and ultimate tensile strength of constructs in the 3% experimental group increased versus iDTS

and were not statistically different from matched FDST. Thus over- or under-stimulation

decreases the efficacy of MSC differentiation toward tendon and/or inhibits ECM anabolism in

our model. The material properties of tendons are dependent on cross-sectional area [430].

Whole tissues in vivo may therefore have elastic moduli and ultimate tensile strengths

approximately an order of magnitude greater than what is reported here [405]. For future tissue

augmentation applications, several layers of constructs may potentially be combined to achieve

the properties desired [431].

GAGs are functional side chains of proteoglycans, the concentrations of which are

quantifiable measures of tendon structure and function [29]. GAGs facilitate collagen fibril

sliding under load [462], and influence microenvironments and mechanotransduction on a

cellular level [31]. Decellularization results in GAG loss, but MSC-seeded DTS regained or

exceeded native composition. Indeed, GAG levels detected in the 3% experimental group may be

considered supraphysiological for equine FDST [448], though similar results in the horse were

observed in response to low-intensity exercise [463]. BGN, with two GAG chains per molecule,

modulates tenogenic signaling pathways and enhances stem cell differentiation toward tendon

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[34]. Thus, supraphysiological concentrations of GAG may naturally coincide with

supraphysiological concentrations of resident stem cells.

Turnover of GAGs is a hallmark of tendon homeostasis [460], hence GAG release into

culture media was observed. However, MSC seeding appeared to prevent the rapid and

substantial GAG loss observed in cDTS, suggesting an immediate and sustained influence of

these cells on maintaining ECM integrity. This corroborates a recent study reporting a cell type-

dependent reduction in GAG loss in equine tendon explants [464].

Histological examination confirmed MSC integration into scaffolds at high density, as

well as the adoption of a tenocytic morphology [440], in line with results observed following

MSC cell therapy in vivo [150] and in the cultivation of other tissue engineered tendons in vitro

[450]. DNA quantification supported subjective microscopic assessment of cellularity, validating

the high cytocompatibility of DTS reported previously [448]. Despite its relatively low porosity,

DTS accommodates cells exceeding physiological concentrations. Morphology, cellularity and

soluble collagen content were not influenced by bioreactor strain amplitude.

The limited extracellular matrix expression and biomechanical properties observed in the

5% group was somewhat surprising considering the similar cellularity between groups. A

potential explanation for this phenotype is that ECM damage resulted in stress-deprived

microenvironments within the 5%-strained constructs [81], resulting in a gene expression profile

that in many ways more closely resembled the 0% group than the 3% group. Differences in

collagen gene expression did not translate to detectable differences in endpoint soluble collagen

after 11 days. It would be interesting to further determine temporal characteristics of the collagen

remodeling process and test for the degradation products in the media. Finally, while

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biomechanical testing confirmed the functionality of incorporating a bioreactor conditioning

protocol, the clinical relevance of these changes in gene expression is unknown.

The results of this study suggest that a tuned, bioreactor-based conditioning protocol may

assist in the cultivation of functional tendon graft material and serve as a platform for in vitro

testing. A protocol applying 3% cyclic strain at 0.33Hz for up to an hour daily for 11 days

promoted tenocytic differentiation of MSCs on DTS and improved construct material properties.

This bioreactor platform could uniquely address questions of relative tenogenic efficacy of

various types of stem cells, or be used to test the influence of small molecule or protein

supplementation on pathways relevant to tendon differentiation and regeneration.

Figure 5.8 – MSC integration into DTS in bioreactor. Scaffolds were successfully decellularized

and reseeded at supraphysiological density relative to FDST. MSCs integrated into DTS and

adopted a tenocytic phenotype which did not change relative to strain amplitude.

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ACKNOWLEDGEMENTS

Partial funding for this study was provided by the Virginia Tech Institute for Critical

Technology and Applied Science (ICTAS), the Morris Animal Foundation, and the Tufts

University Tissue Engineering Resource Center (TERC) [NIH P41 EB002520]. DWY gratefully

acknowledges support from the Grayson-Jockey Club Research Foundation via the 2013 Storm

Cat Career Development Award. Shannon L. Smith, MS and Jonathan A. Kluge, PhD designed

the bioreactor system, provided training and facilitated helpful discussions at Tufts University.

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CHAPTER 6: COMPARISON OF TENOGENIC EFFICACY OF ADULT STEM CELLS

This article has been submitted for peer review as:

Youngstrom DW, LaDow JE, Barrett JG. Relative tendon differentiation of bone

marrow-, adipose- and tendon-derived stem cells in a dynamic bioreactor.

ABSTRACT

Tendons are frequently damaged and fail to regenerate, leading to pain, loss of function

and reduced quality of life. Mesenchymal stem cells (MSCs) possess clinically useful tissue-

regenerative properties and have been exploited for use in tendon tissue engineering and cell

therapy. However, MSCs exhibit phenotypic heterogeneity based on the type of donor tissue

used, and the efficacy of cell-based treatment modalities may be improved by optimizing cell

source based on relative differentiation capacity. Equine MSCs were isolated from bone marrow

(BM), adipose (AD) and tendon (TN), expanded in monolayer prior to seeding on decellularized

tendon scaffolds (DTS), and cell-laden constructs were placed in a bioreactor designed to mimic

the native biophysical environment of tendon. It was hypothesized that TN MSCs would

differentiate toward a tendon cell phenotype better than BM and AD MSCs in response to an 11-

day conditioning period involving cyclic mechanical stimulation for 1 hour per day at 3% strain

and 0.33Hz. All three cell types integrated into DTS, adopted an elongated morphology similar

to tenocytes, expressed tendon marker genes and improved tissue mechanical properties. TN

MSCs expressed the greatest levels of scleraxis, collagen type-I and cartilage oligomeric matrix

protein. Major histocompatibility class-II protein mRNA expression was not detected in any of

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the three MSC types, suggesting low immunogenicity for allogeneic transplantation. The results

suggest that TN MSCs are predisposed to form tendon and are the ideal cell type for regenerative

medicine therapies for tendinopathies. When TN MSCs are unavailable, BM or AD MSCs may

serve as robust alternatives.

INTRODUCTION

Tendons connect muscles to bones, and act as springs to store and transmit force to the

skeletal system during locomotion. Tendon injury leads to decreased quality of life due to pain

and loss of function. Tendons are hypocellular tissues composed of aligned, hierarchically

organized extracellular matrix (ECM): predominantly fibrillar collagens [31]. The biomechanics

of tendon improve efficiency of locomotion [465], and forces on the tendon are translated to the

cellular level where they provide important mechanobiological signals to resident tendon cells

[52]. Tendinopathies are widely believed to result from the progressive buildup of

microstructural damage during overuse [77, 81], leading to abnormal biomechanical signals to

the cells resulting in altered cellular phenotypes [79] and ECM composition [30]. The therapeutic

application of adult mesenchymal stem cells (MSCs) of autologous or allogeneic origin may help

regenerate acute or chronic tendon damage not only by promoting tissue neogenesis, but also by

modulating inflammation [144] and providing trophic support [466].

BM (bone marrow) and AD (adipose) are established MSC sources often studied in

regenerative medicine applications. While they share major phenotypic similarities [467],

transcriptional and proteomic differences suggest preferential pre-commitment to certain

mesenchymal lineages [468]. Thus, tissue-specific stem cells may have better regenerative

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efficacy in their tissue of origin [469, 470]. TN (tendon)-derived MSCs (also known as tendon

stem/progenitor cells) possess similar mesenchymal-lineage mulitipotency to BM or AD MSCs

[34], respond to exercise [471], and have unique features based on their specific origin [472].

Stem cells enhance healing of damaged tendon [148, 473, 474], but a lack of standardization in

pre-transplantation processing techniques complicates controlled evaluation of relative efficacy

across studies [475, 476]. Better in vitro models are needed to characterize the use of these cells.

Tissue-engineered tendon graft material can be manufactured using any of these cell

types [477], but proper selection of cell source and scaffold origin are important decisions in

optimizing graft design [478, 479]. The discovery that tendon tissue contains populations of cells

with characteristics of both MSCs and tenocytes [34, 480] has prompted investigation of their

use in animal models of tendon regeneration, with mixed results [481, 482]. TN MSCs have

however demonstrated improved collagen alignment, stronger graft mechanical strength and a

decreased tendency for ectopic ossification versus BM cells when used to augment surgical

repair of full-sized defects in a rat model [483]. TN cells also promote tenogenesis of allogenic

MSCs via paracrine signaling or cell-cell contact [484], which may enhance extrinsic tendon

healing in vivo. Further investigation into the tissue-regenerative properties of TN MSCs is

required to evaluate their potential use in cell therapy.

Biomimetic tissue culture systems enhance experimental control and decrease the number

of animals used in pre-clinical investigations [392]. Bioreactors to study tendon and ligament cell

behavior have been around for over a decade [485, 486], and it is now evident that mechanical

stimulation dramatically enhances tenogenic differentiation of MSCs [487] and can be used to

precondition graft materials [488]. Isolated components of natural extracellular matrix provide

important cues for in vitro cell culture [382, 450, 489, 490]. Moreover, intact decellularized

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scaffolds provide near-native mechanical properties and provide further opportunities to assess

tissue remodeling ex vivo. A number of studies have demonstrated differentiation and cell-

mediated improvements in tissue mechanical properties in response to tendon-like ultrastructure

and strain [378, 491]. However, at present there are only three other groups working with

decellularized tendon scaffolds in bioreactors [377, 379, 380], and there are no published

experiments comparing stem cell differentiation in these systems. The aim of this study was to

determine to what degree the tendon differentiation and ECM anabolic responses of MSCs are

dependent on their source of origin, and discover whether the use of particular cell types is

advantageous for tendon graft maturation or cell therapy.

Table 6.1 – Surface marker profiles of BM, AD and TN cells. Flow cytometry data (%) of MSC

cell surface markers show similar identities between cells derived from BM, AD and TN.

In order to evaluate the relative regenerative potential of BM, AD and TN MSCs, this study

applied a bioreactor protocol previously used to evaluate amplitude-dependent behavior of MSCs

in response to cyclic strain [294]. The bioreactor is designed to simulate gentle exercise, while

decellularized tendon scaffolds (DTS) provide “biophysical beacons” [492] such as native

topography and force translation to cells. Outcomes were assessed using microscopy, qPCR,

biochemical analysis and tensile testing. It was hypothesized that TN MSCs would integrate into

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DTS, exhibit a more tenocytic gene expression profile compared to BM and AD MSCs and

improve tissue mechanical properties.


Experimental design

Matched DTS and MSC cell lines were obtained from four adult sport horses aged

4.75±1.75 years, euthanized with Institutional Animal Care and Use Committee approval.

Tissues and cell lines were cryopreserved until ready for use in a -80°C chest freezer or in liquid

nitrogen, respectively. Donor tissues for cell lines include sternal bone marrow (BM),

subcutaneous adipose (AD) and flexor digitorum superficialis tendon (FDST) (TN). Matched

FDST tissue and DTS are included as control groups, with the exception of qPCR data, which

references a bank of four unrelated adult sport horses, as suitable syngeneic samples were

unavailable.

Decellularized tendon scaffolds

DTS samples measuring 45mm x 10mm x 400µm were produced from forelimb FDST

obtained at necropsy. The decellularization process has been described in detail elsewhere [448].

Briefly, longitudinally-sliced tendon sections underwent four freeze-thaw cycles, 48 hours of

detergent decellularization in 2% SDS (Sigma), enzymatic cleanup with 0.05% trypsin-EDTA

(Life Technologies) and 10µg/mL DNase-I (STEMCELL Technologies), and washing steps with

95% ethanol (Sigma), H2O and PBS (Lonza). Residual SDS was detected at 71.7±30.7 ng/mg,

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orders of magnitude below cytotoxic levels (data not shown). DTS was frozen until ready for

use, at which point samples were thawed and saturated with tissue culture media.

Cell isolation and culture

MSCs were derived from primary tissue samples using routine isolation protocols reliant

on cell separation techniques and adherence to tissue culture plastic. All cell culture was

conducted at 37°C, 5% CO2 and 90% humidity, with manipulations performed in a BSL2

biosafety cabinet (NuAire), including 50% media changes every 2-3 days. The following media

cocktails were used for monolayer expansion through two passages – BM MSCs: low-glucose

GlutaMAX DMEM with 110µg/mL sodium pyruvate (Gibco), 10% MSC FBS (Sigma) and

100U/mL sodium penicillin, 100µg/mL streptomycin sulfate (Sigma), AD and TN MSCs: high-

glucose GlutaMAX DMEM with 110µg/mL sodium pyruvate (Gibco), 10% Cellect Silver FBS

(MP Biomedicals), 10% Horse serum (Life Technologies) and 100U/mL sodium penicillin,

100µg/mL streptomycin sulfate (Sigma). Colony forming unit (CFU) assays were performed for

each cell line at P2 by plating 1,000 cells on plastic 100mm-diameter cell culture dishes (Thermo

Scientific) in triplicate. After 9 days, cells were fixed in 4% paraformaldehyde (Sigma) and

refrigerated overnight. Colonies were then washed and stained with 0.05% crystal violet (Fisher

Scientific) and photographically counted in ImageJ (National Institutes of Health).

Cell-laden bioreactor constructs

BM, AD and TN MSCs were seeded in suspension directly over the surface of DTS at

250,000 cells per construct in a two-stage 333µL solution transfer procedure separated by 20

minutes. Samples were paired with their technical replicates in 60mm petri dishes (Thermo

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Scientific) and incubated for 24 hours to facilitate MSC adhesion to DTS. All bioreactor

constructs regardless of cell type were cultured in TN MSC media as previously described, with

the exclusion of streptomycin [493] and the addition of 35.7µg/mL L-ascorbic acid (Sigma).

After seeding, individual samples were carefully clamped into custom-fabricated bioreactor

vessels, in which they remained for the following 10 days with regular media changes.

Cyclic strain bioreactor

This study implemented a custom bioreactor (U.S. patent #62050792 pending) (Figure

6.1A) [294]. Aluminum stages anchor three opposed pairs of load cells (Honeywell, Model 31)

and NEMA 11 captive linear actuators (Hayden-Kerk) driven by microstepping chopper drives

(Hayden-Kerk). Aluminum clamps stabilized by polytetrafluoroethylene brackets were built

around T-175 tissue culture flasks. This hardware is run by National Instruments modular units,

including a CompactRIO 9076 controller and chassis, a NI 9237 analog input module, and three

NI 9512 stepper drive interfaces. Custom software for bioreactor culture and tensile testing was

designed in LabVIEW. Cell-laden bioreactor constructs were subject to one hour of daily cyclic

stretching: 3% strain at 0.33Hz (Figure 6.1B). After 10 days, samples were cut from their vessels

and divided for assay (Figure 6.1C).

Quantitation of gene expression

To produce qPCR samples for each tissue type and cell line, 70% of each technical

duplicate construct was pooled. After harvest, these samples were snap-frozen in liquid nitrogen

and immediately ground in a cryomill (SPEX SamplePrep). The resulting homogenates were

suspended in guanidinium thiocyanate lysis buffer before undergoing phenol-chloroform

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separation and an overnight isopropanol precipitation (Life Technologies and Sigma). The pellets

were then purified using RNeasy spin columns (Qiagen) and quantified using a NanoDrop

spectrophotometer (Thermo Scientific) before reverse-transcription with a commercial cDNA kit

(Life Technologies). Duplicate single-plex reactions were conducted in an Applied Biosystems

7500 Real-Time PCR System using custom TaqMan (Life Technologies) primers and probes on

a list of gene targets outlined in Table 6.1, abbreviated as follows: scleraxis (SCX), tenomodulin

(TNMD), collagen type-I (COL-I), collagen type-III (COL-III), decorin (DCN), biglycan (BGN),

elastin (ELN), cartilage oligomeric matrix protein (COMP), and major histocompatibility

complex classes I (MHC-I) and II (MHC-II). Reactions were quantified using the 2-ΔΔCt

method

using glyceraldehyde 3-phosphate dehydrogenase (GAPDH) as a previously validated

housekeeping gene [467, 494].

Figure 6.1 – Bioreactor protocol and experimental timeline. (A) Photograph of sample vessel

mounted in bioreactor (patent pending: US #62050792). (B) Strain versus time protocol

implemented in bioreactor. (C) Graphical representation of experimental timeline.

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Microscopic imaging

Histological samples were placed in solutions of 4% paraformaldehyde (Sigma) and

commercially embedded in paraffin, sectioned into 5µm-thick slices, and stained with

hematoxylin and eosin (Histoserv, Inc.). Histology images were taken using an Olympus IM

inverted microscope and a Moticam 10 CMOS camera. Representative samples for confocal

imaging were collected at the completion of the experiment, labeled with calcein-AM and DAPI,

and freshly mounted for fluorescence microscopy. Confocal images were acquired at the

Microscopy Suite at James Madison University using a Nikon Eclipse TE2000 laser scanning

confocal microscope. Confocal images represent z-stacks approximately 100µm in thickness.

Additional images were acquired using an AMG EVOSFL digital inverted fluorescence

microscope after probing representative samples with a live/dead kit (Invitrogen).

Biochemical composition

A quantitative assay for sulfated glycosaminoglycan content was conducted on all media

aliquots and on samples of all constructs, using 1,9-dimethylmethylene blue (DMMB)

referencing bovine chondroitin sulfate A (Sigma). Solid samples for DMMB analysis were first

dehydrated and digested in papain. A second set of samples from all constructs was digested in

pepsin (Sigma) and analyzed for soluble collagen content using a Sircol kit (Biocolor). DNA

content was also quantified using a NanoDrop spectrophotometer on pepsin digests.

Mechanical testing

Daily secant moduli were calculated from the maximum stress/strain values between

cycles 40-60 during each bioreactor session. After the final day, tensile testing was conducted on

portions of samples from each horse and cell line (17.26±2.73mm length, 2.48±0.33mm width),

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submerged in PBS. These samples underwent 11 priming cycles of 3% strain at 0.33Hz

immediately followed by an extension of 0.1% strain per second to failure. Elastic modulus was

computed as the slope between two points in the linear deformation region of the stress/strain

curve, and failure stress was calculated as the maximum load per cross-sectional area.


All data is represented as mean ± standard error. Statistical significance was determined

via one-way multivariate analyses of variance using standard F-tests. A repeated measures

design was used for all analyses except for qPCR comparisons with FDST, as these samples

were not syngeneically matched as all other sets were. Significance (p≤0.05) is presented

alphabetically in bar graphs and symbolically with asterisks in line graphs. Computations were

performed in JMP 11 (SAS Institute, Inc.) and figures were designed in Excel 14 (Microsoft).

Table 6.2 – List of qPCR primers and probes in Chapter 6. Custom-designed primers and probes

were designed to target: glyceraldehyde 3-phosphate dehydrogenase (GAPDH), scleraxis

(SCX), tenomodulin (TNMD), type-I collagen (COL-I), type-III collagen (COL-III), decorin

(DCN), biglycan (BGN), elastin (ELN), cartilage oligomeric matrix protein (COMP) and

class-I/class-II major histocompatibility complex domains (MHC-I/MHC-II).

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RESULTS

BM MSCs form fewer colonies than AD or TN MSCs

Of TN MSCs plated at P2, 31.4±1.6% established their own colonies (Figure 6.2A),

followed up by AD MSCs (29.9±2.4%) and BM MSCs (23.7±2.4%). BM MSCs formed fewer

colonies than TN MSCs (p=0.0456), while AD MSCs did not differ significantly from either

group (Figure 6.2B). As a proxy for endpoint cellularity, DNA content of bioreactor constructs

did not reveal significant differences across cell types (Figure 6.2C).

Figure 6.2 – Proliferative capacities of MSCs derived from BM, AD and TN. (A) Representative

image from TN CFU assay: photograph converted to binary for automated counting. (B)

Results of CFU assay for BM, AD and TN cells at P2, demonstrating the high proliferative

capacity of TN cells. (C) Final DNA concentrations in bioreactor constructs suggested no

differences in endpoint cellularity between groups.

MSCs integrate into DTS during bioreactor culture

Cells exhibited elongated, tenocytic morphologies in parallel with the axis of scaffold

anisotropy, with extensive cell-cell contacts (Figure 6.3). Confocal microscopy revealed a dense

population of live cells extending 100µm or deeper into all scaffolds. Although all groups had

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similar cellularity, AD MSCs appeared to reside in the more superficial level of DTS, while BM

and TN MSCs tended to integrate more deeply into the scaffold. This correlated with relative

fluorescence intensities plotted from the three representative z-stack images (Figure 6.4).

Confocal images were also used to demonstrate seeding heterogeneity (Figure 6.5A) and to

produce a 3-D reconstruction of a portion of one of the constructs (Figure 6.6B).

Gene expression profiles during tenogenesis differ by MSC source

Relative gene expression data is shown in Figure 6.6. SCX was most highly expressed in

the TN group, reaching 76±11% of the level of FDST, a difference that did not reach

significance (p=0.1336). SCX expression in the BM and TN groups was statistically greater than

in the AD group (p=0.0133 and p=0.0099, respectively). TNMD expression was at least an order

of magnitude less in all groups than FDST, though no differences were detected across cell

types. COL-I expression was greatest in the TN group and lowest in the AD group, with the BM

group falling between. Expression of COL-I was significantly greater than FDST in all groups,

but this effect was most pronounced in the TN group (p=0.0003). Similarly, COL-III expression

was greater than FDST in all groups.

DCN was upregulated in all groups. While DCN expression trended lower in the BM

group, this difference did not reach significance relative to the AD and TN groups (p=0.0815 and

p=0.0797, respectively), and was still greater than FDST (p=0.0306). BGN was expressed at

native levels in all groups. ELN expression was greatest in the AD group, and lower than FDST

in the BM and TN groups (p=0.0056 and p=0.0065, respectively). COMP expression was

2.43±0.52-fold greater than FDST in the TN group (p=0.0035), and was less than FDST in the

BM and AD groups (p=0.0070 and p=0.0015, respectively).

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Figure 6.3 – Histology and confocal microscopy. (A) 5µm-thick sections of DTS and bioreactor

constructs stained with H&E. Cells acquired tenocytic morphologies and aligned along axis

of scaffold anisotropy. (B) Confocal microscopy of bioreactor constructs labelled with DAPI

and calcein, approximately 100µm-thick z-stacks. TN and BM MSCs integrated deeper into

DTS than AD MSCs. (C) Morphology and cell-cell connectivity in a representative single-

plane fluorescence sample from the BM group. (D) Confocal top and side views of a

representative sample from the BM MSC group showing extensive recellularization of DTS.

Figure 6.4 – Confocal cellular infiltration. Fluorescence intensities of z-stacks from Figure 6.3B,

normalized to their maximum values. AD cells resided more superficially.

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Figure 6.5 – Seeding density and 3-D reconstruction. (A) Seeding density of a TN cell-laden

bioreactor construct at day 11, labeled with DAPI. (B) 3-D image of a BM-cell laden

bioreactor construct at day 11, labeled with DAPI and calcein. Binary-converted images.

MHC-I expression was approximately an order of magnitude less in all groups versus

FDST. MHC-I expression in the TN group was lower than in the BM group (p=0.0085), with the

AD group falling between. MHC-II expression was not detected in any of the groups.

Scaffold composition was not heavily influenced by cell type

Differences in ECM gene expression profiles did not contribute to detectable differences

in construct GAG content, as high baseline levels of these components within DTS complicate

identification of cellular contributions. (Figure 6.7A). Soluble collagen content was not

statistically different between groups, but the BM and AD groups experienced small but

significant (p=0.0311 and p=0.0246, respectively) losses in collagen content versus FDST

(Figure 6.7B). Small but significant differences were detected in GAG release into the cell

culture media: GAG release was lower in the BM group on days 6 and 8 and higher in the AD

group on days 8 and 10 (Figure 6.7C). Differences in cell integration patterns may be the source

of these transient changes.

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Construct mechanical properties are altered by resident cells

Daily monitoring of stress/strain via bioreactor load cells did not show notable

differences in construct stress over time (Figure 6.8). No differences in elastic modulus were

observed during failure testing (Figure 6.9A). Notably, cell-laden constructs significantly

increased in tensile strength relative to controls (Figure 6.9B). The BM, AD and TN groups all

failed at higher stresses than FDST (p=0.0473, p=0.0177 and p=0.0440, respectively).

DISCUSSION

BM MSCs formed the fewest colonies, agreeing with the observation that AD and TN

MSCs expand more rapidly in culture at low passage numbers [495]. The relevance of

differences in colony-forming ability in monolayer to the behavior of those cells on three-

dimensional scaffolds is not clear. The observed heterogeneity in colony size may arise from

differences in “stemness” across populations of cells in the same flask, as more differentiated

cells undergo cell cycle arrest after multiple cell divisions. TN MSCs fulfill criteria for

characterization as stem cells [496], but their putative position farther down the differentiation

pathway may ultimately decrease their proliferative capacity if scaled up for large-scale

cultivation.

MSCs in the BM, AD and TN groups all established gene expression profiles consistent

with a tenocyte phenotype and ECM remodeling. However, differences in the relative expression

of a number of target genes suggest that TN MSCs form tendon more readily than BM and AD

MSCs. Furthermore, BM MSCs show phenotypic advantages relative to AD MSCs.

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Figure 6.6 – Relative gene expression profiles. Tendon markers, ECM proteins, and MHC-I/II.

101

Figure 6.7 – Endpoint biochemical analysis and GAG release. Final (A) GAG and (B) soluble

collagen content in bioreactor constructs did not reveal significant differences between cell

types. (C) Accumulation of GAG in media, calculated from aliquots obtained at each media

change, suggests that attenuation of GAG loss from DTS is not cell type-dependent.

SCX is a broadly-acting transcription factor associated with a tenocyte phenotype [497],

playing an essential role in tendon development [498, 499]. The AD group expressed

significantly less SCX than the BM or TN groups, while the TN group was not significantly

different from FDST. All groups expressed more COL-I than FDST; relative trends in COL-I

expression between bioreactor groups mirrored trends in SCX expression, with lower COL-I

102

Differences in the relative expression of these targets suggest that TN MSCs form tendon more

readily than BM and AD MSCs.

SCX is a broadly-acting transcription factor associated with a tenocytic phenotype [497],

playing an essential role in tendon development [498, 499]. The AD group expressed

significantly less SCX than the BM or TN groups, while the TN group was not significantly

different from FDST. Although all groups still expressed more COL-I than FDST, this difference

may be responsible for the decreased expression of COL-I in the AD group relative to the TN

group (p=0.0139). Low SCX expression in the AD group likely also correlates with a lack of

activation of other downstream gene products associated with tenogenesis [500]. COL-III was

upregulated more than COL-I in all groups relative to FDST, a feature associated with early

tendon healing [501]. This result contrasts with our previous findings [294], and may be the

result of the higher seeding density, the slightly more rigorous strain protocol or variation across

different primary cell lines in the present study.

Expression of non-collagenous ECM proteins also differed by cell type. DCN and BGN,

proteoglycans differentially regulated during the tendon healing process [502] and associated

with a differentiated tenocyte phenotype [492], did not significantly vary by group. ELN contributes

Figure 6.8 - Daily secant moduli monitored via load cells. Obtained from bioreactor data.

103

Figure 6.9 – Biomechanical characterization. (A) Elastic modulus and (B) failure stress of

bioreactor constructs obtained by endpoint tensile tests. Failure stresses of cell-laden

constructs were significantly greater following bioreactor culture. Constructs in the TN MSC

group endured 6.1±1.7x greater stresses than matched DTS controls.

to the material properties of many connective tissues but is not present in fetal tendon [503] and

is downregulated in three-dimensional culture [504]. ELN expression was greatest in the AD

group and exceeded FDST, but did not result in observable changes in biomechanical properties.

ELN is commonly associated with an endothelial phenotype, expressed during angiogenesis and

desired in cardiovascular tissue engineering [505]. The implications of this difference for cell

therapy are therefore unclear, though it may simply be a result of the more perivascular origin of

AD MSCs [506]. COMP enhances the kinetics of collagen fibrillogenesis [507] and is correlated

with healthy tendon healing in vivo [508], thus the dramatic upregulation of COMP observed

only in the TN group is potentially of clinical relevance and benefit. Interestingly, this effect is

likely mechanically-mediated, as TN MSCs express COMP below physiological levels when

cultured on static tendon matrix [299].

MSCs are immunomodulatory and survive allogeneic transplantation [144], but are not

necessarily immunoprivileged [509]. There is also concern that immunogenicity may be

104

enhanced following differentiation [510], such as would occur following bioreactor priming or

after transplantation. MHC-I gene expression was lower than FDST but detectable in all groups,

a feature similar to embryonic stem cells [511]. MHC-II expression was not detected in any of

the cell types, a finding that supports their use in allogeneic transplantation [512].

Considering the natural collagenous scaffold, the lack of detectable differences in soluble

collagen content between bioreactor groups is not surprising. The difference detected between

FDST and the BM and AD groups was surprising, and may be associated with a weaker anabolic

response versus the TN group. Endpoint GAG was positively correlated with DNA content.

Although final concentrations of these outcome variables did not differ across groups, the TN

group had 77.9% more GAG than the BM group, and this difference reached significance when

normalized to DNA content (p=0.0061). GAG release from DTS is attenuated by MSCs, but our

results do not suggest this is related to cell type. This contrasts with a recent study suggesting

that tendon-derived cells more effectively prevent ECM release from tendon explants than BM

MSCs [464].

Tensile testing corroborated previous findings that show a cell-mediated increase in

construct failure stress in bioreactor-matured samples [294, 377]. This outcome supports the use

of our bioreactor-DTS system as a tenogenesis assay, as well as for developing tendon graft

material. Elastic modulus was not different from FDST, a desirable feature due to the well-

established dependence of MSC differentiation on substrate elasticity [415].

Bioreactors are powerful tools for research and development, but have some limitations.

Our study examined a single snapshot in time (11 days), and temporal changes in gene

expression likely vary significantly over the differentiation process. Changes in mRNA

expression often serve as sensitive early indicators of changes in cell phenotype. However, these

105

are dynamic changes and we looked at an early time point for differentiation studies. Tendons

may experience a greater range and complexity of deformation in vivo than what was simulated

here, and the healing response may be altered by interactions with the immune system,

vasculature, surrounding tissues, and soluble degradation products from injured tendon. The

differentiation response of MSCs may be even more pronounced in vivo due to paracrine signals

from the recipient tendon cells [513]. Nevertheless, this tendon bioreactor provides a tunable,

scalable platform for evaluating tendon regeneration or cultivating tendon graft material.

In summary, BM, AD and TN MSCs all integrate into DTS, express tenocyte marker

genes and increase construct failure stress in response to cyclic mechanical stimulation in a

bioreactor. TN MSCs proliferate most rapidly, express more SCX, COL-I and COMP, deposit

more GAGs and increase construct failure stress. The donor site morbidity associated with

harvesting TN MSCs may make their application most suitable for allogeneic use, while BM

MSCs followed by AD MSCs remain nearly as effective with fewer patient complications for

autologous use.

ACKNOWLEDGEMENTS

The authors gratefully acknowledge partial funding from the Morris Animal Foundation.

DWY received additional support from the Grayson-Jockey Club Research Foundation.

Kristopher E. Kubow, PhD provided generous support for confocal microscopy. Zijuan Ma

conducted gathered flow cytometry data.

106

CHAPTER 7: CONCLUSIONS AND FUTURE RESEARCH

SUMMARY OF RESULTS

Tendon function is essential for quality of life, yet the pathogenesis and healing of

tendinopathy remains poorly understood compared to other musculoskeletal disorders. The aim

of regenerative medicine is to replace traditional tissue and organ transplantation by harnessing

the developmental potential of stem cells to restore structure and function to damaged tissues.

The recently discovered interdependency of cell phenotype and biophysical environment has

created a paradigm shift in cell biology. Cells respond to mechanical deformation via a complex

set of behaviors involving force-sensitive membrane receptor activity, changes in cytoskeletal

contractility and transcriptional regulation. Effective ex vivo model systems are needed to

emulate the native environment of a tissue and to translate cell-matrix forces with high fidelity.

This dissertation introduces a dynamic in vitro model for tendon function, dysfunction

and development, engineered to characterize the mechanobiological relationships dictating stem

cell fate decisions so that they may be therapeutically exploited for tendon healing. A naturally-

derived decellularized tendon scaffold (DTS) was developed to serve as a biomimetic tissue

culture platform, preserving the structure and function of native extracellular matrix. In the first

study, it was hypothesized that we could identify a detergent-based protocol that would remove

the majority of cellular debris from tendon samples without reducing collagen content or

compromising structural organization. Thus, scaffold topography and mechanical properties of

DTS could be preserved for subsequent tissue culture. The DTS protocol results in an 83%

107

reduction in DNA with only a 33% loss in glycosaminoglycan content, while mechanical

properties replicated those of native tendon tissue with an elastic modulus of 70.3MPa.

In the second study, 0%, 3% or 5% cyclic mechanical strain was applied to MSC-seeded

DTS at 0.33Hz for up to 1 hour per day for 11 days in our custom designed bioreactor. It was

hypothesized that intermediate strain would produce optimal differentiation conditions as

assessed via qPCR and biomechanical characterization. The 3% group induced the most

tenocytic phenotype, with maximum expression of scleraxis (70%), type-I collagen (209%) and

decorin (42%) relative to native tendon. MSCs in the 3% group remodeled their scaffolds and

increased failure stress to 17.7MPa; this is more than double that of unseeded DTS. It was

concluded that MSCs respond to mechanical stimulation in an amplitude-dependent manner, a

useful trait for preconditioning engineered graft material or designing rehabilitation protocols.

In the third study, it was hypothesized that tendon-derived stem cells are predisposed to

form tendon more efficiently than bone marrow- and adipose-derived stem cells. MSCs from

each tissue were cultured for 11 days in the bioreactor, and all cell types differentiated toward

tendon. However, tendon-derived MSCs expressed the most tenocytic aggregate gene expression

profile, including scleraxis and extracellular matrix proteins.

Stem cells have enormous regenerative potential, and bioreactor systems such as this can

be used to promote a particular therapeutic phenotype and improve the efficacy of cell therapies.

The results of the described experiments have demonstrated that adult mesenchymal stem cells

are responsive to mechanical stimulation and, while exhibiting heterogeneity based on donor

tissue, are broadly capable of tenocytic differentiation and tissue neogenesis in response to

particular ultrastructural and biomechanical cues. This knowledge of cellular mechanosensation

has direct clinical implications for how we treat, rehabilitate and engineer tendon tissue.

108

FUTURE DIRECTIONS

DTS accommodates total cell densities far in excess of that which is observed in native

tendon, without evidence of problems related to nutrient diffusion. However, overall cell

densities are heterogeneous throughout the surfaces and interiors of constructs, with localized

regions of low cellularity. This problem persisted after increasing cell density and implementing

a two-step seeding procedure between the first and second sets of bioreactor experiments. A

potential solution to this issue is to incorporate a more elaborate seeding protocol, perhaps

utilizing custom vessels to increase cell suspension coverage while preventing runoff of the cell

suspension. Additional steps, such as application of a vacuum, inclusion of a hydrogel, flow

perfusion, or cell homing and adhesion strategies could be optimized for future use. Further

experiments using this dynamic bioreactor platform are nearly infinite, and include comparison

of different strain protocols, growth factor supplementation and genetic manipulation for

optimization of tendon differentiation.

109

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APPENDIX: SPECIALIZED EXPERIMENTAL PROTOCOLS

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RESEARCH PROTOCOL

DECELLULARIZATION OF EQUINE FLEXOR TENDONS

TECHNICAL NOTES

- Wear scrubs, disposable laboratory gowns, and BSL-2 PPE while handling tissue samples

to protect yourself and the sterility of your samples.

- The dermatome and plastic dissection trays must be sterilized in ethylene oxide. All other

tools are sterilized in a steam autoclave. Solutions are filter-sterilized.

- Use of the dermatome requires 2-3 participants. Plan a time when a group of people can

process several samples in a session. However, do not allow dermatome to overheat.

MATERIALS

Padgett Model B Electric Dermatome: Integra Lifesciences

Sterile surgical instruments, dermatome blades, locking clamps (stored with dermatome)

Phosphate buffered saline (PBS) (1X) without calcium or magnesium: Lonza #17-516Q

Tris base: Fisher #BP152 & Hydrochloric acid: Sigma #H1758

Sodium dodecyl sulfate (SDS): Sigma #L3771

Multi-platform orbital shaker, placed in 4°C refrigerator: Fisher #13-687-700

Trypsin, 0.05% (1X) with EDTA 4Na: Gibco #25300062

Protease inhibitor cocktail for mammalian cell and tissue extracts: Sigma #P8340

Deoxyribonuclease (DNase) I solution (1 mg/mL): Stemcell Technologies # NC9007308

Ethyl alcohol, 200 proof for molecular biology: Sigma #E7023

Sterile, autoclaved, distilled water (made in-house)

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METHODS

Step 1: Collection Day

1. Harvest forelimb FDST tendons from donor animals using sterile surgical technique.

Transfer tendons from necropsy suite to laboratory dissection table in a sterile vessel.

2. Dissect away surrounding tissue, leaving clean FDST for subsequent use. If samples are

desired for RNA, remove a portion and immediately flash-freeze it in liquid nitrogen.

3. Transfer FDST samples to sterile storage containers and place in -80°C chest freezer.

Label containers with accession number, date and initials. Tendons for scaffold

production may be stored in this manner indefinitely prior to use.

Step 2: Dermatome Day

4. For best yield, tendons must remain frozen. If processing multiple tendons, remove only

one sample from the freezer at a time. Thawed tendon will clog the dermatome.

5. Secure both ends of the tendon using locking clamps and firmly hold under tension.

6. Pass electric dermatome over the length of the tendon. Tendon slices are traditionally cut

into 400µm-thick ribbons. This parameter is adjustable on the side of the dermatome. The

first few layers are discarded, and it may be useful to increase thickness for these passes.

7. Collect tendon ribbons in glass dissection tray filled with PBS. As ribbons accumulate or

a new sample is prepared, transfer to 50mL conical tubes and freeze at -20°C.

8. Over the next few days, thaw and re-freeze 50mL conical tubes intended for use as DTS.

Mark tubes with number of freeze/thaw cycles, up to 4 cycles. Reference samples may be

set aside prior to this point in order to avoid further processing. Prepare solutions.

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Step 3: Processing and Decellularization

9. Bring samples, surgical instruments and dissection trays into biosafety cabinet. Trim

tendon ribbons into pieces slightly larger than the provided template. Samples will

expand during decellularization, but this ensures that final size will be sufficient.

10. Tendon pieces are individually decellularized in 6-well plates. Place pieces in wells and

fill each well with 2mL of 2% SDS in 1M tris-HCl, pH 7.8. Label all plates.

11. Seal 6-well plates with paraffin film and transfer to refrigerated (4°C) orbital shaker, set

to 120 RPM. Leave for 24 hours.

12. The next day, flip over all samples. Reseal plates and put back on shaker for 24 hours.

13. Add a small quantity of alcohol to the vacuum trap to prevent the solution from bubbling

over into main line. Aspirate SDS solution out of all wells.

14. Fill each well with 2mL PBS. Place on shaker. After 20 minutes, aspirate solution.

Repeat 5 times. Periodically flip samples over to ensure all areas are covered.

15. Add 1mL trypsin-EDTA to each well. Place on refrigerated shaker for 5 minutes. Then

transfer to incubator (37°C) for another 5 minutes. Aspirate.

16. Wash each well with PBS twice as previously described.

17. Add sufficient volume of 0.05mg/mL DNase I solution in PBS to cover all samples. Place

on refrigerated shaker for 5 minutes, followed by the incubator for 10 minutes. Aspirate.

18. Wash each well with water twice as previously described for PBS.

19. Add 1µl/mL protease inhibitor cocktail in PBS to each well. Place on shaker overnight.

20. Aspirate protease inhibitor cocktail, wash once with water, and replace with ethanol.

Place on shaker for 2 hours.

21. Aspirate, and fill all wells with water. Repeat twice more. Freeze in PBS.

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22. Cut to size using a template. Templates can be printed, laminated, sterilized in alcohol

and placed under a transparent plastic dissection tray in a biosafety cabinet.

Figure A.1 – Scaffold cutting template for bioreactor.

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Figure A.2 – Pictorial tendon decellularization protocol. Printable checklist.

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RESEARCH PROTOCOL

RNA ISOLATION AND PURIFICATION FROM TENDON

TECHNICAL NOTES

- Follow RNase avoidance procedures.

- Wear appropriate PPE when handling liquid nitrogen.

- Schedule liquid nitrogen delivery in advance of cryomilling.

- Impactor tubes have a limited lifespan. Use new tubes when available to avoid cracking

and subsequent sample loss.

- Use pipettes and centrifuge vials that can withstand phenol-chloroform extraction.

MATERIALS

RNase AWAY decontamination reagent: Molecular BioProducts #7003

Water, sterile (for RNA work) DEPC-treated and nuclease-free: Fisher #BP561

Guanidine thiocyanate (GITC): Fisher #BP221

Sodium citrate, for molecular biology ≥99%: Sigma #C8532

Sodium hydroxide (NaOH), reagent grade ≥98% pellets (anhydrous): Sigma #S5881

Sodium lauroyl sarcosinate (sarkosyl), for molecular biology ≥95%: Fisher #BP234

β-mercapoethanol, for molecular biology 99%: Sigma #M3148

Sterile surgical instruments, including forceps and scoopula

SPEX SamplePrep 6770 Freezer/Mill (cryomill)

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Impactor tube, steel end caps, steel impactor set and opening tool for cryomill

Triton X-100: Sigma #T8787

Sorvall Legend Mach 1.6R desktop centrifuge

Sodium acetate buffer solution, pH 5.2±0.1 for molecular biology: Sigma #7899

Saturated phenol: Ambion #AM9712

Chloroform, contains 100-200ppm amylenes, >99.5%: Sigma #2432

Sorvall RC 6 Plus superspeed centrifuge

Sorvall Legend Micro 17 R microcentrifuge

Isopropyl alcohol, for molecular biology >99%: Sigma #I5916

Ethyl alcohol, 200 proof for molecular biology: Sigma #E7023

RNeasy Mini kit: Qiagen #74104

RNase-free DNase set: Qiagen #79254

NanoDrop 2000c spectrophotometer

METHODS

1. Prepare lysis buffer stock: in 500mL total volume of H2O – 250g GITC, 17.6mL 0.75M

sodium citrate buffer (11.03g sodium citrate in 50mL H2O, bring down to pH 7.0 with

NaOH) and 26.4mL 10% sarkosyl (in H2O).

2. Immediately prior to milling, prepare 3mL of lysis buffer in a 50mL conical tube. Add

80µL of β-mercapoethanol.

3. Fill cryomill with liquid nitrogen up to the fill line.

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4. Place sample (which is fresh or retrieved from storage at -80°C) in impactor tube and

assemble. Immerse loaded tube in liquid nitrogen until sample is fully frozen and liquid

nitrogen stops bubbling.

5. Insert sample tube into the cryomill. Run cryomill using the following settings – cycles:

2, precool: 1 min, run time: 1 min, cool time: 1 min, rate: 12 CPS.

6. Remove impactor tube from cryomill and ensure integrity of the vessel. Sample should be

visibly powdered. If larger pieces remain, repeat milling for another cycle.

7. Open impactor tube using opening tool, transfer contents to 50mL conical tube

containing lysis buffer. Mix well and keep on ice.

8. If necessary for low-quantity samples, warm impactor tube to room temperature and rinse

with additional lysis buffer. If not warmed, lysis buffer will freeze on contact.

9. Add 487.5µL H2O and 162.5µL Triton X-100. Mix well. Leave on ice for 10 min.

10. Spin at 2000RPM for 5 min at 4°C in standard benchtop centrifuge.

11. Transfer supernatant to a fresh Teflon 50mL ultracentrifuge tube.

12. Add 8ml of sodium acetate buffer, mix well. Then add 13mL of saturated phenol and

3mL chloroform. Mix by inversion and vortex for 10 sec. Leave on ice for 10 min.

13. Balance tubes and centrifuge at 10000RPM for 15 min at 4°C in high-speed centrifuge.

14. Remove upper aqueous phase and transfer to new ultracentrifuge tube. Do not allow

contamination with interphase. Conduct a second collection close to the interphase and

spin in microcentrifuge at 13000RPM for 10 min to retrieve residual.

15. Repeat extraction on aqueous phase with 8mL saturated phenol and 8mL chloroform until

the aqueous phase is completely transparent.

16. Add 13mL of isopropyl alcohol, mix well, and incubate at -20°C overnight.

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17. Centrifuge at 10000RPM in high-speed centrifuge. Small pellet may be visible, or not.

18. Aspirate supernatant and store open on bench to allow alcohol to evaporate.

19. Dissolve RNA pellet in 1mL of H2O in tube. Add 1ml of lysis buffer RLT (containing

10µL/mL β-mercaptoethanol) and 1mL of ethyl alcohol. Mix by inversion.

20. Apply mixture to RNeasy spin column in batches until all of the solution is run through.

21. Follow spin column protocol from manufacturer. Include 15 min DNase step.

22. Perform final elution in 52µL of H2O. Use 2µL to quantify RNA concentration in

duplicate 1µL samples in spectrophotometer. Store at -80°C or immediately reverse-

transcribe into cDNA.

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