TGF-β3 Signaling in Redifferentiating Passaged Human Articular Chondrocytes

by

Katarina Andrejevic

A thesis submitted in conformity with the requirements for the degree of Master of Science

Laboratory Medicine and Pathobiology

University of Toronto

© Copyright by Katarina Andrejevic (2016)

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TGF-β3 Signaling in Redifferentiating Passaged Human Articular Chondrocytes

Katarina Andrejevic

Master of Science

Laboratory Medicine and Pathobiology

University of Toronto

2016

Abstract

Bioengineering articular cartilage for the repair of damaged cartilage is one approach to biological joint

repair. TGF-β3 has been shown to promote articular cartilage tissue formation by dedifferentiated

passaged human chondrocytes. We hypothesize that TGF-β3 signals through both ALK1 and ALK5 in

passaged human chondrocytes, to induce cartilage tissue formation in vitro. Treatment of passaged

chondrocytes in 3D-culture with 10ng/ml TGF-β3 resulted in increased ALK5 expression and decreased

ALK1 expression. This correlated with chondrocyte redifferentiation, as shown by increased gene

expression of type II collagen and aggrecan, and tissue formation including increased collagen synthesis

and retention of newly synthesized proteoglycans and collagen. This study identified that TGF-β3

promotes redifferentiation and ECM accumulation in passaged OA chondrocytes in part by signaling

through both the canonical (ALK1 and ALK5) and the non-canonical (TAK1 and p38) pathways, though

it is a complex regulation. Collagen production appears to be modulated through the ALK1/ALK5-TAK1-

p38 axis.

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Acknowledgments

I would like to thank Dr. Rita Kandel for her continued support, guidance and provision of time

and resources that made the completion of this work possible. In addition to Dr. Rita Kandel, I

would like to thank the remaining members of my Program Advisory Committee, Dr. Mohit

Kapoor and Dr. Boris Hinz for their esteemed advice and expertise, that was invaluable to the

completion of this project. In addition, I would like to thank Dr. David Backstein for providing

human samples from knee arthoplasty that were used as a cell source for our experiments.

Furthermore, I would like to thank the members of the Kandel lab for their constant assistance

and advice. Lastly, I would like to thank my family for providing me with the support,

encouragement and inspiration that allowed me to complete this project.

Contributions

I would like to thank the following people for their specific contributions that assisted the

completion of this Thesis. I would like to thank Dr. Rita Kandel for the guidance she provided in

regards to experiment design and experiment execution. Furthermore, Dr. David Backstein for

providing human samples from knee arthroplasty that were used as a cell source for all

experiments throughout the thesis. I would also like to thank Laurie De Kroon, for kindly sharing

the primer sequences of ALK5 and SMAD2. Additionally, I would like to thank the members of

the Kandel Lab for their advice and assistance regarding experiment optimization and

completion.

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Table of Contents

Acknowledgments and Contributions ............................................................................................ iii

List of Tables ................................................................................................................................ vii

List of Figures .............................................................................................................................. viii

List of Appendices ......................................................................................................................... ix

List of Abbreviations .......................................................................................................................x

Chapter 1 Introduction .....................................................................................................................1

1.1 Cartilage ...............................................................................................................................1

1.1.1 The Structure and Function of Cartilage Tissue ......................................................1

1.1.2 Cartilage Tissue Injury and Development of OA ....................................................4

1.1.3 Current Therapies for OA ........................................................................................5

1.2 Bioengineering Articular Cartilage Tissue ..........................................................................6

1.2.1 Scaffolds ..................................................................................................................7

1.2.2 Hydrogels .................................................................................................................7

1.2.3 Scaffolds That Direct Cell Ingrowth ........................................................................8

1.2.4 Scaffold-free Cartilage Engineering ........................................................................9

1.2.5 Possible Cell Sources .............................................................................................11

1.2.6 Passaged Autologous Chondrocytes as a Cell Source ...........................................11

1.2.7 Redifferentiation of Passaged Chondrocytes for Cartilage Tissue Bioengineering .......................................................................................................12

1.3 TGF-β .................................................................................................................................14

1.3.1 Characteristics and Function of TGF-β .................................................................14

1.3.2 TGF-β Signaling ....................................................................................................15

1.3.3 Canonical TGF-β Signaling Pathways in Chondrocytes .......................................17

1.3.4 Non-Canonical TGF-β Signaling Pathways in Chondrocytes ...............................18

v

1.3.5 TGF-β Signaling in Osteoarthritis .........................................................................20

1.3.6 Redifferentiation of Passaged Chondrocytes using TGF-β ...................................21

1.3.7 Summary ................................................................................................................23

1.4 Hypothesis and Objectives .................................................................................................24

Chapter 2 TGF-β3 Signaling During Redifferentiation and Tissue Formation by Passaged Human OA Chondrocytes .........................................................................................................25

2.1 Introduction ........................................................................................................................25

2.2 Materials and Methods .......................................................................................................27

2.2.1 Isolation and Culture of Chondrocytes ..................................................................27

2.2.2 siRNA Transfection ...............................................................................................28

2.2.3 Gene Expression Analysis .....................................................................................28

2.2.4 Western Blot Analysis ...........................................................................................29

2.2.5 Analysis of matrix synthesis and retention ............................................................30

2.2.6 Biochemical Analysis ............................................................................................30

2.2.7 Histology and Immunohistochemistry ...................................................................31

2.2.8 Statistics .................................................................................................................31

2.3 Results ................................................................................................................................32

2.3.1 Characterization of ALK1 and ALK5 Expression Throughout Passaging ............32

2.3.2 Characterization of ALK1 and ALK5 Expression during Redifferentiation and Tissue Formation ...................................................................................................34

2.3.3 Characterization of Tissue Formation in Response to TGF-β3 .............................37

2.3.4 Identification of TGF-β3 Activated Signaling Pathway Associated with Matrix Accumulation by Passaged Chondrocytes During Redifferentiation ........40

Chapter 3 Discussion .....................................................................................................................46

3.1 Conclusion .........................................................................................................................54

3.2 Future Directions ...............................................................................................................55

vi

Chapter 4 Appendix .......................................................................................................................57

4.1 Additional Tables ...............................................................................................................57

4.2 Additional Figures .............................................................................................................58

References ......................................................................................................................................73

vii

List of Tables

Table 3-1: Summary of Inhibitors used……………………………….…………………….……………28

Table 3-2: Summary of Primers used………………………………………………………….………….29

Table 4-1: Patient Details………..……...………………………………………………………………...57

Table 4-2: Additional Primers.………..…………………………………………………...……………...57

viii

List of Figures

Figure 3-1: ALK5 Decreases in Chondrocytes During Passaging……………….……………............33

Figure 3-2: TGF-β3 Increases Expression of TGF-β Receptor ALK5 and Decreases Expression of

TGF-β Receptor II (TGF-βRII) During Redifferentiation and Tissue Formation…….……….………35

Figure 3-3: ALK1 and ALK5 are Expressed in Chondrocytes During Redifferentiation and Tissue

Formation…………………….…….....………………………………………………………………..36

Figure 3-4: TGF-β3 Increases Expression of Chondrogenic Genes (ACAN, SOX9, Col2) of Cells During

Redifferentiation…………...………..……………………………………………………………....….38

Figure 3-5: TGF-β3 Promotes Hyaline Cartilage Formation……………………………...……….….39

Figure 3-6: TGF-β3 May Signal Through ALK5, ALK1, TAK1 and p38.…...………………………41

Figure 3-7: TGF-β3 Signals Through ALK5, ALK1, TAK1 and p38 and Results in Increased p38

Phosphorylation.……….……………………………………………………………………………….42

Figure 3-8: p38 and TAK1 Levels were not Significantly Decreased by Silencing with Targeted

siRNA…………….....……………………………………………………………………………….....43

Figure 3-9: TGF-β3 Treatment Results in Increased Collagen Synthesis and Retention, and

Glycosaminoglycan Retention.…………...………..……………….…………………………………..45

ix

List of Appendices

Appendix 4.1 Additional Tables…………………….……..……………...……………….…….….57

Appendix 4.2 Additional Figures.…………………………….……...…………………….…….….58

Figure 4-1: ALK5 Decreases in Chondrocytes During Passaging, Additional Data….....................58

Figure 4-2: ALK1 and ALK5 are Expressed in Chondrocytes During Redifferentiation and Tissue

Formation, Additional Data……….…….……………………………..……………………….…...59

Figure 4-3: TGF-β3 has Variable Effects on TGF-β Signaling Pathway Genes (SMAD 5 and

SMAD2) During Redifferentiation and Tissue Formation……………..…………………………...61

Figure 4-4: TGF-β3 Promotes Hyaline Cartilage Formation, Additional Data….……………...….62

Figure 4-5: TGF-β3 Promotes Hyaline Cartilage Formation, Combined Data From Three Patients

………………………………………………………………………………………………………64

Figure 4-6: TGF-β3 Promotes Hyaline Cartilage Formation to a Lesser Extent in a Select Group of

Patients………………………………………………………………...…………………………….65

Figure 4-7: TGF-β Signaling Pathway Inhibitor Panel Results in Decreased Expression of

Chondrogenic Genes (ACAN, SOX9, Col2)…...…………………………..………………….….....66

Figure 4-8: TGF-β3 May Signal Through ALK5, ALK1, TAK1 and p38, Additional Timepoints

………………………………………………………………………………………………………68

Figure 4-9: TGF-β3 Treatment Results in Decreased ADAM-TS5 and Increased TIMP3 Gene Expression...…………………………………………………………...……………………….……70

Figure 4-10: ROCK Inhibition Results in Increased Chondrogenic Gene Expression and Tissue

Thickness..……………………………………………………………..……………………………71

x

List of Abbreviations

ACAN Aggrecan Gene ADAM-TS A Disintegrin and Metalloproteinase with Thrombospondin Motifs ALK Activin Receptor-like Kinase AMH Anti-mullerian Hormone BMP Bone Morphogenetic Proteins Col1 Type I Collagen Gene Col2 Type II Collagen Gene DAPI 4′,6-diamidino-2-phenylindole ECM Extracellular Matrix ERK Extracellular Signal-regulated Kinases ESCs Embryonic Stem Cells FGF Fibroblast Growth Factor GAG Glycosaminoglycan GDF Growth and Differentiation Factors HRP Horseradish Peroxidase IGF-1 Insulin-like Growth Factor-1 iPSCs Induced Pluripotent Stem Cells ITS Insulin, Transferrin, Selenium JNK c-Jun N-terminal Kinases MAPK MEK1

Mitogen Activated Protein Kinases Mitogen-activated protein kinase kinase 1

miRNA microRNA MMPs Matrix Metalloproteinases MRTF-α Myocardin-Related Transcription Factors MSCs Mesenchymal Stromal Cells NSAIDs NT

Non-steroidal Anti-inflammatory Drugs Non-transfected control

OA Osteoarthritis OATS Osteochondral Autograft Transplantation System P0 Passage 0 Chondrocytes P1 Passage 1 Chondrocytes P2 Passage 2 Chondrocytes PCM Pericellular Matrix PGA Polyglycolic Acid PI3K Phosphatidylinositol-3-kinase PKB Protein Kinase B ROCK Rho-associated Kinase Protein RUNX2 Runt-related Transcription Factor 2 SDS-PAGE Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis

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SFM Serum Free Media shRNA Short Hairpin RNA siRNA Small Interfering RNA SLC Small Latent Complex SMAD Homologue of Sma- and Mothers Against Decapentaplegic SMURF SMAD Ubiquitylation Regulatory Factor SOX Sry-related HMG Box TAK1 TGF-β Activating Kinase TGF-β Transforming Growth Factor-beta TGF-βRI TGF-β Type I Receptor TGF-βRII TGF-β Type II Receptor TIMP3 Tissue Inhibitor of Metalloproteinase 3 WOMAC Western Ontario and McMaster Universities' OA Index

1

Chapter 1 Introduction

1 Introduction Articular cartilage tissue covers the ends of long bones to provide a load-bearing surface for

smooth joint articulation [1]. Articular cartilage is structurally complex and contains zones that consist of

varying extracellular matrix composition that provide it with mechanical strength and facilitate its

function [2]. Damage to articular cartilage initiates inflammatory processes that can lead to catabolism of

healthy cartilage [2]. In many individuals this can result in progressive cartilage degradation over time

and development of osteoarthritis, which causes pain and limited mobility, leading to an overall decrease

in quality of life [3].

Articular cartilage lesions have limited potential for self-repair in part because of the alymphatic

and avascular nature of articular cartilage which allows for limited tissue perfusion [4, 5]. Several

different strategies have been employed in an effort to repair damaged cartilage including stimulation of

the healing response (microfracture, chondroplasty), filling of the defect using donor or autologous tissue

(mosaicplasty, allograft, autologous chondrocyte implantation) and in end stage disease, prosthetic joint

replacement [3]. As the aforementioned approaches do not produce optimal repair of defects, current

research has turned to tissue engineering and biological or cell-based alternatives to regenerate articular

cartilage. The goal of these approaches is to bioengineer tissue that is similar in its composition, structure

and biomechanical properties to that of native articular cartilage tissue. To improve upon current

bioengineered articular cartilage, it is necessary to further elucidate the factors that promote and maintain

the chondrogenic phenotype in both native and bioengineered cartilage.

1.1 Cartilage

1.1.1 The Structure and Function of Cartilage Tissue

Cartilage tissue can be characterized into four distinct types (elastic cartilage, fibrocartilage,

growth plate cartilage and hyaline cartilage) based on their differing extracellular matrix (ECM)

composition and macromolecular organization and specific function [2]. Native cartilage tissue contains a

single differentiated cell type (chondrocytes) embedded in an ECM consisting primarily of proteoglycans

and collagen [1]. Elastic cartilage is composed sparsely of chondrocytes embedded in a dense matrix of

proteoglycan and elastin network, providing it with increased flexibility [6]. It functions mainly to

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provide support and maintain the shape of structures within the ear, larynx, small bronchi and nose [7].

Fibrocartilage contains thick bundles of type I and II collagen and a lower glycosaminoglycan to collagen

ratio than hyaline cartilage [8]. It is composed primarily of type I collagen and is found in the meniscus,

the temporomandibular joint and is present during fracture repair [8-11]. It plays a role in resisting tensile

forces. Growth plate cartilage is present at the ends of bones and is the site where bone growth occurs via

the process of endochondral ossification [12]. Hyaline or articular cartilage lines the surfaces of

articulating bones and provides support, resists compressive forces and allows for smooth nearly

frictionless movement at joint surfaces [3]. As in elastic cartilage, the two major macromolecules found

within hyaline cartilage are type II collagen and proteoglycan [1, 3].

Chondrocytes are the only cell type present within cartilage tissue and are therefore central to the

proper formation and function of cartilage tissue. They regulate cartilage ECM homeostasis by producing

proteoglycan and collagen macromolecules [13], as well as proteases that influence ECM remodeling

[14]. Chondrocytes divide throughout development but their proliferative capabilities diminish with age

[15]. Chondrocytes are generally spherical, though they may take on a more discoidal or elongated

morphology depending, on their location in reference to the depth from the tissue surface [16]. They arise

from interzone cells that develop in the cartilage anlagen [17]. The process of chondrocyte differentiation

from mesenchymal cells is regulated by a variety of transcription factors and growth factors, including the

Sry-related HMG box (SOX) transcription factors and fibroblast growth factors, insulin-like growth

factor-1 (IGF-1) and transforming growth factor-beta (TGF-β) [18].

By wet weight, articular cartilage is composed of 15–25% collagens, 5-10% proteoglycans and

70-80% water [19]. The ECM is composed of collagens and proteoglycans in a 3:1 ratio [20]. Collagen is

a triple-helix structural protein that contains high hydroxyproline content. It forms a network of elongated

fibrils that allow cartilage to resist shear and tensile forces [1]. Approximately 80-95% of collagen found

in cartilage is type II collagen [21]. Proteoglycans are macromolecules that consist of a protein backbone

with glycosaminoglycan (GAG) side-chains [1, 19]. The predominant form of proteoglycan in articular

cartilage is aggrecan [22]. Aggrecans can bind to hyaluronan and aggregate increasing water retention and

allowing cartilage to resist compressive forces [1, 23].

In articular cartilage, the ECM immediately surrounding the chondrocytes is known as the

pericellular matrix (PCM), an entity that is both biochemically and biomechanically distinct from the

surrounding territorial ECM [24]. It contains type VI collagen, type IX collagen, perlecan, aggrecan,

hyaluronan and biglycan [24]. The literature suggests that the PCM is responsible for transduction and

regulation of local mechanical and biochemical cues that influence the chondrocytes [16, 24] and may

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therefore play an important role in influencing the cell phenotype. For example pellet culture of human

osteoarthritic chondrocytes isolated with their pericellular matrix intact resulted in formation of tissue

with increased weight, type II collagen content and a more hyaline-like appearance, in comparison to

pellets composed of chondrocytes isolated without any PCM [25]. This suggests that the PCM may also

influence the production of matrix. Taylor et al. have shown that passaged bovine cells in 3D culture,

which were able to form tissue, differed in the pericellular matrix that they accumulated in comparison to

primary bovine chondrocytes that did not form tissue [26].

Articular cartilage is made up of four zones: the superficial zone, middle zone, deep zone and

calcified zones of cartilage, which differ in their collagen fiber orientation, cell morphology, and ECM

composition [2]. Cartilage is able to absorb the compressive, tensile and shear forces experienced by the

joint due to the complex multi-layer architecture of the tissue [3]. The superficial zone forms 10-20% of

the articular cartilage by volume [19]. It has collagen fibrils that are arranged parallel to the cartilage

surface and contains the highest water content and the lowest proteoglycan content. It is more cellular

than the other zones, containing chondrocytes that are elongated in shape [19, 27]. The superficial zone

allows for smooth joint movement, as it is structured to enable the resistance of shear and tensile forces at

the cartilage surface [1]. Directly below the superficial zone lies the mid zone (40-60% volume), which

contains an oblique distribution (Benninghof’s arcades) of collagen fibrils and has greater proteoglycan

content. It has lower cellularity then the superficial zone and contains more rounded chondrocytes [19].

The deep zone (30-40% volume) can have chondrocytes arranged in a columnar orientation and collagen

fibers perpendicular to the articulating surface, which allows articular cartilage to resist compressive

forces [19, 27]. The deep zone also contains the highest proteoglycan content [19]. The proteoglycan

content increases from superficial to deep zone, whereas the amount of water decreases [1, 27]. The

articular cartilage is anchored to the subchondral bone by the calcified zone of cartilage. A tidemark

represents the boundary between the calcified and non-calcified cartilage. The deep zone and calcified

cartilage contains hypertrophic chondrocytes, which synthesize alkaline phosphatase and express type X

collagen [28, 29].

Recapitulation of the complex structural architecture of articular cartilage tissue is an important

consideration for bioengineering articular cartilage. One of the challenges has been to engineer articular

cartilage tissue similar in biochemical composition and zonal arrangement to that of native cartilage.

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1.1.2 Cartilage Tissue Injury and Development of OA

Injuries to cartilage tissue differ in severity, but can be grouped into two broad categories, partial

and full thickness injuries [30]. Partial thickness injuries can result in cartilaginous flaps and loose pieces

of cartilage that can cause catching and locking of the joint [1]. In full thickness injuries the cartilage is

eroded with changes in the underlying bone and there may be an attempt at repair by fibrocartilage [30].

Previous research has shown that only certain smaller cartilage defects (<3mm in diameter) have the

potential for self-repair, while other defects result in further cartilage degeneration and development of

osteoarthritis [31, 32]. The limited ability for self-repair is in part due to the slow proliferation rate of

chondrocytes [33], and the alymphatic and avascular nature of articular cartilage that allows limited

diffusion of soluble factors and nutrients [4, 5, 34]. Focal cartilage defects are one of many factors that

play a role in the degeneration of articular cartilage and development of osteoarthritis (OA) [35].

Osteoarthritis (OA) is a disease of joints caused by articular cartilage degradation and subchondral

bone remodeling and results in pain, stiffness and limited mobility [36]. The loss of articular cartilage

occurs due to an imbalance of ECM production and degradation [21]. In early OA there is evidence of

chondrocyte clustering, due to enhanced proliferation [21]. In addition, there is an increase in catabolic

activity, including the production of matrix metalloproteinases (MMPs) and A disintegrin and

metalloproteinase with thrombospondin motifs (ADAM-TS, aggrecanases) that degrade the ECM [37-

39]. The early changes are in the superficial zone and are characterized by degradation of proteoglycans,

specifically lubricin (proteoglycan-4) and collagen fibrils [39, 40]. These changes lead to increases in

water content, which jeopardizes the mechanical stability of the tissue [21]. Structural changes develop

and the surface of the cartilage becomes fibrillated in localized areas. The loss of ECM progresses,

leading to loss of the cartilage. In advanced OA, mechanically inferior fibrocartilage rich in type I

Collagen begins to form in the bone marrow and grows up to cover the surface in an attempt to repair the

damage [21]. Due to the inferior mechanical and biochemical properties of fibrocartilage, the repair tissue

generally deteriorates over time. The zone of calcified cartilage also thickens, and multiple tidemarks

develop [21]. The changes in the subchondral bone also are thought to contribute to the pathogenesis of

OA [41]. There is increased remodeling and loss of the subchondral bone present in the beginning stages

of OA [41, 42]. In advanced OA, there is thickening of the subchondral bone plate and increased apparent

bone density and bone volume [41]. In some patients, osteochondral projections form from the joint

margins (osteophytes), while others may develop cysts in the subchondral bone [21]. The net effect is the

progressive degradation of ECM, which leads to diminished mechanical function of the articular cartilage

[37]. The limited regenerative potential of the degraded articular cartilage has led to the development of

many treatment options with the goal of either regenerating or replacing the cartilage [34].

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1.1.3 Current Therapies for OA

Interventions for early OA are limited to lifestyle modifications and physical therapy. These

modifications decrease the rate of disease progression to some extent and may benefit specific patient

populations more then others. In a study of 80 obese patients with knee OA, a 10% decrease in weight

correlated to a 28% improvement in function, as assessed by the Western Ontario and McMaster

Universities' (WOMAC) OA index [43]. Non-steroidal anti-inflammatory drugs (NSAIDs) are also

employed in treatment of short-term OA symptoms, such as pain and inflammation [44]. These

modifications may ameliorate OA symptoms, but there is nothing to prevent disease progression. The loss

of cartilage over time results in the progression of OA that eventually requires surgical intervention.

One strategy used in the repair of damaged cartilage is to elicit a healing response in the joint

through techniques such as microfracture and chondroplasty. Arthroscopic chondroplasty is also used to

decrease joint irritation by removing loose cartilage fragments, but does not prevent OA progression [45].

Microfracture involves the creation of very small fractures in the subchondral bone underlying the defect,

in order to encourage the formation of new cartilage [46]. The mesenchymal cells found in the bone are

recruited to the defect. They then differentiate into chondrocytes and form repair cartilage [46]. The

limitation of microfracture as an OA treatment is that the cartilage formed is in part fibrocartilage, a form

of cartilage that contains higher levels of type I collagen and different proteoglycans than hyaline

cartilage thus exhibiting inferior biomechanical properties [46].

Another strategy for treating early OA and focal cartilage defects is replacing the damaged cartilage

and reconstructing the defective area. Mosaicplasty or osteochondral autograft transplantation system

(OATS) is a technique where multiple small cylindrical osteochondral plugs are taken from non-weight-

bearing regions of the joint and implanted into the defect [47]. There are several issues with this treatment

including difficulty in treating larger lesions due to limited availability of donor-site osteochondral plugs,

poor lateral integration of osteochondral plugs with adjacent cartilage and donor site-morbidity [47].

Allograft plugs overcome some of these issues but donor availability is limited. Autologous chondrocyte

implantation is another method of repair used in the USA and Europe. It is performed in two steps. In the

first step, an arthroscopic biopsy of cartilage is taken from the patient and the cells are isolated and

expanded in monolayer culture to obtain an appropriate number of cells to repair the defect. In the second

step, the defect is debrided and a periosteal flap or collagen-based membrane is placed over it, to create a

pocket into which the cultured cells are injected [48]. As with microfracture, the tissue generated is often

a biomechanically inferior mixture of hyaline and fibrocartilage that degrades over time [49, 50].

The above-mentioned procedures have limited long-term potential, as cartilage degradation

6

continues and progression to end-stage OA ensues. Artificial joint replacements are the current standard

of care for patients suffering from end stage OA. The artificial implants are also not an ideal long-term

solution, as all implants will ultimately fail and revision or replacement surgeries lead to further patient

morbidity [51, 52]. The shortcomings of the treatment options summarized above highlight the need for

the development of biological treatments that effectively repair cartilage and stop the progression of OA.

1.2 Bioengineering Articular Cartilage Tissue In order to overcome the limitations presented by the current treatments of cartilage defects, the

field has moved towards other approaches such as bioengineering articular cartilage to replace the

defective cartilage. The advantage of bioengineering tissue is that the implant can be customized to suit

the specific structural and morphological requirements of each defect [22, 53, 54]. The challenge,

however, has been to bioengineer tissue that has the biochemical and mechanical properties, as well as the

complex multi-zonal 3D architecture of native articular cartilage [53]. To address this challenge, tissue-

engineering strategies have used various cell sources, in addition to scaffolds, substrates and stimulatory

factors that support the growth of tissue, as discussed in the subsequent paragraphs.

Tissue engineering approaches often employ the use of growth factors, mechanical stimulation,

bioreactors and varying oxygen tension to mimic the native microenvironment of the cells, encourage the

chondrogenic phenotype and promote the growth of tissue. Growth factors including the Transforming

growth factor beta (TGF-β) superfamily, Insulin-like growth factor 1 (IGF-1) and Fibroblast Growth

Factor (FGF) have been used to increase production of ECM [55, 56]. IGF-1 has been shown in the

literature to promote the chondrogenic phenotype, as well as regulate proteoglycan synthesis in the

cartilage ECM [57, 58]. FGF-2 has been shown to regulate chondrocyte proliferation [59]. Mechanical

loading through dynamic compressive, shear and hydrodynamic forces has also been shown to stimulate

the growth of tissue [60-64].

Another factor that has been considered in bioengineering articular cartilage is oxygen tension.

Due to the avascular nature of articular cartilage, oxygen and nutrient perfusion occurs by passive

diffusion through the tissue, leading to comparatively lower oxygen content then in other tissues [65].

Placing chondrocytes in hypoxic conditions resulted in increased ECM synthesis and enhanced tissue

formation [65, 66] Some groups have also employed the use of bioreactors, in order to enhance delivery

of growth factors and nutrients, while providing mechanical stimulation [62, 67]. In addition to the

7

stimulatory factors summarized in this paragraph, chondrocytes can be seeded into scaffolds or onto

substrates that support tissue formation and growth.

1.2.1 Scaffolds

Cartilage bioengineering can employ the use of scaffolds to provide a 3D environment that promotes cell attachment and tissue growth. Scaffolds may be constructed from natural or synthetic materials, or both in combination. Natural materials that have been used as scaffolds include agarose, alginate, cellulose, collagen, decellularized tissue matrices, fibrin, and hyaluronic acid, among others [54, 68-70]. Collagen is commonly used as a scaffold for cartilage bioengineering as it is biodegradable and can be cross-linked to provide structural support [69, 71, 72]. Limitations of the collagen scaffold include lack of integration to adjacent tissue and inflammatory response upon implantation [73]. Hyaluronic acid is another commonly used biomaterial that is a component of the ECM [68, 74]. Composites of hyaluronan and collagen are in clinical use in Europe. Hyalograft (Hyaff) seeded with autologous chondrocytes has been successfully used to treat single defects [74, 75]. This technique is limited to early disease as outcomes for patients with advanced OA or multiple defects were less satisfactory [75]. Synthetic materials, including biodegradable polymers Poly(α-hydroxy esters), have also been used as scaffolds for cartilage bioengineering [76, 77]. Synthetic materials allow for better control of scaffold pore size which can influence chondrocyte production of matrix molecules [77, 78], shape and size [79]. Bio-seed®-C is a synthetic scaffold consisting of fibrin, polydioxanone, polylactic acid and polyglycolic acid, that can be used to support the growth of autologous chondrocytes and has been shown to have clinical benefits [80]. The limitations of using synthetic scaffolds include matching the rate of biodegradation to the increasing mechanical strength required during tissue formation, as well as toxicity and inflammation caused by degradation products, and fibrosis [81-83]. Scaffolds in various forms, including fibrous mesh and hydrogels, have also been used in articular cartilage engineering. Fibrous mesh scaffolds consist of a network of fibers that are interlaced to provide structural support [84]. They provide more mechanical support then hydrogels. However, there may not be an equal distribution of cells throughout the mesh, resulting in irregular ECM accumulation [54]. Hydrogels scaffolds consist of 3D polymer chain networks that are capable of retaining large amounts of water due to their hydrophilic nature [54].

1.2.2 Hydrogels

Hydrogels are frequently used as a scaffold for bioengineering articular cartilage, as they can fill defects of various shapes and sizes while encapsulating chondrocytes uniformly in a 3D environment [85]. Hydrogels mimic the high water content present in native articular cartilage [86]. Erych et al. developed long-term stable fibrin hydrogels that were able to support bovine chondrocyte proliferation and production of proteoglycans and type II collagen, during a five-week culture period [87]. Kim et al. were able to recreate the multi-zonal architecture of cartilage by encapsulating chondrocytes derived from

8

the superficial, middle and deeps zones of bovine cartilage in separate hydrogel layers [85]. Hydrogels also support the delivery of nutrients and biological stimulatory factors, and the transport of waste [85]. Mauck and colleagues have shown that dynamic compressive loading and delivery of factors to chondrocytes encapsulated in agarose hydrogel disks, further enhanced sulfated glycosaminoglycan content, hydroxyproline content and tissue formation [62, 88]. The formation and polymerization of hydrogels can also be spatially and temporally controlled through photopolymerization or different temperature gradients [58]. Fisher and colleagues, were able to create a polymer that maintains a liquid form below 25°C and polymerizes above 35°C, allowing for encapsulation of cells at body temperature. The proliferation of bovine chondrocytes encapsulated within the thermo-reversible hydrogel was similar to that of chondrocytes encapsulated in agarose and alginate hydrogels [86].

Despite their frequent use in tissue engineering, there are many limitations that may affect their success in vivo. Hydrogels provide inferior mechanical support in comparison to the other forms of scaffold [54]. It is difficult to match degradation over time to suit the varying levels of biomechanical support need by the tissue throughout formation. The degradation rate must be tightly controlled as it influences chondrocyte survival and ECM production [89]. The degradation products must be biocompatible, to avoid triggering inflammatory responses that may compromise the tissue [54].

1.2.3 Scaffolds That Direct Cell Ingrowth

Another strategy for the repair of osteochondral defects is the implantation of an acellular

scaffold that provides mechanical support in the area of the defect and is porous to allow the ingrowth of

native tissue. These scaffolds must provide an environment that results in the growth of tissue by

promoting cell migration, attachment, proliferation and sometimes differentiation [54]. Lee and

colleagues showed that implanted acellular bio-scaffolds infused with TGF-β3-containing hydrogels

promoted hyaline cartilage growth in rabbits [90]. Four months post surgery, implanted bio-scaffolds

contained cartilage with type II collagen and aggrecan, and mechanical properties similar to native

cartilage [90]. Implants that contained TGF-β3-free collagen bio-scaffolds displayed sparse cartilage

formation [90]. Kon et al. used a collagen and hydroxyapatite biomimetic scaffold to treat chondral

defects in two horses. Two months post surgery chondral defects were filled with fibrocartilaginous tissue

[91]. The same approach has been used to treat 15 defect sites in 13 different patients and resulted in

repair tissue formation at all sites and complete graft attachment in 13 lesions. One of the two tissues that

underwent immunohistochemical analysis showed intense staining for type II collagen throughout and

type I collagen staining confined to the deep zone [92]. The limitation of using acellular scaffolds is

similar to that of microfracture, the tissue formed seems to be, at least in part, fibrocartilaginous.

9

There are many considerations that must be addressed when using scaffolds in cartilage tissue

engineering. Some of these aspects include varying mechanical strength, ability to integrate, rate of

degradation versus tissue formation, cell survival within the scaffold and biocompatibility of the scaffold

and degradation products. The use of scaffolds is associated with limitations in many of these areas.

Approaches that attempt to bioengineer articular cartilage in a scaffold-free environment have the

potential to overcome many of these limitations.

1.2.4 Scaffold-free Cartilage Engineering

In scaffold-free tissue engineering, cells are not seeded into an exogenous 3D scaffold, instead the

cells synthesize the entire ECM themselves to produce tissue. Scaffold-free tissue engineering can be

broadly grouped into two categories self-organization and self-assembly [93]. Self-organization is when

the system forms organized tissue (goes from a state of disorder to order) due to an external energy input

and includes techniques such as aggregate engineering and cell sheet engineering [93]. Self-assembly is

when organized tissue forms spontaneously (the system tends towards order) without any external energy

input [93].

One self-organization approach creates cell aggregates, often through rotational forces applied to

cells in non-adherent culture conditions [93]. Different forms of aggregate culture, including pellet

culture, have been used in cartilage tissue engineering [94, 95]. For example, pellet culture of human bone

marrow stromal cells has been used in combination with growth factors TGF-β3, bone morphogenetic

protein 6 (BMP-6) or IFG-1 to promote differentiation and tissue formation [94]. This method has also

been used to bioengineer human articular cartilage with biomechanical properties similar to that of native

tissue. The cartilage was generated using mesenchymal stromal cells (MSCs) in pellet culture to create

condensed mesenchymal cell bodies that were fused using a demineralized bone support [95]. The

limitations of aggregate culture are that the tissue formed often varies in size, shape and ECM distribution

due to unequal nutrient diffusion [94, 96, 97].

In cell sheet engineering, monolayer culture is used to expand the cells until high confluence

allows them to form a cohesive layer. The entire layer is then lifted from the substrate and in combination

with additional layers, is grown to form tissue [93]. Chondrocyte sheet engineering was successfully used

to fill cartilage defects and showed integration to adjacent cartilage tissue in the mini-pig model, though

repair tissue varied in quality and safranin O staining [98]. Ando et al. were also able to create

“chondrogenic-like tissue” that stained for safranin O and collagen which integrated to adjacent cartilage

upon implantation, using porcine synovial MSCs cultured in high-density monolayer [99]. Generation of

a hyaline-like cartilage tissue ~1cm in diameter from human synovial MSCs, has also been achieved

10

using this strategy in a low oxygen tension environment [100]. In humans, RevaFlex, sheets of expanded

juvenile chondrocytes, have been used in allogeneic repair of focal cartilage defects in phase I and II

clinical trials [101].

The approach of self-assembly relies on the innate ability of cells to organize themselves and

form complex tissues, without external intervention [93]. During development, cells self-assemble to

form tissues and organs [93]. In tissue engineering, attempts to mimic the developmental processes and

environment have been used both to drive stem cells towards differentiation into specific lineages and

encourage the regenerative ability of cells to form tissues.

Waldman et al. have shown that when primary bovine chondrocytes were seeded in high density

onto a bone-like substrate (calcium polyphosphate) a continuous layer of tissue formed, that resembled

native articular cartilage in tissue thickness and proteoglycan content, but contained lower collagen

content [102]. The in vitro formed tissue exhibited biomechanical properties similar to those seen in other

in vitro formed cartilage [102]. Hu and colleagues have also used a self-assembly approach to form tissue

by seeding primary bovine articular chondrocytes in high-density on top of agarose gels, in a 96-well

plate. The cells formed non-attached constructs that by 12 weeks contained type II collagen in the absence

of type I collagen, reached almost 40% of native bovine cartilage stiffness and resembled native articular

cartilage in appearance and upon histological analysis [103]. It was found that certain aspects of native

cartilage development were recapitulated, including the proteoglycan and collagen content, and the

presence of type VI collagen in the pericellular matrix at 4 weeks [104]. Additionally, it was found that

mechanical properties were similar to those seen in development during the first four weeks of culture

[104]. Tissue formation during self-assembly was therefore found to resemble the developmental

processes in native cartilage tissue [104]. Ahmed et al. showed tissue formation through the self-assembly

process using passaged bovine chondrocytes cultured in high density on type II collagen-coated

membrane inserts [105]. This culture system supported the redifferentiation of chondrocytes, as well as

type II collagen and proteoglycan accumulation [105].

An advantage of the self-assembly approach is that it can occur on bone substitutes, which would

allow better treatment of OA-related and trauma-related defects where both bone and cartilage tissues

may need replacement. The Kandel lab has developed a biphasic construct that consists of in vitro formed

articular cartilage tissue integrated to the intended articulating surface of a porous, biodegradable bone

substitute (calcium-polyphosphate). Focal cartilage defects in sheep have been successfully treated using

this biphasic construct [106]. Other groups have also shown tissue formation [107] and repair of focal

cartilage defects in animal models [108] using variations of the biphasic construct. The self-assembly

11

approach also promotes the formation of bioengineered cartilage that mimics the zonal architecture of

native cartilage tissue. Culture of sheep bone marrow stromal cells and predifferentiated chondrocytes on

top of calcium polyphosphate yielded tissue with a hyaline cartilage zone rich in type II collagen and a

mineralized calcified cartilage zone that expressed type X collagen [109]. Culture of human passaged OA

chondrocytes yielded tissue that expressed proteoglycan 4 in the superficial zone of cartilage, as is seen in

native tissue (Bianchi et al., unpublished). The scaffold-free self-assembly approach to cartilage tissue

engineering therefore has the potential to overcome many of the limitations associated with the use of

scaffolds and result in formation of tissue that recapitulates the multi-zonal structure of native cartilage.

1.2.5 Possible Cell Sources

Engineering constructs large enough for clinical use requires a readily available cell source.

Possible cell sources include primary chondrocytes isolated from native articular cartilage or progenitor

cells such as embryonic stem cells (ESCs), induced pluripotent stem cells (iPSCs) and mesenchymal

stromal cells (MSCs). All of these cell sources have limitations. The social stigma associated with ESCs

and their potential tumorigenicity makes them a less than ideal cell source for clinical use [110]. iPSCs

must be differentiated to chondrocytes, a process which can result in genome mutation and epigenetic

changes during the cell culture period and tumour formation post implantation [111]. More research is

required to establish a safe and standardized protocol for both the generation of iPSCs and their

differentiation into chondrocytes [111]. MSCs can also be differentiated into chondrocytes and used to

bioengineer cartilage [95, 112, 113]. During the process of differentiation, MSCs may not maintain

chondrogenic phenotype and instead mature to terminally differentiated chondrocytes [114, 115]. The use

of chondrocytes does not present many of the limitations associated with stem cells and is therefore more

suitable for cartilage bioengineering purposes.

1.2.6 Passaged Autologous Chondrocytes as a Cell Source

Primary chondrocytes have been found to synthesize more proteoglycans and type II collagen

then stem-cell derived chondrocytes in 3D culture [116]. Using autologous cells also eliminates the risk of

developing an immunological response against allogeneic or xenogeneic cells [117, 118]. Lastly,

autologous chondrocytes are already being used clinically for treatment of focal cartilage defects [49].

The limitation of using autologous chondrocytes is that only a limited number of cells are

available. The repair of large cartilage defects therefore requires in vitro monolayer culture of

chondrocytes to produce adequate cell numbers. This culture method results in chondrocyte

dedifferentiation, characterized by a loss of cell phenotype and ability to produce hyaline cartilage [119,

12

120]. The chondrocytes assume a fibroblast-like morphology [121] and exhibit increased type I collagen

(Col1) expression and decreased aggrecan (ACAN) and type II collagen (Col2) expression [122]. We have

recently shown that in bovine chondrocytes, actin stress fibers form and changes in actin polymerization

status (from globular to filamentous actin) occur during dedifferentiation [123]. These changes in

cytoskeletal organization result in the nuclear translocation of Myocardin-Related Transcription Factors

(MRTF-α) and the upregulation of contractile and fibroblastic matrix genes [123]. Functionally, using

dedifferentiated chondrocytes to form bioengineered cartilage tissue results in constructs with inferior

biomechanical properties [124]. It is therefore necessary to redifferentiate these chondrocytes, resetting

their ability to produce the appropriate ECM molecules, in order to bioengineer articular cartilage

functionally similar to native tissue.

1.2.7 Redifferentiation of Passaged Chondrocytes for Cartilage Tissue Bioengineering

Through monolayer expansion, an adequate number of cells can be attained for cartilage

bioengineering. It is necessary to redifferentiate these cells in order to generate superior tissue.

Redifferentiation can be defined as reestablishing the chondrogenic phenotype, including production of an

ECM rich in type II collagen and aggrecan, and low levels of type I collagen [125]. Different approaches

have been used to redifferentiate passaged chondrocytes with varying degrees of success. These

approaches include placing the cells in a 3D environment, seeding the cells in high-density culture and

co-culture with other cell types. The use of defined culture media and supplementation with growth

factors has also been used in combination with these approaches to enhance redifferentiation.

Placing the cells in a 3D environment can encourage the redifferentiation of passaged

chondrocytes. In the literature, hydrogel scaffolds are often used for this purpose. Zeng and colleagues

placed passaged porcine chondrocytes in a microcavity alginate hydrogel and showed enhanced ECM

synthesis, as well as increased expression of type II collagen and decreased expression of type I collagen

compared to cells encapsulated in plain alginate hydrogels [126]. Levett and colleagues explored the

ability of four different hydrogel materials (gelatin, HA, polyethylene glycol and alginate), often used in

cartilage tissue engineering, to promote redifferentiation [127]. The redifferentiation response was found

to depend on the type of hydrogel used to encapsulate the chondrocytes. In short, the best redifferentiation

was seen with HA hydrogels, but newly synthesized ECM was limited to the pericellular regions and the

cells did not from a continuous layer of cartilage [127]. Gelatin hydrogels supported accumulation of

ECM, but not redifferentiation and alginate hydrogels did not promote ECM accumulation [127]. The use

of scaffolds presents limitations discussed in the previous sections, and further supports the use of

scaffold-free techniques for chondrocyte redifferentiation.

13

Scaffold free-attempts to redifferentiate chondrocytes utilize high-density cell culture methods to

re-capitulate the 3D environment to some extent. Primary human chondrocytes expanded in monolayer

for 1 to 4 passages were able to regain a chondrocyte-like phenotype when placed in high-density culture,

at the medium and air interface [128]. The chondrocytes formed cartilage nodules that expressed type II

collagen and cartilage-specific proteoglycan molecules. Cells passaged in monolayer for 5 to 8 passages

did not regain a chondrocyte-like phenotype in high-density culture [128]. Another study compared

redifferentiation of passaged human chondrocytes placed either in monolayer, alginate bead or pellet

culture [129]. Chondrocytes redifferentiated in high-density pellet culture showed increased expression of

chondrogenic mRNA SOX9, aggrecan (ACAN) and type II collagen (COL2), compared to monolayer

control [129]. However though increased glycosaminoglycan content was seen with pellet culture, type II

collagen protein levels were higher in monolayer controls [129].

Placing passaged chondrocytes in co-culture conditions with other cell types is another

redifferentiation strategy. Co-culture is thought to enhance ECM production and redifferentiation due to

soluble factors secreted by these cells. Meretoja et al. placed passaged or primary bovine chondrocytes

and rabbit MSCs onto a porous scaffold and evaluated redifferentiation [130]. Passaged bovine

chondrocytes, showed improved glycosaminoglycan synthesis and Col2 and ACAN gene expression only

when cultured in serum free induction media supplemented with dexamethasone and TGF-β3 [130].

Ahmed et al. found that passaged human chondrocytes in co-culture with primary bovine chondrocytes

showed increased expression of COL2, as well as decreased expression of type I collagen (COL1). These

results were supported by increased ECM production and ability to form hyaline-like tissue, indicating

redifferentiation [131]. Taylor et al. showed that after three weeks in co-culture, accumulation of

proteoglycan and collagen in the extracellular matrix, as well as expression levels of chondrogenic genes

(SOX9, COL2 and ACAN) were similar to those found in primary bovine chondrocyte culture controls.

These results indicate that the optimal co-culture period is approximately three weeks [132].

In addition to the approaches discussed above, defined culture media and supplementation with

growth factors is often used to enhance redifferentiation of passaged chondrocytes. Culture media for

redifferentiation of chondrocytes varies but typically contains a serum substitute, such as Insulin,

Transferrin, and Selenium (ITS), dexamethasone, ascorbic acid and L-proline. Dexamethasone is often

used to direct progenitor cells to a chondrogenic lineage and it may have effects on ECM synthesis [105].

Ascorbic acid and L-proline are both found to support collagen synthesis and cross-linking [133, 134].

Ahmed et al. found that redifferentiation of passaged bovine chondrocytes is promoted by the use of a

defined serum-free media containing high-glucose, insulin, and dexamethasone [105]. This

‘redifferentiation media’ activated an insulin-dependent signal transduction cascade that upregulated

14

chondrogenic transcription factor SOX9 and promoted cartilage tissue formation in passaged cells [105].

However, these conditions were insufficient for formation of articular cartilage using human osteoarthritic

chondrocytes. Supplementation with growth factors has also been used in attempt to redifferentiate

passaged chondrocytes [56, 135, 136]. It has been shown that supplementation with FGF-2 during

passaging and subsequent supplementation with BMP-2 during 3D culture on polyglycolic acid (PGA)

scaffolds enhanced wet-weight GAG fraction present in the cartilage formed [136]. Recently, we have

shown that redifferentiation of OA chondrocytes and subsequent articular cartilage formation is possible

with the addition of TGF-β3 (Bianchi et al., unpublished), a known regulator of chondrogenesis [137].

This culture method resulted in the accumulation of aggrecan and type II collagen by the cells (Bianchi et

al., unpublished). Although we have established that addition of exogenous TGF-β3 is required for

redifferentiation and tissue formation, we have yet to characterize the signaling mechanism through which

redifferentiation occurs.

1.3 TGF-β

1.3.1 Characteristics and Function of TGF-β

The TGF-β superfamily is composed of many factors including BMPs, growth and differentiation

factors (GDF), anti-mullerian hormone (AMH), Activin, Nodal and TGF-β. TGF-β is a secreted growth

factor that is involved in development, growth, homeostasis and apoptosis, in both adult and developing

organisms [138]. In mammals, three isoforms including TGF-β1, TGF-β2 and TGF-β3, have been

identified [138, 139]. The chromosomal locations of the TGF-β1, TGF-β2 and TGF-β3 genes in humans

are 19q13, 1q41 and 14q23-4, respectively [140, 141]. The three isoforms are similar in structure (84-

92% homologous) and size [142, 143]. TGF-β1 contains 390 amino acids, while TGF-β2 contains 414

amino acids and TGF-β3 contains 412 amino acids [143]. Binding affinities for the TGF-β type II

receptor (TGF-βRII) differ between the three TGF-β isoforms. Generally, TGF-β1 and TGF-β3 have a

higher binding affinity for the TGF-βRII [144].

TGF-β is synthesized as a larger precursor molecule. It contains a 20-30 amino acid long N-

terminal region that is crucial for secretion and a C-terminal region that is approximately 112-114 amino

acids long and contains the mature TGF-β molecule [145]. The TGF-β transcript contains a pro-region

(N-terminal region), the latency-associated propeptide, which it binds non-covalently and together forms

a small latent complex (SLC) [138]. The SLC can then bind the latent TGF-β binding protein to form the

large latent complex, which is important for the activation of TGF-β [146, 147]. Its processing occurs in

15

the Golgi apparatus where it is cleaved by the convertase endonucleases [148]. The mature form of TGF-

β is a dimer molecule of approximately 25KDa. TGF-β monomers contain 9 cysteine residues, 8 of which

form internal disulfide bonds [149]. Three of these disulfide bonds are arranged into a cysteine knot that

provides structural stability and is commonly found in growth factors [149, 150]. The ninth cysteine

residue forms a disulfide bond that links the two TGF-β monomers, forming a homodimer [149, 150].

The two main mechanisms of TGF-β activation are proteolytic cleavage and integrin-dependent

activation [151]. It has been shown that the large latent TGF-β1 complex can be proteolytically cleaved to

release active TGF-β by a variety of enzymes including MMPs, plasmin, thrombin, elastase and cathepsin

[151]. It is proposed that integrins avβ8 and avβ3 may also play a part in the activation of TGF-β by

inducing proteolytic cleavage of the large latent TGF-β1 complex [151]. The second proposed mechanism

of integrin-mediated TGF-β activation is through conformational changes in the large latent TGF-β1

complex, that occur due to force transduction by avβ3, avβ5, avβ6 and avβ8 and lead to the release of

active TGF-β1 [152, 153].

TGF-β is involved in pre-chondrogenic condensation, joint formation, growth plate signaling and

chondrocyte homeostasis [154]. In postnatal cartilage, TGF-β promotes the chondrogenic phenotype and

behaves as an inhibitor of terminal hypertrophic differentiation [155]. It has been shown to inhibit

hypertrophy of epiphyseal rat chondrocytes [156]. In human mesenchymal stromal cells, TGF-β3 was

also shown to prevent chondrocyte terminal differentiation in pellet culture [157].

1.3.2 TGF-β Signaling

There exist three main types of TGF-β receptors (I, II and III). TGF-β type I receptor (TGF-βRI)

and TGF-β type II receptor (TGF-βRII) are capable of activating signal cascades as they contain

intracellular kinase domains [149]. TGF-β type I receptors are also known as the activin receptor-like

kinase (ALK) family [138]. TGF-β has a high affinity for the TGF-βRII, but not TGF-βRI, and does not

seem to bind TGF-βRI directly [158]. TGF-βRII binds a single monomer of the TGF-β dimer. Therefore,

the binding of a TGF-β dimer usually results in the formation of a large complex that includes two sets of

receptors, each containing TGF-βRI and TGF-βRII [158].

It has been shown that chimeric receptors containing an extracellular TGF-βRII domain and an

intracellular TGF-βRI form complexes with TGF-βRI, but do not participate in TGF-β signal transduction

suggesting that formation of TGF-βRI/TGF-βRII heterodimers is necessary for signal transduction [159].

Luo et al. also showed that heterodimerization of the cytoplasmic domain of TGF-βRI, with TGF-βRII is

required for intracellular TGF-β signal transduction [160]. Erythropoietin was able to induce formation of

16

heteromeric complexes and downstream signaling only in cells expressing chimeric receptors for both

TGF-βRI and TGF-βRII, but not in cells expressing a single receptor[160]. The cytoplasmic GS domain

of TGF-βRI was also identified as important for activation of the receptor kinase domain and downstream

signaling [160]. Persson et al. showed that TGF-βRI is important for signal specificity using a chimeric

receptor with an extracellular TGF-βRI domain and an intracellular type I BMP receptor domain [161].

The signaling pathway is initiated by the binding of TGF-β to the extracellular domain of TGF-

βRII, which leads to the recruitment, phosphorylation and activation of TGF-βRI [138]. Subsequently,

downstream receptor-regulated SMAD (R-SMAD) proteins are recruited and phosphorylated. The

signaling R-SMADs are SMAD1, SMAD2, SMAD3, SMAD5, and SMAD8. SMAD4 is a common

SMAD that binds to the other SMAD complexes, while SMAD6 and SMAD7 are known as the inhibitory

SMADs [138]. The activated R-SMADs can translocate to the nucleus and regulate the downstream

expression of target genes [138].

The TGF-β type III receptor, alternatively known as Betaglycan, is not capable of signal

transduction [162]. Its extracellular domain binds ligands of the TGF-β family at the cell surface and can

present them to TGF-βRII [162]. Betaglycan may also increase the affinity of TGF-βRII to TGF-β2 [162].

Endoglin is another accessory receptor that is part of the TGF-β receptor complex and can also influence

TGF-β signaling. It promotes binding of TGF-β1 and TGF-β3 to TGF-βRII, as well as recruitment of

ALK1, favoring signaling through the SMAD1/5/8 pathway [163].

Molecules that increase SMAD turnover or disrupt SMAD-dependent transcriptional activation can

also modulate TGF-β signaling. SMAD ubiquitination regulatory factor 1 (SMURF) and SMURF2 are E3

ubiquitin-protein ligases that cause degradation of SMADs through ubiquitination and subsequent

proteosomal degradation [164]. SMURF1 induces the degradation of SMAD1 and SMAD5[164], while

SMURF2 leads to degradation of SMAD1, SMAD2 and SMAD3, depending on the cell type [165, 166].

SMURF2 is also capable of forming a complex with SMAD7 and inducing degradation of both TGF-β

receptors and SMAD7 [167]. TGF-β signaling can also be modulated by molecules that disrupt inhibit

SMAD complexes from activating target genes, such as transcriptional co-repressors c-Ski and SnoN

[168].

17

1.3.3 Canonical TGF-β Signaling Pathways in Chondrocytes

In chondrocytes, the canonical R-SMAD signaling pathway has been well defined. TGF-β signals

primarily through the ALK5 receptor which results in the phosphorylation of SMAD2 and SMAD3 [154].

After its phosphorylation and activation, SMADs 2 and 3 form a complex with SMAD4, which

translocates into the nucleus and affects the transcription of target genes. In mesenchymal stromal cells,

SMAD3 has been shown to form a transcription factor complex with SOX9, resulting in upregulation of

Col2 and induction of primary chondrogenesis [169]. The SMAD2/3 pathway has also been shown to

regulate metalloproteinase inhibitor 3 (TIMP-3) gene expression in chondrocytes [170]. Both human

articular cartilage samples and in vitro formed hyaline-like cartilage that lack expression of type X

collagen and MMP13, have been shown to only express phosphorylated SMAD2/3 [171]. SMAD2/3

signaling through activation of ALK5 by TGF-β therefore seems to promote ECM production and

accumulation, as well as the chondrogenic phenotype.

Recent research has shown that TGF-β is also capable of binding ALK1 and inducing an

intracellular cascade involving SMAD1/5/8 phosphorylation [172]. In primary bovine chondrocytes,

BMP-9 induced phosphorylation of SMAD1/5 and downstream expression of alkaline phosphatase and

type X Collagen gene, markers of hypertrophy [173]. SMAD1/5/8 signaling through activation of ALK1

seems to promote expression of terminal differentiation and hypertrophy markers. It has been shown that

SMAD1/5/8 phosphorylation requires the presence of both ALK1 and ALK5 TGF-β receptors, whereas

SMAD2/3 phosphorylation only requires ALK5 signaling [174].

The literature suggests that the SMAD2/3 and SMAD1/5/8 pathways play opposing roles in

chondrocyte differentiation and regulation of the chondrogenic phenotype [173, 174]. It has been shown

that ALK1 inhibits SMAD3-dependent transcriptional activity and type II collagen expression, induced

by TGF-β[174]. In primary bovine chondrocytes, addition of TGF-β1 led to increased SMAD3

phosphorylation and counteracted the effect of BMP-9 on SMAD1/5/8 phosphorylation and expression of

hypertrophy markers, highlighting the importance of TGF-β in maintaining the chondrogenic phenotype

[173].

The balance between SMAD2/3 and SMAD1/5/8 signaling may also be crucial during

chondrogenesis. Blocking either SMAD2/3 or SMAD1/5/8 phosphorylation through chemical inhibitors

of ALK1 and ALK5 in human MSCs, inhibited expression of type II collagen, suggesting the

involvement of both pathways during chondrogenesis [171]. Blocking SMAD2/3 signaling after the onset

of chondrogenesis resulted in decreased type II collagen expression, while blocking SMAD1/5/8

signaling decreased expression of alkaline phosphatase, type X collagen and MMP13 [171]. These results

18

support that after the onset of chondrogenesis, SMAD2/3 signaling is involved in ECM production while

SMAD1/5/8 signaling is involved in hypertrophy and terminal differentiation.

There is some evidence to suggest that additional molecules in the TGF-β pathway may regulate

ALK5-SMAD2/3 and ALK1-SMAD1/5/8 signaling. The accessory receptor Endoglin was found to

inhibit phosphorylation of SMAD2/3, SMAD3 dependent transcriptional activity and production of ECM;

conversely it enhanced phosphorylation of SMAD1/5 [175]. The oncoprotein Ski, a known inhibitor of

the TGF-β/SMAD3 mediated transcription, was found to increase hypertrophic differentiation including

expression of type X collagen, Runx2 and osteocalcin in chick upper sternal chondrocytes[176].

The regulation of SMAD1/5/8 and SMAD2/3 signaling and the balance between the

chondrogenic and hypertrophic phenotype may also be affected by mechanical stimulation. In human

chondrocytes, cyclic tensile strain was found to result in release of TGF-β1, increased nuclear

translocation of SMAD2/3 and SOX9, and increased expression of COL2 [177]. In MSCs cultured in a 3D

scaffold, dynamic compression decreased phosphorylation of SMAD1/5/8 and decreased expression of

hypertrophic markers [178]. Conversely, dynamic compression increased SMAD2/3 phosphorylation and

expression of chondrogenic markers aggrecan and type II collagen [178]. Treatment with chemical

inhibitor of ALK5 resulted in increased SMAD1/5/8 phosphorylation and expression of hypertrophic

markers, while decreasing expression of chondrogenic markers [178]. This study suggests that there is

crosstalk between the integrin pathway and the TGF-β pathway during chondrogenesis in MSCs, which

regulates the chondrogenic and hypertrophic phenotypes [178].

The studies summarized above suggest that TGF-β signaling through the canonical pathway plays

a role in regulating the chondrocyte phenotype and cartilage homeostasis. There is also evidence that

suggests involvement of the non-canonical signaling pathways. It is quite likely that signaling through

both pathways is involved in regulating the chondrogenic phenotype during redifferentiation.

1.3.4 Non-Canonical TGF-β Signaling Pathways in Chondrocytes

It has also been shown that TGF-β can signal through non-canonical pathways including mitogen

activated protein kinases (MAPK), Rho-GTPases, and phosphatidylinositol-3-kinase (PI3K)/protein

kinase B (PKB) pathways [179, 180]. Much of the current literature focuses on non-canonical signaling

during chondrogenesis, while not much is known about its role in maintaining cartilage homeostasis.

Activation of TGF-β activating kinase (TAK1) occurs through Type I BMP and TGF-β receptors [181]

and TAK1 has previously been shown to take part in the non-canonical signaling pathways. Its

downstream effectors are three MAP kinases: p38, c-Jun N-terminal kinases (JNK) and extracellular

19

signal-regulated kinases (ERK). TAK1 plays an essential role during development, specifically in

differentiation and proliferation of growth plate chondrocytes [154]. BMPs are the main activators of

TAK1 during cartilage development [182]. TAK1 deletion in murine chondrocytes leads to decreased

proliferation, survival, and the delayed onset of chondrocyte hypertrophy [183]. Deletion of TAK1 in a

post-natal murine model led to growth delays and reduced BMP-2-dependent SOX9 promoter activation,

as well as aggrecan and type II Collagen expression and accumulation in the ECM [184]. This suggests

that TAK1 influences SOX9 gene expression and regulates the development of postnatal articular

cartilage.

Downstream of TAK1 the most studied signaling molecule is p38. Studies in mouse models have

shown that p38 signaling inhibits chondrocyte proliferation and terminal differentiation [154]. Articular

cartilage defects develop in the knee joints of one-year-old mice heterozygous for the dominant negative

p38 transgene [185]. In chondrocytes, p38 has also been shown to signal by forming a phosphorylated

complex with SMAD3 [186]. Through this pathway, p38 signaling has been shown to inhibit the

expression of type X collagen and chondrocyte hypertrophy [186]. Conversely, it has also been shown to

induce the expression of MMP13 [187]. MMP13 may not only be a marker of hypertrophy but has been

implicated in matrix remodeling and may be involved in generating tissue with a proper architecture

[188]. There is little known about the interactions between non-canonical and canonical signaling

pathways in chondrocytes.

Chondrocytes also signal through another non-canonical pathway that involves the MAP kinases

ERK1 and ERK2. ERK1 null mice do not exhibit major skeletal defects [189] and ERK2 null mice are

embryonic lethal due to their inability to form the ectoplacental cone and the extra-embryonic ectoderm

[190]. When MEK1 (a kinase upstream of ERK1) is constitutively activated in mice, hypertrophy of

growth plate chondrocytes is inhibited and endochondral ossification is delayed [191]. There is no effect

on chondrocyte proliferation. The chondrocytes also exhibit reduced SOX9 and type X collagen

expression [191].

Though not yet confirmed in chondrocytes, the PI3K/BKB pathway is also capable of crosstalk

with the canonical TGF-β signaling pathways. It forms a complex with SMAD3 in HEK293T cells,

decreasing its phosphorylation and nuclear localization. The formation of this complex is inhibited by

TGF-β treatment and enhanced by insulin treatment [192].

We have shown that Rho-associated kinase protein (ROCK) inhibition in bovine P2 chondrocytes

treated with TGF-β1 results in increased SOX9 and Aggrecan expression, decreased α-smooth muscle

actin expression, and decreased cell contraction in passaged bovine chondrocytes (Parreno et al.,

unpublished). There is also data that suggests RhoA/ROCK signaling may inhibit chondrogenic genes in a

20

context dependent manner. Inhibition of RhoA/ROCK signaling in three-dimensional micromass cultures

of mesenchymal cells led to increased SOX9 expression but decreased transcript levels of Col2 and ACAN

[193]. In contrast, inhibition of RhoA/ROCK signaling in monolayer culture increased both SOX9 activity

and COL2 and ACAN expression [193, 194]. Inhibition of ROCK during passaging has also been shown

to decrease the dedifferentiation of monolayer-cultured human articular chondrocytes and increase their

ability to redifferentiate in subsequent pellet culture [195]. This data suggests that TGF-β may also signal

through the RhoA/ROCK pathway.

1.3.5 TGF-β Signaling in Osteoarthritis

The involvement of TGF-β in OA development has been extensively investigated, due to its

effect on cartilage homeostasis. Immunohistochemical analysis of human articular cartilage revealed

localized changes in expression of TGF-β and its receptors [196]. The articular cartilage exhibited

decreased TGF-β1 expression in degraded cartilage, increased TGF-β1 and TGF-β3 expression in

osteophytes and decreased expression of TGF-βRII in the fibrillated cartilage [196]. Chondrocytes in

osteoarthritic cartilage undergo a phenotypic change that resembles terminal differentiation including

hypertrophy and expression of type X collagen and MMP13 [197]. The literature suggests that changes in

chondrocyte metabolism and articular cartilage homeostasis in OA are partially due to changes in TGF-β

signaling. Age related changes, such as decreased expression of ALK5 and decreased phosphorylation of

SMAD2/3, have been shown in murine articular cartilage [198]. These findings are of interest as OA is a

disease associated with age.

It has been shown that there is an association between development of OA and decreased ALK5

expression in mice [198]. Conversely, both the age-related and OA associated decreases in ALK1

expression are not as great as the decreases in ALK5 expression [172]. It is therefore postulated that in

OA, the increased ratio of ALK1/ALK5 expression causes a shift in TGF-β signaling from SMAD2/3 to

SMAD1/5/8 [154, 172]. Overexpression of ALK5 in murine chondrocytes promotes SMAD2/3 signaling,

leading to an increased expression of ACAN [172]. Increasing signaling through SMAD1/5/8 through

constitutive expression of ALK1 results in increased MMP13 expression [172]. Homozygous murine

knockdowns of SMAD3 also develop a disease similar to human OA [199]. This data suggests that the

decrease in ALK5 signaling may be responsible for decreases in type II collagen and aggrecan expression

and therefore ECM production and homeostasis.

Other components of the TGF-β signaling pathway have also been implicated in the development

of OA. The deletion of TGF-βRII in murine chondrocytes resulted in a phenotype resembling OA,

including increased MMP13 expression [200]. In human OA cartilage, there are increased

21

phosphorylation levels of non-canonical signaling molecules p38 and JNK [201]. Lastly, it has been

shown that when OA is induced in homozygous Akt1-deficient mice, there is decreased osteophyte

formation [202].

The current literature suggests that there is an age-related shift in TGF-β signaling that

contributes to the development of OA. In young, healthy cartilage, TGF-β maintains homeostasis by

stimulating ECM production and inhibiting chondrocyte hypertrophic differentiation through ALK5-

SMAD2/3 signaling. As chondrocytes age, there is a shift towards ALK1-SMAD1/5/8 signaling that

results in the expression of catabolic enzymes leading to degradation of ECM [203]. This may in part

explain the inability of chondrocytes to repair degenerated or damaged cartilage.

1.3.6 Redifferentiation of Passaged Chondrocytes using TGF-β

TGF-β has been shown to promote redifferentiation of passaged chondrocytes in several animal

studies [204-208]. In passaged porcine articular chondrocytes, adenoviral mediated co-transfection of

TGF-β3 and type I collagen short hairpin RNA (shRNA) resulted in decreased expression of type I

collagen and increased expression of type II collagen and aggrecan, at both the gene and protein level

[204]. Similar results were achieved when using a lentiviral vector to co-transfect TGF-β3 and type I

collagen shRNA into chondrocytes [205]. These results suggest that TGF-β3 may promote the

chondrogenic phenotype during redifferentiation [204, 205].

Tekari and colleagues showed that if bovine chondrocytes from young animals are passaged more

then 3 times, the addition of TGF-β1 is required for cartilage tissue formation. Chondrocytes treated with

TGF-β1 generated pellets that showed increased expression of type II collagen and aggrecan, when

compared to non-treated controls [206]. These results suggest that TGF-β may promote the

chondrogeneic phenotype in passaged cells. TGF-β1 was also found to increase hypertrophic marker type

X collagen and fibroblast matrix molecule type I collagen in bovine passaged chondrocytes [206]. They

also showed that transcript levels of TGF-βR2, TGF-βR1 and TGF-β2 decrease with passaging,

suggesting that a decrease in endogenous TGF-β signaling may play a role in the loss of the chondrogenic

phenotype [206]. van Osch and colleagues have shown that in passaged chondrocytes obtained from

young rabbit ears, treatment with IGF-1 and TGF-β2 in serum free 3D monolayer culture conditions lead

to increased GAG synthesis and number of type II collagen positive cells. The results for

glycosaminoglycan were replicated when using human chondrocytes, however type II collagen was

increased by IGF-1 and TGF-β2 to a much lesser extent [208].

To the best of our knowledge, there is limited literature that shows the ability of TGF-β to

22

promote redifferentiation and enhance tissue formation of passaged human OA and non-diseased

chondrocytes [56, 209-212]. Goldberg and colleagues showed that pellet culture of passaged OA

chondrocytes in serum free media (SFM) with TGF-β resulted in increased production of

glycosaminoglycan and type II collagen, though the integrity of tissue formed was not assessed [211]. It

has also been shown that the addition of TGF-β1 or TGF-β2 in combination with the addition of insulin or

IGF-1 is required for increased expression of COL2 and ACAN by passaged adult human articular

chondrocytes in suspension culture [210]. Non-OA human chondrocytes passaged in media supplemented

with FGF-2 and TGF-β, then subsequently cultured as pellets in SFM with TGF-β and Dexamethasone

expressed higher levels of COL2 and ACAN, as well as had greater glycosaminoglycan content, compared

to cells cultured in media supplemented with serum but without TGF-β [56].

Interestingly, treatment with TGF-β seems to have different affects during passaging and

redifferentiation. Depletion of TGF-β during passaging was found to improve the redifferentiation

capacity of passaged OA chondrocytes, in pellet cultures [209]. Narcisi and colleagues found that treating

with a TGF-β antibody during passaging lead to increased expression of COL2 and ACAN, as well as

decreased gene expression of hypertrophic markers type X collagen and MMP-13 in 3D culture [213].

The presence of TGF-β during passaging appears to negatively influence redifferentiation capacity [213],

whereas during redifferentiation it seems to promote the chondrogenic phenotype and tissue formation.

We have demonstrated that exogenous TGF-β3 can promote redifferentiation of passaged human

chondrocytes and tissue formation (Bianchi et al., unpublished). To the best of our knowledge, there is

very limited literature that investigates TGF-β signaling during the redifferentiation of passaged human

chondrocytes. The literature on TGF-β signaling in OA and during chondrogenesis in MSCs suggests the

balance between ALK1 and ALK5 signaling regulates the chondrogenic phenotype and therefore may be

important during redifferentiation. It is necessary to further characterize the redifferentiation process and

elucidate the signaling mechanisms by which redifferentiation enhances tissue formation occur. This will

facilitate identification of ways to further enhance redifferentiation.

23

1.3.7 Summary

Recent research activity has turned to cartilage tissue-engineering techniques, as current biological

treatments are not optimal for the repair of cartilage defects. The challenge has been to bioengineer a

continuous layer of articular cartilage that resembles native cartilage in its multi-zonal organization and

ECM abundant in type II collagen and aggrecan. We have previously shown that a scaffold-free culture

approach using defined serum-free media promotes tissue formation in bovine passaged chondrocytes

[105]. The formation of articular cartilage tissue expressing high levels of aggrecan and type II collagen is

also possible in human OA chondrocytes with the addition of TGF-β3 to this system (Bianchi et al.,

unpublished). This is not unexpected, as TGF-β has been shown to play a role in promoting the

chondrogenic phenotype and the differentiation of MSCs to chondrocytes. The balance between two

TGF-β canonical signaling pathways ALK5-SMAD2/3 and ALK1-SMAD1/5/8 has been shown to

regulate chondrocyte phenotype and cartilage homeostasis. There is also evidence that suggests

involvement of the non-canonical signaling pathways. This project will further characterize the

redifferentiation process by examining changes in TGF-β signaling pathway molecules throughout

redifferentiation and the signaling mechanisms through which TGF-β enhances tissue formation.

24

1.4 Hypothesis and Objectives Characterizing the TGF-β signaling pathway involved in redifferentiation of passaged human

chondrocytes will improve our understanding of this process, which will allow us to better understand and

improve upon cartilage tissue engineering. This will also allow us to better understand how

redifferentiation and tissue formation differ between patients and contribute to our ability to predict the

quality of articular cartilage formation.

Hypothesis: TGF-β3 treatment results in TGF-β receptor level changes and signaling through both

ALK1 and ALK5 in passaged human chondrocytes, to stimulate redifferentiation and accumulation of

extracellular matrix molecules in vitro.

Specific Objectives:

1. Characterize ALK1 and ALK5 receptor levels in human chondrocytes during passaging.

The first objective was to investigate if there were changes in the levels of ALK1 and

ALK5 receptors throughout passaging that may allow passaged cells to respond to TGF-β3 once

they are placed in redifferentiation conditions.

2. Characterize and determine signaling pathways involved during redifferentiation and tissue

formation in response to TGF-β3 treatment.

The second objective was to investigate the changes in the levels of TGF-β signaling

pathway receptors and effector molecules throughout the redifferentiation process.

3. Elucidate the initial signaling pathway(s) activated by TGF-β3 that lead to matrix accumulation

by passaged human chondrocytes during redifferentiation.

The third objective was to identify the signaling pathway involved in ECM accumulation

during early redifferentiation by assessing the effect of specific chemical inhibitors of the TGF-β

pathway and small interfering RNA (siRNA) on ECM molecule gene expression and synthesis.

25

Chapter 2 TGF-β3 Signaling During Redifferentiation and Tissue Formation by Passaged Human OA Chondrocytes 2 TGF-β3 Signaling During Redifferentiation and Tissue Formation by Passaged Human OA Chondrocytes

2.1 Introduction Biological repair of cartilage defects, using methods such as microfracture and autologous

chondrocyte implantation, is one of the current approaches used to treat damaged joints. These therapies

have limited long-term potential as they often result in a biomechanically inferior mixture of hyaline and

fibrocartilage that differs from native cartilage in ECM composition and degrades over time due to

inadequate mechanical properties [49, 50]. To overcome the limitations presented by these treatments,

there has been increasing interest in bioengineering articular cartilage tissue suitable to use for the repair

of cartilage defects. Native articular cartilage has a distinct multi-zonal 3D architecture and biochemical

composition, with an ECM composed primarily of type II collagen and aggrecan. These aspects are

crucial to the biomechanical function of cartilage and have proven difficult to replicate in the in vitro

formed tissue [53]. To support the formation of articular cartilage tissue, strategies have employed the

use of scaffolds, substrates and stimulatory factors, as well as various cell sources. In an attempt to

overcome the limitations associated with scaffolds, there has been a shift towards the use of scaffold-free

approaches, where the cells themselves synthesize ECM without the aid of an exogenous scaffold [93].

We have previously shown that a scaffold-free approach using defined serum-free media

containing high-glucose, insulin, and dexamethasone promotes the formation of hyaline-like cartilage

tissue by passaged bovine chondrocytes [105]. It was shown that an insulin-dependent signal transduction

cascade was responsible for the upregulation of SOX9 and re-expression of the chondrogenic phenotype

in passaged cells [105]. We have subsequently shown that human OA chondrocytes will re-express

aggrecan and type II collagen, as well as form cartilage tissue with a superficial zone when TGF-β3 is

added to this system (Bianchi et al., unpublished).

Previous studies have demonstrated redifferentiation with TGF-β3[56, 209-212]. The use of

aggregate cultures in these studies can result in uneven ECM deposition and cell death at the center of the

tissue due to limited diffusion of nutrients [93]. Furthermore, other studies used multiple growth factors

during passaging/redifferentiation to promote redifferentiation and cartilage tissue formation [56, 209-

211]. Our scaffold-free self-assembly method utilizes only TGF-β3 and ITS (source of insulin) in serum-

free conditions (Bianchi et al., unpublished). With this method, we are able to form a continuous layer of

articular-like cartilage tissue on Millipore® membranes with evenly distributed ECM and evidence of the

26

appropriate zonal architecture (Bianchi et al., unpublished). The use of fewer growth factors in our system

will increase cost-effectiveness and clinical applicability relative to some of the previously described

methods.

TGF-β is a growth factor with pleotropic effects and is involved in chondrogenic differentiation

of MSCs, joint formation, and chondrocyte homeostasis [154]. It has been shown to promote the

chondrogenic phenotype and proteoglycan synthesis and to inhibit terminal hypertrophic differentiation in

rat and human chondrocytes [155-157]. With the involvement of this growth factor in cartilage

homeostasis, it is not surprising that the addition of exogenous TGF-β3 promotes chondrocyte

redifferentiation and tissue formation. What remains to be characterized is the signaling pathway(s)

through which TGF-β3 mediates its effects during the redifferentiation process.

The canonical R-SMAD signaling pathways are well defined in chondrocytes. TGF-β signals

through the ALK5 receptor resulting in SMAD2 and SMAD3 phosphorylation and subsequently, the

formation of a complex with SMAD4 [154]. The complex then translocates into the nucleus and has been

shown to effect the transcription of target genes such as COL2 [169]. TGF-β also signals through the

ALK1 receptor, resulting in the phosphorylation of SMAD1/5/8 [172]. In primary bovine chondrocytes,

SMAD1 and SMAD5 phosphorylation in response to BMP-9 resulted in downstream expression of

alkaline phosphatase and type X collagen, markers of hypertrophy [173]. The two canonical signaling

pathways seem to play opposing roles in maintaining the chondrogenic phenotype as signaling through

ALK5-SMAD2/3 promotes ECM production while signaling through ALK1-SMAD1/5/8 promotes

terminal differentiation.

TGF-β has also been shown to signal through several non-canonical pathways, including TGF-β

activating kinase (TAK1) [181]. In chondrocytes, TAK1 has been implicated in maintaining cartilage

homeostasis. The deletion of TAK1 in murine chondrocytes leads to decreased proliferation and survival,

as well as loss of the elbow joint interzone and delayed onset of chondrocyte hypertrophy [183]. In

another study deletion of TAK1 in a murine model resulted in decreased aggrecan and type II collagen

expression, suggesting that this molecule plays an important role in regulating ECM synthesis [184].

Downstream of TAK1, there are three main effector molecules, p38, JNK and ERK, of which p38 is most

commonly identified in mediating chondrocyte function. In a murine model, animals heterozygous for the

dominant negative p38 transgene develop cartilage defects in their knee joints [185]. Studies in SMAD3

knockout C57BL/6 mice have shown that p38 signaling can inhibit expression of type X collagen and

chondrocyte hypertrophy [186].

27

Given the evidence that both the canonical TGF-β signaling pathways and the non-canonical

signaling pathways play a role in regulating the chondrocyte phenotype and cartilage homeostasis, we

hypothesized that signaling through both ALK1 and ALK5 as well as the non-canonical pathway

involving TAK 1-p38 are involved in the TGF-β3 induction of chondrocyte redifferentiation and cartilage

tissue formation. The specific objectives of this study were to (1) characterize ALK1 and ALK5 receptor

levels in human chondrocytes during passaging; (2) characterize and determine the signaling pathways

involved during redifferentiation and tissue formation; and (3) elucidate the initial signaling pathway(s)

activated by TGF-β3 that leads to matrix accumulation by passaged human chondrocytes during

redifferentiation. Characterizing the signaling pathway(s) may result in the identification of

pharmacological agents that will further enhance formation of bioengineered cartilage.

2.2 Materials and Methods

2.2.1 Isolation and Culture of Chondrocytes

All cartilage was obtained from femoral heads and condyles removed during arthroplasty

following informed consent using a protocol approved by Mount Sinai Hospital Research Ethics Board.

Full thickness articular cartilage was dissected from low-grade OA areas. The cartilage was digested

using 0.5% protease (Sigma-Aldrich) in Dulbecco’s Modified Eagle’s Medium (DMEM) for 30 minutes

and subsequently in 1% collagenase (Roche) in DMEM at 37°C for 18 hours. The chondrocytes were

filtered through a sterilized mesh strainer (100µm pore size) to eliminate bone fragments and were seeded

in monolayer culture at a density of 2000cells/cm2. The cells were cultured in DMEM supplemented with

20% FBS and passaged twice (P2). 1% Trypsin (Gibco) was used to harvest the chondrocytes from the

polystyrene flasks and cells were reseeded at the same density.

P2 chondrocytes were seeded onto Type II Collagen coated Millipore® membranes in 24 well

plates as previously described, at 2 million cells/filter (diameter 12mm; EMD Millipore) [105]. Cells were

cultured in high glucose DMEM (4.5g/L) serum-free media (SFM), containing dexamethasone (0.1nM),

1% ITS+ (BD, Bioscience, MA), L-ascorbic acid (Sigma; 100µg/mL) and L-Proline (40µg/mL) and

pyruvate (110µg/mL), as previously described [105] in the presence or absence of TGF-β3 (10ng/mL)

(R&D Systems). For select experiments, cultures were treated with the following inhibitors 1 hour before

TGF-β3 treatment (see Table 3.1): 10µM Y- 27632 (Cayman Chemical company), 100nM Wortmannin

(Sigma Aldrich), 10µM U0126 (Calbiochem), 10µM SB431542 (Sigma-Aldrich), 10µM SB203580

(Calbiochem), 3µM SIS3 (Sigma-Aldrich), 10µM CCG-1423 (Cayman Chemical Company), 10µM

Dorsomorphin (Abcam) or 5µM (5Z)-7-Oxozeaenol (Sigma-Aldrich). Controls were treated with carrier

28

alone (DMSO). These concentrations were chosen based on previous work that showed that these

concentrations were effective in bovine chondrocytes (data not shown).

Table 3-1: Summary of Inhibitors used.

2.2.2 siRNA Transfection

Chondrocytes were seeded onto Type II Collagen coated Millipore® membranes as described

above. After 24 hours the media was changed; to SFM supplemented with 10ng/mL of TGF-β3 (R&D

Systems) for select tissues. Signal Silence siRNA against TAK1 or p38 MAPK, (15nM, Cell Signaling

Technology), Signal Silence Control siRNA (scrambled) (Cell Signaling Technology) and the

Lipofectamine® 3000 Transfection Kit (Life Technologies) were used to transfect chondrocytes,

according to the manufacturer’s instructions for 24-well plates. Three control groups were utilized

including non-transfected control (NT), as well as, control tissues and TGF-β3 treated tissues both

transfected with scrambled control siRNA.

2.2.3 Gene Expression Analysis

Total RNA was extracted from tissue samples using TRIzol® (Life Technologies) according to

the manufacturer’s instructions, after flash freezing with liquid nitrogen and grinding. RNA was

quantified using the Nanodrop 1000 spectrophotometer (Thermo Scientific). 1µg of RNA was reverse

Inhibitor Concentration Targets

Y-27632 10uM ROCK

Wortmannin 100nM PI3K

U0126 10uM MEK1/MEK2

SB431542 10uM ALK5

SB203580 10uM p38, PKB (Akt)

SIS3 3uM SMAD3

CCG-1432 10uM MRTF-A/SRF

(5Z)-7-oxozeanol 5uM TAK1

Dorsomorphin 10uM ALK1/ALK2/

ALK3/ALK6, AMPK

29

transcribed using SuperScript III Reverse Transcriptase (Life Technologies) and amplified using a

Mastercycler Thermocycler (Eppendorf, AG). Semi-quantification of gene expression was performed

using Fast SYBR Green I Master Mix (Applied Biosystems, Foster City, CA, USA), gene specific

primers (See Table 3.2) and a LightCycler 96 Real- Time PCR System (Roche). Pffafl method was used

to calculate the mean relative quantification values [214]. 18s rRNA levels were used as an endogenous

control[214].

Primer Forward Sequence 5’ to 3’ Reverse Sequence 5’ to 3’

18S GTAACCCGTTGAACCCCATT CCATCCAATCGGTAGTAGCG

Type II Collagen

(Col2) TCTGTGTCTGTGACACTGGG GAGGTCAGTTGGGCAGATGG

Type I Collagen

(Col1) [105] CGGCTCCTGCTCCTCTTAG CACACGTCTCGGTCATGGTA

Aggrecan

(ACAN) [105] TGGGACTGAAGTTCTTGGAGA GCGAGTTGTCATGGTCTGAA

SOX9 [215] GTACCCGCACTTGCACAAC GTGGTCCTTCTTGTGCTGC

TAK1 CAACCACAGGCAACGGAC TGACACTGGGACTGGATGAC

ALK5 [216] CGACGGCGTTACAGTGTTTCT CCCATCTGTCACACAAGTAAA

ALK1 [172] GACTCAAGAGCCCCAATGTG GGTCGGCGATGCAACAC

TGF-βRII ATCACACTCCATGTGGGAGG GACAGAGTAGGGTCCAGACG

SMAD5 CGCGAGACTTGACCCAATGA AAACAAGCTGGCCATTGACG

SMAD2 CCGACACACCGAGATCCTAAC AGGAGGTGGCGTTTCTGGAAT

Table 3-2: Summary of Primers used.

2.2.4 Western Blot Analysis

Protein was extracted for western blot from tissues or cells using RIPA buffer (50 mM Tris HCl,

150 mM NaCl, 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS) with complete mini-protease inhibitor

(Roche), 20µM of sodium fluoride and 10µM sodium orthovanadate. Total protein was quantified using

bicinchoninic acid protein assay (Thermo Scientific). Protein samples were heated for 10 minutes at 98°C

in Laemmli buffer and 60µg of protein sample was separated on a 12% sodium dodecyl sulfate

polyacrylamide gel electrophoresis (SDS–PAGE) gel and wet transferred onto nitrocellulose membrane

(Bio Rad). The membranes were blocked with 5% skim milk (in TBS) and then incubated in primary

antibody (phospho p38 at 1:350, p38 at 1:1000; Life Technologies, ALK5 at 1:350; Santa Cruz

30

Biotechnology Inc.) overnight at 4oC. Membranes were washed in 0.005% Tween/PBS and subsequently

incubated with horseradish peroxidase (HRP)-conjugated secondary antibody (1:20,000; Abcam) for an

hour. AmershamTM ECL SelectTM Western Blotting Detection Reagent (GE Healthcare) was used for

chemiluminescent detection of HRP. Semi-quantification of protein bands was preformed through

densitometry using Image J software.

For immunocytochemistry, cells on glass slides were blocked with 20% goat serum in 0.2% triton

and then incubated with primary antibody reactive to ALK1 (1:50, Santa Cruz Biotechnology Inc.) or

ALK5 (1:100, Santa Cruz Biotechnology Inc.), overnight at 4°C. The cells were then incubated with

Alexa-594 goat anti-rabbit IgG (1:1000, Life Technologies) and 4′,6-diamidino-2- phenylindole (DAPI)

(1:1500, Life technologies) was used to stain the nuclei. Immunoreactivity was visualized using a

fluorescent OptiGrid microscope (Leica).

2.2.5 Analysis of matrix synthesis and retention

24 hours after seeding, cells were incubated with 2µCi/mL each of 3H-proline and 35S-SO4

(Perkin Elmer) to quantify newly synthesized matrix molecules, collagen and proteoglycan. Tissues were

harvested 48 hours after the addition of isotope. The medium was collected and separated into two parts.

To precipitate newly synthesized 3H-proline labeled collagen present in the media, 70% ammonium

sulfate was added slowly (3:4 ratio). For precipitation of 35S-SO4 labeled proteoglycan in the media,

100% ice-cold ethanol was added to the media (3:2 ratio). Samples were left at 4°C overnight, then

centrifuged at 14,000g at 4 °C for 30 minutes. The pellets were washed with 70% ice-cold ethanol three

times, then dried for 15 minutes at room temperature and resuspended in 10% SDS in 0.1 M Tris buffer

(pH 7.0) to solubilize the collagen or 4M guanidinium hydrochloride to solubilize the proteoglycan. To

quantify the newly synthesized collagen (3H-proline) and proteoglycan (35S-SO4) retained in the tissue,

tissue samples were digested in 40µg/mL of papain (Sigma) at 65°C for 48 hours. Aliquots were mixed

with Biosafe II scintillation cocktail (RPI Corp.) and radioactivity counted using a LS6500 Scintillator

(Beckman Coulter) and normalized to DNA content.

2.2.6 Biochemical Analysis

Tissue thickness was measured using calipers, by one observer blinded to the experimental

condition. Tissue samples were digested for 48 hours using 40µg/mL papain (Sigma) in digestion buffer

(pH 6.2, 20mmol/L ammonium acetate, 2mmol DTT and 1mmoL/L EDTA) at 65°C. DNA content was

quantified using Hoechst 33258 dye assay and fluorometry (excitation wavelength 365nm, emission

wavelength 458nm). The standard curve was generated using calf thymus DNA (Sigma), as previously

31

described [217]. Collagen content was quantified using a hydroxyproline assay [218]. Briefly, papain

digest aliquots were acid hydrolyzed at 110°C for 18 hours. Chloramine T/Erlich’s Reagent (Sigma-

Aldrich) assay and spectrophotometry (wavelength 560nm) was used to quantify hydroxyproline content.

L-hydroxyproline (Sigma- Aldrich) was used to produce a standard curve. Sulfated glycosaminoglycan

content was quantified using a dimethylmethylene blue (Sigma-Aldrich) dye binding assay and

spectrophotometry (wavelength 525nm). Chondroitin sulfate (Sigma) was used to generate the standard

curve [219].

2.2.7 Histology and Immunohistochemistry

Tissue samples were fixed in 10% formalin for 24 hours at 4°C, then placed in a 30%

sucrose/PBS solution for an additional 24 hours at 4°C. The tissues were embedded in Tissue-Tek® OCT

(Sakura), sectioned at 7µm thickness and stained with haematoxylin and eosin or toluidine blue. For

immunohistochemistry, tissue sections were digested with 0.4% pepsin (w/v) in TBS-HCL (pH 2.0),

blocked with 20% goat serum in 0.2% triton and then incubated with primary antibody reactive with Type

I collagen (1:300, CL-50111AP-1 Cedarlane) or Type II collagen (1:300, Millipore, MAB8887) overnight

at 4°C. Heat antigen retrieval was used for ALK1 (1:100, Santa Cruz Biotechnology Inc.) and ALK5

(1:200, Santa Cruz Biotechnology Inc.); tissues were boiled for 20 minutes in Dako Target Retrieval

Solution, pH9 (Tris/EDTA buffer; Dako). No antigen retrieval was used for Aggrecan staining (1: 500

AHP0022, ThermoFisher Scientific). Immunoreactivity was visualized by incubating with Alexa-488

goat anti-rabbit IgG (1:1000, Life Technologies), Alexa-488 goat anti-mouse IgM (1:1000, Life

Technologies) or Alexa-594 goat anti-mouse (1:1000, Life Technologies) secondary antibody depending

on the source of the primary antibody. DAPI (1:2000, Life technologies) was used to stain the nuclei.

2.2.8 Statistics

All conditions were done in triplicate. For western blots, protein extracted from three samples

was combined and one blot was performed per patient. All experiments were repeated three times, unless

otherwise stated and results are expressed as mean ± standard error of the mean (SEM). Significance was

determined using ANOVA and Tukey’s post hoc for multiple time points and T-test when two groups

were compared. Significance was assigned at p ≤0.5.

32

2.3 Results

2.3.1 Characterization of ALK1 and ALK5 Expression Throughout Passaging

Levels of TGF-β receptors were determined in chondrocytes at passage 0 (P0), passage 1 (P1)

and passage 2 (P2). There was increased gene expression of ALK1 from P0 to P2 (p=0.050) and a trend

towards TGF-βRII increase between P1 to P2 (p=0.077, Figure 3-1). A significant decrease in ALK5 gene

and protein expression was present between P0 and P2 (p<0.05, Figure 3-1). Chondrocytes from P0

through to P2 in three patients expressed both ALK1 and ALK5 as determined by immunocytochemistry

(Figure 3-1 & Figure 4-1).

33

Figure 3-1: ALK5 Decreases in Chondrocytes During Passaging.

Expression of ALK1 and ALK5 in monolayer cultured chondrocytes, from passage 0 through to passage 2, as characterized by (A) gene expression analysis, (N=4 patients), (B) immunocytochemical staining and (C) western blot analysis (N=3 patients). All data is expressed as fold increase relative to P0. Please note the varying y-axes. Data is expressed as mean of triplicate cultures; the horizontal bar indicates mean ± SEM. Representative immunocytochemical images and western blot images (patient 3) are presented. Green dot indicates patient 1, red dot indicates patient 2, blue dot indicates patient 3 and pink dot indicates patient 4. *, p < 0.05. All images are of equivalent magnification.

34

2.3.2 Characterization of ALK1 and ALK5 Expression during Redifferentiation

and Tissue Formation

Levels of TGF-β signaling pathway molecules were characterized during chondrocyte

redifferentiation and subsequent cartilage tissue formation. In tissues treated with TGF-β3, there was an

increase in ALK5 gene expression during early redifferentiation (24 hours and 48 hours), compared to the

non-treated control group at the same time point (p<0.05, Figure 3-2). There was a trend towards

increased ALK5 gene expression in TGF-β3 treated group of tissues at the 2 week time point, when

compared to the 2 week control group of tissues (p = 0.06, Figure 3-2). The gene expression increase

correlated with a trend towards increased ALK5 protein expression, as demonstrated by western blot

analyses (Figure 3-3 and Figure 4-2). Conversely, TGF-βRII gene expression levels decreased in the

TGF-β3 treatment group, at both the 48 hour and 1 week time points compared to non-treated control

group of tissues (p<0.05, Figure 3-2). TGF-β3 was found to have no consistent effect on gene expression

of SMAD2 and SMAD5 over time (Figure 4-3).

35

Figure 3-2: TGF-β3 Increases Expression of TGF-β Receptor ALK5 and Decreases Expression of TGF-β Receptor II (TGF-βRII) During Redifferentiation and Tissue Formation.

Gene expression of TGF-β receptors ALK1, ALK5 and TGF-βRII were determined for up to 3 weeks of 3D culture in the presence or absence of TGF-β3. Data expressed as a fold change relative to P2. Please note the varying y-axes. Data is expressed as mean of triplicate cultures; the horizontal bar indicates mean ±SEM (N=3 patients). Red dot indicates patient 2, orange dot indicates patient 5, purple dot indicates patient 6, CON = non-treated control. *, p < 0.05 in comparison to non-treated controls at each time point.

36

Figure 3-3: ALK1 and ALK5 are Expressed in Chondrocytes During Redifferentiation and Tissue Formation.

ALK1 and ALK5 levels were determined following 1, 2, or 3 weeks of 3D culture in the presence or absence of TGF-β3 by (A) immunostaining of histological sections and (B) western blot analysis. The horizontal bar indicates mean ±SEM (N=3 patients). Green dot indicates patient 1, blue dot indicates patient 3 and pink dot indicates patient 4. Representative immunohistochemical images (patient 3) and representative western blot images (patient 4) are presented. CON = non-treated control.

37

2.3.3 Characterization of Tissue Formation in Response to TGF-β3

The effect on TGF-β3 on tissue formation was assessed by examining expression of downstream

target genes, including the chondrogenic genes (Aggrecan (ACAN), SOX9 and Type II Collagen (Col2))

and the fibroblast matrix gene (Type I Collagen (Col1)). TGF-β3 treatment resulted in increased

expression of the chondrogenic genes. SOX9, ACAN and Col1 gene expression as early as 24 hours

(p<0.05, Figure 3-4). By one week ACAN expression was approximately 10 fold higher than the untreated

controls at that time point (Figure 3-4) ACAN expression was increased significantly in TGF-β3 treated

tissues at 2 weeks (p<0.05), compared to the untreated controls at that time point. Col2 expression

increased significantly in TGF-β3 treated tissues by 1 week (p<0.05) of redifferentiation culture whereas

the 3 week time-point only showed a trend to increased expression (p=0.063), compared to the untreated

controls. It should be noted that the magnitude of response to TGF-β3 treatment varied between the three

patients. For example, SOX9 expression remained increased at 48 hours, 2 weeks and 3 weeks for each

individual, but due to the variable magnitudes in response, this effect was not significant when the mean

of three patients was compared to the mean of the controls.

The ability of the passaged chondrocytes to form tissue was also assessed throughout the 3-week

culture period. Tissues formed in the presence of TGF-β3 had significantly increased thickness when

compared to non-treated control tissues (p<0.05), at each time-point in all 3 patients (Figure 3-5 and

Figure 4-4). Tissues treated with TGF-β3 appeared to stain more with Toluidine Blue than non-treated

controls, suggesting presence of chondroid matrix (Figure 3-5 and Figure 4-4). Immunohistochemical

staining of aggrecan and type II collagen also appears more prominent in the TGF-β3 treated tissues

compared to the controls, especially at the 2 and 3 week time points (Figure 3-5 and Figure 4-4).

Biochemical analysis of the tissue confirmed that there is significantly more glycosaminoglycan content

and collagen content, in TGF-β3 treated tissues at 3 weeks compared to untreated controls at the same

time point (p<0.05) (Figure 3-5 and Figure 4-4). Despite the increase in Col1 gene expression, tissues

treated with TGF-β3 showed less prominent staining for Type I collagen, throughout the 3-week culture

period (Figure 3-5 and Figure 4-4).

38

Figure 3-4: TGF-β3 Increases Expression of Chondrogenic Genes (ACAN, SOX9 and Col2) of Cells During Redifferentiation.

Gene expression of chondrogenic genes (ACAN, SOX9 and Col2) and fibroblast matrix gene Col1 was determined for up to 3 weeks of 3D culture in the presence or absence of TGF-β3. Data expressed as a fold change relative to P2, Please note the varying y-axes. Data is expressed as mean of triplicate cultures; the horizontal bar indicates mean ±SEM (N=3 patients). Red dot indicates patient 2, orange dot indicates patient 5, purple dot indicates patient 6. CON = non-treated control. *, p < 0.05, in comparison to non-treated controls at each time point.

39

Figure 3-5: TGF-β3 Promotes Hyaline Cartilage Formation.

P2 cells were grown in the presence or absence of TGF-β3 for up to 3 weeks in 3D membrane culture. Evaluation of (A) tissue thickness, (B) histological morphology (haematoxylin and eosin or toluidine blue staining) and (C) tissue composition by immunohistochemical staining for aggrecan (green), Type II collagen (green), Type I collagen (red) and cell nuclei (blue). (D) Biochemical quantification of DNA, sulphated GAG accumulation and collagen accumulation. *, p <0.05 within time point, #, p <0.05 between time points. n = 3 membranes, Data shown is from patient 2. All images are of equivalent magnification.

40

2.3.4 Identification of TGF-β3 Activated Signaling Pathway Associated with Matrix Accumulation by Passaged Chondrocytes During Redifferentiation

A screen of inhibitors specific to the TGF-β pathway molecules was conducted to identify

possible candidate pathways for TGF-β3 signaling in chondrocytes during redifferentiation (Figure 4-7).

The ALK5-TAK1-p38 pathway was identified as the most promising candidate and a time course was

then conducted to investigate the ability of specific chemical inhibitors to decrease the TGF-β3 mediated

effects on chondrogenic (SOX9, Col2, ACAN) and fibroblast matrix (Col1) genes. A significant increase

in all four genes was visible by 24 hours post TGF-β3 treatment, compared to the corresponding non-

treated control (Figure 3-6, Figure 4-8). The ALK5 (SB431542), ALK1 (Dorsomorphin), p38

(SB203580) and TAK1 ((5Z)-7-oxozeanol) inhibitors resulted in decreased expression of SOX9 and

ACAN by 12 hours post treatment and Col2 by 24 hours post treatment (p<0.05, Figure 3-6). The ALK5,

ALK1 and TAK1 inhibitors also significantly downregulated expression of Col1 by 24 hours post

treatment (p<0.05), while the p38 inhibitor did not (Figure 4-7). To confirm the results, p38 siRNA and

TAK1 siRNA were used to inhibit the downstream signaling effects of TGF-β3 (Figure 3-8). TAK1 and

p38 siRNA resulted in a slight decrease of TGF-β3 mediated increases in SOX9, ACAN and Col2 gene

expression, but efficiency of knockdown was minimal as demonstrated by evaluating gene expression

levels for TAK1 and protein levels for p38 (Figure 3-8).

The levels of phospho-p38 were quantified using western blot, to confirm activation of the

ALK5-TAK1-p38 axis in response to TGF-β3 treatment. At 1 hour (Figure 3-7) post-treatment, TGF-β3

led to an increase in the levels of phosphorylated p38 (p<0.05), which was sustained at 3-hours post-TGF-

β3 treatment (p<0.05), compared to non-treated control tissues. p38 phosphorylation was decreased in

tissues treated with ALK5 inhibitor (p<0.05), ALK1 inhibitor (p<0.05), p38 inhibitor (p<0.05) and TAK1

inhibitor (p<0.05) at 1 hour post-TGF-β3 treatment. This trend towards decreased p38 phosphorylation

was sustained in tissues treated with TAK1 inhibitor in 2 out of the 3 patients at both 3 hours (patient 2

and patient 4) and 24 hours (patient 2 and patient 3). However, p38 phosphorylation was increased in

tissues treated with ALK1 inhibitor (p<0.05) and p38 inhibitor (p<0.05) at both 3 hours and 24 hours.

To determine the effect of TGF-β3 on ECM synthesis and retention and to identify the signaling

pathways that influence this response, the amounts of newly synthesized GAG and collagen were

quantified. There was a significant increase in newly synthesized collagen (3H-proline) and significantly

increased retention of newly synthesized collagen and proteoglycans (3H-proline and 35S-SO4) 48 hours

after TGF-β3 treatment (p<0.05), which were decreased by ALK5, ALK1 and TAK1 chemical inhibition

(p<0.05) (Figure 3-9). The p38 inhibitor only significantly inhibited total collagen synthesis (p<0.05).

41

Figure 3-6: TGF-β3 May Signal Through ALK5, ALK1, TAK1 and p38.

P2 chondrocytes were grown in 3D culture and TGF-β3 in the presence or absence of various signaling inhibitors for up to 48 hours. Gene expression for (A) SOX9, at 12 hours; (B) ACAN, at 12 hours; (C) Col2, at 24 hours; (D) Col1, at 24 hours post-treatment with TGF–β3. Data is presented as fold change relative to the TGF-β3 only treated group. Data is expressed as mean of triplicate cultures; the horizontal bar indicates mean ±SEM (N=3 patients). Orange dot indicates patient 5, purple dot indicates patient 6, and teal dot indicates patient 7. CON = non-treated control. Dotted line indicates the mean of the TGF-β3 only treated group. Time points were selected based on most pronounced inhibition. *, p <0.05 relative to the TGF–β3 treated group.

42

Figure 3-7: TGF-β3 Signals Through ALK5, ALK1, TAK1 and p38 and Results in Increased Downstream p38 Phosphorylation.

P2 chondrocytes were grown in 3D culture and TGF-β3 in the presence or absence of various signaling inhibitors for up to 24 hours. Western blot and respective densitometry of phospho-p38 levels at (A) 1 hour, (B) 3 hours and (C) 24 hours post-treatment with TGF–β3. The horizontal bar indicates mean ±SEM (N=3 patients). Data is presented as fold change relative to the TGF-β3 only treated group. Please note the varying y-axes. Red dot indicates patient 2, blue dot indicates patient 3, and pink dot indicates patient 4. CON = non-treated control. Dotted line indicates the mean of the TGF-β3 only treated group. *, p <0.05 relative to the TGF–β3 only treated group.

43

Figure 3-8: p38 and TAK1 Levels were not Significantly Decreased by Silencing with Targeted siRNA.

Evaluation of (A) TAK1 silencing through gene expression and (B) p38 silencing through western blot following siRNA transfection after 2 days. Gene expression profile of P2 chondrocytes treated with TGF-β3 and p38 siRNA or TAK1 siRNA in 3D culture at (C) 2 days and (D) 3 days. Data presented as fold change relative to the scrambled siRNA + TGF-β3 group (N=2 patients). Blue dot indicates Patient 3 and pink dot indicates Patient 4. Dotted line indicates the mean of the scrambled siRNA + TGF-β3 group.

44

Figure 3-8: p38 and TAK1 Levels were not Significantly Decreased by Silencing with Targeted siRNA, Continued.

45

Figure 3-9: TGF-β3 Results in Increased Collagen Synthesis and Retention, and Glycosaminoglycan Retention.

Quantification of newly synthesized and retained (A) collagen (3H-Proline) and (B) proteoglycan (35S-SO4) by P2 chondrocytes grown in 3D culture and TGF–β3 in the presence or absence of their respective inhibitors, at 48hours post-treatment. Counts/minute are normalized to DNA and the data is presented as fold change relative to TGF-β3 only treated group. Data is expressed as mean of triplicate cultures; the horizontal bar indicates mean ±SEM (N=3 patients). Red dot indicates patient 2, blue dot indicates patient 3 and pink dot indicates patient 4. Dotted line indicates the mean of the TGF-β3 only treated group. CON = non-treated control. *, p <0.05 relative to the TGF–β3 only treated group.

46

Chapter 3 Discussion

3 Discussion

TGF-β has profound effects on passaged human chondrocytes and their ability to form cartilage

tissue. This study shows that monolayer culture of chondrocytes leads to changes in the expression levels

of TGF-β receptors ALK1, ALK5 and TGF-βRII. It is possible that these changes may allow the

chondrocytes to respond to TGF-β3, as primary cells are unable to respond and make cartilage tissue. The

changes in TGF-β receptor expression continue throughout the 3-week redifferentiation period upon

TGF-β3 treatment. TGF-β3 promoted chondrocyte redifferentiation as early as 24 hours after

supplementation as indicated by enhanced expression of the chondrogenic genes SOX9, Col2, and ACAN.

Redifferentiation continues over several days as Col2 and ACAN gene expression continues to rise until it

plateaus at two weeks. The presence of Type II Collagen and Aggrecan were evident in the cartilage

tissue by 2 weeks, further supporting the claim that the chondrocytes have redifferentiated. Additionally,

TGF-β3 increased synthesis and retention of collagen and retention of proteoglycan, which likely

contributed to cartilage tissue formation. Many signaling pathways, including ALK1, ALK5, TAK1 and

p38, appear to be involved in the complex process of differentiation and tissue formation. Here, we show

that the regulation of collagen synthesis appears to be at least partially through the ALK1/ALK5-TAK1-

p38 axis. Proteoglycan retention, however, seems to be regulated by an alternate signaling pathway.

Together, the results presented here suggest that TGF-β3 promotes redifferentiation and tissue formation

in part through both the canonical (ALK1 and ALK5) and the non-canonical (TAK1 and p38) pathways.

However, many other signaling pathways and/or environmental cues likely play a role in this complex

process.

Although the effect of TGF-β on redifferentiating chondrocytes has not been extensively studied,

the profound effect of TGF-β on chondrocyte redifferentiation and ECM accumulation is not unexpected

for several reasons. Firstly TGF-β has been identified as an important factor in chondrogenesis, cartilage

maintenance and homeostasis [154]. Secondly, aging chondrocytes as well as OA chondrocytes lose their

ability to regenerate cartilage as the disease progresses and these cells have been shown to undergo

phenotypic changes in TGF-β receptors such as an increase in the ratio of ALK1/ALK5 expression [172].

Knockout of TGF-β and/or its receptors leads to development of OA [200]. Lastly, TGF-β has been

shown to enhance redifferentiation of passaged human chondrocytes when grown in pellet culture [211].

To the best of our knowledge, we are the first to show the involvement of the non-canonical

signaling pathway during redifferentiation of passaged OA chondrocytes. Though p38 and TAK1 have

been show to influence chondrocyte phenotype [185, 187], the involvement of this pathway during

47

redifferentiation is unexpected as the majority of the literature has focused on canonical TGF-β signaling

pathways for cartilage maintenance. We are also the first to characterize this pathway in relation to matrix

synthesis during redifferentiation of passaged chondrocytes. TGF-β3 was shown to result specifically in

increased collagen synthesis during early redifferentiation, which was decreased by inhibitors of ALK5,

ALK1, TAK1 and p38. This finding is supported by studies using bovine chondrocytes, which

demonstrated increased synthesis and retention of collagen (specifically Type XII) during the early stages

of redifferentiation [26]. This study also showed an increase in the proteoglycan versican, which together

with Types I, II, III, and/or XII collagen, may provide the necessary microenvironment to promote

redifferentiation [26]. Further study is required to determine if these specific molecules are present during

early redifferentiation of passaged human OA chondrocytes.

In contrast to passaged chondrocytes, TGF-β treatment of primary chondrocytes does not enhance

in vitro formation of tissue (Bianchi et al., unpublished). Monolayer expansion of human chondrocytes

was shown to result in changes in ALK5, ALK1 and TGF-βRII expression. It is possible that the changes

in TGF-β receptor levels during passaging may therefore allow passaged chondrocytes to respond to

TGF-β once placed in 3D culture, to generate cartilage tissue. During passaging, we showed an increase

in ALK1 and a decrease in ALK5. This was not entirely unexpected, as there is a shift towards increased

ALK1/ALK5 with OA and aging [203], and monolayer expansion of chondrocytes has been shown to age

chondrocytes as many as 30 years [220]. It was surprising, however, that dedifferentiated chondrocytes

(with increased ALK1 and decreased ALK5) could redifferentiate and form cartilage tissue in response to

TGF-β3. It has been well established that ALK1 signals through the SMAD1/5/8 pathway to induce

catabolism and hypertrophy, while ALK5 signals through the SMAD2/3 pathway to promote anabolism

in chondrocytes [174, 221]. Furthermore, overexpression of ALK5 has been shown to decrease ALK1

gene expression in human fetal BMSCs undergoing chondrogenesis [216]. Conversely, TGF-β1 decreased

BMP-9-induced (ALK1 signaling) SMAD1/5/8 phosphorylation and expression of hypertrophic markers

in primary bovine chondrocytes [173]. This suggests that cross-talk exists between the two signaling

pathways, and that one pathway can have an inhibitory effect on the other [173, 174, 216]. Thus, it is

crucial to have the appropriate balance between ALK1 and ALK5 signaling, and this may be more

important than the overall receptor levels to promote redifferentiation. The proper ratio of ALK1:ALK5

would provide a balance between SMAD2/3 and SMAD1/5/8 signaling. Alternatively, the ability of

dedifferentiated cells to redifferentiate and form tissue may suggest that TGF-β signaling in

dedifferentiated chondrocytes differs from that of primary cells, at least during early redifferentiation. It

has been previously shown that both ALK1 and ALK5 are required for chondrogenic differentiation of

mesenchymal stem cells (MSCs) [216], suggesting that signaling through both receptors may be required

for redifferentiation.

48

To our knowledge, this is the first study to examine ALK1 signaling during redifferentiation of

passaged OA chondrocytes. Interestingly, our data raises the possibility that during redifferentiation,

TGF-β3 signals, at least in part, through the ALK1 receptor. Treatment with ALK1 inhibitor decreased

the TGF-β3 induced increases in SOX9, Col2 and ACAN gene expression. In contrast, Finnson et al have

shown that ALK1 signaling inhibits TGF-β induced phosphorylation of SMAD3 and downstream

expression of Type II Collagen in human chondrocytes [174]. In our system, we are suggesting that

signaling during redifferentiation may be different than signaling in primary cells, and that a balance

between ALK1 and ALK5 signaling may be required for redifferentiation, as both are present during early

redifferentiation. The potentially different signaling mechanism in redifferentiating cells may explain the

differences in ECM gene expression between our study and the study by Finnson et al.

It is not surprising that treatment with TGF-β3 results in changes of TGF-β receptors. We showed

that ALK1 gene expression levels decrease while ALK5 gene and protein levels increase in high density

SFM culture. As mentioned above, the progression of OA results in an increased ALK1/ALK5 ratio

compared to healthy tissue [172]. It is suggested that the relative decrease in ALK5 results in the

decreased ability of OA chondrocytes to produce the appropriate ECM [221]. In our system, a reversion

in the ALK1/ALK5 ratio expressed by the chondrocytes during redifferentiation may influence their

ability to respond to TGF-β and produce tissue with more hyaline-like properties. Interestingly,

immunohistochemical staining and western blot analysis shows that increased expression of ALK5 is

present in both control and TGF-β treated tissues. This suggests that the 3D environment and

chondrogenic media may promote re-expression of ALK5. An additional signal, exogenous TGF-β is

needed to induce the cells to form tissue.

This study also showed an increase in TGF-βRII (in select patients) during passaging. TGF-βRII

binds TGF-β and recruits either ALK1 or ALK5 [222-224]. The importance of TGF-βRII for chondrocyte

redifferentiation has been demonstrated in human articular chondrocytes [225]. Contrary to our results,

Bauge et al. showed a decrease of TGF-βRII during passaging (dedifferentiation) and an increase during

redifferentiation [225]. The contradictory results may be due to the use of healthy articular chondrocytes

(from femoral head fractures or cadavers with no known joint disease) by Bauge et al. compared with the

OA chondrocytes that were used in this study [225]. Additionally, we showed a significant decrease in

TGF-βRII gene expression at the 48 hour and 1 week time points during redifferentiation. Again, this

contradicts what has been previously reported. Of note, a decrease in TGF-βRII may not have negative

consequences in our system, as the presence of the high concentration of exogenous TGF-β3, may be

sufficient to promote redifferentiation. This requires further study. Exogenous TGF-β3 supplementation

in our system (which was not used in the initial study by Bauge et al.), may promote a negative feedback

49

loop preventing the transcription of TGF-βRII as shown in a more recent study by Bauge et al. [226]. In

response to TGF-β1 treatment, they showed that a negative feedback loop results in the down-regulation

of TGF-βRII and ALK5 gene expression over 24 hours, due at least in part to decreased mRNA stability

[226]. This corresponds to an initial, brief increase in gene expression of Col2 and SOX9 (in response to

TGF-β), which diminishes along with ACAN expression as the expression of the TGF-β receptors

decreases [226]. It is important to note, however, that this study was conducted in 2-dimensional culture,

which does not mimic a chondrocyte’s native environment. Here, we showed decreased expression of

ALK1 and TGF-βRII, but not ALK5 in response to TGF-β3 treatment. It is possible that the 3-

dimensional environment prevents the down-regulation of ALK5 in response to TGF-β3 treatment,

allowing for prolonged signaling through ALK5 and subsequent ECM synthesis. Bauge et al. have also

shown that the overexpression of transcription factor specificity protein 1 (SP1) is able to recover the

expression of the TGF-β receptors in chondrocytes in 2D culture [226]. SP1 expression in human

chondrocytes decreases with passaging, however it has been shown that it can be restored when the

dedifferentiated cells are placed in 3D alginate bead culture [227]. It has also been shown that during

chondrogenic differentiation of MSCs in 3D culture, there is both an early (within 72hours) and late (at 2

and 3 weeks) increase in gene expression of SP1 [225]. Interestingly, we have shown a similar gene

expression pattern of ALK5 in redifferentiating chondrocytes, with early (24 hours) and late (2 and 3

week) increases. Recent studies have suggested that passaged chondrocytes can attain a phenotype that is

similar to MSCs [228]. Therefore, it is possible that the passaged chondrocytes presented here may

exhibit stem/progenitor cell-like properties, similar to the MSCs studied by Bauge et al. [225]. The

culture system used in this study (3D) may promote expression of SP1 (or similar transcription factors),

leading to increased expression of ALK5 and a prolonged response to TGF-β3, despite continuous TGF-β

treatment.

Based on our findings, it appears that TGF-β3 signals downstream, in part, through ALK5 to

regulate chondrogenic gene expression. Although many studies suggest that ALK5 signals through the

canonical SMAD3 pathway [174, 175, 229, 230], our findings suggest that it may also signal through the

TAK1 pathway during redifferentiation of passaged chondrocytes. Even though treatment with SMAD3

phosphorylation inhibitor (SIS3) does result in inhibition of gene expression in some patients, more

consistent inhibition is achieved after treatment with TAK1 or p38 chemical inhibitors. There are several

important differences between previous studies investigating TGF-β signaling in chondrocytes and our

current study. Firstly, there are differences in cell source; animal cells, such as murine [172] and rat [230]

chondrocytes, may have different signaling mechanisms than human chondrocytes. The use of primary

versus passaged cells may also contribute to these differences [172]. During redifferentiation of

chondrocytes, TGF-β may signal through the TAK1 axis, differing from canonical signaling in native

50

chondrocytes. Additionally, there may be cross talk between SMAD3 and TAK1, as has been described to

occur in murine chondrocytes [186]. In SMAD3(-/-) mice, TAK1 signaling is disrupted due to decreased

p38 MAP kinase (MAPK) activation [186], suggesting the importance of both pathways for TGF-β

mediated signaling.

Our results suggest that p38 may be one of the mediators downstream of TAK1, as the increase in

phosphorylation with TGF-β3 treatment was reduced by inhibitors of ALK5, ALK1 and TAK1 (p<0.05)

at the 1 hour timepoint. However, the regulation of p38 seems to be complex as p38 phosphorylation was

increased in tissues treated with ALK1 and p38 inhibitors (p<0.05) at later timepoints. p38 has been

shown to be involved in regulating chondrogenesis, cell proliferation, cell survival and hypertrophy [185-

187]. Increased levels of p38 phosphorylation have been shown to promote hypertrophy [231], while

reduced levels of p38 activity have led to inhibition of chondrogenesis in chick and mouse limb bud

MSCs [232, 233]. Due to the array of cellular processes that p38 is involved in, it is not surprising that its

expression and activity must be tightly regulated. There are likely additional regulatory mechanisms such

as feedback loops or cross-talk with other molecules that may influence the expression and activity of

p38, which may explain our results

The study presented here does not show any consistent significant changes in SMAD2 or

SMAD5 gene expression. A trend towards slightly decreased expression of SMAD2 was present with

TGF-β3 treatment, however this down-regulation may be due to negative feedback from high exogenous

levels of TGF-β3. It is likely that changes in phosphorylation levels and activation of SMAD2/3 and

SMAD1/5/8 are more indicative of signaling occurring within the chondrocytes during redifferentiation.

These changes should be further characterized as it is possible that a TGF-β3 induced increase in SMAD3

activity may have an effect on TAK1/p38 signaling in dedifferentiated/redifferentiated human

chondrocytes as well [186].

A possible mechanism for the TGF-β3 mediated increase in ECM retention may be due to changes

in expression of molecules that regulate ECM turnover, such as a disintegrin and metalloproteinase with

thrombospondin motifs (ADAM-TS5) and tissue metalloproteinase inhibitor 3 (TIMP3). In addition to

changes in synthesis and retention of matrix molecules, collagen and proteoglycan, there was down-

regulation of ADAM-TS5 and up-regulation of TIMP3 gene expression, at early time points with TGF-β3

treatment. ADAM-TS5 is an enzyme that degrades aggrecan [234], and TIMP3 is a protein that inhibits

matrix metalloproteinases (MMPs; enzymes that can remodel ECM through degradation) [235, 236]. To

our knowledge, there are no studies examining the expression of ADAM-TS5 or TIMP3 during

chondrocyte redifferentiation. TIMP3(-/-) mice have been shown to display an increase in collagen and

51

aggrecan degradation [237]. Sahebjam et al. also showed increased Type II collagen cleavage neoepitopes

and collagen type X expression in the superficial zone of the articular cartilage in these mice [237]. The

degradation of the superficial zone in that study parallels OA progression in humans, which highlights the

importance of TIMP3 expression in cartilage homeostasis. Therefore during redifferentiation, TGF-β3

treatment may further inhibit ADAM-TS5 expression and increase TIMP3 expression, resulting in overall

increased matrix retention. Further study is required to determine if the gene changes correlate with

increased protein production and/or activity.

Inhibitors of ALK5 and TAK1 reverse the TGF-β3 mediated increase in TIMP3 expression, suggesting that TGF-β3 may signal through this pathway to promote TIMP3 expression. Interestingly, inhibitors of the ALK1/ALK5-TAK1-p38 axis do not reverse the decrease seen in ADAM-TS5 gene expression, suggesting that TGF-β3 likely signals through another pathway to mediate ADAM-TS5. These changes may be regulated by the transcription factor SOX9, which was up-regulated by TGF-β3. Inactivation of SOX9 in adult mice has been shown to decrease the expression of genes involved in involved cartilage maintenance and extracellular organization [238]. Specifically, an increase in ADAM-TS5 was seen [238, 239]. Therefore, it is possible that increases in SOX9 seen with TGF-β3 treatment could lead to opposite effects, such as decreases in extracellular matrix degrading enzymes, as shown in our study.

One of the limitations of our study was the use of chemical inhibitors to inhibit TGF-β signaling

molecules, as they can have non-specific effects. A more specific method of inhibition (p38 siRNA and

TAK1 siRNA) was used (Figure 3-8) to confirm the results obtained by the chemical inhibitors (Figure 3-

6). Unfortunately, TAK1 and p38 siRNA did not significantly decrease TGF-β3 mediated increases in

SOX9, ACAN and Col2 gene expression. This is likely due to the inefficient knockdown of TAK1 and p38

with siRNA (Figure 3-8). Chondrocytes are known for being difficult to transfect, partly due to the

ECM/pericellular matrix that surrounds them [240-242]. Transfection protocols must be further optimized

to overcome this challenge, and is an important consideration when planning future experiments.

Other studies have explored the use of TGF-β to promote redifferentiation of human passaged

chondrocytes and subsequent tissue formation in vitro. Yeager et al. have shown that treatment of

passaged adult human chondrocytes with a combination of TGF-β1 or TGF-β2 in addition to IGF-1 in

serum free media (SFM) alginate bead culture results in increases in Col2 and ACAN gene expression in

comparison to non growth-factor treated controls [210]. Goldberg et al. showed that pellet culture of

passaged OA chondrocytes in SFM with TGF-β1 and ITS+ Premix (insulin, transferrin, selenous acid)

resulted in increased production of glycosaminoglycan and DNA compared to pellet culture in 10% serum

with and no TGF-β1 supplementation [211]. Additionally, Jakob et al. have shown that supplementation

with FGF-2 and TGF-β during monolayer culture (dedifferentiation) of non-OA human chondrocytes

52

resulted in enhanced tissue formation in pellet culture SFM conditions containing dexamethasone and

TGF-β [56]. This included higher levels ACAN and Col2 gene expression, as well as Type II collagen

expression and glycosaminoglycan content compared to cells expanded without growth factor

supplementation[56]. In contrast, Narcisi et al. have shown that TGF-β depletion of media during

monolayer culture resulted in enhanced chondrogenic potential and tissue formation by human OA cells

when they were placed in pellet culture [209]. This was shown by demonstrating increased expression of

Type II collagen, increased glycosaminoglycan content and a reduction in hypertrophic markers,

compared to cells that were not TGF-β depleted during monolayer culture [209]. This is not surprising, as

TGF-β exposure during cell number expansion pushes the cell’s phenotype towards hypertrophy [213].

These studies provide valuable information regarding chondrocyte redifferentiation and subsequent tissue

formation, however they are associated with several limitations. The use of scaffold materials or

aggregate cultures may result in an uneven distribution of matrix deposition and cell death in the center of

the tissue due to uneven diffusion of nutrients [93]. The scaffold-free self-assembly method presented in

this study forms a continuous layer of articular-like cartilage tissue with an even distribution of ECM and

development of a superficial zone (Bianchi et al., unpublished). Furthermore, others have used multiple

growth factors during passaging/redifferentiation to promote cartilage tissue formation [56, 209-211],

which adds additional complexity to the system. Here, we present a simpler method that utilizes only

TGF-β3 and ITS (source of insulin) in serum-free conditions during redifferentiation. Insulin has been

shown to promote type II collagen expression via SOX9-Col2a1 binding [105]. Unfortunately, the process

of redifferentiation takes time (ranging from 2-4 weeks) [56, 210]. The results presented in this study

confirm this, as there are signaling changes throughout the first week, and changes in gene expression up

to 3 weeks. To use this approach clinically it would be more cost effective if the time needed for

redifferentiation and subsequent tissue formation could be decreased. The addition of other factors such

as mechanical loading may accelerate this process. Mechanical stimulation of chondrocytes has been

shown to accelerate matrix accumulation [64], and may promote superior tissue formation in our model.

Additionally, it may be possible to design a more potent chemical activator of the non-canonical TGF-β

pathway, which could promote redifferentiation over a shorter time period. It will be crucial to optimize

this procedure to ensure cost-effectiveness, and future studies should address this.

One of the observations made over the course of these experiments is that the response to TGF-β3

differs between patients in both timing and magnitude. Specifically, there were differing receptor levels,

levels of p38 phosphorylation, gene expression and matrix synthesis between patients. This variation in

response to TGF-β3 may be one of the factors that contributes to differences in tissue formation between

patients, as shown by quantification of tissue thickness, as well as glycosaminoglycan and collagen

accumulation. A small number of patients formed inferior tissue, however TGF-β3 treatment was still

53

able to induce significant increases in ECM accumulation compared to untreated cells. The variation in

response may be due to patient specific differences in age, grade of OA [203], and/or epigenetic

modifications [243]. Although we harvested cells from “healthier” tissue, OA is a disease of the joint and

these cells may have various degrees of changes associated with the grade of disease. In OA, expression

of TGF-βRII and TGF-β1 protein is decreased in fibrillated and degraded articular cartilage, respectively,

when compared to normal cartilage [196], suggesting that there may be changes in TGF-β signaling with

disease progression. OA and age-associated decreases in ALK5 protein expression are more marked than

changes in ALK1 protein expression in mice [172]. Therefore, age may also influence the ALK1/ALK5

expression ratio on the cells, affecting their ability to respond to TGF-β [203]. In this study, the patients

we collected cells from ranged in age from 28-73 years. This variable may account for, at least in part, the

varied response to TGF-β3 shown here. Madej et al. showed a decreased activation of SMAD2/3 in

response to mechanical compression by aged chondrocytes, further suggesting the possibility that aged

chondrocytes are unable to respond as effectively to external stimuli [229]. Moreover, there are epigenetic

changes in chondrocytes with age and OA progression, such as histone modifications and DNA

methylation/demethylation, which have been shown to regulate both catabolic (ADAM-TS/MMP) and

anabolic (SOX9/Col2) gene expression [244-246]. These epigenetic changes may not be reversed with

passaging and persist during redifferentiation, affecting the chondrocyte’s ability to generate cartilage

tissue. Additionally, the microRNA (miRNA) profile of chondrocytes has been shown to change during

osteoarthritis, influencing the expression of both catabolic and anabolic proteins [247-249]. The effect of

monolayer culture on miRNAs expression is unknown, and whether or not cell passaging can revert this

miRNA profile has not yet been studied. Differing miRNA profiles between patients may lead to

variability in response to TGF-β3 as seen in our study. Specific miRNA profiles may be able to predict

whether or not the cells from a specific patient will create superior tissue, however this needs to be further

explored. Interestingly, TGF-β has been shown to regulate miRNA in OA, and may have similar effects in

our system [250]. The impact of these factors on redifferentiation and tissue formation will become

clearer as studies with larger sample sizes and more complete investigation of these different entities are

done.

TGF-β3 treatment resulted in a significant increase in total DNA content in the in vitro formed

tissue. This suggests that there are more cells in the TGF-β3 treated tissues that have the potential to

secrete ECM molecules and enhance tissue formation. TGF-β has been shown to enhance proliferation of

growth plate chondrocytes [251], and survival and viability of intervertebral disc cells [252]. In our

system, TGF-β3 may stimulate the chondrocytes to divide or promote cell survival, leading to the increase

in DNA and greater number of cells. Alternatively, TGF-β3 may prevent cell death [253]. The patients

that produce inferior tissue generally have lower values of total DNA in the tissue, which may suggest

54

that cells from these tissues do not divide (or survive) in response to TGF-β3, and thus do not generate

superior tissue.

3.1 Conclusion The use of autologous chondrocytes for bioengineering purposes is favorable as there are no risks

of allogeneic or xenogeneic response if grown serum free, however generating an adequate cell number

with the appropriate phenotype remains a challenge [254]. We have established that TGF-β3 and a 3-

dimensional environment promote redifferentiation of passaged human OA chondrocytes and formation

of bioengineered tissue. Here we show further characterization of the dedifferentiation and

redifferentiation processes.

Monolayer culture (passaging) of human chondrocytes was shown to result in changes in ALK5,

ALK1 and TGF-βRII expression. We have also shown that during redifferentiation there are changes in

the ALK1 and ALK5 expression levels. These changes may allow the chondrocytes to respond to TGF-β3

stimulation and form superior tissue through synthesis and accumulation of the appropriate chondrogenic

ECM molecules (namely aggrecan and type II collagen, although other important molecules are also

involved). Furthermore, we have identified that TGF-β3 promotes redifferentiation and ECM

accumulation in passaged OA chondrocytes in part by signaling through both the canonical (ALK1 and

ALK5) and the non-canonical (TAK1 and p38) pathways, though it is a complex regulation. Collagen

production and retention in particular appear to be modulated through the ALK1/ALK5-TAK1-p38 axis.

This may be critical, as previous studies in our lab have shown that collagen accumulation occurs before

proteoglycan accumulation during redifferentiation[26, 105].

It is well established that TGF-β is critical for cartilage development and homeostasis and that

changes in TGF-β signaling can have profound impacts on cartilage composition and structure [199, 221,

255]. Characterization of the changes in receptor molecules expression and identification of TGF-β

signaling pathway(s) contributes to our knowledge on both the redifferentiation process and in vitro tissue

formation by passaged chondrocytes. Understanding these processes will allow for the identification of

pharmacological agents and other techniques to further enhance formation of bioengineered cartilage.

55

3.2 Future Directions In this study we identify changes in TGF-β signaling during passaging and redifferentiation, as

well as one of the TGF-β signaling pathways that results in increased ECM accumulation by passaged

human OA chondrocytes. Further research is required to translate the application of these findings to

enhance in vitro cartilage formation. Understanding how patient specific differences influence the

redifferentiation process is important in being able to predict the quality of cartilage formation and the

potential success of this model in the general population. It should be noted that our results may have

been influenced by the variability between patients and the small sample size. Future experiments should

focus on characterizing the redifferentiation process, in a patient specific manner for a larger group of

patients to confirm the significance of the results. Of specific interest are how differences in expression of

ALK1, ALK5 and TGF-βRII correlate to TGF-β3 responsiveness, ECM synthesis and tissue formation.

We have shown that ALK5 protein expression decreases with passaging and increases with

redifferentiation. Therefore, what remains to be characterized is the levels of protein expression for ALK1

and TGF-βRII. Additionally, in this study patients were not sorted according to any particular variable. It

is conceivable that both age and grade of OA are factors that influence changes in TGF-β receptor level

expression and therefore redifferentiation ability [172, 196]. Increasing the number of patients studied

might provide more insight on how variables such as age, gender and grade of OA affects the ability of

chondrocytes to respond to TGF-β3 and form tissue. This should be considered when planning future

studies.

Our study used full thickness articular cartilage dissected from low-grade OA areas, with no

visible fibrillation or damage. Future experiments should compare the abilities of cells sampled from

“normal” and more fibrillated or damaged areas of the same joint, to respond to TGF-β3 and form

cartilage tissue. This analysis may further confirm the importance of the TGF-β receptor profile, which

could be used as a screening tool to determine whether a population of cells will generate hyaline

cartilage tissue.

Characterizing the levels of downstream R-SMADs (SMAD2/3 and SMAD1/5/8) is also

important as the literature suggests that they play opposing roles in regulation of the chondrogenic

phenotype [173, 174]. We have shown that there are no sustained significant changes in gene expression

of SMAD2 and SMAD5. There may, however, be changes in SMAD2 or SMAD5 phosphorylation and

activation. Levels of phospho-protein expression should therefore be quantified in future experiments.

This is especially relevant as it has been shown that there may be crosstalk between the canonical

SMAD2/3 and non-canonical signaling pathways, which could potentially influence signaling in our

56

system [186]. Future experiments should also focus on confirming the proposed TGF-β3 signaling

pathway through quantification of phosphorylation changes in TAK1 by western blot.

This study has shown that downstream of TAK1, p38 is involved in TGF-β signaling during

redifferentiation. Further experiments should confirm p38 and TAK1 interaction and focus on identifying

downstream targets of p38. TAK1 also has two other downstream effectors, c-Jun N-terminal kinases

(JNK) and extracellular signal-regulated kinases (ERK). TGF-β has been shown to activate JNK via

TAK1 in chondrocytes [256]. It has been shown that TGF-β induced activation of ERK promotes both a

mitogenic response and induction of TIMP3 in chondrocytes, suggesting that it affects cell proliferation

and ECM accumulation [257, 258]. The potential involvement of JNK and ERK in redifferentiation

should be characterized through measuring levels of phosphorylated JNK and ERK and by investigating

if inhibition of these molecules results in changes of accumulation of newly synthesized ECM molecules.

Characterizing the signaling pathways involved in redifferentiation, may aid in the design and

development of a synthetic peptide that activate these pathways and result in enhanced in vitro tissue

formation. Ideally, the synthetic peptide would be a cheaper alternative to biologically derived TGF-β3. It

could also be designed to induce stronger activation of the redifferentiation pathways and perhaps reduce

the culture period required for redifferentiation. This requires further investigation.

57

Chapter 4 Appendix

4 Appendix

4.1 Additional Tables

Table 4-1: Patient Details

Patient Age Sex

Patient 1 74 M

Patient 2 65 F

Patient 3 64 F

Patient 4 73 M

Patient 5 28 F

Patient 6 56 F

Patient 7 73 M

Patient 8 70 F

Patient 9 60 F

Patient 10 67 F

Table 4-2: Additional Primers.

Primer Forward Sequence 5’ to 3’ Reverse Sequence 5’ to 3’

TIMP3 [122] ACCTGCCTTGCTTTGTGACT GGCGTAGTGTTTGGACTGGT

ADAM-TS5

[259] GCTCACGAAATCGGACATTTACTT ACCAAAGGTCTCTTCACAGAATTTG

58

4.2 Additional Figures

Figure 4-1: ALK5 Decreases in Chondrocytes During Passaging, Additional Data.

Expression of ALK1 and ALK5 in monolayer cultured chondrocytes, from passage 0 through to passage 2, as characterized by immunocytochemical staining for (A) patient 1 and patient 2, and (B) western blot analysis for patient 1 and patient 4. All images are of equivalent magnification.

59

Figure 4-2: ALK1 and ALK5 are Expressed in Chondrocytes During Redifferentiation and Tissue

Formation, Additional Data.

ALK1 and ALK5 levels were determined following 1, 2, or 3 weeks of 3D culture in the presence or absence of TGF-β3 by immunohistochemical staining for (A) patient 1 and (B) patient 2, and (C) western blot analysis for patient 1 and patient 3. CON= non-treated control. Figure continued on next page.

60

Figure 4-2: ALK1 and ALK5 are Expressed in Chondrocytes During Redifferentiation and Tissue Formation, Additional Data Continued.

61

Figure 4-2: ALK1 and ALK5 are Expressed in Chondrocytes During Redifferentiation and Tissue Formation, Additional Data Continued.

Figure 4-3: TGF-β3 has Variable Effects on TGF-β Signaling Pathway Genes (SMAD5 and

SMAD2) During Redifferentiation and Tissue formation.

SMAD5 and SMAD2 gene expression levels were determined for up to 3 weeks of 3D culture in the presence or absence of TGF-β3. Data is expressed as mean of triplicate cultures; the horizontal bar indicates mean ±SEM (N=3 patients). Red dot indicates patient 2, orange dot indicates patient 5 and purple dot indicates patient 6. CON = non-treated control. *, p < 0.05 in comparison to non-treated controls at each time point.

62

Figure 4-4: TGF-β3 Promotes Hyaline Cartilage Formation, Additional Data.

P2 cells were grown in the presence or absence of TGF-β3 for up to 3 weeks in 3D membrane culture. Evaluation of (A, E) tissue thickness, (B, F) histological morphology (haematoxylin and eosin or toluidine blue staining) and (C, G) tissue composition by immunohistochemical staining for aggrecan (green), Type II collagen (green), Type I collagen (red) and cell nuclei (blue). (D, H) Biochemical quantification of DNA, sulphated GAG accumulation and collagen accumulation. *, p <0.05 within time point, #, p <0.05 between time points. n= 3 membranes. All images are of equivalent magnification.

Figure 4-4: TGF-β3 Promotes Hyaline Cartilage

Formation, Additional Data Continued.

Patient 1 is presented in Figure 4-4A-D and Patient 3

is presented in Figure 4-4E-H.

64

Figure 4-5: TGF-β3 Promotes Hyaline Cartilage Formation, Combined Data From Three Patients.

P2 cells were grown in the presence or absence of TGF-β3 for up to 3 weeks in 3D membrane culture. Combined data for patient 1, patient 2 and patient 3. Evaluation of (A) tissue thickness and biochemical quantification of (B) DNA, (C) sulphated GAG accumulation and (D) collagen accumulation. Data is expressed as mean of triplicate cultures; the horizontal bar indicates mean ±SEM (N=3 patients). CON = non-treated control. *, p <0.05 within time point, #, p <0.05 between time points.

65

Figure 4-6: TGF-β3 Promotes Hyaline Cartilage Formation to a Lesser Extent in a Select Group of

Patients.

P2 cells were grown in the presence or absence of TGF-β3 for up to 3 weeks in 3D membrane culture. Evaluation of (A) tissue thickness, (B) histological morphology (haematoxylin and eosin or toluidine blue staining) and (C) Biochemical quantification of DNA, sulphated GAG accumulation and collagen accumulation. CON = non-treated control. *, p <0.05 within time point, #, p <0.05 between time points. n= 3 membranes, data shown is from patient 8. All images are of equivalent magnification.

66

Figure 4-7: TGF-β Signaling Pathway Inhibitor Panel Results in Decreased Expression of

67

Chondrogenic Genes (ACAN, SOX9, Col2).

P2 chondrocytes were grown in 3D culture and TGF-β3 in the presence or absence of various signaling

inhibitors for up to 48 hours. Gene expression for tissues harvested at (A) 24 hours and (B) 48 hours. The

horizontal bar indicates mean ±SEM (n=6 membranes). Brown dot indicates patient 9, light blue dot

indicates patient 10. Data is presented as fold change relative to TGF–β3 only treated group. Please note

the varying y-axes. Dotted line indicates the mean of the TGF-β3 only treated group. *, p < 0.05 relative

to the TGF–β3 treated group.

68

Figure 4-8: TGF-β3 May Signal Through ALK5, ALK1, TAK1 and p38, Additional Timepoints.

P2 chondrocytes were grown in 3D culture and TGF-β3 in the presence or absence of various signaling inhibitors for up to 48 hours. Gene expression for remaining timepoints not shown in Figure 3-6, at (A) 6 hours, (B) 12 hours, (C) 24hours and (D) 48hours post-treatment with TGF–β3. Data is presented as fold change relative to the TGF-β3 only treated group. Please note the varying y-axes. Data is expressed as mean of triplicate cultures; the horizontal bar indicates mean ±SEM (N=3 patients). Orange dot indicates patient 5, purple dot indicates patient 6, and teal dot indicates patient 7. CON = non-treated control. Dotted line indicates the mean of the TGF-β3 only treated group. *, p <0.05 relative to the TGF–β3

69

treated group.

Figure 4-8: TGF-β3 May Signal Through ALK5, ALK1, TAK1 and p38, Additional Timepoints Continued.

70

Figure 4-9: TGF-β3 Treatment Results in Decreased ADAM-TS5 and Increased TIMP3 Gene Expression.

P2 chondrocytes were grown in 3D culture and TGF-β3 in the presence or absence of various signaling inhibitors for up to 48 hours. Gene expression for ADAM-TS5 at (A) 6 hours and (B) 24 hours, and TIMP3 at (C) 6 hours, and (D) 24 hours post-treatment with TGF–β3. Data is presented as fold change relative to the TGF-β3 only treated group. Please note the varying y-axes. Data is expressed as mean of triplicate cultures; the horizontal bar indicates mean ±SEM (N=3 patients). Orange dot indicates patient 5, purple dot indicates patient 6 and teal dot indicates patient 7. CON = non-treated control. Dotted line indicates the mean of the TGF-β3 only treated group. *, p <0.05 relative to the TGF–β3 treated group.

71

Set1 Set2 Average

A

B C

µg O

HPr

o/µg

DN

A

21

µg G

AG

/µg

DN

A

µg D

NA

72

Figure 4-10: ROCK Inhibition Results in Increased

Chondrogenic Gene Expression and Tissue Thickness

P2 chondrocytes were grown in 3D culture and TGF-β3 in the presence or absence of Y-2763 (ROCKi), for up to 3 weeks. (A) Gene expression for ACAN, SOX9, Col1 and Col2, at 48 hours, (B) histological morphology (haematoxylin and eosin or toluidine blue staining), (C) Biochemical quantification of DNA, sulphated GAG accumulation and collagen accumulation (D) and tissue thickness. The mean ±SEM was calculated and the mean is indicated by the horizontal bar (N= 6 membranes) *, p <0.05.

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