A MULTISCALE APPROACH TO STEM CELL-BASED
CHONDROGENESIS FOR CARTILAGE REPAIR
by
CHIH-LING CHOU
Submitted in partial fulfillment of the requirements
For the degree of Doctor of Philosophy
Dissertation Advisor: Dr. Harihara Baskaran
Department of Chemical Engineering
CASE WESTERN RESERVE UNIVERSITY
May, 2013 CASE WESTERN RESERVE UNIVERSITY
SCHOOL OF GRADUATE STUDIES
We hereby approve the thesis/dissertation of
Chih-Ling Chou
candidate for the Doctor of Philosophy degree *.
(signed) Dr. Harihara Baskaran . (chair of the committee)
Dr. Jean F. Welter .
Dr. Chung-Chiun Liu .
Dr. Heidi B. Martin .
Date of Defense March 14, 2013.
*We also certify that written approval has been obtained for any proprietary material
contained therein.
TABLE OF CONTENTS
Table of contents ...... I
List of tables ...... VI
List of figures ...... VII
Acknowledgements ...... XI
Abstract ...... XIII
Chapter 1. Introduction ...... 1
1.1 Purpose of the Study ...... 1
1.2 Significance and Motivation of the study ...... 2
1.2.1 Articular Cartilage Injury ...... 2
1.2.2 Limitations of Current Treatment Options ...... 2
1.2.3 Challenges for Tissue-Engineered (TE) Cartilage ...... 3
1.3 Overall Research Goals ...... 6
1.4 Thesis Outline ...... 8
1.5 Summary ...... 9
1.6 References ...... 9
Chapter 2. Background ...... 12
2.1 Articular Cartilage ...... 12
2.1.1 Knee joint and Cartilage ...... 12
I
2.1.2 Ultrastructure and Constituents of Articular cartilage ...... 13
2.1.3 Articular Cartilage Injury and Repair ...... 17
2.1.4 Current Treatment for Articular Cartilage Injury ...... 20
2.2 Cartilage Tissue engineering ...... 25
2.2.1 Cell Sourcing for Cartilage Tissue engineering ...... 27
2.2.2 Biomaterials for Cartilage Tissue Engineering ...... 29
2.2.3 Challenges in Cartilage Tissue Engineering ...... 31
2.3 Cartilage Structure and Guidance ...... 34
2.3.1 Contact Guidance ...... 34
2.3.2 Biomechanics of Articular Cartilage ...... 36
2.3.3 Structure and Guidance ...... 38
2.4 References ...... 39
Chapter 3. Control of human mesenchymal stem selective adhesion on collagen- glycosaminoglycan and Poly(dimethyl siloxane) surfaces ...... 51
3.1 Introduction ...... 51
3.2 Materials and Methods ...... 54
3.2.1 Materials ...... 54
3.2.2 CG Solution and EDC Solution Formation ...... 55
3.2.3 Design of Microchannels ...... 55
3.2.4 Fabrication of Silicon Substrate Template with Microchannels ...... 56
II
3.2.5 Fabrication of Collagen and PDMS Microchannels ...... 59
3.2.6 Selective Cell Attachment in Collagen Microchannels ...... 63
3.2.7 Selective Cell Attachment in PDMS Microchannels ...... 65
3.2.8 Cell Culture...... 66
3.2.9 MSC Viability, and Adhesion Assessment...... 68
3.3 Results ...... 68
3.3.1 CG Membrane Patterning ...... 68
3.3.2 MSC Viability and Adhesion Assessment...... 70
3.2.3 Selective Cell Attachment ...... 71
3.4 Discussion ...... 74
3.5 Conclusion ...... 78
3.6 References ...... 78
Chapter 4. Investigate the effect of microscale guidance on mesenchymal stem cell-based chondrogenesis ...... 81
4.1 Introduction ...... 81
4.2 Materials and Methods ...... 89
4.2.1 Materials ...... 89
4.2.2 Fabrication of Collagen-hMSC construct and PDMS-hMSC construct ...... 90
4.2.3 Image Acquisition...... 91
4.2.4 Histology and Immunohistochemistry...... 91
III
4.2.5 Cell alignment analysis ...... 92
4.2.6 Mechanical Property Testing ...... 94
4.2.7 Biochemical Measurements ...... 96
4.3 Result ...... 98
4.3.1 Cellular Growth and Organization in Microchannels ...... 98
4.3.2 Microscale Guidance on Cell alignment ...... 99
4.3.3 ECM Production ...... 105
4.3.4 Mechanical Properties ...... 108
4.4 Discussion ...... 111
4.5 Conclusion ...... 115
4.6 References ...... 116
Chapter 5. Design of large-scale 3-dimensional cartilage constructs with microchannels for preclinical studies ...... 123
5.1 Introduction ...... 123
5.2 Materials & Methods ...... 127
5.2.4 Autonomous Rolled-up CG-hMSC Construct (ARCGs) Formation ...... 130
5.2.5 Induced Rolled-up CG-hMSC construct (IRCGs) Formation ...... 131
5.2.6 Compression Testing ...... 132
5.2.7 Histological and Immunohistochemistry Analysis ...... 134
5.2.8 Collagen Fibers Alignment Analysis ...... 135
IV
5.2.9 Metabolism Analysis ...... 135
5.2.10 Statistical Methods ...... 136
5.3 Result ...... 136
5.3.1 Autonomous Rolled-Up CG-hMSC constructs (ARCGs) ...... 136
5.3.2 Induced Roll-up CG-hMSC constructs (IRCGs) ...... 140
5.3.3 Collagen Fibrils Alignment Analysis ...... 148
5.3.4 Metabolism Analysis ...... 153
5.4 Discussion ...... 155
5.5 Conclusion ...... 163
5.6 Reference ...... 164
Chapter 6. Conclusions ...... 169
6.1 Conclusions ...... 169
6.2 Future Directions ...... 173
6.3 References ...... 175
Bibliography ...... 176
V
LIST OF TABLES
Table 3-1. Accuracy of channel reproduction in CG membranes...... 69
Table 4-1. Cell alignment angle during contact guidance ...... 105
Table 4-2. GAG and DNA content of cartilage tissue grown in the channels ...... 108
VI
LIST OF FIGURES
Figure 1-1. MSC seeded CG sponge TE construct after 3 weeks in bioreactor chondrogenic culture. DAPI staining on day 1 ...... 5
Figure 1-2. Orientation of the collagen fibrils throughout the depth of native articular cartilage (A) and Ultrastructure of human articular cartilage after freeze fracture processing(B) ...... 6
Figure 2-1. Schematic of a Synovial Joint ...... 13
Figure 2-2. Ultrastructure and composition of articular cartilage ...... 16
Figure 2-3. Schematic of normal and arthritic knee Joint...... 18
Figure 2-4. Illustration of normal cartilage and osteoarthritic cartilage ...... 18
Figure 2-5. Schematic of cartilage defects with different depths. Partial-thickness defects and full-thickness defects...... 20
Figure 2-6. Schematic of autologous chondrocyte implantation (ACI) procedure ...... 24
Figure 2-7. Schematic diagram of typical tissue engineering strategy for cartilage repair.
...... 26
Figure 2-8. Multilineage differentiation potentials of adult human Mesenchymal stem cell
...... 29
Figure 2-9. Contact guidance of neurons on polymer ε-polycaprolactone microgrooves of different dimensions...... 35
Figure 2-10. Native hyaline cartilage ultrastructure and schematic of ultrastructure of articular cartilage...... 37
Figure 2-11. Depth-dependent compressive properties of human articular cartilage...... 37
VII
Figure 3-1. Microchannel design ...... 56
Figure 3-2. Schematic of standard UV light lithography ...... 59
Figure 3-3. Schematic of collagen soft lithography method ...... 61
Figure 3-4. Schematic of standard soft lithography ...... 62
Figure 3-5. Schematic of Pluronic mechanism...... 64
Figure 3-6. A schematic of the technique used to obtain selective seeding of mesenchymal stem cells ...... 64
Figure 3-7. Schematic of selective cell attachment technique for PDMS membrane...... 66
Figure 3-8. (A) Microchannel design created in AutoCAD (B) Digtal image of patterend
CG membrane (C) Phase Contrast image of collagen microchannel ...... 69
Figure 3-9. Scanning electron micrographs of linear collagen channels ...... 70
Figure 3-10. (A) Scanning electron micrographs of MSCs in collagen channel of width 50
µm after 24 hours culture. (C)-(D) Live/Dead Staining of hMSCs in linear channels. .... 71
Figure 3-11. Effect of F108 on selective seeding of human MSCs in polydimethylsiloxane (PDMS) microchannels ...... 72
Figure 3-12. Effect of F108 selective seeding of human MSCs in collagen microchannels.
...... 73
Figure 3-13. Fraction of MSCs attached to the PDMS and Collagen microchannels after
F108 treatment...... 74
Figure 4-1. Schematic of ultrastructure of articular cartilage...... 88
Figure 4-2. Illustration of experiment design...... 88
Figure 4-3. Schematic of nuclear alignment angle measurement...... 94
Figure 4-4. Microchannel pattern on silicon wafer for tensile testing...... 95
VIII
Figure 4-5. Tensile testing of Collagen-hMSC constructs...... 96
Figure 4-6. MSCs morphology and organization as a function of time in guidance channels...... 99
Figure 4-7. Chondrogenesis under microscale guidance after 21 days culture...... 101
Figure 4-8. Alignment of actin microfilaments in guidance channels...... 102
Figure 4-9. Actin Fiber Orientation Angle under Microscale Guidance ...... 102
Figure 4-10. Cell nucleus orientation under microscale guidance...... 104
Figure 4-11. Chondrogenesis under microscale guidance...... 106
Figure 4-12. Confocal fluorescent images of MSCs in guidance channels after 21 days culture ...... 107
Figure 4-13. Effect of microscale guidance on mechanical properties...... 110
Figure 5-1. Schematic of hMSC-CG Based rolled-up large scale construct formation .. 127
Figure 5-2. Microchannel pattern on a silicon wafer for the rolled-up constructs ...... 130
Figure 5-3. Tissue tensile modulus of elasticity under microscale guidance...... 133
Figure 5-4. Schematic of rolled-up construct for compressive testing...... 134
Figure 5-5. Histology result of Autonomous rolled-up collagen-hMSC constructs ...... 138
Figure 5-6. Autonomous rolled-up CG-hMSC constructs. 2 hours after seeding (A). 7 days in culture (B-C). 21 days in culture (D-F)...... 139
Figure 5-7. Toluidine blue staining of Autonomous rolled-up CG-hMSC constructs with
250 μm thick CG membranes after 3 weeks in culture...... 139
Figure 5-8. Toluidine blue staining of IRCG250 constructs after 3 weeks in culture ... 140
IX
Figure 5-9. Patterned 100 μm thick CG membrane scaffolds featuring 100 μm channels with 100 spacings (A-C). Multi-layered IRCG100-100C_100S immersed in chondrogenic medium (E) and after 21 days chondrogenic culture (D) ...... 142
Figure 5-10. Histological appearance of induced rolled-up CG-hMSC constructs
(IRCS100)...... 143
Figure 5-11. Histology result of the induced rolled-up CG-hMSC construct (IRCG100-
100C_100S) with guidance channles ...... 145
Figure 5-12. Histology and Immunohistochemistry an induced rolled-up CG-hMSC construct. IRCG100-100C_50S construct cultured in chondrogenic medium for 3 weeks
...... 146
Figure 5-13. Histological appearance of the induced rolled-up CG sponge-hMSC construct after 3 weeks in culture...... 147
Figure 5-14. SHG Images of ECM collagen produced by hMSCs within linear channels of widths 100µm (A) and hMSCs randomly seeded on constructs without channels (B)
...... 148
Figure 5-15. Sirius-red stained longitudinal section of IRCG100 constructs ...... 149
Figure 5-16. Effect of microscale guidance on mechanical properties of large scale
IRCG100 constructs. IRCG100 constructs were subject to compressive testing ...... 152
Figure 5-17. Glucose consumption rate (A) and lactate production rate (B) per cell for the
IRCG100 constructs during 21 days in chondrogenic culture...... 154
X
ACKNOWLEDGEMENTS
On the journey of pursuing my PhD in the past years, I have been blessed and helped by many people. Without them, this dissertation would not be complete and I wouldn’t be who I am today. I would like to express my sincere gratitude to Professor Harihara
Baskaran who is an intelligent, open-minded, and supportive advisor. I am very grateful to have been part of his lab. Throughout my Ph.D. studies, he has mentored me by guiding my Ph.D. research, acting as a role model in the scientific field, and helping me to develop as an independent scientist. I appreciate his support and encouragement during my time in his laboratory.
I am very thankful to my committee member, Professor Jean Welter, for his valuable guidance in the biology field and for making the facilities available in his laboratory for my research. Additionally, I would like to thank my committee member, Professor Heidi
Martin, for her helpful advice that improved my research. Finally, I would like to express my special thanks to Professor C.C. Liu who is not only a great researcher but also a great caretaker for helping me settle down in Cleveland in the beginning and always showing his care about me during my Ph.D. studies.
I would like to acknowledge the members of Skeletal Research Center and Electronics
Design Center, especially Lori Duesler who is a friendly lady who helped me perform biochemical assays and provided assistance whenever necessary. Additionally, I would like to thank Amad Awadallah for helping in the histology process. Moreover, I would like to thank Professor Joseph Mansor of the Department of Mechanical and Aerospace
XI
Engineering for mechanical testing, Professor Takao Sakai of the Department of
Biomedical Engineering at the Learner Research Institute for his help in polarized light microscopy, and Dr. Judith Drazba of the Imaging Core at the Lerner Research Institute for helping me in second harmonic generation microscopy.
I would like to express my gratitude for all the members in Biotransport group including former members Wan-Hsiang Liang and Saheli Sarkar and current members Alexander L.
Rivera, Valencia Williams, and Kuo-Chen Wang who accompanied with me in my research life supported each other in times of need, and made my stay in Cleveland a memorable and wonderful time. I would particularly like to express my special appreciation to Alexander L. Rivera, who has been working in the same project with me during the past five years, for supporting me in the project and helping me with the manuscript writing and lab management.
In the end, I would like to show my greatest appreciation to my family－my parents, my grandfather, my sister, and my brothers for their support, unconditional love, and encouragement. I would also to thank all my good friends in Cleveland for their support and for making my life in Cleveland happy and memorable.
XII
A Multiscale Approach to Stem Cell-based Chondrogenesis
for Cartilage Repair
Abstract
by
CHIH-LING CHOU
Tissue engineering is a possible method for long-term repair of cartilage lesions, but current tissue-engineered cartilage constructs have inferior mechanical properties compared to native cartilage. This problem may be due to the lack of an oriented structure in the constructs at the microscale that is present in the native tissue. In this study, we utilize contact guidance to develop constructs with microscale architecture for improved chondrogenesis and function. Stable channels of varying microscale dimensions were formed in collagen-based and polydimethylsiloxane membranes via a combination of microfabrication and soft-lithography. Human mesenchymal stem cells
(hMSCs) were selectively seeded in these channels. The chondrogenic potential of hMSCs seeded in these channels was investigated. We demonstrate selective seeding of viable hMSCs within the channels. hMSC aligned and produced mature collagen fibrils along the length of the channel in smaller linear channels of widths 25-100 µm compared to larger linear channels of widths 500-1000 µm. Further, substrates with microchannels that led to cell alignment also led to superior mechanical properties compared to constructs with randomly seeded cells or selectively seeded cells in larger channels. The ultimate stress and modulus of elasticity of constructs with cells seeded in smaller
XIII channels increased by as much as four folds. Furthermore, we extended the 2- dimensional finding and successfully created 3-dimensional large scale constructs (3.5 mm in diameter × 18 mm in length) with microscale architecture for in vivo applications.
Histology and immunohistochemistry indicated extensive GAG and collagen type II production in 3-dimensional construct, which are both indicative of chondrogenesis. Our results show that the microscale guidance channels incorporated within the 3-dimensional cartilage constructs lead to the production of aligned cell-produced collagenous matrix and enhanced mechanical function. The tissue modulus of elasticity of 3-dimensional cartilage constructs containing guidance increased by as much as six times compared to constructs without channels. Overall, these findings offer new insight into how microscale guidance channel regulates matrix deposition and long term construct development.
XIV
Chapter 1
INTRODUCTION
1.1 Purpose of the Study
Tissue engineering has tremendous potential for long‐term repair of cartilage lesions but current tissue engineered (TE) cartilage constructs have inferior mechanical properties compared to native hyaline cartilage. We hypothesized that the suboptimal mechanical properties of cartilage constructs obtained via these approaches are the result of their lack of the organized ultrastructure present in native cartilage tissue. When compared to the highly organized structure of native hyaline cartilage, TE cartilage displays randomly oriented microscale architecture. In this project, we investigated methods to produce structurally oriented TE cartilage constructs. Therefore, the aim of this project was to direct differentiating human mesenchymal stem cells (hMSCs) to create microscale oriented extracellular matrix (ECM) similar to the native tissue structure. We tested the hypotheses that the microscale topography of TE scaffolds can be used to guide MSCs to form self-organizing and oriented 2- and 3-dimensional structures in vitro. This study provided us with fundamental knowledge of the controlled chondrogenesis of MSCs that can be used for rational engineering of the next-generation of scaffolds for structured chondrogenesis.
1
1.2 Significance and Motivation of the study
1.2.1 Articular Cartilage Injury
Over 49.9 million individuals suffer from arthritis, which is one of the major causes of disability in the United States (MMWR 2010). This disease usually results from age- related degeneration, trauma, or sports injuries which lead to severe joint pain. Articular cartilage, as an avascular tissue that lacks a blood supply and the subsequent wound healing response, has very little intrinsic repair capacity. It cannot mobilize or recruit sufficient numbers of reparative cells into the damaged area nor can it establish the microenvironment for the repair process (Owen and Friedenstein 1988; Buckwalter 1998;
Bhosale and Richardson 2008). Partial –thickness articular cartilage defects are typically not repaired and deteriorate with time. Full-thickness defects that penetrate the subchondral bone have limited healing capacity and are usually restored with fibrocartilage, which has inferior biological and mechanical properties compared to hyaline cartilage tissue. Therefore, measures must be taken to repair cartilage lesions
(Detterline, Goldberg et al. 2005).
1.2.2 Limitations of Current Treatment Options
Current treatments of cartilage defects aim to relieve the symptoms temporarily and prevent further degeneration rather than offering a long-term solution for the problem.
Pharmacological treatments such as analgesics, cox-2 inhibitors, and hyaluronic acid injections are symptom relief therapies, which aim to minimize the pain and swelling of the joints.(Adams, Atkinson et al. 1995; Shimizu, Yoshioka et al. 1998; Peloso and
Scheiman 2001)..Current surgical techniques including subchondral drilling, abrasive
2 chondroplasty, and microfracture aim to stimulate the bone marrow stem cells below the subchondral bone to enter the cartilage defect (Simon 1999; Bhosale and Richardson
2008; Ahmed and Hincke 2010), which encourages the native repair process. However, these surgical procedures ultimately result in the formation of fibrocartilagenous scar tissue that lacks the mechanical properties of native articular cartilage (Curl, Krome et al.
1997; Nehrer, Spector et al. 1999; Kasemkijwattana, Kesprayura et al. 2009). Despite considerable efforts, these cartilage repair techniques provide less than ideal repair tissue, and none of these techniques have been proven to successfully regenerate long lasting hyaline cartilage tissue to replace damaged cartilage.
1.2.3 Challenges for Tissue-Engineered (TE) Cartilage
Tissue engineering has been proposed as a promising method for the development of cartilage constructs to treat articular cartilage injuries but has thus far failed to yield optimal cartilage repair (Temenoff and Mikos 2000). One of the major challanges for current cartilage tissue engineering is to develop TE constructs that emulate the exquisite tissue microstructure and the subsequent unique mechanical properties of native articular cartilage (Song, Baksh et al. 2004). Cartilage tissue engineering approaches usually involve the traditional tissue engineering strategy of culturing scaffolds seeded with mesenchymal stem cells (MSCs) or chondrocytes and growth factors to produce cartilage constructs in vitro (Temenoff and Mikos 2000). Using in vitro bioreactor culture, our group has shown that we can produce TE constructs that are biochemically similar to native articular cartilage [Figure1-1] (Liang, Kienitz et al. 2010). The toluidine blue staining and type II collagen staining showed that the extracellular matrix (ECM) produced by the differentiating MSCs within the Collagen - glycosaminoglycans sponge
3
TE construct was rich in glycosaminoglycans (GAG) and collagen type II, the two primary ECM components of cartilage.. Despite this considerable success, these TE cartilage constructs were structurally amorphous and thus clearly failed to duplicate articular cartilage, which possesses an exquisite architecture [Figure1-2]. Therefore, these TE cartilages constructs displayed inferior mechanical strength when compared to native cartilage. The mechanical properties of native articular cartilage are depth- depenedant and an increase in compressive strength from the surface of the cartilage to the deep zone is attributed to the ECM alignment within these regions (Wilson, Huyghe et al. 2007; Shirazi, Shirazi-Adl et al. 2008). In this study, we addressed the major limitation of current cartilage tissue engineering: the inferior mechanical properties of TE constructs. To produce structurally oriented and mechanically enhanced tissue, we fabricated constructs with microscale guidance channels that guide the differentiating
MSCs to produce aligned ECM with enhanced mechanical properties. This study provides us with fundamental knowledge for controlled and structured chondrogenesis by
MSCs. The design parameters for rational tissue engineering in this work can be utilized for designing next-generation scaffolds for structured chondrogenesis of cartilage constructs as well as used for other stem cell-based tissue engineering research where spatial cellular architecture plays a key role such as nerves and blood vessel tissue.
4
Figure 1-1. MSC seeded CG sponge TE construct after 3 weeks in bioreactor chondrogenic culture. DAPI staining on day 1 (A). Toludine blue staining (B)
Immunohistochemistry staining (C) Scale bar: 1000 µm. (Liang, Kienitz et al. 2010)
5
Figure 1-2 Orientation of the collagen type II fibrils throughout the depth of native articular cartilage (A) [Figure from (Wilson, van Donkelaar et al. 2004)]. Ultrastructure of human articular cartilage after freeze fracture processing (B) [Figure from (C. M. Jack, S. S. Rajaratnam et al. 2012)].
1.3 Overall Research Goals
The underlying hypothesis of this study is that functional survival of TE cartilage within the joint environment will require a 3-dimensional structural organization of the TE constructs which emulates that of native cartilage. Therefore, this study addressed a critical but under-investigated issue in scaffold engineering for tissue engineering approaches to treat cartilage injuries, which is to determine whether physical features of a biodegradable scaffold can be used to influence and direct the structure of the tissue formed by cells. Therefore, the ultimate goal for this study was to test the hypothesis regarding the impact of microscale architectural properties of TE scaffolds on MSC differentiation, specifically on their ability to form self-organizing and oriented 2- and 3- dimensional structures in vitro.
We utilized the method of contact guidance in which we used microscale physical features (guidance channels) on the biomaterial scaffold surface to control spatial cellular organization, cellular morphology, and cellular function of hMSCs. Native articular cartilage has a highly organized structure [Figure 1-2]. The ECM of cartilage contains collagen fibrils that are highly organized in different orientations depending on the particular cartilage zone. In the deep zone of native articular cartilage, the perpendicular arrangement of the collagen fibrils relative to the subchondral bone [Figure 1-2: (A)-(B))]
6
(Wilson, van Donkelaar et al. 2004; C. M. Jack, S. S. Rajaratnam et al. 2012) has been shown via mathematical modeling to enhance its compressive properties (Wilson,
Huyghe et al. 2007; Shirazi, Shirazi-Adl et al. 2008). For improved mechanical function of TE cartilage constructs, we aimed to direct the MSCs to produce ECM aligned perpendicular to the diameter of the construct in order to achieve a structure similar to the deep zone of native articular cartilage. To achieve this goal, the first objective of this study was to develop a technique that allows for greater control of MSC adhesion and spatial organization over the scaffolds. Specific microfabrication techniques were developed for embedding microscale patterns (guidance channels) onto the two scaffolds:
Collagen-Glycosaminoglycan (CG) and poly(dimethylsiloxane) (PDMS) membranes.
Thin membranes containing microchannel designs that include various channel dimensions were fabricated. Our first objective was achieved by tuning surface chemistry and topography of the scaffolds to direct cell selective attachment and spreading within the guidance channels of the scaffolds.
The second objective of this study was to investigate the effects of contact guidance on hMSC-based chondrogenesis. Microscale guidance channels were designed and produced on the scaffold surface and were used to guide differentiating MSCs to produce oriented
ECM. The microscale guidance effect on cell alignment and ECM function in a hMSC- based chondrogenesis model was investigated. This study provided us information on designing parameters for rational tissue engineering of cartilage constructs.
However, these 2-dimensional constructs do not provide a solution for 3-dimensional cartilage defects, and therefore, lack clinical relevance. Hence, the final objective of this research project was to develop a method of scaling these 2-dimensional constructs to 3-
7 dimensional TE constructs to overcome this limitation. Based on the design parameters in our findings for 2–dimensional constructs, we optimized the scaffold design and assembled a 3-dimensional structure mimicking an ultrastructure component of native cartilage. These 3-dimensional constructs were developed by rolling-up 2-dimensional scaffolds containing guidance channels and seeded with MSCs. The microscale guidance effect on cell alignment and ECM function in this 3-dimensional hMSC-based chondrogenesis model was investigated.
1.4 Thesis Outline
In chapter 2, we introduced the background knowledge for this dissertation includes the current clinic treatment for articular cartilage injury, challenges in current cartilage tissue engineering and the relationship between cartilage structure and guidance. To control differentiating hMSCs to synthesize ECM that emulates the aligned ECM structure of native tissue, the cells need to be localized within the channels; contact guidance and its efficacy assessment require the cells to be located inside the channels. In chapter 3, we described our first objective of this study which was to localize the hMSCs within the guidance channels.We developed a technique that allows for greater control MSC adhesion and spatial organization over the two micropatterned scaffolds: CG and PDMS membranes.
After MSCs selectively adhering within the channels, in chapter 4, the microscale guidance effect on cell alignment and ECM function in an hMSC-based chondrogenesis model was investigated. The chondrogenic potential of MSCs seeded in these channels was investigated by culturing them for 3 weeks under differentiating conditions, and then
8 evaluating the subsequent synthesized tissue for mechanical function and by type II collagen immunohistochemistry.
Furthermore, in chapter 5, we extended the 2-dimensional construct findings to a 3- dimensional construct for translational studies. We successfully created 3-dimensional constructs for in vivo applications by rolling up 2-dimensional hMSC-seeded CG based scaffold and formed 3-dimensional rolled-up large scale (3.5 mm in diameter × 18 mm in length) cartilage constructs with microscale architecture to guide the differentiating hMSCs to produce oriented ECM. ECM function of microscale guidance channels incorporated within the 3-dimensional large scale cartilage constructs was investigated in this chapter. Lastly, we concluded the findings in this work in chapter 6. The limitations and possible improvements are summarized in this section.
1.5 Summary
This study addresses a critical but under-investigated issue in scaffold engineering for tissue engineering approaches to treat cartilage injuries. We describe a basic science approach to determine whether microscale guidance features can be used to influence the architecture of the differentiating tissue. This study provided us with fundamental knowledge for controlled chondrogenesis by MSCs in 2 and 3-dimensional constructs that can be utilized for rational engineering of next generation scaffolds for structured chondrogenesis that displays superior mechanical properties.
1.6 References
Adams, M. E., M. H. Atkinson, et al. (1995). "The role of viscosupplenentation with hylan G-F 20 (Synvisc(R)) in the treatment of osteoarthritis of the knee: A Canadian
9 multicenter trial comparing hylan G-F 20 alone, hylan G-F 20 with non-steroidal anti- inflammatory drugs (NSAIDs) and NSAIDs alone." Osteoarthritis and Cartilage 3(4): 213-225.
Ahmed, T. A. E. and M. T. Hincke (2010). "Strategies for articular cartilage lesion repair and functional restoration." Tissue Engineering Part B-Reviews 16(3): 305-329.
Bhosale, A. M. and J. B. Richardson (2008). "Articular cartilage: structure, injuries and review of management." British Medical Bulletin 87(1): 77-95.
Buckwalter, J. A. (1998). "Articular cartilage: Injuries and potential for healing." Journal of Orthopaedic & Sports Physical Therapy 28(4): 192-202.
C. M. Jack, S. S. Rajaratnam, et al. (2012). "The modified tibial tubercle osteotomy for anterior knee pain due to chondromalacia patellae in adults.A five-year prospective study." Bone Joint Res 1(8): 167-173.
Curl, W. W., J. Krome, et al. (1997). "Cartilage injuries: A review of 31,516 knee arthroscopies." Arthroscopy 13(4): 456-460.
Detterline, A. J., S. Goldberg, et al. (2005). "Treatment options for articular cartilage defects of the knee." Orthopaedic Nursing 24(5): 361-366.
Kasemkijwattana, C., S. Kesprayura, et al. (2009). "Autologous chondrocytes implantation with three-dimensional collagen scaffold." Journal of the Medical Association of Thailand 92(10): 1282-1286.
Liang, W.-H., B. L. Kienitz, et al. (2010). "Concentrated collagen-chondroitin sulfate scaffolds for tissue engineering applications." Journal of Biomedical Materials Research Part A 94A(4): 1050-1060.
MMWR (2010). "Prevalence of doctor-diagnosed arthritis and arthritis-attributable activity limitation --- United States, 2007-2009." MMWR. Morbidity and mortality weekly report 59(39): 1261-5.
Nehrer, S., M. Spector, et al. (1999). "Histologic analysis of tissue after failed cartilage repair procedures." Clinical Orthopaedics and Related Research(365): 149-162.
Owen, M. and A. J. Friedenstein (1988). "Stromal stem-cells - marrow-derived osteogenic precursors." Ciba Foundation Symposia 136: 42-60.
Peloso, P. M. and J. M. Scheiman (2001). "The economic implications of cyclooxygenase-2-specific inhibitors." American Journal of Medicine 110(Supplement 3A): 50S-54S.
10
Shimizu, C., M. Yoshioka, et al. (1998). "Long-term effects of hyaluronan on experimental osteoarthritis in the rabbit knee." Osteoarthritis and Cartilage 6(1): 1-9.
Shirazi, R., A. Shirazi-Adl, et al. (2008). "Role of cartilage collagen fibrils networks in knee joint biomechanics under compression." Journal of Biomechanics 41(16): 3340- 3348.
Simon, L. S. (1999). "Osteoarthritis: a review." Clinical cornerstone 2(2): 26-37.
Song, L., D. Baksh, et al. (2004). "Mesenchymal stem cell-based cartilage tissue engineering: cells, scaffold and biology." Cytotherapy 6(6): 596-601.
Temenoff, J. S. and A. G. Mikos (2000). "Review: tissue engineering for regeneration of articular cartilage." Biomaterials 21(5): 431-440.
Wilson, W., J. M. Huyghe, et al. (2007). "Depth-dependent compressive equilibrium properties of articular cartilage explained by its composition." Biomechanics and Modeling in Mechanobiology 6(1-2): 43-53.
Wilson, W., C. C. van Donkelaar, et al. (2004). "Stresses in the local collagen network of articular cartilage: a poroviscoelastic fibril-reinforced finite element study." Journal of Biomechanics 37(3): 357-366.
11
Chapter 2
BACKGROUND
2.1 Articular Cartilage
2.1.1 Knee joint and Cartilage
Cartilage, as an internal cellular support tissue rich in fibrous protein and mucopolysaccharides (Cole and Hall 2004; Cole 2011) can be classified into 3 types in mammals: hyaline cartilage, fibrocartilage and elastic cartilage. All of them are composed of chondrocytes and extracellular matrix (ECM) macromolecules and are classified depending on the composition and distribution of the fibers in the ECM
(Temenoff and Mikos 2000). Hyaline cartilage, a semi-transparent white tissue, is found in the knee. Hyaline cartilage is the typical type of cartilage that people generally refer as cartilage and is also called articular cartilage. Elastic cartilage, which is generally characterized by the elastin fiber presented in the ECM, is found in the ear and nose.
Elastic cartilage supports the shape and flexibility for these organs. Fibrocartilage is usually found in intervertebral discs, tendons, and ligaments (Stockwell 1979; Temenoff and Mikos 2000; Cole 2011). Fibrocartilage contains fibrous tissue and a higher ratio of collagen type I to collagen type II in the ECM than the articular cartilage. (Stockwell
1979; Temenoff and Mikos 2000; Cole 2011).
The Knee is the largest joint in the body and is composed of cartilage, bones, ligaments, tendons and muscles. Articular cartilage covers the bones (the end of femur and the top of
12 the tibia) at the joint surface in the knee (Temenoff and Mikos 2000) [Figure 2-1].
Articular cartilage is a low friction and load-bearing soft tissue which provides a smooth, low friction gliding surface that allows movement of the joint surface by stress absorption, stress distribution, and joint lubrication. The synovial membranes in the joint form a barrier to retain the synovial fluid in the knee [Figure 2-1]. Due to the avascular nature of articular cartilage, cartilage lacks blood vessels and nerves. Hence, the synovial fluid plays an important role for providing lubrication and nutrients in cartilage (Temenoff and
Mikos 2000).
Figure 2-1. Schematic of a Synovial Joint [Figure from (Setton 2008)]
2.1.2 Ultrastructure and Constituents of Articular cartilage
Chondrocytes, very specialized cells, represent only 1% to 5 % of the volume of hyaline cartilage and are responsible for synthesizing the ECM and replacing the degraded matrix molecules to maintain the correct composition of the matrix and the mechanical
13 properties of cartilage tissue (Bhosale and Richardson 2008). Some chondrocytes have been shown to sense their mechanical environment via cilia and further adjust matrix properties in response to loading (Buckwalter and Mankin 1998; Temenoff and Mikos
2000). Due to the avascular nature, chondrocytes obtain nutrition supply by diffusion through the ECM (Temenoff and Mikos 2000; Bhosale and Richardson 2008).
Chondrocytes have been reported to survive in a low oxygen concentration environment and live on anaerobic metabolism (Pfander and Gelse 2007).
Cartilage is largely (60–80% of wet weight) water, which helps nutrient transport, low friction interface generation, and load-dependent deformation of the cartilage (Temenoff and Mikos 2000). A total of 90% of the dry weight of the tissue is ECM (Linn and
Sokoloff 1965; Hardingham and Fosang 1992), which is principally composed of type II collagen, proteoglycans, and non-collagenous proteins [Figure 2-2 (A)]. The major component (90-95%) of the macrofibrilar network in articular cartilage is type II collagen
(Bhosale and Richardson 2008) but types VI and XI are also present, as are very small amounts of types I ,III, V, and XIV. Proteoglycans constitute 10 to 20 % of the wet weight of hyaline cartilage and are composed of 95% polysaccharides and 5 % proteins.
The subunit of the proteoglycans is glycosaminoglycan (GAG) chains. Chondroitin sulphate and keratin sulphate are the two major disaccharide molecules of GAG chains
[Figure 2-2 (B)]. GAGs are negatively charged which assistance the interaction with water and therefore proteoglycan have the ability to retain the fluid and electrode balance
(Buckwalter and Mankin 1998; Temenoff and Mikos 2000; Bhosale and Richardson
2008). Proteoglycans fill the space between the fibrilar network and contribute to the stress absorption and distribution in articular cartilage because of highly negative charge
14 which facilitates to attract water. Tissue fluid in the matrix contains a lot of cations which balance the negative charge on the GAGs. The interactions between tissue fluid and ECM provide cartilage unique ability to resist compression and back to normal shape after loading (Temenoff and Mikos 2000).
Based on morphology of the matrix, from top to bottom, articular cartilage can be classified into four zones: superficial zone, radial zone, deep zone and calcified zone.
Chondrocytes regulate the matrix that contains collagen, proteoglycan and noncollagenous proteins in each zone and form a greatly specialized tissue [Figure 2-2
(D)]. The matrix in each zone is divided into three zones: the pericellular zone, territorial zone and inter-territoirial zone [Figure 2-2 (B)]. The pericellular zone is the area close to cell membrane rich in proteoglycan. The territorial zone is the outer layer surrounding the pericllular and chondrocytes. The collagen fibers in this area are organized cross to reach other as a backet structure surrounding the chondrocytes. The inter-territoirial zone is the principal region contributing the mechanical properties for the cartilage. It presents largest volume and largest diameter of collagen fibrils. The collagen fibrils oriented parallel to the joint surface in superficial zone and perpendicular to the joint surface at deep zone (Temenoff and Mikos 2000; Bhosale and Richardson 2008).
Superficial zone is the thinnest zone among all the layers and contains higher amount of collagen and lower amount of proteoglycan than other zones. Collagen fibrils are oriented parallel to joint surface which provide tensile strength and help resist shear force in the joint surface. In addition, superficial zone also has been reported responsible for isolating the cartilage from synovial immune system (Buckwalter and Mankin 1998; Temenoff and
Mikos 2000; Bhosale and Richardson 2008). In radial zone, proteoglycan content is
15 higher and collagen fibers are thicker than superficial zone. The collagen fibers arranged randomly in the radial zone. Chondrocytes in deep zone are in round shape and stacking in column perpendicular to the joint surface. The collagen fibril aligned perpendicular to the joint surface and chondrocytes in this area have high biosynthesis ability (Wong,
Wuethrich et al. 1996). Calcified zone is the area closet to subchondroal bone and plays as an transition area transforming from soft cartilage to stiff bone. The cells possess low metabolism in this zone (Buckwalter and Mankin 1998; Temenoff and Mikos 2000;
Bhosale and Richardson 2008).
Figure 2-2. Ultrastructure and composition of articular cartilage [Figure from (Bhosale and Richardson 2008)]
16
2.1.3 Articular Cartilage Injury and Repair
Osteoarthritis (OA) [Figure 2-4], known as the fifth leading cause of disability (Michaud,
McKenna et al. 2006) affects over 27 million people in the United States in 2008
(Helmick, Felson et al. 2008). This degenerative joint disease is generally characterized by the loss of hyaline cartilage [Figure 2-3, middile] and cause serious sharp joint pain.
OA lesion is usually found in load-bearing cartilage and the cause of OA may result from trauma, abnormal mechanical stress of the joint, degradation and age-related disease (15).
The disruption typically start from the loss of proteoglycan follows by the disruption of the collagenous fibrillar network. It may start from a focal small matrix disruption and gradually become the destruction of the thick defect or even extend to the whole layer of the articular cartilage (Bullough 1981; Goldring 2000; Hunziker 2002). Water content of the osteoarthritic cartilage tissue increases to more than 90% due to the matrix disruption which result in the reduction in load bearing function of the cartilage (Bhosale and
Richardson 2008).
17
Figure 2-3. Schematic of Normal (left) and Arthritic Knee Joint (right) (Figures from MedicineNet. Inc. http://www.medicinenet.com/rheumatoid_arthritis/article.htm).
Figure 2-4. Illustration of Normal Cartilage (left) and Osteoarthritic Cartilage (right) [Figure from(Kuo, Li et al. 2006)]
It has been believed that whether the cartilage defect will gradually disrupt and lead to
OA may contribute to some important factors: the size of defect, depth of defect, limb alignment and preinjury joint degeneration (P.K. Bos 2010). Injuries in articular cartilage generally either do not repair or partially repair under certain circumstances (Hunziker
2002). The repair results were influenced by depth depending on if the injury discrution extending to subchondral vascular bone marrow (Bhosale and Richardson 2008). Based on the depth of the defects, articular cartilage defects can be classified into two types: partial thickness defect and full thickness defects [Figure 2-5]. Partial thickness defect are the disruption on the articular cartilage surface without penetrate to subcondral bone
[Figure 2-5 (A)]. The partial thickness defects do not heal spontaneously and may
18 increase in size and depth over time (Buckwalter, Rosenberg et al. 1990; Bhosale and
Richardson 2008). This is due to the lack of access to the bone marrow mesenchymal progenitor cells in the bone marrow area. Full thickness defect are the disruptions down to the subcondral bone [Figure 2-3 (B)]. While the partial thickness defects do not heal on their own, full thickness defect have certain form of repair capacity because bone marrow mesenchymal progenitor cells are able to migrate and fill the lesion. Once the injury penetrates to bone and bone marrow area, the defect area will undergo spontaneously wound healing response and locally formed fibrin clot. (Hunziker 2002;
Bhosale and Richardson 2008). The clot matrix that contains proteins, glycolproteins and blood cells are able to completely fill the defect with specific size (1mm to 2mm in diameter). The repair is limited in the lesions that are not completely filled by blood clot and may be necessary to insert biomaterials for complete repair (Jackson, Lalor et al.
2001; Hunziker 2002). After going through spontaneously repair in full thickness defect, mesenchymal progenitor cells start migrating to the matrix. After a few weeks, the defect will be substituted by a fibrocartilagous scar tissue. However, the fibrocartilagous scar tissue that formed in this repair process has been showed structurally inferior to hyaline cartilage and yield poor mechanical property (Simon 1999; Temenoff and Mikos 2000;
Hunziker 2002; Redman, Oldfield et al. 2005; Bhosale and Richardson 2008).
19
Figure 2-5. Schematic of cartilage defects with different depths. Partial-thickness defects (A) and full-thickness defects (B).
The other factor affecting the cartilage repair response is the size of the defects. Several studies showed that defects less than a critical size of 3 mm in diameter may lead to complete spontaneous repair of articular cartilage while larger defects do not repair completely (Convery, Akeson et al. 1972; Shapiro, Koide et al. 1993; ButnariuEphrat,
Robinson et al. 1996; Jackson, Lalor et al. 2001). Larger defects result in the damage to the ECM and chondrocytes that cannot be self-repaired. Defects smaller than 1 cm2 are may not affect the load-bearing function (Bhosale and Richardson 2008). In addition, age is another important factor while the depth of injury has been shown age-related in both animal and human study (Cahill 1995; Johnstone and Yoo 1999; Bhosale and Richardson
2008). It has been reported that juvenile are found to have more osteochondral defect whereas adults are found to have more chondral defects (Cahill 1995).
2.1.4 Current Treatment for Articular Cartilage Injury
Defect filling and regeneration of cartilage surface with the best possible tissue is principal task for cartilage injury repair (P.K. Bos 2010). Current treatment of the cartilage defects aim to relieve the symptoms temporarily and prevent further
20 degeneration rather than offering a long-term solution for the problem. Pharmacological treatment such as analgesics, cox-2 inhibitors, or hyaluronic acid injections are symptom relief which aim to minimize the pain and swelling of the joints (Linn and Sokoloff 1965;
Hardingham and Fosang 1992; Buckwalter and Mankin 1998; Cole and Hall 2004;
Bhosale and Richardson 2008). Since articular cartilage defects do not heal spontaneously, it will be a benefit if the defect downward to the subchondral bone. Once the bone and bone marrow area are involved in the defect, classical wound healing response (bleeding and hematoma formation) will be initiated which lead to the spontaneously repair process. The blood clot with various kinds of proteins and growth factors serves as an optimal environment for mesenchymal progenitor cells to differentiate Current surgical treatment strategies for cartilage defects are mostly based on bone marrow stimulation penetrating to the subcondral bone, disrupts the blood vessels and induce the spontaneously healing process (Hunziker 2002; Bhosale and
Richardson 2008). However these methods usually result in the formation of a fibrocartilage scar tissue and not capable to provide long lasting biomechanical properties that hyaline cartilage possess. These methods are usually applied once the other has failed.
Current treatment options for cartilage repair include symptomatic treatments such as drilling, joint debridement, spongialization, microfracture, mosaicplasty and tissue engineering strategy such as autologous chondrocyte implantation (ACI) (McLaren,
Blokker et al. 1991; O'Driscoll 1998; Hunziker 2002; Redman, Oldfield et al. 2005;
Bhosale and Richardson 2008). Joint debridement is a procedure used to relieve the pain which involves articular trimming ,articular abrasions and other procedure (McLaren,
Blokker et al. 1991). The debridement aim to remove the damage tissue form the articular
21 cartilage and ease the pain. It has been showed successful in treating early stage OA
(Jackson and Dietrichs 2003; Redman, Oldfield et al. 2005). However, this method showed offering palliative effect rather than inducing the self-repair of the cartilage
(Redman, Oldfield et al. 2005). The spongialization was described as a modified method of debridement which further removes the involved subchondral bone along with the damage cartilage to stimulate the repair process (Ficat, Ficat et al. 1979; Bhosale and
Richardson 2008). Pride drilling is a method based on stimulation the spontaneously repair response which involves removing the loose pieces of cartilage, drilling a hole to subchondral bone of the damaged tissue and induce bleeding (Insall 1967; Hunziker
2002). Thought it has been reported to successfully help the patients suffering from the joint pain for short term period (Schmidt, Schulze et al. 1988), the fibrocartilage scar tissue was formed in the repair tissue and can only last for a few years (Hice, Freedman et al. 1990). Microfracture technique is regarded as a modified method of Pride drilling. In microfacrture process, the debridement is first performed in the damaged tissue to remove the loose cartilage pieces follows by a bone marrow stimulation technique.
Multiple holes with 0.5-1mm in diameter were perforated in subchondral bone by a specialized awl with a distance of 3 to 4 mm apart which induce the bleeding and spontaneously repair. The advantages of this method including avoiding the overheating of the drilling method and producing rough surface facilitate new-generated tissue attached. Thought it has been reported this method can only last for 5 to 7 years depending on patient’s (Hunziker 2002; Steadman, Briggs et al. 2003) and the inferior biomechanical scar tissue is accompany with repair articular surface (Mankin 1974;
Mankin 1974), it is still one of the most popular surgical treatment in current stage due to
22 the technical accessible and minus invasive (Bhosale and Richardson 2008).
Mosaicplasty is a technique that involves removing osteochondral bone plug from a low- loading bearing area in the joint for filling the chondral defect of the patient. The advantage of this technique is it provides a stable surface for load bearing and it’s been reported repaired cartilage last for up to 7 years after treatment for small and medium sizes lesions (Hangody, Feczko et al. 2001; Hangody, Vasarhelyi et al. 2008). However, the gap of these plug are filled with fibrocartilage scar tissue that provide lower stability for the plugs and therefore lack of lateral integration to the native cartilage tissue
(Bentley, Biant et al. 2003; Bhosale and Richardson 2008).
Autologous Chondrocyte Implantation (ACI) [Figure 2-6] (Redman, Oldfield et al. 2005), a cell-based therapy, has been developed for cartilage repair since 1994 (Redman,
Oldfield et al. 2005). The first step of this method is excising the autologous chondrocyte from a healthy relatively non-weight loading site of the articular cartilage. The cells were then cultured in vitro for proliferating under best possible environment and the cultured autologous chondrocyte was further injected into the defects (Brittberg, Lindahl et al.
1994; Chanlalit, Kasemkijwattana et al. 2007). More than 70% of good result in ACI repair method has been reported and several studies showed ACI yield hyaline-like tissue and the repair tissue last for more than 11 years (Minas 2001; Brittberg M 2003; Bhosale and Richardson 2008).
23
Figure 2-6. Schematic of Autologous Chondrocyte Implantation (ACI) procedure. [Figure from (Redman, Oldfield et al. 2005) ]
Although ACI has been shown yield better results over the conventional surgical treatment, the drawback of this method includes small amount of available cells, slow expansion rate and the inadequate ability of the cells to proliferate (Cancedda, Dozin et al.
2003). In addition, several studies showed fibrocartilagenous tissue was still found after
ACI treatment (Clar, Cummins et al. 2005). A variety of modified tissue engineering methods for ACI are studied which combines ACI method with biomaterial scaffold and bioactive factors (Bhosale and Richardson 2008).
Current clinic treatment for cartilage repair often depends on the size and severity of the defects. Microfracture has been used to heal focal defects smaller than 2 cm2.
Mosaicplasty and ACI treatment are used for defect size of 2-3 cm2. For the defects more than 4 cm2 or multiple defects, osteochondral allograft are commonly employed. The
24 patients with end-stage osteoarthritic symptom, are typically treated with the total joint replacement (Williams, Chan et al. 2010).
2.2 Cartilage Tissue engineering
Tissue loss and organ failure raise a major health issue in the United States. With the lack of available source of suitable donors, the access for direct transplantation procedure for the patients is very limited (Chapekar 2000). With the increasing need of the organ donation and transplantation for the past decade (Wynn and Alexander), tissue engineering transplant serve as an optimal alternative which solve the major issue in direct transplantation : pathogen transmission, rejection and limited source. Tissue
Engineering is generally defined as the application combines the principles and methods of engineering life sciences and medicine to fundamentally understand and develop biological substitutes to repair, maintain or improve tissue or organ (Chapekar 2000).
Tissue engineering process can be performed either completely in vitro which fully functional mature tissue are developed in vitro and transplanted to the body or partially culture in vitro and completely matured in vivo [Figure 2-5] (Hunziker 2002). The
Principal of Tissue Engineering combines three key components: cells, supportive scaffold and signaling biomolecules to restore the pathologically damaged tissues or organ. Despite of some promising results for current medical treatments, most surgical treatments offers temporary symptom relief and company with the formation of inferior fibrous tissue which gradually disruption. Tissue Engineering serves as promising alternatives for cartilage repair. In typical cartilage tissue engineering [Figure 2-5] strategy, biocompatible and mechanical supportive scaffold were combined with suitable source of the cells (chondrocyte or mesenchymal stem cell) and appropriate biomolecules
25
(such as TGF-β1 and TGF-β3) to form tissue-engineered constructs in vitro for cartilage tissue repair [Figure 2-7].
Figure 2-7 Schematic diagram of typical tissue engineering strategy for cartilage repair. [Figure from (Chen, Rousche et al. 2006) ]
26
2.2.1 Cell Sourcing for Cartilage Tissue engineering
Cell selection plays a crucial role in tissue engineering. Generally, the ideal selected cells in tissue engineering should be renewable and non-immunogenic (Jason R. Fritz 2009).
Three major cell types have been extensively studied for cartilage tissue engineering: chondrocytes, embryonic stem (ES) cells, and adult mesenchymal stem cells (MSC)
(Song, Baksh et al. 2004). While chondrocyte is the only cell type responsible maintaining the tissue functions in cartilage, autologous chondrocytes directly from donors supposed to be the best choice for cartilage tissue engineering. Autologous chondrocytes are usually isolated from articular, auricular, costal, and nasoseptal cartilages (Nayyer, Patel et al.). Fully differentiated autologous chondrocytes have been reported to be an ideal cell source candidates for cartilage repair (Peterson, Minas et al.
2000). However, there are some downfall for using chondrocyte and limited it’s application. First, chondrocytes have to be extracted from healthy cartilage otherwise the result tissue showed exhibiting limited mechanical function than normal cartilage and therefore the source of autologous chondrocyte is very limited. In addition, the harvested chondrocytes may lose their phenotype during the in vitro expansion culture (Schnabel,
Marlovits et al. 2002; Jason R. Fritz 2009). Therefore, the optimal cell candidate for cartilage tissue engineering should possess easily accessible source and high expansion
/differentiating potentials. Stem cells have been proposed as a promising source for cartilage tissue engineering because of their self-renewal and multilineage differentiation potentials. Embryonic stem cells (ESCs) and mesenchymal stem cell (MSC) are regarded as two potential sources for cartilage repair (Lee and Hui 2006).
27
Human Embryonic stem cells (hESCs) are able to differentiate into all cell types in the body with its pluripotent potential and have almost unlimited self-renewal ability
(Thomson, Itskovitz-Eldor et al. 1998). Chondrogenic potential of hESCs for cartilage tissue engineering has been investigated via combining growth factors, biomaterials in different approaches (Toh, Lee et al. 2011). Though hESCs possess the potential of providing unlimited cells and the ability of differentiating into any type of the cells in embryo, ethical and sociological issues limits its development in tissue engineering application (Jason R. Fritz 2009). The other issue is ESCs have been reported have a high risk of teratoma formation due to the spontaneous differentiation (Heng, Cao et al. 2004;
Jason R. Fritz 2009). On the other hand, adult stem cells don’t have these issues. Adult
Mesenchymal stem cell (hMSC) were found in various type of tissue in the body such as bone marrow, synovia, muscle and adipose (Pittenger, Mackay et al. 1999; Toh, Lee et al.
2011). MSCs have promising ability of self-renewal and differentiation into various cell/tissue lineages including chondrocytes, osteoblasts and adipocytes under proper stimulation factors and in vitro culture (Pittenger, Mackay et al. 1999) [Figure 2-6] and therefore has been regarded as an excellent source for regenerative medicine. MSCs have been combined with biodegradable scaffolds and growth factors to produce cartilage constructs in vitro (Solchaga 2002; Welter 2002; Baskaran 2003; Solchaga L 2003;
Welter 2003; Baskaran 2004; Solchaga 2004; Welter 2004) and have shown great promises for cartilage tissue engineering applications.
28
Figure 2-8. Multilineage differentiation potentials of adult human Mesenchymal stem cell. [Figure from (Tuan, Boland et al. 2003)]
2.2.2 Biomaterials for Cartilage Tissue Engineering
The selection and design of ideal scaffolds plays an important role in tissue engineering.
Considering in the creation of TE constructs, the optimal scaffolds for cartilage tissue engineering should have the following properties. The scaffold must be biodegradable
29 possessing optimal mechanical property for supporting cells proliferation and migration; and excellent biocompatibility (Risbud and Sittinger 2002). A variety of scaffolds includes natural or synthetic materials have been employed for cartilage tissue engineering (Hunziker 1999; Hunziker 2002). The natural material includes protein-based polymers such as collagen (Speer, Chvapil et al. 1979), fibrin (Pelaez, Huang et al. 2009) and gelatin (Ponticiello, Schinagl et al. 2000); carbohydrate-based polymers such as hyaluronan (Erickson, Kestle et al.; Erickson, Huang et al. 2009), agarose (Awad,
Wickham et al. 2004), and alginate (Wei, Zeng et al. 2012) are widely and successfully used for this purpose (Hunziker 2002). Although natural biomaterials have been widely used in tissue engineering and showed promising results in many studies, there are issues and downfalls that limit their use. The first issue is their uncontrollable mechanical properties and degradation rate. In addition, these materials may cause serious immune response because they are derived from lives (Schmidt and Baier 2000; Lee, Singla et al.
2001).
Among all the natural materials, collagen is the most broadly distributed proteins in the body. The benefit using collagen in tissue engineering application is they are biodegradable, easily accessible and versatile (Parenteau-Bareil, Gauvin et al. 2010).
Collagen-based biomaterials have been used for implantation to cartilage defects (Harley,
Lynn et al. 2010). Among all the collagen, collagen type I have been reported as the relatively suitable biomaterials for in vitro use and has been tested no immune response issue in most human body (Charriere, Bejot et al. 1989; Eaglstein, Alvarez et al. 1999;
Parenteau-Bareil, Gauvin et al. 2010). Type I collagen are widely used in cartilage tissue engineering because of their flexibility and compatible with type II collagen which is the
30 major component of ECM in cartilage (Parenteau-Bareil, Gauvin et al. 2010). Type I collagen seeded matrix has been used for modify convention autologous chondrocyte implantation and have shown great promises (Gooding, Bartlett et al. 2006).
Synthetic polymer scaffolds includes poly (L-lacticacid) (PLLA), poly (L-glycolic acid)
(PLGA) and their derivatives are used for cartilage tissue engineering applications and have shown some promising results for the chondrogenesis potential (Banu, Banu et al.
2005; Williams and Gamradt 2008). One of the advantages for the synthetic materials is their biological and physical properties are controllable and can be tailored made to meet the specific needs. In addition, they are able to be degraded through chemical hydrolysis and the degradation degree is more consistent and predicable than natural material.
However, the biocompatibility is always the major concern for employing the synthetic scaffolds.
2.2.3 Challenges in Cartilage Tissue Engineering
Articular cartilage has little self-repair ability which contributes to the serious disruption and degeneration once injured. Current clinical treatment for cartilage lesions aim to help patients relieve the symptom and replace the lost tissue with best possible substitutes to allow damaged cartilage function as normal as possible instead of offering long term solutions (Jason R. Fritz 2009). Tissue engineering has great potential to contribute to innovative treatment for cartilage repair. The ultimate goal for cartilage tissue engineering is to generate replacement TE constructs similar to the native cartilage tissue in both structure and function which is stable in the long term. For cell associate issue includes chondrogenesis enhancement,
31
One of the major issues for current cartilage tissue engineering is developing TE constructs that emulate the exquisite multiphasic microstructure and the subsequent unique mechanical properties of native articular cartilage (Song, Baksh et al. 2004).
Current TE cartilage cultured with various types of scaffolds typically displays a compressive elasticity modulus between 10-15 KPa (Awad, Wickham et al. 2004; Bian,
Zhai et al. 2011; Huang, Farrell et al. 2011) and a tensile modulus between 0.23 MPa to
5 MPa (Fedewa, Oegema et al. 1998; Williams, Klein et al. 2005; Gemmiti and Guldberg
2006; Gratz, Wong et al. 2006; Huang, Baker et al. 2012) whereas native human hyaline cartilage possess a significant superior mechanical property with a compressive elasticity modulus of 1.16 to 7.75 MPa (Chen, Bae et al. 2001) and 0.7 to 12.5 MPa for tensile strength (Almarza and Athanasiou 2004; Huang, Stankiewicz et al. 2005).
Therefore, the inferior mechanical property of current TE construct is a major issue which prevents current constructs from reaching a clinical setting. The inadequate mechanical properties is contributed by two factors. First is the chondrogenesis potential of these constructs which determine the production of GAGs and collagen matrix. To enhance the chondrogenesis, recent studies have developed methods that greatly accelerate chondrogenesis and cartilaginous matrix production to produce constructs biochemically similar to native cartilage (Welter, Solchaga et al. 2007; Erickson, Kestle et al. 2012). With the advances for these studies, the major issue responsible for inferior mechanical property is attributed to the lack of an oriented microscale structure in TE constructs, which is present in the native tissue (Bhosale and Richardson 2008).
Scaffold associated issue includes bioactive factor and nutrient transport in the scaffold tissue interface, degradation rate management and scaffold-tissue integration (Sharma
32
2011). It has been well known that cartilage repair without vertical and lateral integration will lead to failure (Hunziker 2002). Another major challenge for current cartilage tissue engineering is the integration of TE constructs to the surrounding host tissue, in particular lateral integration to the host cartilage (Hunziker 2002).. There are many factors that directly or indirectly affect the tissue integration including biomaterial and tissue integration, proteoglycans, growth factors, cellular differentiation (Khan, Gilbert et al.
2008). TE constructs have been displayed better integration with bone than cartilage
(Tognana, Borrione et al. 2007). While the vertical integration of the TE constructs and host tissue are often observed (Tognana, Borrione et al. 2007), the lateral integration has become a serious issue in cartilage tissue engineering (Wakitani, Goto et al. 1994). TE implants have been showed to disrupt with time due to the lack of lateral integration between the host tissue and donor cartilage (Smith, Richardson et al. 2003; Lane, Massie et al. 2004). In ACI treatment, chondrocyte viability between the graft and the host edges and cell phenotype plays a crucial role for defect repair quality. While the density and phenotype of the chondrocytes determine the cartilage function, it has been reported that the area with dead cell hampers the integration between the tissue-engineered cartilage grafts and native cartilage and lead to the formation of inferior biomechanical fibrous scar tissue and cause treatment failure (Khan, Gilbert et al. 2008). Previous studies have showed that inhibition of chondrocyte death lead to enhance the bovine cartilage integrative repair (Archer, Redman et al. 2006; Khan, Gilbert et al. 2008). As most current TE constructs are investigated is in vitro or animal studies, the most difficult final challenge in cartilage tissue engineering is utilizing these results and transfer from in vitro and animal models to practical clinic applications.
33
2.3 Cartilage Structure and Guidance
2.3.1 Contact Guidance
Contact guidance is generally described as the response of cells to physical features of their environment. The response can be in terms of replication, migration, alignment,
ECM formation, or any other cell function (Hoffman-Kim, Mitchel et al. 2010). The concept of contact guidance were first been introduced to study the orientation of neurons and nerve fibers (Weiss 1934; Weiss 1945). Since then, it has been extensively used for nervous system studies [Figure 2-9] (Sorensen, Alekseeva et al. 2007). Since then, a variety of different types of cells have been demonstrated aligned using the strategy of contact guidance (Wang, Jia et al. 2003; Manwaring, Walsh et al. 2004; Hwang, Park et al. 2009; Aubin, Nichol et al. 2010; Huang, Lee et al. 2010; Kapoor, Caporali et al. 2010;
Roach, Parker et al. 2010). While controlling the interactions between cells and scaffold is plays a crucial role in many tissue engineering applications, most of these studies utilize the topography of the substrate to regulate the cell morphologies and further control the specific mechanical, physiological functions of the cells. Initial studies are mostly performed with rigid substrate such as quartz, glass, silicon wafer (Dunn and
Brown 1986; Hirono, Torimitsu et al. 1988; Meyle, Gultig et al. 1995). With the advances in microfabrication and surface modification techniques, substrate options for contact guidance extended to various natural and synthesis polymers (Alaerts, De Cupere et al. 2001). Sørensen et al. employed microgroove of polymer ε-polycaprolactone to control orientation of neuronal cells. Richard et al. used contact guidance to investigate the migration of fibroblasts in oriented collagen gels (Dickinson, Guido et al. 1994).
34
Figure 2-9. Contact guidance of neurons on polymer ε-polycaprolactone (PCL) microgrooves of different dimensions. [Figure from (Sorensen, Alekseeva et al. 2007)]
While the effect of “contact guidance” is mostly seen on cell adhesion and migration
(Dickinson, Guido et al. 1994), some studies suggests that textured surfaces guide the aligment of the cells and further produced aligned ECM proteins (Manwaring, Walsh et al. 2004). Manwaring et al. proposed alignment of meningeal cell align ECM proteins
(Manwaring, Walsh et al. 2004). Wang et al. found the orientation of MC3T3-E1 cells determines the alignment of cell-produced collagenous matrix (Wang, Jia et al. 2003).
Via micropatterneing myoblast on polydimethylsiloxane substrate, Huanget al. generated aligned tissue-like muscle constructs (Huang, Lee et al. 2010). These examples suggest that through contact, the alignments of the cell and cell-produced ECM are able to be controlled and may further regelate the specific mechanical, physiological functions of the TE constructs.
35
2.3.2 Biomechanics of Articular Cartilage
Most tissues found in the body are structured at the microscopic level. Hyaline cartilage, however has been known to have high degree of structural anisotropy [Figure 2-7]
(Hunziker, Michel et al. 1997). The bulk of the cartilage mass is made up of intercellular substance, and it is this component which is largely responsible for the special mechanical properties (Mankin and Thrasher 1975; Maroudas 1979). The insoluble collagenous components in the inter-territorial matrix are further organized into the classical, highly anisotropic, arcade-like architectural arrangement [Figure 2-7, right].
Collagen fibrils about 70% of the tissue dry weight (Rieppo, Rieppo et al.) and mainly made of collagen type II. The fibrils are oriented parallel to the articular surface in the superficial zone and perpendicular to the joint surface in deep zone. This organization is indispensable for the biomechanical functions of cartilage (Lai, Hou et al. 1991; Lu and
Mow 2008) and is the result of chondrocyte activity. Mathematical models suggest that, within native articular cartilage, the well-organized depth dependent structure and composition of the ECM [Figure 2-10 ] provides the unique depth-dependent mechanical strength necessary for everyday load bearing properties (Kempson, Freeman et al. 1968;
Schinagl, Gurskis et al. 1997; Chen, Bae et al. 2001) (Schinagl, Gurskis et al. 1997).
Previous studies showed the composition and structure of articular cartilage vary with the depth of the tissue. As a consequence, biomechanical parameters of Adult human hyaline cartilage have been reported to be strongly depth dependent with a compressive elasticity modulus that varies from 1.16±0.20 MPa in superficial zone to 7.75±1.45 MPa in deep zone [Figure 2-9]. (Chen, Bae et al. 2001)
36
Figure 2-10. Native hyaline cartilage ultrastructure (on the left) [Figure from (Tat, Pelletier et al. 2009) ] and schematic of ultrastructure of articular cartilage adapted from Hunziker et al.(on the right) [Figure from (Hunziker, Michel et al. 1997)]: Blue circular shapes are chondrocytes and the black bands are collagen fibrils.
Figure 2-11. Depth-dependent compressive properties of human Articular Cartilage. Variation of compressive modulus depth from articular surface [Figure from (Chen, Bae et al. 2001)]
37
2.3.3 Structure and Guidance
Based on the arcade model of Benninghof (Benninghoff 1925), Wilson et al. developed a poroviscoelastic fibril-reinforced FEA model to study collagen network behavior. With this model, he suggest that local stresses and strains in the articular cartilage are highly influenced by the local morphology of the collagen fibril network (Wilson, van
Donkelaar et al. 2005; Wilson, van Donkelaar et al. 2005) which suggest that structure leads to improve mechanical properties. The increase in mechanical strength from the surface of the cartilage to the deep zone is attributed to the ECM alignment (Wilson,
Huyghe et al. 2007; Shirazi, Shirazi-Adl et al. 2008). Despite numerous studies investigate cell adhesion and migration via contact guidance, the effects of contact guidance on stem cell differentiation and chondrogenesis are poorly understood. To our knowledge, no study has shown the effect of contact guidance on human MSC (hMSC)- based chondrogenesis; specifically, the effects of contact guidance on the synthesized type II collagen and on the mechanical properties of the resulting constructs are not known. There is evidence that mechanisms at various levels of scales result in structural organization of tissue. For example, Kapoor et al. showed that microtopography affects cell density and alignment of tenocytes microtopographical features and leads to the deposition of an aligned collagen matrix (Kapoor, Caporali et al. 2010). Wang et al. found the orientation of MC3T3-E1 cells determines the alignment of cell-produced collagenous matrix (Wang, Jia et al. 2003). However, there is very little information on guidance mechanisms in cartilage tissue engineering, especially MSC-based cartilage TE.
In this process, we typically begin with an extremely cellular construct; as the cells differentiate, progressively more ECM is deposited. Once deposited, the turnover of this
38
ECM is minimal; it is thus clear that, if we are going to influence the local architecture of the nascent ECM by external cues, these cues will have to be present from the initiation of differentiation. We are not suggesting that it is possible to achieve the level of structural complexity present in natural hyaline cartilage in a single step.
Embryologically, the cells progress from condensation to the differentiation of chondrocytes in parallel with the replacement of an initial matrix of fibronectin and versican by type II collagen and aggrecan. However, although there are excellent studies describing the final architecture of normal cartilage (Hunziker, Michel et al. 1997;
Bhosale and Richardson 2008), there is little information on how the tissue arrives at this final organization. Furthermore, while it may be scientifically interesting to understand how native cartilage arrives at its adult architecture, this may or may not be useful on a practical level. For clinically applicable TE, the outer bounds on the time available to create an implant will be short; months at best, but certainly not years. Thus, influencing tissue development on an accelerated time-scale is important. There is evidence that substrate features can be used to guide extra-cellular matrix (ECM) synthesis into oriented structures. Therefore, the goal of this project is to design external templates for cartilage TE, which will direct differentiating cells to create oriented structures with features that emulate those found in native cartilage
2.4 References
Alaerts, J. A., V. M. De Cupere, et al. (2001). "Surface characterization of poly(methyl methacrylate) microgrooved for contact guidance of mammalian cells." Biomaterials 22(12): 1635-1642.
Almarza, A. J. and K. A. Athanasiou (2004). "Design characteristics for the tissue engineering of cartilaginous tissues." Annals of Biomedical Engineering 32(1): 2-17.
39
Archer, C. W., S. Redman, et al. (2006). "Enhancing tissue integration in cartilage repair procedures." Journal of Anatomy 209(4): 481-493.
Aubin, H., J. W. Nichol, et al. (2010). "Directed 3D cell alignment and elongation in microengineered hydrogels." Biomaterials 31(27): 6941-6951.
Awad, H. A., M. Q. Wickham, et al. (2004). "Chondrogenic differentiation of adipose- derived adult stem cells in agarose, alginate, and gelatin scaffolds." Biomaterials 25(16): 3211-3222.
Banu, N., Y. Banu, et al. (2005). "Biodegradable polymers in chondrogenesis of human articular chondrocytes." Journal of artificial organs : the official journal of the Japanese Society for Artificial Organs 8(3): 184-91.
Baskaran, H. S., L.A.; Berilla, J.; and Welter, J.F (2004). Assessment of mass transport limitations during cartilage tissue engineering. Transactions of the Orthopæ dic Research Society. 29(2): 721.
Baskaran, H. S., L.A.; Penick, K.; Berilla, J.; and Welter, J.F (2003). Substrate mass transport in tissue engineered cartilage. Proceedings of the Arthritis Research Conference. Keystone, CO: 70.
Benninghoff, A. (1925). "Form un bau der gelenkknorpel in ihren beziehungen zur funktion." Zeitschrift für Zellforschung und Mikroskopische Anatomie 2(5): 783-862.
Bentley, G., L. C. Biant, et al. (2003). "A prospective, randomised comparison of autologous chondrocyte implantation versus mosaicplasty for osteochondral defects in the knee." Journal of Bone and Joint Surgery-British Volume 85B(2): 223-230.
Bhosale, A. M. and J. B. Richardson (2008). "Articular cartilage: structure, injuries and review of management." British Medical Bulletin 87(1): 77-95.
Bian, L., D. Y. Zhai, et al. (2011). "Enhanced MSC chondrogenesis following delivery of TGF-beta 3 from alginate microspheres within hyaluronic acid hydrogels in vitro and in vivo." Biomaterials 32(27): 6425-6434.
Brittberg, M., A. Lindahl, et al. (1994). " Treatment of deep cartilage defects in the knee with autologous chondrocyte transplantation." New England Journal of Medicine 331(14): 889-895.
Brittberg M, P. L., Sjogren-Jansson E, Tallheden T, Lindahl A (2003). "Articular cartilage engineering with autologous chondrocytes transplantation. A review of recent developments." J Bone Joint Surg Am 85-A (3): 109–115.
Buckwalter, J. A. and H. J. Mankin (1998). "Articular cartilage: tissue design and chondrocyte-matrix interactions." Instructional course lectures 47: 477-86.
40
Buckwalter, J. A., L. C. Rosenberg, et al. (1990). Articular cartilage composition, structure, response to injury, and methods of facilitating repair. Articular Cartilage and Knee Joint Function: Basic Science and Arthroscopy: 19-56.
Bullough, P. G. (1981). "The geometry of diarthrodial joints, its physiologic maintenance, and the possible significance of age-related-changes in geometry-to-load distribution and the development of osteo-arthritis." Clinical Orthopaedics and Related Research(156): 61-66.
ButnariuEphrat, M., D. Robinson, et al. (1996). "Resurfacing of goat articular cartilage by chondrocytes derived from bone marrow." Clinical orthopaedics and related research(330): 234-243.
Cahill (1995). "Osteochondritis dissecans of the knee: treatment of juvenile and adult forms." J Am Acad Orthop Surg 3(4): 237-247.
Cancedda, R., B. Dozin, et al. (2003). "Tissue engineering and cell therapy of cartilage and bone." Matrix Biology 22(1): 81-91.
Chanlalit, C., C. Kasemkijwattana, et al. (2007). "Autologous chondrocyte implantation for traumatic large cartilage defect." Journal of the Medical Association of Thailand 90(7): 1435-1442.
Chapekar, M. S. (2000). "Tissue engineering: Challenges and opportunities." Journal of Biomedical Materials Research 53(6): 617-620.
Charriere, G., M. Bejot, et al. (1989). "Reactions to a bovine collagen implant - clinical and immunological study in 705 patients." Journal of the American Academy of Dermatology 21(6): 1203-1208.
Chen, A. C., W. C. Bae, et al. (2001). "Depth- and strain-dependent mechanical and electromechanical properties of full-thickness bovine articular cartilage in confined compression." Journal of Biomechanics 34(1): 1-12.
Chen, F. H., K. T. Rousche, et al. (2006). "Technology Insight: adult stem cells in cartilage regeneration and tissue engineering." Nat Clin Pract Rheum 2(7): 373-382.
Clar, C., E. Cummins, et al. (2005). "Clinical and cost-effectiveness of autologous chondrocyte implantation for cartilage defects in knee joints: systematic review and economic evaluation." Health Technology Assessment 9(47).
Cole, A. G. (2011). "A review of diversity in the evolution and development of cartilage: the search for the origin of the chondrocyte." European Cells & Materials 21: 122-129.
41
Cole, A. G. and B. K. Hall (2004). "Cartilage is a metazoan tissue; integrating data from nonvertebrate sources." Acta Zoologica 85(2): 69-80.
Convery, F. R., W. H. Akeson, et al. (1972). "The repair of large osteochondral defects. An experimental study in horses." Clinical orthopaedics and related research 82: 253-62.
Dickinson, R. B., S. Guido, et al. (1994). "Biased cell-migration of fibroblasts exhibiting contact guidance in oriented collagen gels." Annals of Biomedical Engineering 22(4): 342-356.
Dunn, G. A. and A. F. Brown (1986). "Alignment of fibroblasts on grooved surfaces described by a simple geometric transformation." Journal of Cell Science 83: 313-340.
Eaglstein, W. H., O. M. Alvarez, et al. (1999). "Acute excisional wounds treated with a tissue-engineered skin (Apligraf)." Dermatologic Surgery 25(3): 195-201.
Erickson, I. E., A. H. Huang, et al. (2009). "Macromer density influences mesenchymal stem cell chondrogenesis and maturation in photocrosslinked hyaluronic acid hydrogels." Osteoarthritis and Cartilage 17(12): 1639-1648.
Erickson, I. E., S. R. Kestle, et al. "High mesenchyrnal stem cell seeding densities in hyaluronic acid hydrogels produce engineered cartilage with native tissue properties." Acta Biomaterialia 8(8): 3027-3034.
Erickson, I. E., S. R. Kestle, et al. (2012). "High mesenchyrnal stem cell seeding densities in hyaluronic acid hydrogels produce engineered cartilage with native tissue properties." Acta Biomaterialia 8(8): 3027-3034.
Fedewa, M. M., T. R. Oegema, et al. (1998). "Chondrocytes in culture produce a mechanically functional tissue." Journal of Orthopaedic Research 16(2): 227-236.
Ficat, R. P., C. Ficat, et al. (1979). "Spongialization - new treatment for diseased patellae." Clinical Orthopaedics and Related Research(144): 74-83.
Gemmiti, C. V. and R. E. Guldberg (2006). "Fluid flow increases type II collagen deposition and tensile mechanical properties in bioreactor-grown tissue-engineered cartilage." Tissue Engineering 12(3): 469-479.
Goldring, M. B. (2000). "The role of the chondrocyte in osteoarthritis." Arthritis and Rheumatism 43(9): 1916-1926.
Gooding, C. R., W. Bartlett, et al. (2006). "A prospective, ranomised study comparing two techniques of autologous chondrocyte implantation for osteochondral defects in the knee: Periosteum covered versus type I/III collagen covered." Knee 13(3): 203-210.
42
Gratz, K. R., V. W. Wong, et al. (2006). "Biomechanical assessment of tissue retrieved after in vivo cartilage defect repair: tensile modulus of repair tissue and integration with host cartilage." Journal of Biomechanics 39(1): 138-146.
Hangody, L., P. Feczko, et al. (2001). "Mosaicplasty for the treatment of articular defects of the knee and ankle." Clinical Orthopaedics and Related Research(391): S328-S336.
Hangody, L., G. Vasarhelyi, et al. (2008). "Autologous osteochondral grafting-technique and long-term results." Injury-International Journal of the Care of the Injured 39: S32- S39.
Hardingham, T. E. and A. J. Fosang (1992). "Proteoglycans - many forms and many functions." Faseb Journal 6(3): 861-870.
Harley, B. A., A. K. Lynn, et al. (2010). "Design of a multiphase osteochondral scaffold. II. Fabrication of a mineralized collagen-glycosaminoglycan scaffold." Journal of Biomedical Materials Research Part A 92A(3): 1066-1077.
Helmick, C. G., D. T. Felson, et al. (2008). "Estimates of the prevalence of arthritis and other rheumatic conditions in the United States." Arthritis and Rheumatism 58(1): 15-25.
Heng, B. C., T. Cao, et al. (2004). "Directing stem cell differentiation into the chondrogenic lineage in vitro." Stem Cells 22(7): 1152-1167.
Hice, G., D. Freedman, et al. (1990). "Scanning and light microscopic study of irrigated and nonirrigated joints following burr surgery performed through a small incision." Journal of Foot Surgery 29(4): 337-344.
Hirono, T., K. Torimitsu, et al. (1988). "Recognition of artificial microstructures by sensory nerve-fibers in culture." Brain Research 446(1): 189-194.
Hoffman-Kim, D., J. A. Mitchel, et al. (2010). Topography, Cell Response, and Nerve Regeneration. Annual Review of Biomedical Engineering, Vol 12. 12: 203-231.
Huang, A. H., B. M. Baker, et al. (2012). "Sliding contact loading enhances the tensile properties of mesenchymal stem cell-seeded hydrogels." European Cells & Materials 24: 29-45.
Huang, A. H., M. J. Farrell, et al. (2011). "Mechanics and mechanobiology of mesenchymal stem cell-based engineered cartilage." Journal of Biomechanics 43(1): 128- 136.
Huang, C. Y., A. Stankiewicz, et al. (2005). "Anisotropy, inhomogeneity, and tension- compression nonlinearity of human glenohumeral cartilage in finite deformation." Journal of Biomechanics 38(4): 799-809.
43
Huang, N. F., R. J. Lee, et al. (2010). "Engineering of aligned skeletal muscle by micropatterning." American journal of translational research 2(1): 43-55.
Hunziker, E. B. (1999). "Articular cartilage repair: are the intrinsic biological constraints undermining this process insuperable?" Osteoarthritis and Cartilage 7(1): 15-28.
Hunziker, E. B. (2002). "Articular cartilage repair: basic science and clinical progress. A review of the current status and prospects." Osteoarthritis and Cartilage 10(6): 432-463.
Hunziker, E. B., M. Michel, et al. (1997). "Ultrastructure of adult human articular cartilage matrix after cryotechnical processing." Microsc Res Tech 37(4): 271-84.
Hwang, C. M., Y. Park, et al. (2009). "Controlled cellular orientation on PLGA microfibers with defined diameters." Biomedical Microdevices 11(4): 739-746.
Insall, J. N. (1967). "Intra-articular surgery for degenerative arthritis of the knee. A report of the work of the late K. H. Pridie." The Journal of bone and joint surgery. British volume 49(2): 211-28.
Jackson, D. W., P. A. Lalor, et al. (2001). "Spontaneous repair of full-thickness defects of articular cartilage in a goat model - A preliminary study." Journal of Bone and Joint Surgery-American Volume 83A(1): 53-64.
Jackson, R. W. and C. Dietrichs (2003). "The results of arthroscopic lavage and debridement of osteoarthritic knees based on the severity of degeneration: A 4-to 6-year symptomatic follow-up." Arthroscopy-the Journal of Arthroscopic and Related Surgery 19(1): 13-20.
Jason R. Fritz, D. P. a. H. S. C. (2009). "Current challenges in cartilage tissue engineering: a review of current cellular-based therapies." Current Rheumatology Reviews 5(1): 8-14 (7)
Johnstone, B. and J. U. Yoo (1999). "Autologous mesenchymal progenitor cells in articular cartilage repair." Clinical Orthopaedics and Related Research(367): S156-S162.
Kapoor, A., E. H. G. Caporali, et al. (2010). "Microtopographically patterned surfaces promote the alignment of tenocytes and extracellular collagen." Acta Biomaterialia 6(7): 2580-2589.
Kempson, G. E., M. A. R. Freeman, et al. (1968). "Tensile properties of articular cartilage." Nature 220(5172): 1127-&.
Khan, I. M., S. J. Gilbert, et al. (2008). "Cartilage integration: evaluation of the reasons for failure of integration during cartilage repair. A review." European Cells & Materials 16: 26-39.
44
Kuo, C. K., W. J. Li, et al. (2006). "Cartilage tissue engineering: its potential and uses." Current Opinion in Rheumatology 18(1): 64-73.
Lai, W. M., J. S. Hou, et al. (1991). "A triphasic theory for the swelling and deformation behaviors of articular-cartilage." Journal of Biomechanical Engineering-Transactions of the Asme 113(3): 245-258.
Lane, J. G., J. B. Massie, et al. (2004). "Follow-up of osteochondral plug transfers in a goat model - A 6-month study." American Journal of Sports Medicine 32(6): 1440-1450.
Lee, C. H., A. Singla, et al. (2001). "Biomedical applications of collagen." International Journal of Pharmaceutics 221(1-2): 1-22.
Lee, E. H. and J. H. P. Hui (2006). "The potential of stem cells in orthopaedic surgery." Journal of Bone and Joint Surgery-British Volume 88B(7): 841-851.
Linn, F. C. and L. Sokoloff (1965). "Movement and composition of interstitial fluid of cartilage." Arthritis and Rheumatism 8(4): 481-&.
Lu, X. L. and V. C. Mow (2008). "Biomechanics of articular cartilage and determination of material properties." Medicine and Science in Sports and Exercise 40(2): 193-199.
Mankin, H. J. (1974). "Reaction of articular-cartilage to injury and osteoarthritis .1." New England Journal of Medicine 291(24): 1285-1292.
Mankin, H. J. (1974). "Reaction of articular-cartilage to injury and osteoarthritis .2." New England Journal of Medicine 291(25): 1335-1340.
Mankin, H. J. and A. Z. Thrasher (1975). "Water-content and binding in normal and osteoarthritic human cartilage." Journal of Bone and Joint Surgery-American Volume A 57(1): 76-80.
Manwaring, M. E., J. F. Walsh, et al. (2004). "Contact guidance induced organization of extracellular matrix." Biomaterials 25(17): 3631-3638.
Maroudas, A. (1979). "Physico-chemical properties of articular cartilage." In: Freeman M.A.R., editor. Adult Articular Cartilage. Tunbridge Wells, England: Pitman Medical: 215-290.
McLaren, A. C., C. P. Blokker, et al. (1991). "Arthroscopic debridement of the knee for osteoarthrosis." Canadian Journal of Surgery 34(6): 595-598.
Meyle, J., K. Gultig, et al. (1995). "Variation in contact guidance by human-cells on a microstructured surface (vol 29, PG 81, 1995)." Journal of Biomedical Materials Research 29(7): 905-905.
45
Michaud, C. M., M. T. McKenna, et al. (2006). "The burden of disease and injury in the United States 1996." Population health metrics 4: 11-11.
Minas, T. (2001). "Autologous chondrocyte implantation for focal chondral defects of the knee." Clinical Orthopaedics and Related Research(391): S349-S361.
Nayyer, L., K. H. Patel, et al. "Tissue Engineering: Revolution and Challenge in Auricular Cartilage Reconstruction." Plastic and Reconstructive Surgery 129(5): 1123- 1137.
O'Driscoll, S. W. (1998). "The healing and regeneration of articular cartilage." Journal of Bone and Joint Surgery-American Volume 80A(12): 1795-1812.
P.K. Bos, M. L. v. M., G.J.v. Osch (2010). "Articular cartilage repair and the evolving role of regenerative medicine " Open Access Surgery 3(1): 109-122.
Parenteau-Bareil, R., R. Gauvin, et al. (2010). "Collagen-based biomaterials for tissue engineering applications." Materials 3(3): 1863-1887.
Pelaez, D., C.-Y. C. Huang, et al. (2009). "Cyclic compression maintains viability and induces chondrogenesis of human mesenchymal stem cells in fibrin gel scaffolds." Stem Cells and Development 18(1): 93-102.
Peterson, L., T. Minas, et al. (2000). "Two- to 9-year outcome after autologous chondrocyte transplantation of the knee." Clinical Orthopaedics and Related Research(374): 212-234.
Pfander, D. and K. Gelse (2007). "Hypoxia and osteoarthritis: how chondrocytes survive hypoxic environments." Current Opinion in Rheumatology 19(5): 457-462.
Pittenger, M. F., A. M. Mackay, et al. (1999). "Multilineage potential of adult human mesenchymal stem cells." Science 284(5411): 143-147.
Ponticiello, M. S., R. M. Schinagl, et al. (2000). "Gelatin-based resorbable sponge as a carrier matrix for human mesenchymal stem cells in cartilage regeneration therapy." Journal of Biomedical Materials Research 52(2): 246-255.
Redman, S. N., S. F. Oldfield, et al. (2005). "Current strategies for articular cartilage repair." European Cells & Materials 9: 23-32; discussion 23-32.
Rieppo, L., J. Rieppo, et al. (2012). Fourier transform infrared spectroscopic imaging and multivariate regression for prediction of proteoglycan content of articular cartilage. PLoS One. 7 (2):e32344
Risbud, M. V. and M. Sittinger (2002). "Tissue engineering: advances in in vitro cartilage generation." Trends in Biotechnology 20(8): 351-356.
46
Roach, P., T. Parker, et al. (2010). "Surface strategies for control of neuronal cell adhesion: A review." Surface Science Reports 65(6): 145-173.
Schinagl, R. M., D. Gurskis, et al. (1997). "Depth-dependent confined compression modulus of full-thickness bovine articular cartilage." Journal of Orthopaedic Research 15(4): 499-506.
Schmidt, C. E. and J. M. Baier (2000). "Acellular vascular tissues: natural biomaterials for tissue repair and tissue engineering." Biomaterials 21(22): 2215-2231.
Schmidt, H., K. J. Schulze, et al. (1988). "Results in the treatment of cartilage lesions by drilling according to the pridie method." Beitraege zur Orthopaedie und Traumatologie 35(3): 117-122.
Schnabel, M., S. Marlovits, et al. (2002). "Dedifferentiation-associated changes in morphology and gene expression in primary human articular chondrocytes in cell culture." Osteoarthritis and Cartilage 10(1): 62-70.
Setton, L. (2008). "Polymer therapeutics - Reservoir drugs." Nature Materials 7(3): 172- 174.
Shapiro, F., S. Koide, et al. (1993). "Cell origin and differentiation in the repair of full- thickness defects of articular-cartilage." Journal of Bone and Joint Surgery-American Volume 75A(4): 532-553.
Sharma, C. G., Sneh; Dinda, Amit K.; Mishra, Narayan C (2011). "Cartilage tissue engineering: current scenario and challenges." Advanced Materials Letters 2(2): 90.
Shirazi, R., A. Shirazi-Adl, et al. (2008). "Role of cartilage collagen fibrils networks in knee joint biomechanics under compression." J Biomech 41(16): 3340-8.
Simon, L. S. (1999). "Osteoarthritis: a review." Clinical cornerstone 2(2): 26-37.
Smith, G. D., J. B. Richardson, et al. (2003). "Autologous chondrocyte implantation and osteochondral cylinder transplantation in cartilage repair of the knee joint." Journal of Bone and Joint Surgery-American Volume 85A(12): 2487-2488.
Solchaga L, T. E., Goldberg V, Caplan A, Welter J (2003). Tissue engineered cartilage implants: in vitro chondrogenic preconditioning. Proceedings of the 4th Symposium of the International Cartilage Repair Society 271.
Solchaga, L. A. (2002). Tissue engineered cartilage implants: optimizing construct assembly. Transactions of the Orthopaedic Research Society. 27: 483.
47
Solchaga, L. A. (2004). FGF-2 modifies the gene expression profile of bone marrow- derived human mesenchymal stem cells. Transactions of the Orthopaedic Research Society. 29: 777.
Song, L., D. Baksh, et al. (2004). "Mesenchymal stem cell-based cartilage tissue engineering: cells, scaffold and biology." Cytotherapy 6(6): 596-601.
Sorensen, A., T. Alekseeva, et al. (2007). "Long-term neurite orientation on astrocyte monolayers. aligned by microtopography." Biomaterials 28(36): 5498-5508.
Speer, D. P., M. Chvapil, et al. (1979). "Enhancement of healing in osteochondral defects by collagen sponge implants." Clinical Orthopaedics and Related Research(144): 326- 335.
Steadman, J. R., K. K. Briggs, et al. (2003). "Outcomes of microfracture for traumatic chondral defects of the knee: Average 11-year follow-up." Arthroscopy: The Journal of Arthroscopic & Related Surgery 19(5): 477-484.
Stockwell, R. A. (1979). Biological structure and function vol. 7 biology of cartilage cells.
Tat, S. K., J.-P. Pelletier, et al. (2009). "Treatment with ephrin B2 positively impacts the abnormal metabolism of human osteoarthritic chondrocytes." Arthritis Research & Therapy 11(4).
Temenoff, J. S. and A. G. Mikos (2000). "Review: tissue engineering for regeneration of articular cartilage." Biomaterials 21(5): 431-440.
Thomson, J. A., J. Itskovitz-Eldor, et al. (1998). "Embryonic stem cell lines derived from human blastocysts." Science 282(5391): 1145-1147.
Tognana, E., A. Borrione, et al. (2007). "Hyalograft (R) C: Hyaluronan-based scaffolds in tissue-engineered cartilage." Cells Tissues Organs 186(2): 97-103.
Toh, W. S., E. H. Lee, et al. (2011). "Potential of human embryonic stem cells in cartilage tissue engineering and regenerative medicine." Stem Cell Reviews and Reports 7(3): 544-559.
Tuan, R. S., G. Boland, et al. (2003). "Adult mesenchymal stem cells and cell-based tissue engineering." Arthritis Research & Therapy 5(1): 32-45.
Wakitani, S., T. Goto, et al. (1994). "Mesenchymal cell-based repair of large, full- thickness defects of articular-cartilage." Journal of Bone and Joint Surgery-American Volume 76A(4): 579-592.
Wang, J. H. C., F. Y. Jia, et al. (2003). "Cell orientation determines the alignment of cell- produced collagenous matrix." Journal of Biomechanics 36(1): 97-102.
48
Wei, Y., W. Zeng, et al. (2012). "Chondrogenic differentiation of induced pluripotent stem cells from osteoarthritic chondrocytes in alginate matrix." European Cells & Materials 23: 1-12.
Weiss, P. (1934). "In vitro experiments on the factors determining the course of the outgrowing nerve fiber." Journal of Experimental Zoology 68(3): 393-448.
Weiss, P. (1945). "Experiments on cell and axon orientation in vitro; the role of colloidal exudates in tissue organization." The Journal of experimental zoology 100: 353-86.
Welter, J. (2002). Tissue engineered cartilage implants: In vitro chondrogenic preconditioning. Proceedings of the 2nd Annual workshop of the Midwestern Tiss. Eng. Consortium: 21.
Welter, J. (2003). MSC-based Tissue engineered cartilage: Effects of construct assembly method and perfusion rates. Arthritis Research Conference. Altanta, GA.
Welter, J. (2004). Modulation of growth and differentiation conditions enhances chondrogenesis in large tissue engineered constructs. Transactions of the Orthopaedic Research Society. 29(2): 719.
Welter, J. F., L. A. Solchaga, et al. (2007). "Simplification of aggregate culture of human mesenchymal stem cells as a chondrogenic screening assay." Biotechniques 42(6): 732-+.
Williams, G. M., E. F. Chan, et al. (2010). "Shape, loading, and motion in the bioengineering design, fabrication, and testing of personalized synovial joints." Journal of Biomechanics 43(1): 156-165.
Williams, G. M., T. J. Klein, et al. (2005). "Cell density alters matrix accumulation in two distinct fractions and the mechanical integrity of alginate-chondrocyte constructs." Acta Biomaterialia 1(6): 625-633.
Williams, R. J. and S. C. Gamradt (2008). "Articular cartilage repair using a resorbable matrix scaffold." Instructional course lectures 57: 563-71.
Wilson, W., J. M. Huyghe, et al. (2007). "Depth-dependent compressive equilibrium properties of articular cartilage explained by its composition." Biomechanics and Modeling in Mechanobiology 6(1-2): 43-53.
Wilson, W., C. C. van Donkelaar, et al. (2005). "A fibril-reinforced poroviscoelastic swelling model for articular cartilage." Journal of Biomechanics 38(6): 1195-1204.
Wilson, W., C. C. van Donkelaar, et al. (2005). "Stresses in the local collagen network of articular cartilage: a poroviscoelastic fibril-reinforced finite element study (vol 37, pg 357, 2004)." Journal of Biomechanics 38(10): 2138-2140.
49
Wong, M., P. Wuethrich, et al. (1996). "Zone-specific cell biosynthetic activity in mature bovine articular cartilage: A new method using confocal microscopic stereology and quantitative autoradiography." Journal of Orthopaedic Research 14(3): 424-432.
Wynn, J. J. and C. E. Alexander "Increasing organ donation and transplantation: the US experience over the past decade." Transplant International 24(4): 324-332.
50
Chapter 3
CONTROL OF HUMAN MESENCHYMAL STEM SELECTIVE ADHESION ON COLLAGEN-GLYCOSAMINOGLYCAN AND POLY(DIMETHYL SILOXANE) SURFACES
This work is published in Tissue Engineering, Part A, 2012. (Chou, Rivera et al. 2012)
3.1 Introduction
One of the major challenges for current tissue engineering in vitro is to generate engineered tissues that emulates native tissue structure and function (Patel, Padera et al.
1998; Shin, Jo et al. 2003). To gain control over the engineered tissue’s structure and function, one must investigate methods that allow for regulation over organization, proliferation, and differentiation of the cells. In this study, we utilized the method of contact guidance (See Chapter 2), in which we utilized microscale physical features
(guidance channels) on the biomaterial scaffold surface to control spatial cellular organization, cellular morphology, and cellular function of human mesenchymal stem cells (hMSCs). To control differentiating hMSCs to synthesize ECM that emulates the aligned ECM structure of native tissue, the cells need to be localized within the channels; contact guidance and its efficacy assessment require the cells to be located inside the channels.
Surface engineering has been extensively used as a tool to control and study cellular response and cell-surface interaction (Roach, Parker et al.). Controlling cell-surface
51 interaction is very important in understanding host-biomaterial interaction, investigating cell behavior as well as studying tissue development and tissue organization. Directing cell growth and motility have been reported in tissue engineering applications such as nerve reservation (Lu, Simionescu et al. 2005; Schmalenberg and Uhrich 2005) and endothelialization of microvasculature (Wang and Ho 2004). Many tissue engineering investigations that require precise spatial cellular architectures for function such as nerves and blood vessels tissue have been limited due to the lack of proper technique to pattern the cell to the defined culture surface.
Various surface modification techniques have been developed to pattern all types of cells on different substrates. Some researchers employed passive chemical modification to coat the bioresistant material such as poly(ethylene glycol) (PEG), bovine serum albumin, phospholipid, and polyacrylamide (Kingshott and Griesser 1999), which will impart non- fouling properties to the specific area of cell-adhesive surface which facilitate directing the cell or protein to the defined area. Others used active methods such as surface chemistry to chemically modify the surface via self-assembled mono-layers. For example,
Mrksich et al. employed thiolated PEGs on a gold surface (Mrksich, Dike et al. 1997;
Chen, Mrksich et al. 1998) and Csucs. et al. utilized poly-L-lysine grafted PEG on glass, polystyrene or metal-oxides (Csucs, Michel et al. 2003). These studies demonstrate nanometer precision to control the patterning process. While the above studies utilized patterned surfaces that are in non-biodegradable in nature (glass, gold and polystyrene), it is necessary to achieve similar patterning precision on biodegradable polymer substrate for practical tissue engineering applications.
52
Poly(dimethyl siloxane) PDMS is a non-toxic, biocompatible gas permeable, and optically transparent material amenable to surface treatment and therefore ideally suited for micropatterning via soft lithography (Kane, Takayama et al. 1999). In addition, surface chemistry can be used to obtain cell-adhesive and non-adhesive patterns on
PDMS. Collagen-Glycosaminoglycan (CG) is a natural biodegradable material and has been used extensively as a scaffold material for cartilage tissue engineering. In contrast to most synthetic materials, the surface of natural biodegradable material present cell adhesoion ligands that can stimulate cell attachment and differentiation (Ruoslahti 1996;
Palecek, Loftus et al. 1997; Patel, Padera et al. 1998). CG membranes can be micropatterned to generate guidance channels (Janakiraman, Kienitz et al. 2007)
However, to our knowledge, there have been no methods to pattern adhesive and non- adhesive domains on collagen-based surfaces..
The pluronic family of compounds are commercially available, amphiphilic, non-ionic triblock polymers consisting of hydrophobic poly(propylene oxide) (PPO) centers bounded by two hydrophilic poly(ethyelene oxide) (PEO) tails. Pluronics are widely used in biomedical applications such as drug delivery and as non-fouling chemicals (Marin,
Sun et al. 2002; Wang and Ho 2004). Adeel et al. mixed Pluronic F127 and D-α- tocopheryl polyethylene glycol 1000 succinate to form micelles for anticancer drug delivery (Butt, Amin et al.). Tharmalingam et al. has used Pluronic F-68 to reduce the cell attachment of mammalian cells (Tharmalingam, Ghebeh et al. 2008). The PPO hydrophobic center in Pluronics can attach to an hydrophobic surface while the PEO hydrophilic tails can make the surface exhibit hydrophilic characteristics, and thus prevent the cell and protein attachment. Li et al. studied cell–ligand interactions for NIH
53
3T3 fibroblasts on Pluronic F108 treated polystyrene substrate and reported it as the superior polymer among Pluronic family to resist cell and protein adhesion because it is stable and can absorbed to substrate with highly irregular geometry (Li and Caldwell
1996; Liu, Jastromb et al. 2002).
The goal of this study is to develop a technique that allows for control of hMSC adhesion and spatial organization on collagen–glycosaminoglycan (CG) membrane and PDMS membranes. In this study, we improve the collagen soft lithography technique developed earlier (Janakiraman, Kienitz et al. 2007) to micropattern the CG membrane for use with differentiating MSCs, and develop a new method to selectively adhere MSCs to the defined area. We utilize selective surface modification via Pluronic F108 and fibronectin, an adhesive protein, and microfluidics to control hMSC adhesion on the well-defined topography of micropatterned CG and PDMS membranes. We further demonstrate the potential of thistechnique for use in cartilage tissue engineering.
3.2 Materials and Methods
3.2.1 Materials
Collagen type I from bovine Achilles tendon, chondroitin-6-sulfate sodium salt from shark cartilage, 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDC),
N-hydroxy-succinimide, fetal bovine serum (FBS), 4', 6-diamidino-2-phenylindole
(DAPI), and phalloidin were purchased from Sigma Chemical Co. (St. Louis, MO).
Antibiotic-antimycotic cocktail, low glucose Dulbecco’s Modified Eagle Medium
(DMEM), high glucose DMEM, ITS (insulin, transferrin, selenium), ascorbate 2- phosphate (A2P), dexamethasone, sodium pyruvate, and human plasma fibronectin were
54 obtained from Invitrogen (Carlsbad, CA). Sylgard® silicone elastomer was purchased from Dow Corning Corporation (Midland, MI). Fibroblast growth factor-2 (FGF) and transforming growth factor β1 (TGF- β1) were purchased from Peprotech (Rocky Hill,
NJ). ITS (insulin, transferrin, selenium) + Premix Tissue Culture Supplement were obtained from Becton Dickinson (Franklin Lakes, NJ). Carboxyfluoro diacetate succinimidyl ester (CFDA-SE,Vybrant® ), Pluronic F108 was purchased from BASF
(Whitehouse, OH).
3.2.2 CG Solution and EDC Solution Formation
The base CG solution was made by using a method adopted from Yannas, et al. (Yannas,
Burke et al. 1980). Type I collagen of 2.2 g from bovine Achilles tendon (Sigma, St.
Louis, MO) was added to 800 ml of 0.5% v/v acetic acid solution (Sigma). The solution was then blended for 20 minutes in an ice-bath cooled vessel using an overhead homogenizer (IKA Works, Wilmington, NC) at a speed of 13,500 rpm. Then, 0.22 g
(10% by weight collagen) of chondroitin 6-sulfate from shark cartilage (Sigma), dissolved in 40 ml of 0.5% v/v acetic acid solution, was added drop-wise to the CG solution. The solution was homogenized for an additional 20 minutes at 22,000 rpm. The final concentrations of collagen and chondroitin sulfate in the CG solutions were 2.6 and
0.26 g/liter, respectively. A crosslinking EDC solution was made by dissolving 0.27 g of
EDC and 63 mg of N-hydroxy-succinimide in 100 ml of diH2O.
3.2.3 Design of Microchannels
Templates for rectangular channels of a constant length and varying widths were created using AutoCAD (Autodesk, Inc., San Rafael, CA) [Figure 3-1]. We tested channel widths
55
25, 50, 100, 500 and 1000 μm. The 25 μm width was chosen to represent the size of a single cell, while the 1000 μm width was chosen as the largest dimension to act as a featureless growth surface. The spacing between channels of the same width was varied from 50 μm to 250 μm for different designs to allow testing the effect of channel density in future experiments unrelated to the current investigation.
Figure 3-1. Microchannel design created in AutoCAD. The channels ranged from a minimum width of 25 μm to a maximum width of 1000 µm. The spacing between channels of the same width was varied from 50 to 250 µm for different designs to allow us to test the effect of channel density in future experiments.
3.2.4 Fabrication of Silicon Substrate Template with Microchannels
Prior to patterning the CG and PDMS membranes with microchannel features, silicon substrates with microchannels design were fabricated utilizing SU-8 2075 negative photoresist (PR) (MicroChem Corp., MA, USA) by standard UV photolithography. First, photomasks were obtained from Advance Reproductions Corp., MA. The photomasks were made by printing AutoCAD designs of channels [Figure 3-1] on transparent photofilms in a high resolution (>3000 dpi) laser printer. The plastic photomasks provide low-cost alternatives to traditional chrome masks and have a resolution of 2 μm. Standard
UV lithography was then performed in the Electronics Design Center at CWRU to obtain
56 silicon templates for subsequent soft lithography process. The process for fabricating the negative templates of PR in a silicon substrate is shown in Figure 3-2 and is described in detail below.
(a) Oxide Removal:
In the beginning of microfabrication process, the native oxide layer and contamination in the oxide on the silicon substrate (University Wafers) surface was removed by dipping the wafer in buffered oxide etch (BOE) (MicroChem Corp., MA, USA) for 2 minutes.
The wafer is then rinsed in diH2O for 10 minutes.
(b) SU-8 photoresist spin coating:
A uniformly thick film of PR was coated on the silicon substrate (University Wafer) using a spin coater (Laurell Technologies Co, PA). The thickness of the PR film is controlled by the spin speed and spin conditions which needed to be optimized to achieve the desired thickness. PR was dispensed carefully onto the substrate with volume ratio of
4 mL for 4 inch silicon wafer. We determined the optimal conditions for spin coating to generate PR films of thickness: 85 μm. They lead to four steps in the spinning cycle, including spinning at 500 rpm for 10 s with acceleration of 100 rpm/s to spread PR on the substrate, and spinning at 1000 rpm for 30s with acceleration of 200 rpm/s to obtain the desired thickness.
(c) Soft bake: To evaporate the PR solvent and to achieve PR self-planarization after spin coating, soft baking was performed by baking on a level hotplate. Before the baking process, the hotplate was carefully adjusted to a horizontal position since gravity will affect the flatness of the PR film. Baking time and conditions were determined based on
57 the PR thickness. For 85 μm PR film, the temperature was held at 75oC for 3 min and then ramped gradually to 95oC and held for 20 min. The substrate was then cooled down to room temperature gradually by leaving the substrates on the hotplate for 1 h after turning off the hotplate.
(d) UV lithography: To obtain vertical sidewalls, UV radiation with wavelength below
350 nm was eliminated using a long pass filter from Omega Optical, VT. The UV lithography was processed using a mask aligner (ABM, Inc., CA) with exposure intensity of 15 mW/cm2 for 35 s. With optimal exposure, a visible latent image showed up in the film within 5-15 s after placing on the hotplate for post-exposure bake (PEB).
(e) Post Exposure Bake (PEB): After exposure, PEB was carried out directly using a level hotplate to selectively cross-link the exposed portions of the film. During PEB, PR cross-linking can result in a highly stressed film. Thus, a two-step heating process followed by a slow cooling step was used to minimize stress. For 85 μm PR film, the
PEB temperature was held at 75oC for 3 min and then ramped gradually to 95oC and held for 10 min. The substrates were then cooled down gradually to room temperature by leaving the substrates on the hotplate for 1 h after turning off hotplate.
(f) Development: The development process was carried out by immersing substrates in
SU8 developer (MicroChem Corp., MA, USA) with a gentle agitation initially for 12 min followed by inspection to determine if it was sufficient development. The uncross-linked
PR was dissolved during development. Development was continued for an additional 5 min using a fresh batch of developer if white residue of photoresist was present. This step eliminated the uncrosslinked photoresist.
58
(g) Post-Develop Hard Bake:
The final hard baking step was performed by heating the wafers to 150 oC for 5 min on a hot plate. This process ensured that the photoresist patterns did not alter during continued usage.
Figure 3-2. Schematic of standard UV light lithography
3.2.5 Fabrication of Collagen and PDMS Microchannels
CG membranes were produced using a previously published filtration method
(Janakiraman, Kienitz et al. 2007). The CG solution is poured onto a glass filter attached to a vacuum flask. Care is taken to level the filter so that a membrane of uniform thickness can be obtained.
The silicon wafers containing the patterns in Figure 3-2 were then used to produce channels on the surface of the CG membranes by collagen soft-lithography that involved selective solubilization, patterning, and pattern stabilization through EDC crosslinking
59
[Figure 3-3]. Prior to patterning, silicon wafer templates with the microchannels were diced to 2.5x 6 cm. A CG membrane of size 2.5 cm (width) x 6 cm (length) x 0.5 cm
(depth) was placed in a 150 mm Petri-dish (BD, Franklin Lakes, NJ). Aacetic acid solution 0.5 ml of 0.5% was then spread on top of the membrane to initial selective dissolution. A diced silicon template with microchannels was placed on top of the membrane with pattern side down and in contact with the membrane. A glass slide was placed on top of silicon wafer followed by an aluminum bar (1 x 3x 2 inch) to ensure good contact between the template and the membrane. Freshly-made EDC solution (Lee et al. 2001) was subsequently added to submerge the scaffolds, to stabilize and crosslink the dissolved collagen network around the PR features. After 48 h, CG scaffolds were removed, cross-linked with fresh EDC solution for an additional 24 h. Excess EDC was rinsed off using PBS. For cell culture, the membranes were incubated in 10x antibiotic- antimycotic cocktail for one day and 1x antibiotic-antimycotic cocktail for another day, and then washed thoroughly in sterile PBS. The CG scaffolds were stored in PBS at 4°C until use.
To verify the channel width, phase contrast images of the CG microchannels were were taken with an Olympus IX71 inverted microscope (Olympus, Japan) using UPlanFl 10X and LCPlanFl 20X objectives equipped with a digital camera (SPOT-RT, Diagnostic
Instruments Sterling Heights, MI). The channel dimensions were obtained through image analysis using Image-Pro® Plus 6.2 software (Media Cybernetics, Rockville, MD).
Collagen microchannels were also investigated using scanning electron microscopy
(SEM) (Quanta 200 3D, FEI, Hillsboro, Oregon). The CG membrane were dehydrated in a series of ethanol-water solutions with the increasing ethanol concentration of 30%, 50%,
60
70%, 80%, 90%, 95% and 100%, for at least 1 hour in each solution. The 100% fresh ethanol was changed twice to ensure all water in the samples was exchanged with ethanol.
Samples were then dried using hexamethyldisilazane (HMDS) (Nation et al. 1983). After dehydration, the samples were immersed in HMDS for 5 min at room temperature, and air dried in a fume hood. Dried samples were Pd-sputter coated prior to SEM imaging.
Figure 3-3. Schematic of Collagen soft lithography method: Acetic acid was used to dissolve the surface of the CG membrane. A weight was placed on the silicon wafer containing the channels to pattern the collagen. EDC crosslinking solution was then used to stabilize the channels.
61
We used soft lithography to obtain microchannels in PDMS (Xia and Whiteside, 1998).
In this process, Polydimethylsiloxane (PDMS) membranes were obtained using a standard combination of photolithography and soft lithography techniques. Using the photolithography method describedin section3.2.4 [Figure3-4], we obtained a silicon wafer template containing negative design features of the microchannels. To prevent
PDMS binding to the photoresist, the wafer was exposed to (tridecafluoro-1,1,2,2- tetrahydrooctyl)-1-trichlorosilane for 30 min at room temperature under vacuum. The
Sylgard® elastomer (Sylgard 184, Dow corning, Midland, MI) base was mixed 10:1
(w/w) with Sylgard® silicone curing agent. This mixture was poured over the patterned wafer, degassed under vacuum and cured at 80 °C overnight. After cooling, the membranes were carefully peeled off from the wafer surface. A schematic of this process is shown here [Figure 3-4].
Figure 3-4. Schematic of standard Soft Lithography
62
3.2.6 Selective Cell Attachment in Collagen Microchannels
To attach cells selectively to the channels, the plateaus on the surface of the patterned collagen surface (spaces between the channels) were selectively modified with Pluronic
F108, ((PEO)129-(PPO)56-(PEO)129, molecular weight 14600 g/mol) (BASF, Whitehouse,
OH). Figure 3-5 shows the schematic of the mechanism of action of F108; the PPO chains of Pluronic F108 attached to hydrophobic surface and the free PEO chains converted the surface to hydrophilic in nature. A solution of 10 mg/ml Pluronic F108 in diH2O was prepared. 2.5x 6 cm cover slides were coatedwith F108 by incubating them in the solution in a 100 mm Petri-dishe for 3 hours and air dried in hood for overnight.
Patterned CG membranes were placed in a fresh 100 mm Petri-dishe and the F108 coated glass slide were placed on top on the CG membrane. A metal weight was immediately placed on the glass slide to ensure good contact between the slide and the membrane. By contacting the top surface of the patterned CG membranes with F108 coated glass slides for 3 hours, F108was transferred to the plateaus of the CG membrane while the interior of the collagen channels remained uncoated [Figure 3-6]. hMSCs were stained with Vybrant
CFDA SE dye (excitation/emission wavelengths of 492/517 nm) at a concentration 10
μM. Vybrant-stained hMSCs and 2 million cells were seeded per the membrane and were allowed to attach for two hours, after which the membranes were washed with 1× PBS 3 times to remove unattached cells. Fluorescent images of the cells and the collagen microchannels were taken.
63
Figure 3-5. Schematic of Pluronic mechanism. Black chains are PPO chain and yellow chains are PEO chains. The PPO chains of Pluronic F108 will absorb to hydrophobic surface and the surface will be modified with PEO(yellow chain) chains and exhibit hydrophilic
Figure 3-6. A schematic of the technique used to obtain selective seeding of mesenchymal stem cells (MSCs). The steps are uniform coating of F108 on a glass
64 coverslip, selective transfer of F108 onto nonchannel surfaces through contact with F108 coated glass, and selective seeding of MSCs in channels
3.2.7 Selective Cell Attachment in PDMS Microchannels
To confer hydrophobicity, PDMS membranes with microchannels were exposed to an oxygen plasma using an SPI Plasma Prep™ II plasma etcher for 30 seconds and sterilized in 200-proof ethanol overnight. The substrates were then transferred to a 60 mm Petri- dishes with the microchannels facing down and allowed to adhere to the dish surface. A fibronectin (10 μg/ml) in PBS solution was added to the Petri dish, and the dish placed under vacuum for 20 minutes to allow infiltration of the fibronectin solution into the channels. The fibronectin solution was then removed and the channels air dried in the hood for 3 hours. Then the plateaus on the patterned PDMS membrane surface were selectively modified with F108 as described above for the CG membranes [Figure 3-7].
The modified PDMS membranes were kept in 4°C until use.
65
Figure 3-7. Schematic of Selective Cell Attachment for PDMS membrane.
3.2.8 Cell Culture
Human MSCs were prepared from bone marrow aspirates of healthy volunteer donors through the Stem Cell Core Facility of the Case Comprehensive Cancer Center as described previously. The aspirates were harvested after informed consent was obtained under the terms of an IRB-approved protocol. The isolation and purification of MSCs were carried out by trained personnel at the Skeletal Research Center (SRC) at the Case
Western Reserve University. In this study, cells from three individual donors were used.
Adult human MSCs obtained from the SRC (typically in frozen condition) were cultured to confluency in Dulbecco’s Modified Eagle’s Medium, Low (1.5 g/l) Glucose (DMEM-
LG) containing 10% FBS and 10 ng/ml FGF-2. The MSCs were then trypsinized and
66 resuspended at concentrations of approximately 50 × 106 cells/ml. The cell suspension was applied to the channels (2 million cell per construct) using a micropipette. After a 2- hour period to allow for cell attachment at 37°C and 5% CO2, the seeded membranes were submerged in chondrogenic medium containing TFG-β1 10 ng/ml. The constructs were cultured in a standard incubator at 37 °C and 5% CO2 in air for 21 days. Medium was changed every 2-3 days.
Human MSCs were prepared as previously described from bone marrow aspirates obtained from healthy volunteer donors through the Stem Cell Core Facility of the Case
Comprehensive Cancer Center (Welter, Solchaga et al. 2007). The aspirates were harvested after informed consent under the terms of an IRB-approved protocol. We derived hMSCs from four donors; however the cells from one donor failed to differentiate in preliminary tests, and were excluded from further experiments. Thus, cells from three individual donors were used in this study. Adult human MSCs were cultured to confluence in Dulbecco’s Modified Eagle’s Medium, Low (1.5 g/l) Glucose (DMEM-
LG) containing 10% FBS and 10 ng/ml FGF-2. During the MSC culture, medium were changed every other day until cultures reach 80 to 90% confluence. The cell layer were rinsed by PBS with an appropriate volume (15 ml for 150 cm2 tissue culture dish) for two times. The MSCs were then trypsinized with trypsin-EDTA (10ml for 150 cm2 tissue culture dish) and cultures returned to the incubator for 5-8 mins. To keep the culture in trypsin-EDTA as short as possible, once the majority of the cells have detached from the flask surface and became round shape, bovine serum was added with an equal amount of trypsin-EDTA (10ml for T150 tissue culture dish) and resuspended at approximately 50 ×
106 cells/ml. The cell suspension was applied to the channels using a micropipette. After
67
2 hours at 37 °C and 5% CO2 to allow for cell attachment, the seeded membranes were submerged in chondrogenic differentiation medium: Dulbecco’s Modified Eagle’s
Medium, High (4.5 g/l) Glucose (DMEM-HG) supplemented with 10% ITS+ Premix
Tissue Culture Supplement, 10-7 M dexamethasone , 1 μM ascorbate-2-phosphate , 1% sodium pyruvate, and 10 ng/ml transforming growth factor-beta 1 (TGF-β1).The constructs were cultured in a standard incubator at 37 °C and 5% CO2 in humidified air.
Medium was changed every 2-3 days.
3.2.9 MSC Viability, and Adhesion Assessment
To evaluate cell viability within the microchannels, Live/Dead staining with calcein-AM
(Sigma, 17783) and EthD-1 (Molecular Probes, E-1169) were performed 1 day after seeding. 2 μM calcein AM and 4 μM EthD-1 solution in the FGF-2 culture medium were prepared and then added to the membrane in a 100 mm Petri-dish. The dish was incubated at 37 °C and 5% CO2 in humidified air for 30 min after which the medium was removed and the membranes washed three times with PBS. Labeled cells were imaged under the fluorescent microscopy. Viability was quantified by evaluating the percentage of the total number of cells stained positive for calcein-AM.
3.3 Results
3.3.1 CG Membrane Patterning
Figure 3-8 shows images of the surface of the patterned membrane (B) along with the corresponding AutoCAD design of microchannels (A), and a typical phase contrast image of the CG membrane with microchannels.
68
Figure 3-8. (A) Microchannel design created in AutoCAD (B) Digital image of patterned CG membrane (C) Phase Contrast image of collagen microchannel (left to right, 25 , 50 and 100µm)
Table 3-1. Accuracy of channel reproduction in CG membranes. Template width indicates the width of channels in the mask used for photolithography. Mean values of measured width ± standard deviations are shown. Ten channels from three different membranes for each channel size were used.
Table 3-1 shows the accuracy of the collagen soft-lithography method in reproducing the channel designs. The measured widths of the features in the template and the collagen membrane are shown along with the expected design values. The relative error between the membrane channel widths and the template widths ranged from 1 to 15 %, indicating that our method for forming patterned CG membranes is robust and accurate for forming channels as small as 25 μm.
69
Figure 3-9 shows SEM images of the 25 to 500 µm wide collagen microchannels. The original design [Figure 3-8 (A)] was reproduced well on the CG membrane. The image of
25 µm channels [Figure 3-9 (A)] show that the channels have sharp edges indicative of high resolution which further confirms the robustness and accuracy of EDC-based collagen soft-lithography technique.
Figure 3-9. Scanning electron micrographs of linear channels of width 25 (A), 50 (B), 100 (C), and 500 (D) µm formed in CG membranes. Channel Depth is 70 µm. Arrows point to the channels and arrowheads point to spaces between the channels. Scale bars are 50 µm.
3.3.2 MSC Viability and Adhesion Assessment
Figure 3-10 (A) shows an SEM image of MSCs in a 50 µm collagen channel 1 day after seeding. The results show that MSCs attached and spread well forming cell-matrix and cell-cell bonding. Cell viability results show that collagen lithography does not lead to cell death; the MSC viability in the collagen microchannel was more than 95%.
70
Figure 3-10. (A) Scanning electron micrographs of MSCs in a collagen channel of width 50 µm after 24 hours culture. Live/Dead Staining of hMSCs in linear channels of width 100 (C) and 500 (D) µm. Live cells are stained green, and dead cells are stained red.
3.2.3 Selective Cell Attachment
Without Pluronic treatment, cells adhered both inside and outside the channels in both
CG and PDMS substrates [Figure 3-11 (D) and Figure 3-12 (F)]. Using the Pluronic treatment on collagen and PDMS led to significantly more selective adhesion of cells within the channels [Figure 3-11 (A) to (D) and Figure 3-12 (A) to (E)]. These results indicate the importance of the Pluronic modification to achieve selective cell attachment within the channels.
71
Figure 3-11. Effect of F108 on selective seeding of human MSCs in polydimethylsiloxane (PDMS) microchannels. MSCs prestained with Vybrant were seeded onto F108-treated PDMS linear channels overnight. Fluorescent images of cells in linear channels of widths 25 µm (A), 50 µm (B), and 100 µm (C) show selective seeding compared to untreated channel surfaces (D). Scale bars: 50 µm.
72
Figure 3-12. Effect of F108 selective seeding of human MSCs in collagen microchannels. MSCs prestained with Vybrant were seeded onto F108-treated collagen linear channels overnight. Fluorescent images of cells in linear channels of widths 25 µm (A), 50 µm (B), 100 µm (C), 500 µm (D), and 1000 µm (E) show selective seeding compared to untreated channel surfaces (F). Scale bars: 50 µm (A–C, F), 100 µm (D), 200 µm (E).
73
When comparing selective attachment to CG vs. PDMS membranes, the latter yielded better inchannel attachment [Figure 3-13]. More than 90% of MSCs attached to the
PDMS micorchannels and more than 70% for Collagen microchannels after F108 treatment. Complete cell coverage of the 25 µm and 100 µm width channel was achieved on PDMS membrane [Figure 3-11 (A) and (C)].
Figure 3-13. Fraction (cells inside channel/outside of the channel) of MSCs attached to the PDMS and Collagen microchannels after F108 treatment.
3.4 Discussion
To control the ECM microstructure of differentiating MSCs, microscale channels in
PDMS and CG substrates were produced. While PDMS soft-lithography is a well- established technique, in this study, a process of collagen soft-lithography combined with
EDC crosslinking to create channels within CG membranes is presented. The method employs computer-generated designs; any two-dimensional (2D) microdesigns can be
74 replicated in CG substrates. This method allowed for the production of stable physical features and last for a year (Schmidt and Baier 2000). To test the adhesion of MSCs and biocompatibility in the EDC crosslinked CG membranes, we seeded the MSCs on these with a microchannel pattern and hMSCs attached to these microchannels very well with
95% viability suggesting that EDC crosslinked CG membranes are compatible substrates for MSCs adhesion and can be used as an optimal scaffold to investigate human MSC- based chondrogenesis. Collagen Soft Lithograpy was developed previously by using glutaraldehyde as the crosslinker (Janakiraman, Kienitz et al. 2007). While glutaraldehyde have been reported for itd use as crosslinker in tissue engineering. We further employed EDC crosslinking in this work. Calcification is known as one of the side effect for glutaraldehyde processing which cause the failure of bioprosthetic heart valves (Schmidt and Baier 2000). Cytotoxicity has also been found in glutaraldehyde- treated tissues (Gendler, Gendler et al. 1984). While these side effect severely limits the lifetime of glutaraldehyde-treated tissues. In this work, we use EDC crosslinker for CG membrane, the cytotoxic effects of the typical glutaraldehyde crosslinkers, cytotoxic effect as well as glutaraldehyde-induced autofluorescence, a major problem in applying fluorescencebased cell labeling and tracking, are avoided (Gendler, Gendler et al. 1984).
To study the effects of the guidance channels on cellular orientation and function for human MSC-based chondrogenesis, the hMSCs must be localized within the channels.
Our protocol allows us to localize cells within the physical features of a substrate by using Pluronic F108 to resist cell attachment. Pluronic F108 is nontoxic to cells, and the new technique allows for highly improved selectivity. Pluronic has been reported to reduce the overall cell hydrophobicity of the cell which causes enhanced robustness and
75 reduces the surface attachment (Tharmalingam, Ghebeh et al. 2008). Since Pluronic will spontaneously assemble to the hydrophobic surface as we mentioned in section 3.2.4.
Many researchers have taken advantage of this hydrophobic-hydrophobic interaction to regulate cell organization. Liu et.al used pluronic F108 to control 3T3-J2 fibroblasts adhesion on glass and polystyrene and (Liu, Jastromb et al. 2002) regulate NIH 3T3 adhesion on F108-RGD modified polystyrene substrate (Tan, Liu et al. 2004) micropatterned cells onto glass, silicone rubber, and polystyrene by utilizing Pluronics to block the cell adhesion on these substrates. While most researchers utilize Pluronics to micropattern cells on non-biodegradable substrate (glass, metal–oxides, polystyrene), knowing the importance of biodegradable materials in long-term tissue engineering application (Langer and Vacanti 1993; Freed, Vunjaknovakovic et al. 1994; Hutmacher
2000), we presented a new method to localized hMSCs and control cell spatial organization on a natural biodegradable material: collagen–glycosaminoglycan (CG) membrane. Biodegradable polymeric scaffolds have been widely used in tissue engineering application for spatial microenvironment for cellular growth and tissue in- growth (Langer and Vacanti 1993; Freed, Vunjaknovakovic et al. 1994; Hutmacher
2000).
Contacting the Pluronic F108-coated glass slides with the patterned collagen membranes coats only the plateau regions with F108. The F108 was washed off the membrane after cell attached for two hours and won’t affect for 21 days long term chondrogenesis culture.
For the use on collagen substrates, we provide a simple and efficient method to localize the cells within the channels, TE constructs can be created that use the channels
(Described in chapter 4) to guide the cell structure and function, while avoiding cell
76 attachment outside of the channels. Due to hydrophobic nature of PDMS, besides F108 treatment, we utilized microfluidic techniques to selectively coat microchannels with fibronectin in is to ensure the cell attachment for 3 weeks during chondrogenic culture.
The hydrophobic nature of PDMS membrane inhibits the fibronectin solution flow through the spacing of the channels, so an oxygen plasma technique was employed to render the channels hydrophilic. This treatment increased the hydrophobicity of the
PDMS surface that is not in contact with the petri-dishes and enabled fibronectin solution flow all the way through the channels. Combining the microfluidic technique and
Plutonic treatment, selective cell attachment was ensured to larger than 90%.
We have developed a new technique to control topographical and chemical properties of biocompatible (PDMS membrane) and biodegradable biomaterial surface (Collagen membrane). Excellent channel-selective cell attachment was achieved within PDMS membranes and collagen membranes. However, the collagen channels showed less cellular attachment selectivity when compared to the PDMS channels. We speculate that this difference occurred primarily due to the superior ability of the collagen to facilitate cellular attachment, which makes the selective seeding process through F108 somewhat less effective. In addition, Pluronic adsorption required a low degree of wettability of the surface (Tan, Liu et al. 2004). Though we dried the surface of the collagen membrane prior to the F108 treatment, PDMS still had a relatively lower wetting surface which may promote better adsorption of Pluronic.
77
3.5 Conclusion
In this work, we presented a technique to embed microscale patterns onto a EDC crosslinking CG membrane and further reported a simple means of tuning surface chemistry and topography to direct cell selective attachment and spreading on two biomaterials- PDMS and CG membranes. We develop these two robust techniques as a first step towards the creation of TE constructs with built-in cellular architecture.
Through microfluidic techniques, CG membrane soft lithography and F108 modification, selective attachment and spreading of hMSCs within the channels was ensured. The chondrogenic potential of MSCs seeded in these channels will be investigated in the following chapter.
3.6 References
Butt, A. M., M. C. I. M. Amin, et al. "In vitro characterization of pluronic f127 and d- alpha-tocopheryl polyethylene glycol 1000 succinate mixed micelles as nanocarriers for targeted anticancer-drug delivery." Journal of Nanomaterials.
Chen, C. S., M. Mrksich, et al. (1998). "Micropatterned surfaces for control of cell shape, position, and function." Biotechnology Progress 14(3): 356-363.
Chou, C.-L., A. L. Rivera, et al. (2012). “Micrometer scale guidance of mesenchymal stem cells to form structurally oriented cartilage extracellular matrix. Tissue Engineering Part A, 19(9-10): 1081-1090.
Csucs, G., R. Michel, et al. (2003). "Microcontact printing of novel co-polymers in combination with proteins for cell-biological applications." Biomaterials 24(10): 1713- 1720.
Freed, L. E., G. Vunjaknovakovic, et al. (1994). "Biodegradable polymer scaffolds for tissue engineering." Bio-Technology 12(7): 689-693.
Gendler, E., S. Gendler, et al. (1984). "Toxic reactions evoked by glutaraldehyde-fixed pericardium and cardiac-valve tissue bioprosthesis." Journal of Biomedical Materials Research 18(7): 727-736.
78
Hutmacher, D. W. (2000). "Scaffolds in tissue engineering bone and cartilage." Biomaterials 21(24): 2529-2543.
Janakiraman, V., B. L. Kienitz, et al. (2007). "Lithography technique for topographical micropatterning of collagen-glycosaminoglycan membranes for tissue engineering applications." J Med Device 1(3): 233-237.
Kane, R. S., S. Takayama, et al. (1999). "Patterning proteins and cells using soft lithography." Biomaterials 20(23-24): 2363-2376.
Kingshott, P. and H. J. Griesser (1999). "Surfaces that resist bioadhesion." Current Opinion in Solid State & Materials Science 4(4): 403-412.
Langer, R. and J. P. Vacanti (1993). "Tissue engineering." Science 260(5110): 920-926.
Li, J. T. and K. D. Caldwell (1996). "Plasma protein interactions with Pluronic(TM)- treated colloids." Colloids and Surfaces B-Biointerfaces 7(1-2): 9-22.
Liu, V. A., W. E. Jastromb, et al. (2002). "Engineering protein and cell adhesivity using PEO-terminated triblock polymers." Journal of Biomedical Materials Research 60(1): 126-134.
Lu, Q. J., A. Simionescu, et al. (2005). "Novel capillary channel fiber scaffolds for guided tissue engineering." Acta Biomaterialia 1(6): 607-614.
Marin, A., H. Sun, et al. (2002). Drug delivery in pluronic micelles: effect of high- frequency ultrasound on drug release from micelles and intracellular uptake.
Mrksich, M., L. E. Dike, et al. (1997). "Using microcontact printing to pattern the attachment of mammalian cells to self-assembled monolayers of alkanethiolates on transparent films of gold and silver." Experimental Cell Research 235(2): 305-313.
Nation JL. (1983). "A new method using hexamethyldisilazane for preparation of soft insecttissues for scanning electron microscopy." Stain Technology 58(6):347-351
Palecek, S. P., J. C. Loftus, et al. (1997). "Integrin-ligand binding properties govern cell migration speed through cell-substratum adhesiveness." Nature 385(6616): 537-540.
Patel, N., R. Padera, et al. (1998). "Spatially controlled cell engineering on biodegradable polymer surfaces." Faseb Journal 12(14): 1447-1454.
Roach, P., T. Parker, et al. "Surface strategies for control of neuronal cell adhesion: A review." Surface Science Reports 65(6): 145-173.
79
Ruoslahti, E. (1996). "RGD and other recognition sequences for integrins." Annual Review of Cell and Developmental Biology 12: 697-715.
Schmalenberg, K. E. and K. E. Uhrich (2005). "Micropatterned polymer substrates control alignment of proliferating Schwann cells to direct neuronal regeneration." Biomaterials 26(12): 1423-1430.
Schmidt, C. E. and J. M. Baier (2000). "Acellular vascular tissues: natural biomaterials for tissue repair and tissue engineering." Biomaterials 21(22): 2215-2231.
Shin, H., S. Jo, et al. (2003)." Biomimetic materials for tissue engineering". Biomaterials 24 (24):4353–4364
Tan, J. L., W. Liu, et al. (2004). "Simple approach to micropattern cells on common culture substrates by tuning substrate wettability." Tissue Engineering 10(5-6): 865-872.
Tharmalingam, T., H. Ghebeh, et al. (2008). "Pluronic enhances the robustness and reduces the cell attachment of mammalian cells." Molecular Biotechnology 39(2): 167- 177.
Wang, Y. C. and C.-C. Ho (2004). Micropatterning of proteins and mammalian cells on biomaterials.
Welter, J. F., L. A. Solchaga, et al. (2007). "Simplification of aggregate culture of human mesenchymal stem cells as a chondrogenic screening assay." Biotechniques 42(6): 732.
Yannas, I. V., J. F. Burke, et al. (1980). "Design of an artificial skin .2. Control of chemical-composition." Journal of Biomedical Materials Research 14(2): 107-132.
80
Chapter 4
INVESTIGATE THE EFFECT OF MICROSCALE GUIDANCE ON MESENCHYMAL STEM CELL-BASED CHONDROGENESIS
This work is published in Tissue Engineering, Part A, 2012. (Chou, Rivera et al. 2012)
4.1 Introduction
Arthritis currently affects over 49.9 million individuals in the United States, of which osteoarthritis is the most common cause of disability among U.S. adults (MMWR 2010).
One in five U.S. adults reported doctor-diagnosed arthritis and 128 billion dollars were used to treat arthritis and other rheumatic conditions, which equaled 1.2% of the U.S. gross domestic product (Buckwalter 2002; MMWR 2007). Due to the avascular nature of the tissue, articular cartilage has a very limited capacity for spontaneous healing once damaged. It cannot recruit the necessary regenerative cells to repair these lesions(Friedenstein 1995; Caplan, Elyaderani et al. 1997; Caplan and Goldberg 1999)
Therefore, measures must be taken to repair cartilage lesions; however, present cartilage repair techniques appear to provide less than ideal repair tissue (Gilbert 1998) and none was proved successfully to regenerate long lasting hyaline cartilage tissue to replace damaged cartilage (Cohen, Foster et al. 1998). Current repair techniques including subchondral drilling, abrasive chondroplasty and microfracture, only relieve the symptoms temporarily rather than offering a long-term solution for the problem (Simon
1999; Bhosale and Richardson 2008; Ahmed and Hincke 2010). These conventional
81 repair techniques aim to stimulate the bone marrow stem cells migrating from the subchondral bone to the cartilage defect. However, due to the poor repair capability of cartilage, it ultimately results in a fibrocartilagenous scar tissue that lacks the mechanical properties of native articular cartilage and will eventually undergo pre-mature degeneration (Curl, Krome et al. 1997; Nehrer, Spector et al. 1999; Kasemkijwattana,
Kesprayura et al. 2009). Fibrocartilage is fibrous tissue and biochemically contains more type I collagen and significantly less proteoglycan than hyaline cartilage. Therefore, it has inferior mechanical properties and been reported yielded unsatisfied outcomes in clinical use (Hunziker 2002).
Tissue engineering (TE) is viewed as a promising method for long-term repair of cartilage lesions. For cartilage repair, various materials based on both natural and synthetic polymers have been fabricated to be used as scaffolds in a variety forms, including fibrous structures, porous sponges and hydrogels. Collagen-based biomaterials are widely used in today’s clinical practice such as haemostasis, duramater replacement and cosmetic surgery and have been used experimentally as carriers for chondrocytes
(Frenkel, Toolan et al. 1997) and mesenchymal stem cells (MSCs) (Im, Kim et al. 2001).
Several clinical studies have used collagen membranes (Bentley 2004) as a replacement for perichondral membrane to close the defect. The combination of collagen with glycosaminoglycan (GAG) in scaffolds showed a positive effect on chondrocyte phenotype (van Susante, Pieper et al. 2001). Based on these, collagen-based scaffolds were used for this study. Chondrocytes, highly specialized cells, synthesize all the ECM components and regulate the metabolism of the cartilage tissue (Bhosale and Richardson
2008). Autologous Chondrocyte Implantation (ACI) has been developed for cartilage
82 repair which use autologous chondrocyte from normal, low weight loading site of the damage join and re-implant into the defects (Brittberg, Lindahl et al. 1994; Chanlalit,
Kasemkijwattana et al. 2007). Although ACI have shown positive effect over the conventional method in cartilage repair, several issues limited its application which include small amount of available cells, slow expansion rate and the inadequate ability of the cells to proliferate(Cancedda, Dozin et al. 2003). Some research further showed ACI still result in the formation of fibrocartilagenous scar tissue (Clar, Cummins et al. 2005).
On the other hand, MSCs serve as a promising alternative in cartilage tissue engineering due to their availability and pluripotent potential to differentiate into cell lineages including chondrocytes (Magne, Vinatier et al. 2005). In this approach, mesenchymal stem cells (MSCs) have been combined with biodegradable scaffolds and growth factor to produce cartilage constructs in vitro (Solchaga 2002; Welter 2002; Baskaran 2003;
Solchaga L 2003; Welter 2003; Baskaran 2004; Solchaga 2004; Welter 2004). TE constructs have been shown to be biochemically similar to native cartilage, however, they have inferior mechanical strength compared to native cartilage. Human articular cartilage has been reported to have a depth–dependent compressive modulus of 1.16±0.20 MPa
(0–125μm from the articular surface) to 7.75±1.45 MPa (1250–1500 μm from the articular surface) (Chen, Falcovitz et al. 2001) while the average tensile modulus for native articular cartilage is about 0.7–12.5 MPa (Almarza and Athanasiou 2004; Huang,
Stankiewicz et al. 2005). Huang et. al showed MSC-seeded TE constructs exhibited compressive equilibrium modulus of 0.15MPa after 6 weeks culture (Huang, Farrell et al.
2010). MSC-seeded agarose constructs TE construct has been reported retain a compressive modulus of 0.01MPa and 0.34 MPa for tensile modulus after 3 weeks
83
(Huang, Baker et al. 2012). In addition, autologous mesenchymal stem cells implantation has been showed to successfully repair the defects in rabbits, however the mechanical strength are lower than normal value (Wakitani, Goto et al. 1994; Caplan, Elyaderani et al. 1997). Mechanical property of current TE construct is a major issue which prevents current constructs from reaching a clinical setting. This has been attributed to a lack of an oriented microscale structure in TE constructs, which is present in the native tissue
(Bhosale and Richardson 2008).
Articular cartilage is composed of chondrocytes embedded in a hydrated extracellular matrix (ECM) which is made of collagen, proteoglycan and other non-collagenous proteins. These macromolecules hold a large amount of water (60% to 80% by weight) within the ECM which(Cohen, Foster et al. 1998) provides the unique mechanical strength necessary for everyday load-bearing that occurs within the body. Articular cartilage has a very ordered structure which is usually organized into four zones: the superficial zone, the radial zone, the deep zone and the calcified cartilage zone (Figure 4-
1). The extracellular matrix of cartilage contains collagen fibrils that are highly organized in different orientations depending on the particular cartilage zone. Collagen fibrils make up about 70% of the tissue dry weight (Rieppo, Rieppo et al.) and mainly made of collagen type II. These fibrils are oriented parallel to the articular surface in the superficial zone of cartilage which for providing the greatest tensile and shear strength(Bullough and Goodfellow 1968; Temenoff and Mikos 2000; Bhosale and
Richardson 2008). In the radial (middle) zone, collagen fibrils are randomly. In the deep zone, collagen fibrils are perpendicular to the joint surface and the largest diameter of collagen fibrils and highest amount of proteoglycan are in in this zone (Benninghoff
84
1925; Clarke 1971; Clark 1991; Hasler, Herzog et al. 1999; Bhosale and Richardson
2008). Mathematical models suggest that, within native articular cartilage, the well- organized depth dependent structure and composition of the ECM [Figure 4-1] provides the unique depth-dependent mechanical strength necessary for everyday load bearing properties (Kempson, Freeman et al. 1968; Schinagl, Gurskis et al. 1997; Chen, Bae et al.
2001) (Schinagl, Gurskis et al. 1997). Based on the arcade model of Benninghof
(Benninghoff 1925), Wilson et al. developed a poroviscoelastic fibril-reinforced (FEA) model to study collagen network behavior and concluded that local stresses and strains in the articular cartilage are highly influenced by the local morphology of the collagen fibril network (Wilson, van Donkelaar et al. 2005; Wilson, van Donkelaar et al. 2005) which suggest that structure leads to improve mechanical properties. The increase in mechanical strength from the surface of the cartilage to the deep zone is attributed to the ECM alignment (Wilson, Huyghe et al. 2007; Shirazi, Shirazi-Adl et al. 2008). The inferior mechanical property is the major issue for current tissue engineerned cartilage. This has been attributed to a lack of an oriented microscale structure in TE construct. Based on these, in this study, we test the hypothesis that microfabricated matrices would induce differentiating hMSCs to form oriented collagen microstructures, mimicking the ultrastructure of the native cartilage and further increase the mechanical function of the tissue engineered construct. To reach this goal, we used contact guidance, which in previous studies, has been shown to influence cell orientation, migration, and functions
(Dickinson, Guido et al. 1994; Tessier-Lavigne and Goodman 1996). The concept of contact guidance were first been introduced to study the orientation of neurons and nerve fibers (Weiss 1934; Weiss 1945). Since then, a variety of different types of cells have
85 been demonstrated aligned using the strategy of contact guidance. By naturally control the topography of the substrate, one can regulate the cell morphologies and further control the specific mechanical, physiological functions in tissue construct (Wang, Jia et al. 2003; Manwaring, Walsh et al. 2004; Hwang, Park et al. 2009; Aubin, Nichol et al.
2010; Huang, Lee et al. 2010; Kapoor, Caporali et al. 2010; Roach, Parker et al. 2010).
For example, Manwaring et al. employed contact guidance to direct the organization of meningeal cell and showed that substrate features caused alignment of cells, which in turn could align ECM proteins(Manwaring, Walsh et al. 2004). In addition, Kapoor et al. showed that microtopography affects cell density and alignment of tenocytes microtopographical features and leads to the deposition of an aligned collagen matrix
(Kapoor, Caporali et al. 2010). Wang et al. found the orientation of MC3T3-E1 cells determines the alignment of cell-produced collagenous matrix (mostly collagen type I)
(Wang, Jia et al. 2003). Furthermore, Huanget al. micropatterned on myoblast on polydimethylsiloxane substrate and generated aligned tissue-like muscle constructs
(Huang, Lee et al. 2010). These researches indicated that micro scale guidance lead to cell alignments and result in the deposition of oriented ECM and may further control the specific mechanical, physiological functions for different tissue construct.
Despite these studies, the effects of contact guidance on stem cell differentiation and chondrogenesis are poorly understood. To our knowledge, no study has shown the effect of contact guidance on human MSC (hMSC)-based chondrogenesis; specifically, the effects of contact guidance on the synthesized type II collagen and on the mechanical properties of the resulting constructs are not known. Based on this, therefore, the goal of this study is to test the hypothesis that microscale substrate features would cause
86 differentiating MSCs to preferentially arrange themselves and to deposit an oriented cartilage ECM on a scale similar to native articular cartilage. Using contact guidance, we form constructs with microscale architecture for improved structure and function [Figure
4-2]. The first step of this project, we would like to understand the biology of cartilage ultrastructure formation by MSCs as a result of guidance channels. We developed oriented collagen microstructures and further design external templates for cartilage TE.
Channels of varying microscale dimensions in either collagen-based or polydimethylsiloxane (PDMS) membranes are formed via a combination of microfabrication and soft-lithography as detailed in chapter 3. Selective seeding of viable
MSCs within the channels and subsequent chondrogenic differentiation leads to alignment of mature type II collagen fibrils along the length of the channel. The quality of alignment is significantly dependent on the channel dimensions; further, our results show that alignment is correlated with a significant improvement in mechanical properties.
87
Figure 4-1. (A) Schematic of ultrastructure of articular cartilage adapted from Hunziker et al. (Hunziker, Michel et al. 1997): Blue circular shapes are chondrocytes and the black bands are collagen fibrils. The figure shows orthogonal arrangement of collagen fibrils in the central and near subchondral bone regions of the cartilage and a parallel arrangement near the superficial region of the cartilage. This ultrastructure has been shown to play a role in cartilage mechanical properties (Shirazi, Shirazi-Adl et al. 2008).
Figure 4-2. Illustration of experiment design.Test the hypothesis that microscale substrate features would cause differentiating MSCs to preferentially arrange themselves and to deposit an oriented cartilage ECM on a scale similar to native articular cartilage. Blue represents MSC cells and green represents ECM
88
4.2 Materials and Methods
4.2.1 Materials
Collagen type I from bovine Achilles tendon, chondroitin-6-sulfate sodium salt from shark cartilage, 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDC),
N-hydroxy-succinimide, fetal bovine serum (FBS), 4', 6-diamidino-2-phenylindole
(DAPI), and phalloidin were purchased from Sigma Chemical Co. (St. Louis, MO).
Antibiotic-antimycotic cocktail, low glucose Dulbecco’s Modified Eagle Medium
(DMEM), high glucose DMEM, ITS (insulin, transferrin, selenium), ascorbate 2- phosphate (A2P), dexamethasone, sodium pyruvate, and human plasma fibronectin were obtained from Gibco, Invitrogen (Carlsbad, CA). Sylgard® silicone elastomer was purchased from Dow Corning Corporation (Midland, MI). Fibroblast growth factor-2
(FGF) and transforming growth factor β1 (TGF- β1) were purchased from Peprotech
(Rocky Hill, NJ). ITS (insulin, transferrin, selenium)+ Premix Tissue Culture Supplement were obtained from Becton Dickinson (Franklin Lakes, NJ). Carboxyfluoro diacetate succinimidyl ester (CFDA-SE,Vybrant® ), Pluronic F108 was purchased from BASF
(Whitehouse, OH). 1,1'-dioctadecyl-3,3,3'3'-tetramethylindocarbocyanine perchlorate
(DiI) cell labeling solution, calcein AM, and ethidium homodimer-1 (EthD-1) were obtained from Molecular Probes (Invitrogen, Carlsbad, CA). Pluronic F108 was purchased from BASF (Whitehouse, OH). Fluorescein isothiocyanate (FITC)-conjugated goat anti-mouse IgG secondary antibody was from MP Biomedicals (Irvine, CA,), while the collagen Type II primary antibody was obtained from the Developmental Studies
Hybridoma Bank (University of Iowa). A SPOT RT digital camera was obtained from
89
Diagnostic Instruments (Sterling Heights, MI), while a Leica fluorescence microscope was from Leica Microsystems GmbH (Wetzlar, Germany).
4.2.2 Fabrication of Collagen-hMSC construct and PDMS-hMSC construct
Collagen–GAG membranes were produced using standard filtration method (Janakiraman,
Kienitz et al. 2007). CG membrane scaffolds of size 1.5 (width) x 1.8 (length) x 0.4
(depth) cm were patterned with microchannel feature using the collagen soft lithograpy method while PDMS membranes were patterned with standard soft lithograpy, detailed in
Chapter 3. The patterned membrane were stored in 1x PBS at 4°C prior to use. The channel design we used for studying guidance effect ranged from a minimum width of 25
μm to a maximum width of 1000 μm as shown in Figure 3-1. The spacing between channels of the same width was varied from 50 to 250 μm for different designs to allow us to test the effect of channel density in future experiments. On the other hand, single channel size was used for mechanical property and biochemical measurements. The spacing between channels was equivalent to the channel width for tensile testing and biochemical measurements experiments. PDMS channel area were treated with
Fibronectin using microfluidic technique and both PDMS and collagen membrane were treated with F108 for the gap area. hMSCs are subcultured to 80 to 90% confluence to avid contact inhibition and spontaneous differentiation (Solchaga, Welter et al. 2004) .
Membranes were incubated at 37°C for 1 hour prior to cell seeding. Approximately 50 ×
106 cells/ml cell suspension was applied to the channels using a micropipette. After 2 hours at 37 °C and 5% CO2 to allow for cell attachment, the seeded membranes were submerged in chondrogenic differentiation medium: Dulbecco’s Modified Eagle’s
Medium, High (4.5 g/l) Glucose (DMEM-HG) supplemented with 10% ITS+ Premix
90
Tissue Culture Supplement, 10-7 M dexamethasone , 1 μM ascorbate-2-phosphate , 1% sodium pyruvate, and 10 ng/ml transforming growth factor-beta 1 (TGF-β1).The constructs were cultured in a standard incubator at 37 °C and 5% CO2 in humidified air for 21 days. Medium was changed every 2-3 days.
4.2.3 Image Acquisition
Phase contrast images of cells in constructs were taken with an Olympus IX71 inverted microscope (Olympus, Japan) using UPlanFl 10x and LCPlanFl 20x objectives.
Immunofluorescence microscopy images were obtained either using a SPOT RT digital camera attached to a Leica fluorescence microscope (Leica Microsystems, Gmbh,
Wetzlar, Germany) or by confocal microscopy (Zeiss LSM 510, Zeiss Axiovert 200M;
Carl Zeiss, Thornwood, NY) with HCX PL APO 40x and 20x water onjective. The dyes used for labeling included DAPI (absorption 358 nm, emission 461 nm), FITC (495 nm and 519 nm respectively), Texas-Red-X (608nm and 561nm respectively) and 1,1'- dioctadecyl-3,3,3'3'-tetramethylindocarbocyanine perchlorate (DiI) (549 nm and 565 nm respectively).
4.2.4 Histology and Immunohistochemistry
After culturing for 21 days, some samples were fixed in 4% neutral-buffered formaldehyde (Fisher Scientific, Pittsburgh, PA). Whole-mounts were surface-stained for type II collagen by immunohistochemistry. Antigen unmasking was performed with
1mg/mL pronase (Sigma-Aldrich, St. Louis, MO) in PBS for 15 min at room temperature.
After washing the samples with 1x PBS twice for 30 min, they were then blocked with
10% normal goat serum (NGS) (Sigma-Aldrich, St. Louis, MO) in PBS for 30 min.
91
Anticollagen type II antibody (Developmental Studies Hybridoma bank, University of
Iowa, Iowa City, IA) diluted 1:50 in 1% NGS in PBS, was applied to the samples for 1 h to stain for collagen type II. The samples were then washed with PBS for 30 mins two times. The FITC-conjugated goat anti-mouse IgG secondary antibody (Cappel 55499, detects IgG, IgA, and IgM isotypes. MP Biomedicals, Irvine, CA, USA), diluted 1:500 in
1% NGS in PBS, was then applied to the samples for 45 min follows by 2 times 30 mins wash in PBS. All samples were counterstained with DAPI (Sigma-Aldrich, St. Louis, MO) at 300 nM final concentration for 10 mins to visualize the cell nuclei.
Some samples were counterstained with 1,1'-dioctadecyl-3,3,3'3'- tetramethylindocarbocyanine perchlorate (DiI) cell labeling solution (Molecular Probes,
Invitrogen, Carlsbad, CA) for membrane staining. Cell membrane visualization was done incubating with DiI solution the samples for 20 minutes. Some samples were stained with
Texas Red® -X Phalloidin (Molecular Probes, Invitrogen, Carlsbad, CA) for actin. F- actin visualization was done by diluting phalloidin in PBS (1:40) and incubating with the samples for 20 minutes. The samples were again washed with PBS for 1 h, counterstained, and wet-mounted using 5% N-propyl gallate in glycerol. N-propyl gallate. Prepared as a
5% solution in glycerol (Sigma-Aldrich, St. Louis, MO). All procedures were performed at room temperature. The samples were imaged either using a SPOT RT digital camera attached to a Leica fluorescence microscope (Wetzlar, Germany) or by confocal microscopy (Zeiss LSM 510, Zeiss Axiovert 200M; Carl Zeiss, Thornwood, NY).
4.2.5 Cell alignment analysis
To obtain a quantitative measure of alignment, after culturing for 21 days, Collagen- hMSC constructs and PDMS-hMSC constructs were were fixed with 4% buffered-
92 paraformaldehyde solution for 1hr at room temperature and 8 h at 4°C. F-actin visualization was done by diluting phalloidin in PBS (1:40) and incubating with the cells for 20 minutes. The cell nuclei were labeled with DAPI at 300 nM final concentration for
10 minutes. The constructs were visualized and imaged for fluorescence images using a
SPOT RT digital camera attached to a Leica fluorescence microscope (Wetzlar, Germany) and by confocal microscopy (Zeiss LSM 510, Zeiss Axiovert 200M; Carl Zeiss,
Thornwood, NY). The ImagePro software (Media Cybernetics, Inc., Bethesda, MD) was used to quantify the nuclear alignment angle. To obtain a quantitative measure of alignment, the acute angle of the nuclei or cell outline was calculated relative to the direction of the channel length as shown in Figure 4-3. Nuclear aligment angle is obtained by drawing a straight line along the orientation of the major axis of the nuclei and measuring the angle between the direction of channel length. Actin fiber alignment is obtained by measuring the angle between the drawn line along the major axis of the cell outline with respect to the channel length direction. Angle between the linear channel dimension and cell major axis through the cell nucleus or cell outline was quantified and shown as mean ± SEM for channels of various widths. At least three images from three separate experiments were used and frequency histograms of the angles were plotted.
Alignment angles were subsequently normalized and grouped in 15° angle bins. An alignment angle of 0° indicates perfect alignment with the long dimension of the channels and an angle of 90 °indicates perfect alignment with the short dimension of the channels.
93
Figure 4-3. Schematic of nuclear alignment angle measurement. The angle between the orientation of the major elliptic axis of individual nuclei and the horizontal axis parallel to the channel length.
4.2.6 Mechanical Property Testing
To determine whether alignment led to improved function, the CG membrane samples were subjected to tensile testing. For each mechanical testing sample, only a single channel size was used and the spacing between channels was equivalent to the channel width (projected channel area is 50% of the total surface area for all channel sizes) as shown in Figure 4-4. Therefore, the seeding areas of the channels and the seeding densities are identical for each sample. In addition, the ratio of the channel cross- sectional area containing tissue to the total sample cross-sectional area is constant for every condition and is 0.1 based on a membrane depth of 400 μm. CG membranes with hMSCs cultured for 21 days under differentiating conditions were cut to half using a razor blade to rectangular dimensions of approximate 9mm in length × 5mm in width ×
0.4mm in thickness. The CG membranes are not degradable in the cell culture medium; therefore, the base membranes of all samples subjected to mechanical testing are
94 consistent. The edge of the sample were gripped with film tension clamps and subjected to tensile strain in a dynamic mechanical analysis apparatus (QA800, TA Instruments,
New Castle, DE) along the direction of channel length until failure [Figure 4-5]. Strain was ramped from 20%/min to 200%/min. Stress (σ) vs. strain (ε) data were obtained and fitted to the following equation
( ) to yield the modulus of elasticity (ab) (Woo, Akeson et al. 1976; Liang, Kienitz et al.
2010). Ultimate stress values were calculated at the failure point. At least seven samples were used per condition tested.
Figure 4-4. Microchannel pattern on silicon wafer for tensile testing. The channels ranged from a minimum width of 25 μm to a maximum width of 1000 μm. Single channel size was used and the spacing between channels was equivalent to the channel width. 25 µm (A), 50 µm (B), 100 µm (C), 250 µm (D), 500 µm (E), and 1000 µm (F).
95
Figure 4-5. Tensile testing of Collagen-hMSC constructs. More than 7 samples were used for each condition. The samples were tested to failure by DMA.
4.2.7 Biochemical Measurements
We determined the GAG and DNA content of the tissue formed during chondrogenesis using previously published methods (Welter, Solchaga et al. 2007). The cartilaginous tissue formed for each condition (n=3) was transferred to 1.5-ml microcentrifuge tubes and digested using pH6.5 papain buffer which consist of 25 μg/ml papain, 2 mM cysteine,
2 mM EDTA (all from Sigma-Aldrich, St. Louis, MO) and 50 mM sodium phosphate
(Fisher Scientific, Pittsburgh, PA) in nuclease-free water (Sigma-Aldrich, St. Louis, MO) at 65°C. Check every 30 mins until sample digested completely. 400 μl of 0.1 N NaOH were added to each sample for 20 mins at room temperature. After which, samples were neutralized with 400ml pH 7.2 neutralizing solution consisting of 4 M NaCl (Fisher), 100 mM Na2PO4 (Fisher Scientific, Pittsburgh, PA) and 0.1N HCl. Samples were centrifuged for 5 mins and supernatant were transferred to fresh microcentrifuge tubes which used for
DNA and GAG content measurement.
96
4.2.7.1 DNA content quantification Calf thymus DNA standards were used (Amersham Biosciences, Piscataway, NJ) in this assay. 100 μl of each standard and 100 μl supernatant of each sample were placed to four replicate wells of a 96 well plate. 100 μl of 0.7 μg/ml Hoechst 33258 in water (Sigma
Chemical Co., excitation wavelength 340nm and emission wavelength 465 nm) were added to each well. After mixing well and remove the bubble in the wells, fluorescence was read in a GENios Pro Visible/UV spectrophotometer (Tecan, Durham, NC, USA).
4.2.7.2 Glycosaminoglycan (GAG) content quantification A standard curve was made with Shark cartilage chondroitin sulfate C (Seikagaku
America, East Falmouth, MA) in this method. 0.45 μm nitrocellulose were cut and place in distilled water in a 150 mm2 Petri-Dishes for 30 mins. The moisture nitrocellulose were then placed to the the dot-blot apparatus. 250 μl of 0.02% Safranin-O in 50 mM
(pH 4.8) sodium acetate were added to the wells. 25 μl supernatant of each sample were added to the Safranin-O reagent in these wells. After 1minute, the used well were covered by transparent tap and the vacuum was applied to the dot-blot apparatus. The wells were rinsed by water for 2 to 3 times to allow it to filter through the nitrocellulose membrane under vacuum. The nitrocellulose membrane were then removed from the apparatus and air dried. Nitrocellulose dots in the membrane were punch out with a hole punch and placed in fresh microcentrifuge tubes. 1 ml of 10% cetylpyridinium chloride (CPC)
(Sigma-Aldrich, St. Louis, MO) in water was added to each microcentrifuge tubes to to elute the dye from the nitrocellulose dots. Samples were incubated at 37°C for 20 mins following by vortexing10 mins and reading the absorbance at 530 nm in a GENios Pro
Visible/UV spectrophotometer (Tecan, Durham, NC, USA).
97
4.2.8 Statistical methods
Statistical analysis was carried out using the Origin 8.5.1 (Origin Lab, Northampton, MA) software package. Pairwise comparisons were made using the Tukey’s test with a p value of less than 0.05 considered statistically significant. Sample sizes are indicated in the respective figure legends
4.3 Result
4.3.1 Cellular Growth and Organization in Microchannels
Figure 4-6 shows hMSCs morphology and organization in various guidance channels at different culture period. Phase contrast images shows three hours after seeding, hMSCs attached and spread in the guidance channels [Figure 4-6: A (a)-(c)]. After 7 days culture
[Figure 4-6: B (a)-(c)], hMSCs showed elongated and differentiated in the various dimension of guidance channels. Cells in smaller channels (50 and 100 μm) aligned along the length of the channel [Figure 4-6: B (a)-(b)]. hMSC organization in smaller channels shown self-organized into oriented constructs consisting of elongated and aligned cells.
Although in the early stage of chondrogenesis, the guidance effect may be limited to the cells near the edge of the channel, after 7 days culture, the guidance effect shown in the entire small channels [Figure 4-6: B (a)-(b)]. On the other hand, cells in larger channels
(500 μm) showed random orientation and tended to aggregate into clumps after 7 days culture [Figure 4-6: A (c) and B (c)]. This result indicated the cellular behavior of hMSCs can be controlled and directed by guidance channels which corresponding the first part of our hypothesis that microscale substrate features would cause differentiating MSCs to preferentially arrange themselves into oriented cellular organization.
98
Figure 4-6. MSCs morphology and organization as a function of time in guidance channels. Phase contrast image of hMSCs on guidance channels. Three hours after seeding (A) and Day 7 (B). Small guidance channels induced hMSCs elongated and aligned along the channel length while cells in larger channel remain random orientation. 50 μm (a), 100 μm (b), 500 μm (c). Scale bar: 100 μm
4.3.2 Microscale Guidance on Cell alignment
After 21 days of Chondrogenic culture, fluorescent images of linear channels showed that more hMSC alignment along the length of the channel occurred in the smaller channels
99
[Figure 4-7 (A)-(C)] compared to the larger channels [Figure 4-7 (D)-(E)]. DAPI staining showed that the nuclei of the MSCs elongated into an oval shape in the smaller channels
(25 to 100μm) and aligned along the channel length. In addition, our data also showed that micro guidance channels also affect the actin fiber orientation [Figure 4-8]. Actin fibers of hMSCs aligned with the long dimension of the smaller channels [Figure 4-8 (A),
(Ci)]. Within the larger channels, MSCs showed random spatial orientation; less nuclear elongation and actin fibers extended in all directions outward from the cell nuclear
[Figure 4-8 (B), (Cii)]. To obtain a quantitative measure of alignment, the acute angle of the nuclei or cell outline was calculated relative to the direction of the channel length. By measurement the angle of actin fiber orientation of hMSCs in various guidance channels, our data showed guidance channels highly affect the actin microfilaments orientation.
Figure 4-9 shows the angle of actin alignment within the channels (mean ± SEM). An alignment angle of 0° indicates perfect alignment with the long dimension of the channels and an angle of 90 °indicates perfect alignment with the short dimension of the channels.
The value for actin alignment angle were 2.61±0.46 for 25 μm, 3.11±0.38 for 50 μm,
3.89±0.55 for 100 μm and 31.24±3.01 for 500 μm. The actin angle measurements result indicate that the smaller channels (25–100 μm) aided in microfilaments alignment whereas the larger (500μm) channels did not. In addition, there was significant difference in alignment as measured by the angle, between 25 to 100 μm wide substrates when compared with 500 wide substrates (the Tukey’s test, p < 0.05).
100
Figure 4-7. Chondrogenesis under microscale guidance after 21 days culture. Fluorescent images of linear channels of widths 25 μm (A), 50 μm (B), 100 μm (C), 500 μm (D), and 1000 μm (E) show alignment in smaller 25–100 channels. Scale bar: 50μm (A–C), 100 μm (D), 200 μm (E). Green stained type II collagen and blue stained DAPI.
101
Figure 4-8. Alignment of actin microfilaments in guidance channels. Cell nuclei alignment (blue) is correlated with actin (red) alignment in smaller channels. [A, C(i)] show actin alignment in linear channel of width 50 μm, which is contrasted with random alignment of actin filaments in linear channels of width 500 μm [B,C(ii)].
Figure 4-9. Actin Fiber Orientation Angle under Microscale Guidance. Angle between the linear channel dimension and cell major axis through the cell nucleus was quantified and shown as mean ± SEM for channels of various widths. An alignment angle of 0° indicates perfect alignment with the long dimension of the channels and an angle of 90° indicates perfect alignment with the short dimension of the channels. For smaller
102 channels (25, 50, 100 μm), six channels chosen from three different experiments were used for quantitation. For larger channels (500 μm), three channels from three different experiments were used. Cells from three donors were used in the experiments. Corresponding immunofluorescence data are shown in Figure 4-8 A–C. aStatistically significant (p < 0.05) compared to 500 and 1000 μm wide channels.
Furthermore, by measurement the angle of nuclear of hMSCs in these guidance channels, our data showed guidance channels highly affect the nuclei orientation. Table 4-1 shows the angle of nuclear alignment within the channels (mean ± SEM). Figure 4-10 shows the distribution of angles for all conditions. The nuclear angle measurements suggest that the smaller channels (25–100 μm) aided in cell alignment (3.7° to 12.7° for PDMS substrate and 4.5° to 8.6° for collagen substrate), whereas the larger (500°–1000° μm) channels
(29.2° to 32.2° for PDMS substrate and 41.8° to 42.3° for collagen substrate) did not.
Both collagen and PDMS substrates followed this trend. There was significant difference in alignment as measured by the angle, between 25 to 100 μm wide substrates when compared with 500 and 1000μm wide substrates (the Tukey’s test, p < 0.05). In addition, there was no significant difference in alignment between 500 and 1000 μm wide substrates. In the larger channels, small regions in which the MSCs aligned to each other were noted [Figure 4-8 (B)], but the alignment was not in reference to the channel dimensions.
103
Figure 4-10. Cell nucleus orientation under microscale guidance. Distribution of angles of orientation of human MSCs formed against the longer dimension of the Collagen [(A) and (C)] and PDMS [(B) and (D)] channels of widths 25–1000 μm. (C) and (D) are the normalized versions of (A) and (B) with respect to total cell number. An alignment angle of 0° indicates perfect alignment with the long dimension of the channels and an angle of 90° indicates perfect alignment with the short dimension of the channels. For smaller channels (25, 50, 100 μm), six channels from three different samples were used. For larger channels (500, 1000 μm), three channels from three different samples were used. Cells from three donors were used.
104
Table 4-1. Cell Alignment Angle During Contact Guidance Angle between the linear channel dimension and cell major axis through the cell nucleus was quantified and shown as mean ± SEM for channels of various widths. For smaller channels (25, 50, 100 μm), six channels chosen from three different experiments were used for quantitation. For larger channels (500, 1000 μm), three channels from three different experiments were used. Cells from three donors were used in the experiments. Corresponding immunofluorescence data are shown in Figure 4-7 A–E and Figure 4-11 A-B. aStatistically significant (p < 0.05) compared to 500 and 1000 μm wide channels. PDMS, polydimethylsiloxane.
Channel Width 25 50 100 500 1000 (μm) PDMS 3.7±1.4a 12.7±2.4a 4.6±1.0a 29.2±2.8 32.2±4.2
Collagen 4.5±0.8a 4.1±0.6a 8.6±1.0a 41.8±2.5 42.3±3.0
4.3.3 ECM Production
Our results [Figure 4-7 (A)-(E), Figure 4-8 (A)-(B), Figure 4-11 (A)-(B), Figure 4-12
(A)-(E)] show that mature extracellular type II collagen is deposited and that production closely follows the aligned cell in the channels. Longer type II collagen fibers are seen farther from the cell, indicating that filament assembly occurs. In the 25, 50, and 100 μm channels [Figure 4-7 (A)-(C), Figure 4-11 (A), Figure 4-12 (A)-(C)], the type II collagen aligned along the direction of the channel, while in 500 and 1000 μm channels, the type II collagen was randomly aligned [Figure 4-7 (D)-(E), Figure 4-11 (B), Figure 4-12 (D)-
(E)].
105
Figure 4-11. Chondrogenesis under microscale guidance. Fluorescent images of linear channels of widths 25 μm (A), 500 μm channels show mature aligned extracellular matrix (arrowheads) in the smaller channel. Scale bar: 50μm (A), 100μm (B). Green is type II collagen, blue is DAPI and red is DiI.
106
Figure 4-12. Confocal fluorescent images of linear channels of widths 25 μm (A)-(B), 50 μm (C), 500 μm (D), and 1000 μm (E) channels show mature aligned extracellular matrix (arrowheads) in the smaller channel after 21days culture. Scale bar: 10μm. Green is type II collagen, blue is DAPI and red is DiI.
107
Biochemical assay results show that the GAG/DNA content in microchannels was significantly greater than that when cells were seeded on membranes with no channels
[Table 4-2]. There was, however, no difference in the GAG/DNA content between membranes of different channel widths.
Table 4-2. GAG and DNA Content of Cartilage Tissue Grown in the Channels
Data from three samples from three different experiments are presented as mean±SEM. Cells from three donors were used in the experiments. aStatistical significant difference at p < 0.05. GAG, glycosaminoglycan
Channel Width GAG(μg) DNA(μg) GAG/DNA (μm) 25 4.3±0.5 2.1±0.03 2.0±0.27a
50 4.4±0.4 2.1±0.01 2.1±0.17a
100 4.0±0.1 2.0±0.01 2.0±0.07a
250 3.4±0.2 2.0±0.05 1.7±0.09a
500 3.3±0.1 2.1±0.03 1.6±0.03a
1000 3.3±0.1 2.1±0.02 1.6±0.05a
No Channel 3.0±0.3 2.4±0.04 1.2±0.09
4.3.4 Mechanical Properties
Figure 4-10 shows the modulus of elasticity [Figure 4-13 (A)] and ultimate stress [Figure
4-13 (B)] of various collagen samples when they were subjected to tension along the
108 direction of the channel length. CG membranes containing hMSCs in the smaller channels [Figure. 4-13: 25, 50, 100, 250] cultured for 21 days under differentiating conditions had significantly superior mechanical properties compared to corresponding membranes with larger channels [Figure 4-13: 500, 1000]. Importantly, membranes with randomly seeded cells [Figure 4-13. (A):Ran], while better than membranes with no cells
[Figure 4-13 (A): EDC], had inferior modulus of elasticity compared to membranes containing cells in channels of 25, 50, 100, and 250 μm widths [Figure 4-13 (A): 25, 50,
100, 250]. The ultimate stress data for membranes containing cells in channels of 25, 50, and 100 mm widths [Figure 4-13 (B): 25, 50, 100] are significantly greater compared with membranes with no channels, 250, 500 and 1000 mm-wide channels [Figure 4-13
(B): Ran, 250, 500, 1000]. There was no difference in the mechanical properties between cell-free membranes with 25 and 100 μm channels [Figure 4-13: C25 and C100] and cell- free membranes with no channels [Figure 4-13: EDC]
109
Figure 4-13. Effect of microscale guidance on mechanical properties. CG membranes were subject to tensile testing until failure. Mean values of elasticity modulus (A), a typical force–displacement curve (A: upper left corner), and ultimate stress (B) are shown. BK and EDC represent, respectively, uncrosslinked and EDC crosslinked CG membranes without microchannels or MSCs. C25 and C100 represent CG membranes consisting of 25 and 100 μm linear channels only, without MSCs. All other channels were seeded with MSCs. Ran: EDC crosslinked CG membranes without
110 channels; 25, 50, 100, 250, 500, and 1000 represent widths (μm) of linear channels in the EDC crosslinked CG membranes. n = 4 for 1000, n = 5 for Ran, C25 and C100 and n = 6 for all other conditions. (A) *Statistically significant difference ( p < 0.05) compared to data with no channels (RAN). #Statistically significant difference ( p < 0.05) compared to data with 1000-mm channels (1000). (B) *Statistically significant difference ( p < 0.05) compared to data with no channels (RAN), 250 μm (250), 500 μm (500), and 1000 μm (1000) wide channels. Cells from two donors used in the above experiments.
4.4 Discussion
We hypothesized that channels in the CG and PDMS membranes would control cellular orientation and ECM production via contact guidance. Our results indicate that this is the case in the smaller channels (25, 50, and 100 μm), as these lead to overall MSC alignment along the channel length. The larger channels, on the other hand, had small regions in which the MSCs are aligned to each other [Figure.4-7 and Figure 4-11], but not in reference to the channel dimensions. Overall, the distribution of the MSCs was random in these larger channels. The smaller scale features (25–50 μm) were close to the dimensions of the cells, and can thus guide adhesion and spreading at the single-cell level.
Since EDC crosslinked collagen has been shown to degrade substantially slower than uncrosslinked collagen substrates, the guidance effect can not only persist throughout the
3-week chondrogenesis period in vitro, but potentially also in vivo (Yahyouche, Zhidao et al. 2011). Phase contrast images of hMSCs in guidances channel (Figure 4-6) showed cell migrated and elongated along the channel length in smaller channels (25 to 100μm) after 7 days of culture whereas cell arranged randomly in larger channel (500 and 1000
μm) tended to aggregate into clumps after 7 days culture [Figure 4-6: A (c) and B (c)].
111
Cell clumps were observed in some area on larger channel as well as un-patterned substrate. Another manifestation of the cellular alignment was the orientation of actin fibers [Figure.4-8 and Figure 4-9] . The actin–myosin network is implicated in cell- spreading and migration. Alignment of actin fibers in the small channels thus indicates guidance at the cytoskeletal level. A further indication of alignment was the elongation and orientation of the nuclei parallel to the channel axis [Figure.4-7]. If the cell alignment is principally due to contact guidance through confinement by the substrate surfaces that are in contact with the cell, as suggested by our findings in the 25-50 μm wide channels, it is possible that the alignment can be propagated from cell to cell for some distance.
Kapoor et al used various dimension of microgroove (50,100 and 250 μm) on glass substrate to pattern tenocytes and he found cells in 50 and 100 μm grooves were aligned along the channel axis after 3 days while cells in 250 μm exhibited near-random orientation (Kapoor, Caporali et al. 2010) which is consistent with our result which suggest that the channel dimension of 25 to 100 μm have better guidance effect.
The effect of guidance channels on the ECM assembly was examined using type II collagen immunohistochemistry. Our results indicate that the type II collagen observed was indeed extracellular in nature. On CG membranes, the type II collagen immunoreactivity was clearly outside of the cell membrane, and even 100 μm channels
(which are up to approximately five cell bodies wide) showed alignment (angles of alignment 4.6° for PDMS and 8.6° for collagen substrates, Table 4-1) with the principal axis of the channel. The fiber structure of the type II collagen present suggests that it is also extracellular in nature [Figure 4-12]. Type II collagen was observed aligned along the length of the channels in smaller guidance channels (25 to 100μm) which corresponds
112 with cellular alignment. Similar experiments using contact guidance on tenocytes also found the collagen matrices produced by tenocytes were observed sinusoidal collagen fiber organization in 50 μm microgrooves whereas no collagen alignment showed 250
μm microgrooves (Kapoor, Caporali et al. 2010). which also suggest smaller guidance channel lead to induce oriented ECM. Our result shows the aligned cell secret aligned collagen matrix and this result confirm our hypothesis that microscale substrate features would cause differentiating MSCs to preferentially arrange themselves and to deposit an oriented cartilage ECM. Previous studies have shown that aligned cells may produce aligned collagen fibers (Hata, Hori et al. 1984; Wang, Grood et al. 2000; Wang, Jia et al.
2003). Here, we have demonstrated that MSCs align and elongate along the length of the guidance channels. Other studies have indicated that similarly elongated cells exhibited contractile forces along the direction of elongation(Hata, Hori et al. 1984; Wang, Grood et al. 2000; Wang, Jia et al. 2003) These types of forces have been shown to lead to the alignment of collagen fibersb (Eastwood, Mudera et al. 1998; Wang, Jia et al. 2003). In addition, other studies have demonstrated that collagen produced by moving cells is aligned with respect to the cell movements (Birk and Trelstad 1986; Wang, Jia et al.
2003). In this study, the guidance channels cause the MSCs to physically move and align via contact guidance. As the cells move to align and elongate along the channel length, we expect that they produce aligned collagen due to this movement and the contractile forces established along the direction of cell elongation.
Native cartilage has been shown to have a collagen architecture that imparts mechanical strength under compression and shear. In the 2D architecture of CG membranes, we were able to demonstrate alignment, but the geometry of the samples was unsuitable for
113 compressive testing. The mechanical properties of the samples under tension, however, indicate a strong dependence on alignment. Samples containing guidance features at the cellular scale led to superior mechanical function in the presence of cells. It should be noted [Figure 4-13: C25 and C100] that the channels themselves did not, at any channel width, improve the mechanical properties of the membranes. In addition, the GAG/DNA values [Table 4-2] for constructs of different channel widths were similar indicating that differences in the GAG content between conditions were not responsible for changes in mechanical properties of tissue cultured in different channels. This suggests that the cell- derived ECM under guidance accounts for the superior function.
Current TE construct have a tensile modulus value vary from 0.23 MPa to 5 MPa. For example, Fedewa et al have shown chondrocyte monolayer TE construct with a tensile modulus of 1 to 5 MPa after 8 weeks culture (Fedewa, Oegema et al. 1998); Gemmiti and
Guldberg et al. obtained 2.3 MPa for Chondrocyte monolayerafter after 17 days static culture (Gemmiti and Guldberg 2006); Williams et al. attained 0.01 MPa after 2 weeks culture for chondrocyte-seeded alginate (Williams, Klein et al. 2005); Gratz et al. reported that chondrocyte-seeded fibrin reached 0.64 MPa after 8 month in vivo repair
(Gratz, Wong et al. 2006) and Huang et al. showed chondrocyte or MSC-seeded agarose have a tensile modulus of 0.4 MPa after 56 days culture (Huang, Baker et al. 2012). In this work, we demonstrate tensile modulus for the construct containing smaller channels
(25 to 250 μm) of approximate 9.6 to 12.8 MPa which is higher than current TE construct listed above. The average tensile modulus for native articular cartilage is about 0.7–12.5
MPa (Almarza and Athanasiou 2004; Huang, Stankiewicz et al. 2005). The elasticity modulus for the construct containing smaller channels (25 to 250 μm) was approximate
114
9.6 to 12.8 MPa which is comparable to the highest value found in the literature for native articular cartilage (12.5 MPa). On the other hand, the elasticity modulus of the construct with randomly seeded MSCs was around 5 MPa, which is comparable to the inner annulus fibrosus (3 MPa). Thus, using these guidance channels, we can produce cartilage constructs with a range of tensile properties similar to native articular cartilage.
While the above study demonstrates that microscale guidance has an effect on hMSC differentiation and on the mechanical function of the resulting tissue, there are certain limitations. First, even though we have imposed some organization on the newly synthesized ECM, we do not yet replicate the fine structure or the mechanical function of native articular cartilage. Further, more work is needed to extend the current 2D findings to a 3D construct for preclinical experiments. To create a 3D construct for in vivo applications, one can either roll the current 2D constructs into 3D cylinders with appropriate microfeatures. In addition, one can also stack individual 2D constructs with
MSCs in between the layers. These 3D constructs can then be implanted into the site of articular cartilage injury in vivo. The Study for 3D construct will be detailed introduced in Chapter 4.
4.5 Conclusion
In this work, we test the hypothesis that microscale substrate features would cause differentiating MSCs to preferentially arrange themselves and to deposit an oriented cartilage ECM on a scale similar to native articular cartilage. The effect of microscale guidance on cell alignment and ECM function in a hMSC-based chondrogenesis model was investigated. Microchannels formed in collagen and PDMS were utilized to show
115 that MSCs cultured within the linear channels of smaller widths show alignment. Further, the alignment in cells and ECM leads to significant improvement in mechanical function.
These smaller channels provide a means of controlling cellular orientation and ECM production, which leads to enhanced mechanical properties in tissue engineered constructs. We demonstrate that microfabricated matrices can induce differentiating hMSCs to form oriented collagen microstructures and we test the mechanical strength of the ECM produced within the channels of various dimensions and proved that observed increase in cellular and collagen type II alignment within the smaller channels leads to enhanced mechanical strength of the resultant ECM. These results can be used to develop cartilage constructs with an aligned ECM and superior mechanical properties.
4.6 References
Ahmed, T. A. E. and M. T. Hincke (2010). "Strategies for articular cartilage lesion repair and functional restoration." Tissue Engineering Part B-Reviews 16(3): 305-329.
Almarza, A. J. and K. A. Athanasiou (2004). "Design characteristics for the tissue engineering of cartilaginous tissues." Annals of Biomedical Engineering 32(1): 2-17.
Aubin, H., J. W. Nichol, et al. (2010). "Directed 3D cell alignment and elongation in microengineered hydrogels." Biomaterials 31(27): 6941-6951.
Baskaran, H. S., L.A.; Berilla, J.; and Welter, J.F (2004). Assessment of mass transport limitations during cartilage tissue engineering. Transactions of the Orthopæ dic Research Society. 29(2): 721.
Baskaran, H. S., L.A.; Penick, K.; Berilla, J.; and Welter, J.F (2003). Substrate mass transport in tissue engineered cartilage. Proceedings of the Arthritis Research Conference. Keystone, CO: 70.
Benninghoff, A. (1925). "Form un bau der gelenkknorpel in ihren beziehungen zur funktion." Zeitschrift für Zellforschung und Mikroskopische Anatomie 2(5): 783-862.
116
Bentley, G. (2004). "A prospective, randomised comparison of autologous chondrocyte implantation versus mosaicplasty for osteochondral defects in the knee - Reply." Journal of Bone and Joint Surgery-British Volume 86B(4): 619-620.
Bhosale, A. M. and J. B. Richardson (2008). "Articular cartilage: structure, injuries and review of management." British Medical Bulletin 87(1): 77-95.
Birk, D. E. and R. L. Trelstad (1986). "Extracellular compartments in tendon morphogenesis - collagen fibril, bundle, and macroaggregate formation." Journal of Cell Biology 103(1): 231-240.
Brittberg, M., A. Lindahl, et al. (1994). "Treatment of deep cartilage defects in the knee with autologous chondrocyte transplantation." New England Journal of Medicine 331(14): 889-895.
Buckwalter, J. A. (2002). "Articular cartilage injuries." Clinical Orthopaedics and Related Research(402): 21-37.
Bullough, P. and J. Goodfellow (1968). "The significance of the fine structure of articular cartilage." The Journal of bone and joint surgery. British volume 50(4): 852-7.
Cancedda, R., B. Dozin, et al. (2003). "Tissue engineering and cell therapy of cartilage and bone." Matrix Biology 22(1): 81-91.
Caplan, A. I., M. Elyaderani, et al. (1997). "Principles of cartilage repair and regeneration." Clinical Orthopaedics and Related Research(342): 254-269.
Caplan, A. I. and V. M. Goldberg (1999). "Orthopaedic tissue engineering - Editorial comment." Clinical Orthopaedics and Related Research(367): S2-S4.
Chanlalit, C., C. Kasemkijwattana, et al. (2007). "Autologous chondrocyte implantation for traumatic large cartilage defect." Journal of the Medical Association of Thailand 90(7): 1435-1442.
Chen, A. C., W. C. Bae, et al. (2001). "Depth- and strain-dependent mechanical and electromechanical properties of full-thickness bovine articular cartilage in confined compression." Journal of Biomechanics 34(1): 1-12.
Chen, S. S., Y. H. Falcovitz, et al. (2001). "Depth-dependent compressive properties of normal aged human femoral head articular cartilage: relationship to fixed charge density." Osteoarthritis and Cartilage 9(6): 561-569.
Chou, C.-L., A. L. Rivera, et al. (2012). “Micrometer scale guidance of mesenchymal stem cells to form structurally oriented cartilage extracellular matrix. Tissue Engineering Part A, 19(9-10): 1081-1090.
117
Clar, C., E. Cummins, et al. (2005). "Clinical and cost-effectiveness of autologous chondrocyte implantation for cartilage defects in knee joints: systematic review and economic evaluation." Health Technology Assessment 9(47): III-+.
Clark, J. M. (1991). "Variation of collagen fiber alignment in a joint surface - a scanning electron-microscope study of the tibial plateau in dog, rabbit, and man." Journal of Orthopaedic Research 9(2): 246-257.
Clarke, I. C. (1971). "Articular cartilage a review and scanning electron microscope study part 1 the inter territorial fibrillar architecture." Journal of Bone and Joint Surgery British Volume 53(4): 732-750.
Cohen, N. P., R. J. Foster, et al. (1998). "Composition and dynamics of articular cartilage: Structure, function, and maintaining healthy state." Journal of Orthopaedic & Sports Physical Therapy 28(4): 203-215.
Curl, W. W., J. Krome, et al. (1997). "Cartilage injuries: A review of 31,516 knee arthroscopies." Arthroscopy 13(4): 456-460.
Dickinson, R. B., S. Guido, et al. (1994). "Biased cell-migration of fibroblasts exhibiting contact guidance in oriented collagen gels." Annals of Biomedical Engineering 22(4): 342-356.
Eastwood, M., V. C. Mudera, et al. (1998). "Effect of precise mechanical loading on fibroblast populated collagen lattices: Morphological changes." Cell Motility and the Cytoskeleton 40(1): 13-21.
Fedewa, M. M., T. R. Oegema, et al. (1998). "Chondrocytes in culture produce a mechanically functional tissue." Journal of Orthopaedic Research 16(2): 227-236.
Frenkel, S. R., B. Toolan, et al. (1997). "Chondrocyte transplantation using a collagen bilayer matrix for cartilage repair." Journal of Bone and Joint Surgery-British Volume 79B(5): 831-836.
Friedenstein, A. J. (1995). "Marrow stromal fibroblasts." Calcified Tissue International 56: S17-S17.
Gemmiti, C. V. and R. E. Guldberg (2006). "Fluid flow increases type II collagen deposition and tensile mechanical properties in bioreactor-grown tissue-engineered cartilage." Tissue Engineering 12(3): 469-479.
Gilbert, J. E. (1998). "Current treatment options for the restoration of articular cartilage." Am J Knee Surg 11(1): 42-6.
118
Gratz, K. R., V. W. Wong, et al. (2006). "Biomechanical assessment of tissue retrieved after in vivo cartilage defect repair: tensile modulus of repair tissue and integration with host cartilage." Journal of Biomechanics 39(1): 138-146.
Hasler, E. M., W. Herzog, et al. (1999). "Articular cartilage biomechanics: Theoretical models, material properties, and biosynthetic response." Critical Reviews in Biomedical Engineering 27(6): 415-488.
Hata, R. I., H. Hori, et al. (1984). "Selective-inhibition of type-i collagen-synthesis in osteoblastic cells by epidermal growth-FACTOR." Endocrinology 115(3): 867-876.
Huang, A. H., B. M. Baker, et al. (2012). "Sliding contact loading enhances the tensile properties of mesenchymal stem cell-seeded hydrogels." European Cells & Materials 24: 29-45.
Huang, A. H., M. J. Farrell, et al. (2010). "Long-term dynamic loading improves the mechanical properties of chondrogenic mesenchymal stem cell-laden hydrogels." European Cells & Materials 19: 72-85.
Huang, C. Y., A. Stankiewicz, et al. (2005). "Anisotropy, inhomogeneity, and tension- compression nonlinearity of human glenohumeral cartilage in finite deformation." Journal of Biomechanics 38(4): 799-809.
Huang, N. F., R. J. Lee, et al. (2010). "Engineering of aligned skeletal muscle by micropatterning." American journal of translational research 2(1): 43-55.
Hunziker, E. B. (2002). "Articular cartilage repair: basic science and clinical progress. A review of the current status and prospects." Osteoarthritis and Cartilage 10(6): 432-463.
Hunziker, E. B., M. Michel, et al. (1997). "Ultrastructure of adult human articular cartilage matrix after cryotechnical processing." Microsc Res Tech 37(4): 271-84.
Hwang, C. M., Y. Park, et al. (2009). "Controlled cellular orientation on PLGA microfibers with defined diameters." Biomedical Microdevices 11(4): 739-746.
Im, G. I., D. Y. Kim, et al. (2001). "Repair of cartilage defect in the rabbit with cultured mesenchymal stem cells from bone marrow." Journal of Bone and Joint Surgery-British Volume 83B(2): 289-294.
Janakiraman, V., B. L. Kienitz, et al. (2007). "Lithography technique for topographical micropatterning of collagen-glycosaminoglycan membranes for tissue engineering applications." J Med Device 1(3): 233-237.
Kapoor, A., E. H. G. Caporali, et al. (2010). "Microtopographically patterned surfaces promote the alignment of tenocytes and extracellular collagen." Acta Biomaterialia 6(7): 2580-2589.
119
Kasemkijwattana, C., S. Kesprayura, et al. (2009). "Autologous chondrocytes implantation with three-dimensional collagen scaffold." Journal of the Medical Association of Thailand 92(10): 1282-1286.
Kempson, G. E., M. A. R. Freeman, et al. (1968). "Tensile properties of articular cartilage." Nature 220(5172): 1127-&.
Liang, W. H., B. L. Kienitz, et al. (2010). "Concentrated collagen-chondroitin sulfate scaffolds for tissue engineering applications." J Biomed Mater Res A 94(4): 1050-60.
Magne, D., C. Vinatier, et al. (2005). "Mesenchymal stem cell therapy to rebuild cartilage." Trends in Molecular Medicine 11(11): 519-526.
Manwaring, M. E., J. F. Walsh, et al. (2004). "Contact guidance induced organization of extracellular matrix." Biomaterials 25(17): 3631-3638.
MMWR (2007). "National and state medical expenditures and lost earnings attributable to arthritis and other rheumatic conditions--United States, 2003." MMWR. Morbidity and mortality weekly report 56(1): 4-7.
MMWR (2010). "Prevalence of doctor-diagnosed arthritis and arthritis-attributable activity limitation --- United States, 2007-2009." MMWR. Morbidity and mortality weekly report 59(39): 1261-5.
Nehrer, S., M. Spector, et al. (1999). "Histologic analysis of tissue after failed cartilage repair procedures." Clinical Orthopaedics and Related Research(365): 149-162.
Rieppo, L., J. Rieppo, et al. (2012). Fourier transform infrared spectroscopic imaging and multivariate regression for prediction of proteoglycan content of articular cartilage. PLoS One. 7 (2):e32344
Roach, P., T. Parker, et al. (2010). "Surface strategies for control of neuronal cell adhesion: A review." Surface Science Reports 65(6): 145-173.
Schinagl, R. M., D. Gurskis, et al. (1997). "Depth-dependent confined compression modulus of full-thickness bovine articular cartilage." Journal of Orthopaedic Research 15(4): 499-506.
Shirazi, R., A. Shirazi-Adl, et al. (2008). "Role of cartilage collagen fibrils networks in knee joint biomechanics under compression." J Biomech 41(16): 3340-8.
Simon, L. S. (1999). "Osteoarthritis: a review." Clinical cornerstone 2(2): 26-37.
120
Solchaga L, T. E., Goldberg V, Caplan A, Welter J (2003). Tissue engineered cartilage implants: in vitro chondrogenic preconditioning. Proceedings of the 4th Symposium of the International Cartilage Repair Society 271.
Solchaga, L. A. (2002). Tissue engineered cartilage implants: optimizing construct assembly. Transactions of the Orthopaedic Research Society. 27: 483.
Solchaga, L. A. (2004). FGF-2 modifies the gene expression profile of bone marrow- derived human mesenchymal stem cells. Transactions of the Orthopaedic Research Society. 29: 777.
Solchaga, L. A., J. F. Welter, et al. (2004). "Generation of pluripotent stem cells and their differentiation to the chondrocytic phenotype." Cartilage and Osteoarthritis, Vol 1: Cellular and Molecular Tools 100: 53-67.
Temenoff, J. S. and A. G. Mikos (2000). "Review: tissue engineering for regeneration of articular cartilage." Biomaterials 21(5): 431-440.
Tessier-Lavigne, M. and C. S. Goodman (1996). "The molecular biology of axon guidance." Science 274(5290): 1123-33. van Susante, J. L. C., J. Pieper, et al. (2001). "Linkage of chondroitin-sulfate to type I collagen scaffolds stimulates the bioactivity of seeded chondrocytes in vitro." Biomaterials 22(17): 2359-2369.
Wakitani, S., T. Goto, et al. (1994). "Mesenchymal cell-based repair of large, full- thickness defects of articular-cartilage." Journal of Bone and Joint Surgery-American Volume 76A(4): 579-592.
Wang, J. H. C., E. S. Grood, et al. (2000). "Alignment and proliferation of MC3T3-E1 osteoblasts in microgrooved silicone substrata subjected to cyclic stretching." Journal of Biomechanics 33(6): 729-735.
Wang, J. H. C., F. Y. Jia, et al. (2003). "Cell orientation determines the alignment of cell- produced collagenous matrix." Journal of Biomechanics 36(1): 97-102.
Weiss, P. (1934). "In vitro experiments on the factors determining the course of the outgrowing nerve fiber." Journal of Experimental Zoology 68(3): 393-448.
Weiss, P. (1945). "Experiments on cell and axon orientation in vitro; the role of colloidal exudates in tissue organization." The Journal of experimental zoology 100: 353-86.
Welter, J. (2002). Tissue engineered cartilage implants: In vitro chondrogenic preconditioning. Proceedings of the 2nd Annual workshop of the Midwestern Tiss. Eng. Consortium: 21.
121
Welter, J. (2003). MSC-based Tissue engineered cartilage: Effects of construct assembly method and perfusion rates. Arthritis Research Conference. Altanta, GA.
Welter, J. (2004). Modulation of growth and differentiation conditions enhances chondrogenesis in large tissue engineered constructs. Transactions of the Orthopaedic Research Society. 29(2): 719.
Williams, G. M., T. J. Klein, et al. (2005). "Cell density alters matrix accumulation in two distinct fractions and the mechanical integrity of alginate-chondrocyte constructs." Acta Biomaterialia 1(6): 625-633.
Wilson, W., J. M. Huyghe, et al. (2007). "Depth-dependent compressive equilibrium properties of articular cartilage explained by its composition." Biomechanics and Modeling in Mechanobiology 6(1-2): 43-53.
Wilson, W., C. C. van Donkelaar, et al. (2005). "A fibril-reinforced poroviscoelastic swelling model for articular cartilage." Journal of Biomechanics 38(6): 1195-1204.
Wilson, W., C. C. van Donkelaar, et al. (2005). "Stresses in the local collagen network of articular cartilage: a poroviscoelastic fibril-reinforced finite element study (vol 37, pg 357, 2004)." Journal of Biomechanics 38(10): 2138-2140.
Woo, S. L. Y., W. H. Akeson, et al. (1976). "Measurements of nonhomogeneous, directional mechanical properties of articular cartilage in tension." Journal of Biomechanics 9(12): 785-791.
Yahyouche, A., X. Zhidao, et al. (2011). "Macrophage-mediated degradation of crosslinked collagen scaffolds." Acta Biomater 7(1): 278-86.
122
Chapter 5
DESIGN OF LARGE-SCALE 3-DIMENSIONAL CARTILAGE CONSTRUCTS WITH MICROCHANNELS FOR PRECLINICAL STUDIES
5.1 Introduction
Tissue engineering has been proposed as a promising method for the development of cartilage constructs to treat articular cartilage injuries. Most of these approaches involve the traditional tissue engineering strategy of culturing scaffolds seeded with mesenchymal stem cells (MSCs) or chondrocytes and growth factors to produce cartilage constructs in vitro (Wang, Kim et al. 2005; Djouad, Mrugala et al. 2006; Wang, Blasioli et al. 2006; Zwingmann, Mehlhorn et al. 2007; Dickhut, Gottwald et al. 2008; Janjanin,
Li et al. 2008; Yang, Peng et al. 2008; Chung and Burdick 2009; Spadaccio, Rainer et al.
2009; Jakobsen, Shandadfar et al. 2010; Liang, Kienitz et al. 2010; Gong, Xue et al. 2011;
Xue, Gong et al. 2012). These tissue engineered constructs may show biochemical similarities to native cartilage. Many studies have shown that when exposed to chondrogenic factors, MSCs produce extracellular matrix (ECM) rich in collagen type II and glycosaminoglycans (GAG), which are two of the primary ECM components of native articular cartilage. However, these tissue engineered cartilage constructs are biomechanically inferior to native cartilage and are not suitable for physiological load bearing function. This limitation has been attributed to a lack of an oriented microscale
123 structure in tissue engineered constructs, which is present in the native tissue and prevents these constructs from reaching a clinically viable tissue. In our previously published work (Chou, Rivera et al. 2012), we showed that microscale guidance channels can be used to align hMSCs, which then deposited aligned ECM with enhanced mechanical properties. These two-dimensional constructs containing guidance channels with oriented hMSCs and aligned ECM show superior biomechanical properties compared to constructs lacking guidance channels that have randomly-seeded cells. The alignment of cells and ECM lead to significantly higher modulus of elasticity values under tension when compared to randomly seeded constructs. However, these two- dimensional constructs do not provide a solution for a 3-dimensional cartilage defect, and therefore, lack clinical relevance. In this study, we investigated a method of scaling these two-dimensional constructs to 3-dimensional tissue engineered constructs to overcome this limitation.
Several studies showed that defects less than a critical size of 3 mm in diameter may lead to complete spontaneous repair of articular cartilage while larger defects do not repair completely (Convery, Akeson et al. 1972; Shapiro, Koide et al. 1993; ButnariuEphrat,
Robinson et al. 1996; Jackson, Lalor et al. 2001). Larger defects result in damage to the
ECM and chondrocytes that cannot be self-repaired. Any self-repair attempts for larger defects ultimately result in a fibrocartilage repair tissue that lacks the mechanical properties of native articular cartilage and eventually fails (Hunziker 2002). Tissue engineering serves as a promising method to provide 3-dimensional cartilage constructs to fill defects larger than this critical size; however, the creation of tissue engineered cartilage greater than this size remains a challenge. Other studies have investigated the
124 creation of 3-dimensional tissue engineered cartilage constructs by using stem cells and various types of scaffolds including poly(e-caprolactone), poly(L-lactic acid), collagen type I, hyaluronic acid hydrogels, gelatin, alginate, and agarose (Awad, Wickham et al.
2004; Li, Tuli et al. 2005; Janjanin, Li et al. 2008; Bian, Zhai et al. 2011; Huang, Farrell et al. 2011; Ousema, Moutos et al. 2012). However, none of these studies have succeeded in creating engineered cartilage constructs that display the compressive properties of native hyaline cartilage (Awad, Wickham et al. 2004; Janjanin, Li et al. 2008; Bian, Zhai et al. 2011; Huang, Farrell et al. 2011; Ousema, Moutos et al. 2012). While adult human hyaline cartilage shows depth dependant mechanical properties and a compressive elasticity modulus that varies between 1.16-7.75 MPa, tissue engineered cartilage typically displays a compressive elasticity modulus between 10-15 KPa (Chen, Falcovitz et al. 2001; Awad, Wickham et al. 2004). For example, Awad et al. investigated the creation of tissue engineered cartilage constructs using adipose derived adult stem cells within gelatin, agarose, or alginate scaffolds under chondrogenic conditions for 28 days.
They performed compression testing on the samples to determine their equilibrium
Young’s modulus at different time points in culture, and found that for all three scaffolds, the resulting tissue construct’s equilibrium Young’s modulus was approximately 10-15 kPa (Awad, Wickham et al. 2004). In one study, Chung et al. achieved an equilibrium compressive modulus of 60 kPa for tissue engineered cartilage constructs developed from human MSCs within hyaluronic acid hydrogels; however, this value is still far from the value for native articular human cartilage (Chung and Burdick 2009). With this limitation in mind, the objective of this study was to develop a new method that translates the results of the basic micro-scale science we investigated in our previous study (Chou,
125
Rivera et al. 2012) into large-scale, clinically useful cartilage constructs with an ultrastructure similar to that of the native tissue shown in Figure 5-1.
As described in the previous study (chapter 4), channels of varying microscale dimensions in type I collagen–based membranes were formed via a combination of microfabrication and soft-lithography. By rolling up the 2-dimensional MSC seeded collagen-GAG (CG) based scaffold and allowing it to undergo subsequent chondrogenic differentiation, we formed 3-dimensional rolled-up cell layered cartilage constructs with microscale architecture to guide the differentiating MSCs to produce oriented ECM. Via this method, we aimed to direct the MSCs to produce ECM aligned perpendicular to the diameter of the construct in order to achieve a structure similar to the deep zone of native articular cartilage. In the deep zone of native articular cartilage, the perpendicular arrangement of collagen fibrils relative to the subchondral bone has been shown via mathematical modeling to enhance its compressive properties (Wilson, Huyghe et al.
2007; Shirazi, Shirazi-Adl et al. 2008). After culturing these rolled 3-dimensional constructs in TGF-β1 containing chondrogenic medium for 21 days, histology and immunohistochemistry indicated extensive GAG and collagen type II production, which are both indicative of chondrogenesis. Our results show that the microscale guidance channels incorporated within the 3-demensional cartilage constructs lead to the production of aligned cell-produced collagenous matrix and enhanced mechanical function.
126
Figure 5-1. Schematic of hMSC-CG Based rolled-up large scale construct formation
5.2 Materials & Methods
5.2.1 Materials
Collagen type I from bovine Achilles tendon, chondroitin-6-sulfate sodium salt from shark cartilage, 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDC),
N-hydroxy-succinimide, fetal bovine serum (FBS), 4', 6-diamidino-2-phenylindole
127
(DAPI), and phalloidin were purchased from Sigma Chemical Co. (St. Louis, MO).
Antibiotic-antimycotic cocktail, low glucose Dulbecco’s Modified Eagle Medium
(DMEM), high glucose DMEM, ascorbate 2-phosphate (A2P), dexamethasone, sodium pyruvate, and human plasma fibronectin were obtained from Gibco, Invitrogen (Carlsbad,
CA). Fibroblast growth factor-2 (FGF) and transforming growth factor β1 (TGF- β1) were purchased from Peprotech (Rocky Hill, NJ). ITS(insulin, transferrin, selenium)+
Premix Tissue Culture Supplement were obtained from Becton Dickinson (Franklin
Lakes, NJ), calcein AM, and ethidium homodimer-1 (EthD-1) were obtained from
Molecular Probes (Invitrogen, Carlsbad, CA). Fluorescein isothiocyanate (FITC)- conjugated goat anti-mouse IgG secondary antibody was from MP Biomedicals (Irvine,
CA,) and texas red conjugated goat anti-mouse IgG secondary antibody was from
Molecular Probes (Invitrogen, Carlsbad, CA) while the collagen Type I and Type II primary antibodies were obtained from the Developmental Studies Hybridoma Bank
(University of Iowa). Sterile 27" poly- glycolic acid (PGA) absorbable sutures was obtained from Integra Miltex (York, PA )
5.2.2 CG Scaffold fabrication
5.2.2.1 CG solution and CG membrane fabrication
The base CG solution was made using a method adopted from Yannas, et al. (Yannas,
Burke et al. 1980) and CG membranes were fabricated using a previously published filtration method (Janakiraman, Kienitz et al. 2007) as described in chapter 3.2.2. Various thicknesses of CG membranes were produced by controlling the amount of CG solution and the filtration time. Membranes of 100-800 µm were fabricated in this study. The silicon wafers containing the microchannle patterns were then used to produce channels
128 on the surface of the CG membranes by the technique of collagen soft-lithography, which involved selective solubilization, patterning, and pattern stabilization through EDC crosslinking as detailed in chapter 3.2.5.
5.2.2.2 Concentrated CG solution and CG Sponge fabrication
CG solution was made using the previously described homogenization method.. The concentrated CG solution was produced using a published technique that our laboratory developed (Liang, Kienitz et al. 2010). Briefly, 40 ml of collagen solution was centrifuged in an Oak Ridge centrifugal tube (Nalge, Rochester, NY) for 30 mins at
38720 g (r.c.f) in the superspeed refrigerated centrifuge (RC-5C, Sorvall Instruments,
DuPont, Wilmington, DE). A total of 30ml of supernatant fluid was removed from the each tube and the remaining supernatant was remixed thoroughly with collagen sediment.
The remixed solution was placed in a 100 mm2 petri-dishes, degassed and placed in a flat aluminum container. The concentrated solution was frozen from 4°C to -40°C for 120 minutes and at -40°C overnight. The samples were then lyophilized (VirTis AdVantage
EL, SP Industries, Inc., Warminster, PA) for 24 hours and were physically crosslinked via a dehydrothermal process of dehydration at 120°C under a vacuum of 25 mTorr for one day. CG sponges were then stored under sterile conditions at 4°C.
5.2.3 Design of Microchannels
Templates for rectangular channels of constant length, width, and spacing were created using AutoCAD LT (AutoDesk, San Rafael, CA) (Figure 5-2). Channels of 100 µm width,r 50 or 100 µm spacing, and 1.5 cm length were used in this study. The depth of the channels was maintained constant at 70 µm. To form these channels on a silicon template, standard microfabrication methods were used as detailed in Chapter 3.2.4 .
129
Figure 5-2. Microchannel pattern on a silicon wafer for the rolled-up constructs. A single channel width of 100 µm with the spacings of 50 or 100 µm were used in this study
5.2.4 Autonomous Rolled-up CG-hMSC Construct (ARCGs) Formation
CG membranes were cut to 18 mm length x 15 mm width and sterilized in 10x antibiotic- antimycotic cocktail (Gibco) for one day and 1x antibiotic-antimycotic cocktail (Gibco) for another day prior to cellular seeding. CG membranes were rinsed with PBS twice and
2.1 million hMSCs were seeded on each construct. CG membrane Thicknesses of 250 and 800 µm were used in this experiment. After 2 hours at 37 °C and 5% CO2 to allow for cell attachment, the seeded constructs were submerged in chondrogenic differentiation medium: Dulbecco’s Modified Eagle’s Medium, High (4.5 g/L) Glucose
(DMEM-HG) supplemented with 10% ITS+ Premix Tissue Culture Supplement, 10-7 M dexamethasone, 1 μM ascorbate-2-phosphate, 1% sodium pyruvate, and 10 ng/ml transforming growth factor-beta 1 (TGF-β1). The constructs were cultured in a standard incubator at 37 °C and 5% CO2 in humidified air. Medium was changed every 2-3 days.
130
5.2.5 Induced Rolled-up CG-hMSC construct (IRCGs) Formation
CG membranes of 100 and 250 µm thicknesses and CG sponges of 100 and 500 µm thicknesses were used in this experiment. The silicon wafers containing the patterns in
Figure 5-2 were then used to produce channels on the surface of the CG membranes and sponges by a technique of collagen soft-lithography, which involved selective solubilization, patterning, and pattern stabilization through EDC crosslinking as detailed in chapter 3.2.5. Prior to cellular seeding, the CG membranes and sponges were then sterilized in 10x antibiotic-antimycotic cocktail (Gibco) for one day and 1x antibiotic- antimycotic cocktail (Gibco) for another day, rinsed with PBS twice, and incubated at 37
°C and 5% CO2 in humidified air for 3 hours. Adult human MSCs were cultured to 90% confluence in Dulbecco’s Modified Eagle’s Medium, Low (1.5 g/l) Glucose (DMEM-LG) containing 10% FBS and 10 ng/ml FGF-2. The MSCs were then trypsinized and resuspended at approximately 50 × 106 cells/ml. The cell suspension was applied to the patterned surface of the collagen membrane (1.8 cm width by 7.2 cm length) using a micropipette. A total of 8.4 million cells were seeded per construct. After a 2 hour period to allow for cell attachment, the patterned collagen membrane seeded with MSCs was subsequently rolled- up using two forceps to create a multi-layer spiral cylinder construct.
Two sterile polyglycolic acid (PGA) absorbable sutures were tied around the construct to maintain the rolled structure by preventing unfolding of the layers as shown in Figure 5-9
(E) . The dimensions for the rolled-up construct were approximately 1.8 cm (length) x 3.5 cm (diameter). All samples were cultured in DMEM containing TGF-β1 (10 ng/ml) for
21 days.
131
5.2.6 Compression Testing
To determine whether alignment led to improved mechanical function, IRCGs were subjected to compressive testing. Collagen membranes with hMSCs cultured for 21 days under differentiating conditions were cut to four sections using a razor blade, and the thickness and diameter of the samples were measured [Figure. 5-4]. The compressive modulus of each sample was determined in a Rheometrics Solids Analyzer II compression device (Piscataway, NJ). Each sample was compressed for 10 s at a strain rate of 1% s-1, and compression tests were performed on 6samples per condition. Stress (σ) vs. strain (ε) data were recorded by a computer data acquisition system and transferred to
Excel and fitted to the equation to yield the modulus of elasticity ( ) of the
IRCG constructs (Woo, Akeson et al. 1976; Liang, Kienitz et al. 2010).
Furthermore, we calculated the compressive modulus of elasticity ( ) of the tissue of the
IRCG constructs. To do this, we found the ratio of tissue to construct tensile modulus of elasticity from our previous study and assumed that the compressive modulus of elasticity would display a similar ratio. From the data we obtained from the tensile testing in chapter 3, we calculate tissue tensile modulus according to the following equation:
EAEAEAc c t t s s ………………………………………………………………………(1)
Where Ec :Construct tensile modulus of elasticity; Et :Tissue tensile modulus of elasticity;
Es :Scaffold modulus of elasticity, Ac : Construct cross-sectional area; At : Tissue cross-
sectional area; As : Scaffold cross-sectional area
The relationship between the tensile modulus of elasticity of the tissue and the guidance channel width was obtained [ Figure 5-3]. From these values, for each channel width
132 used, we could calculate the ratio of the tissue tensile modulus of elasticity to the tensile modulus of elasticity of the scaffold alone. Membranes with 100µm channels and without channels were used for these IRCG constructs. From tensile testing data and equation (1), we obtained the ratio of the tensile modulus of the tissue to the scaffold, which was 20.58 MPa for the 100 µm guidance channel constructs and 5.34 MPa for the randomly seeded (without channels) constructs. The ratio of the compressive modulus of the tissue to the scaffold for the IRCG constructs was assumed to be the same as tensile modulus ratio. The ratios were applied to equation (1) combined with the constructs’ compressive modulus data to obtain the compressive modulus of elasticity for the tissue.
50
40
30 (MPa)
20 Modulus of Elasticity of Modulus Elasticity 10 0 200 400 600 800 1000 Guidance Length (µm)
Figure 5-3. Tissue tensile modulus of elasticity under microscale guidance. Tissue tensile modulus under contact guidance was quantified and shown as the mean for channels of various widths.
133
Figure 5-4. Schematic of rolled-up construct for compressive testing. hMSC- collagen rolled-up constructs of 1.8 cm (length) x 3.5 mm (diameter) were cut to four sections. Each sample was compressed for 10 s at a strain rate of 10% s-1
5.2.7 Histological and Immunohistochemistry Analysis
Collagen-hMSC constructs were fixed with 4% paraformaldehyde overnight at 4°C.
Samples were rinsed with PBS 3 times and kept in PBS at 4°C. Tissues samples were embedded in Paraplast, sectioned to 5 µm slices, and stained with toluidine blue at the
Case Skeletal Research Center for histology analysis. Five μm thick sections were deparaffinized and stained for type I and type II collagen by immunohistochemistry.
Antigen unmasking was performed with 1 mg/mL pronase in PBS for 15 minutes at room temperature. After washing the samples with PBS twice for 30 minutes, they were then blocked with 10% normal goat serum (NGS) in PBS for 30 minutes. The primary antibodies were diluted 1:50 for type I and 1: 50 for type II in 1% NGS in PBS. These antibodies were applied to the samples for 1 hour to stain for collagen type I and type II.
The samples were then washed with PBS (2 × 30 minutes). Fluorescein isothiocyanate
(FITC)-conjugated goat anti-mouse IgG secondary antibody, diluted 1:500 in 1 % NGS in PBS, was then applied to all samples for 45 minutes. The samples were again washed
134 with PBS for 1 hour and wet-mounted using 5% N-propyl gallate in glycerol. The samples were imaged using a SPOT RT digital camera attached to a Leica fluorescence microscope (Wetzlar, Germany).
5.2.8 Collagen Fibers Alignment Analysis
Cell produced ECM inside the guidance channels of the constructs was observed by polarized light microscopy and second harmonic generation (SHG) microscopy (Leica
Microsystems GmbH, Wetzlar, Germany) to observe the collagen fiber alignment. For polarized light microscopy, the IRCG100 constructs were fixed with 4% paraformaldehyde overnight and embedded in paraffin. Sections of 15 µm thickness were stained with Sirius-Red. Tissue in the channel was imaged under polarized light microscopy using a polarized 20X field objective.
5.2.9 Metabolism Analysis
Aliquots of culture medium (n=5 per condition) were sampled every other day during medium change and were stored at -20°C for glucose and lactate content measurement.
Glucose concentration in the medium was determined using an Accu-Chek Aviva meter
(Roche Diagnostics, Indianapolis, IN), which measures a range from 10 to 600 mg/dL.
Lactate concentration was obtained using a Lactate Scout meter (SensLab GmbH Leipzig,
Germany), which measures a range from 0.5 to 25 mmol/l. Ten µl of medium were applied to the glucose strip and 20 µl of medium were applied to the lactate strip on the meter to read the value.
135
5.2.10 Statistical Methods
For all quantitative results, statistical analysis was carried out using the Origin 8.5.1
(Origin Lab, Northampton, MA) software package. Pairwise comparisons were performed by Tukey’s test to compare data groups. A p value of less than 0.05 was used to determine statistical significance. Sample sizes are indicated in the respective figure legends.
5.3 Result
5.3.1 Autonomous Rolled-Up CG-hMSC constructs (ARCGs)
After 4 days in culture, an interesting phenomenon was observed for hMSCs seeded on
1000 μm thick CG membranes (ARCG1000). After hMSC adhesion on the CG scaffold and exposure to chondrogenic medium for 4 days, the contructs rolled-up on their own, which we feel is caused by the contractile forces produced by hMSCs during chondrogenesis.. We believe a slight mismatch in forces at the different ends leads to the roll-up of the membranes. [Figure 5-5 (A)]. The ARCG1000 constructs were subjected to histological analysis for ECM distribution and cell morphology after 3 weeks in chondrogenic culture . Toluidine blue stained sections showed the internal structure of the ARCG1000 constructs and the presence of GAG within the tissue [Figure 5-5 (B)-
(C)]. Mature cartilage tissue with an average thickness of 248.93μm was formed within the ARCG1000 constructs. Longitudinal sections werestained via immunohistochemistry for collagen type I and type II [Figure 5-5 (D)-(E)]. Immunohistochemistry data showed that type II collagen (green) [Figure 5-5 (D)] was extensively expressed in the tissue layers while type I collagen (red) [Figure 5-5 (E)] was expressed in the scaffold layer,
136 suggesting dense ECM indicative of chondrogenesis is deposited within the scaffold of the ARCG1000 constructs. However, the ARCG 1000 constructs showed poor integration between the tissue and the scaffold [Figure 5-5 (B)-(C)] and only formed one spiral in the construct, which may limit the use of thick scaffold for clinical use.
137
Figure 5-5. Autonomous rolled-up collagen-hMSC constructs, ARCG1000 (A) cultured in chondrogenic medium for 3 weeks. Thickness of the CG membrane: 1000 μm. Toluidine blue stained cross section (B) and longitudinal section (C). Longitudinal section (C) was immunohistochemistry stained with collagen types I (D) and II (E). Green is type II collagen and red is type I collagen. Scale Bar: 100 µm (D-E)
To improve tissue-scaffold integration, 250 μm thick CG membrane scaffolds were used for the autonomous roll-up procedure. Our data showed that ARCG250 constructs with thinner scaffolds (250 µm) yielded better tissue and scaffold integration [Figure 5-7 (A)-
(B)] with a higher tissue/scaffold ratio (1.11). In addition, ARCG250 constructs contracted significantly more over time [Figure 5-6 (B/E) and (C/F)] when compared to the ARCG1000 constructs [Figure 5-5 (A)]. ARCG250 constructs were halfway folded
[Figure 5-6 (B)] by day 7 and completely folded by day 21 [Figure 5-6 (E)]. However, the distribution of ECM within the ARCG250 constructs was not evenly deposited
[Figure 5-7 (A)-(B)] and only formed 2 layers of spiral tissue within the construct [Figure
5-6 (D), Figure 5-7 (A)]. With the autonomous roll-up method, the size and shape of the
ARCG 250 constructs were not controllable [Figure 5-6: (E) is 1.5 cm (length) and (F) is
2 cm (length)], and only a maximum of 2 rolled-up cell layers can be achieved in these
ARCG constructs.
138
Figure 5-6. Autonomous rolled-up CG-hMSC constructs. 2 hours after seeding (A). 7 days in culture (B-C). 21 days in culture (D-F). B, D and E images were taken from the same construct while C and F images were taken from another single construct.
Figure 5-7. Autonomous rolled-up CG-hMSC constructs with 250 μm thick CG membranes after 3 weeks in culture. Toluidine blue-stained cross section (A) and longitudinal section (B). Scale Bar: 100 µm (A-B). Corresponding construct images are shown in Figure 5-6 (D)
139
5.3.2 Induced Roll-up CG-hMSC constructs (IRCGs)
To overcome the limitations of the ARCG constructs and to create a multi-layered (more than 2 layers) large hMSC-collagen construct, we developed a technique to manually roll-up CG thin membranes (100 and 250µm) and CG sponges (100µm and 500 µm) to create multi-layered induced rolled-up CG-hMSC constructs (IRCG). We first successfully formed multi-layer IRCG constructs with the 250µm CG membrane scaffolds (IRCG250) [Figure 5-8 (A)]; however, the histology data of the IRCG250 constructs revealed sparsely distributed GAG production (as stained by Toluidine blue ), which suggests that the ECM only deposited in very small regions of the construct
[Figure 5-8 (B-C)]. Overall, 95% of the IRCG250 constructs were not positively stained for GAG after 21 days in chondrogenic culture [Figure. 5-8 (C)]. We attribute this lack of
GAG production to the large thickness (250 µm) of the membrane, which resulted in poor mass transport of nutrients and signaling molecules such as TGF-ß1 to the MSCs within the IRCG250 constructs.
Figure 5-8. IRCG250 constructs after 3 weeks in culture (A). Toluidine blue stained cross section (B) and longitudinal section (C). Scale Bar: 1mm (A) and 100 µm (B-C)
140
To solve the nutrient transport limitation caused by the thick membrane, we further optimized the thickness of CG membrane to 100 μm for forming IRCG constructs
(IRCG100). The thinner 100 µm CG membrane facilitates nutrient transport in the
IRCG100 while still supporting cell attachment during the roll-up procedure. Two different features were used for patterning the CG scaffolds for different constructs as shown in Figure 5-2. One construct is the IRCG100 construct with a CG membrane scaffold featuring 100µm channels with spacings of 50µm(IRCGs100-100C_50S), while the other construct is the ICRS construct with a CG membrane scaffold featuring 100µm channels with spacings of 100µm (IRCG100-100C_100S). A CG membrane patterned with 100µm channels and 100 spacings is shown in Figure. 5-9 (B). hMSCs were seeded on the CG scaffolds, rolled-up, and cultured in chondrogenic medium with TGF-β1 (10 ng/ml) for 3 weeks. A IRCG100-100C_100S construct under static culture conditions is shown in Figure. 5-9 (E). After 21 days, multi-layered IRCG100-100C_100S constructs were formed, and up to 7 l tissue-scaffold layers were formed in the IRCG100-
100C_100S constructs [Figure 5-9 (D)].
141
Figure 5-9. Patterned 100 μm thick CG membrane scaffolds featuring 100 μm channels with 100 spacings (A-C). Multi-layered IRCG100-100C_100S immersed in chondrogenic medium (E) and after 21 days chondrogenic culture (D)
For Histological analysis, longitudinal sections [Figure 5-10 (A)-(C)] and cross sections
[Figure 5-10 (a)-(c)] of IRCG100 constructs were first stained with toluidine blue.
Intense toluidine blue staining was demonstrated in the outer 3 tissue layers of the
IRCG100 constructs [Figure 5-10(A)-(C), (a)-(c)]. Cells were distributed throughout the first 3-4 layers of the scaffold. Robust deposition of GAG (as stained by toluidine blue) was present in the first 3 scaffold layers and parts of the fourth layer. Similar results were found in all constructs (IRCG100-100C_100S, IRCG100-100C_50S and IRCG100 without channels) [Figure 5-10]. A Sparse number of cells and amount of ECM were distributed after the fourth layer. ECM deposition in constructs with microchannels
(IRCG100-100C_100S, IRCG100-100C_50S) were more homogenously distributed in the outer 3 tissue layers. In contrast, constructs without the channels displayed non- uniform ECM distribution. Similar results were observed in both cross section (outer to
142 inner layer) [Figure 5-10 (a)-(c)] and longitudinal section (edge to center layer) [Figure 5-
10 (A)-(C)].
Figure 5-10. Histological appearance of induced rolled-up CG-hMSC constructs (IRCS100). IRCG100-100C_50S (A and a), IRCG100-100C_100S (B and b) and Control IRCGs (C and c) after 3 weeks chondrogenic culture. Toluidine blue-stained longitudinal sections (A-C) and cross sections (a-c). Scale Bar: 100 µm (A-C) and 1mm
143
(a-c). IRCG100-100C_50S: IRCG constructs composed of 100 μm thick scaffolds with 100 μm channels and spacings of 50 μm (A and a); IRCG100-100C_100S: IRCG constructs composed of 100 μm thick scaffolds with 100 μm channels and spacings of 100 μm (B and b); Control IRCG100: IRCG constructs composed of 100 μm thick scaffolds without channels.
Immunohistochemistry data of the IRCG100 constructs cultured for 21 days in chondrogenic medium showed strong positive staining for collagen type II [Figure 5-11
(D); Figure 5-12 (B)]. Dense ECM rich in both GAG (as shown by toluidine blue staining)
[Figure 5-11 (B-C); Figure 5-12 (A)] and type II collagen (as shown by type II collagen immnohistochemistry staining) [Figure 5-11 (D); Figure 5-12 (B)] throughout the outer 3 layers of the constructswere displayed. ECM accumulated and filled the gap between the
CG scaffold layers, displaying excellent tissue-scaffold integration.
144
Figure 5-11. Cross section of the induced rolled-up CG-hMSC construct (IRCG100- 100C_100S) with guidance channles. Toluidine blue-stained cross section (A-C). Immunohistochemistryl staining of type II collagen (D). Arrowheads point to the guidance channels. Scale Bar: 1mm (A) and 100 µm (B-D)
145
Figure 5-12. Histology and Immunohistochemistry an induced rolled-up CG-hMSC construct. IRCG100-100C_50S construct cultured in chondrogenic medium for 3 weeks. Toluidine blue stained longitudinal section (A); immunohistochemistryl collagen type II stained longitudinal section (B) . Green is type II collagen. Scale Bar: 100 µm (A-B)
We also used CG sponges as another type of scaffold for the induced roll-up experiments.
Human MSCs were seeded in CG sponges of 100µm (IRCS100) and 500 µm (IRCS500) thickness and formed induced rolled-up CG Sponge-hMSC constructs. These IRCS constructs were stained with Toluidine blue. For the IRCS500 constructs, intense toluidine blue staining was observed in the outer first tissue layer and weak staining was seen in the second layer [Figure 5-13 (B), (b)]. ECM of an average thickness of 748 µm
146 was deposited in the first sponge scaffold layer and sparse tissue was observed in the second layer of the IRCS500 constructs [Figure 5-13 (B), (b)]. On the other hand, ECM with an average thickness of 104 µm was distributed throughout the outer 2 layers of the sponge scaffolds in the IRCS100 constructs [Figure 5-13 (A), (a)], which suggests that thinner CG sponge scaffolds yield thinner and more evenly-distributed ECM. In addition, the central region of the IRCS constructs contained relatively thicker ECM when compared to the outer the edge regions. However, the CG sponges are soft biomaterials with lower mechanical properties than the CG membranes and, only 2 layer constructs can be fabricated after the roll-up process.
Figure 5-13. Histological appearance of the induced rolled-up CG sponge-hMSC construct after 3 weeks in culture. Toluidine blue stained longitudinal section (A) and cross section (a) of a IRCS100 construct; toluidine blue stained longitudinal section (B) and cross section (b) of a IRCS500 construct. Scale Bar: 100 µm (A), 250 µm (B) and 1mm (a-b).
147
5.3.3 Collagen Fibrils Alignment Analysis
Collagen fibril structure in the 100 µm linear guidance channels was observed under second harmonic generation (SHG) microscopy after 3 weeks in chondrogenic culture.
Our data showed that dense collagen fibrils were produced by the cells after 3 weeks in culture and were aligned along the channel length [Figure 5-14 (A)] within guidance channels, in contrast to randomly oriented collagen fibrils produced by hMSCs on constructs without channels [Figure 5-14 (B)].
Figure 5-14. Images of ECM collagen produced by hMSCs within linear channels of widths 100µm (A) and hMSCs randomly seeded on constructs without channels (B) using SHG microscopy after 3 weeks in chondrogenic culture. Dense collagen fibrils were produced by the cells and aligned along the channel length direction (A) within the guidance channels, which is contrasted with random alignment of collagen fibrils on constructs without channels (B). Arrowheads point to the aligned collagen fibrils within the guidance channels. Scale bar: 100µm
148
Figure 5-15 shows longitudinal sections (A-B) and a cross section (C) of cell produced collagenous matrix of the IRCG100 constructs. Figure 5-15 (A) and (B) shows Sirius-red stained longitudinal sections of the IRCG100 constructs. Cell produced ECM within the guidance channels was observed using polarized light microscopy [Figure 5-15 (A)].
Thick collagen fibrils (orange and yellow) [Arrowheads in Figure 5-15 (A) (B)] and thin fibrils (green or yellow green) [Figure 5-15 (A)] were produced in the IRCG100 constructs. Importantly, cell produced collagen fibrils were aligned along the length of the 100 µm guidance channels [Figure 5-15 (B)]. Figure 5-15 (C) shows the type II collagen immunohistochemistry staining of a cross section of an IRCG100-100C_50S construct (containing 100 µm channels and 50 µm spacings). Dense cell produced type II collagen was displayed within the 100 µm guidance channels [Figure 5-15 (C)].
Figure 5-15. Sirius-red stained longitudinal section of IRCG100 constructs (A-B). Guidance channels within IRCG100 constructs were observed using polarized light microscopy(A-B). Type II collagen immunohistochemistry staining of a cross section of
149 an IRCG100-100C_50S construct (C). Orange and yellow regions are thick collagen fibrils; green and yellow green regions are thin fibrils; black dots are cell nuclei; * represents the CG membrane. Arrowheads point to the thick cell-produced collagen fibrils. Thick collagen fibrils were produced and aligned along the guidance channel (A- B). Cell produced type II collagen within guidance channels is shown (C). Scale bar: 50µm.
5.2.5 Mechanical Properties of IRCG100 Constructs
Figure 5-16 shows the compressive modulus of the IRCG100 constructs [Figure 5-16
(A)] and the compressive modulus of the tissue [Figure 5-16 (B)] within the constructs.
IRCG100 constructs were subjected to compression along the direction of the channel length. Each sample was compressed for 10 s at a strain rate of 10% s-1, and compression tests were performed on 6 samples per condition. Each sample was cut to four sections, and both the edge and center regions of the sample were tested [Figure 5-16 right]. After
21 days in chondrogenic culture, the tissue compressive modulus of the IRCG100 constructs, from edge to center, ranged from 1.17 to 1.99 MPa for the 100C_100S constructs, 1.69 to 2.07 MPa for the 100C_50S constructs, and 0.35 to 0.39 MPa for the constructs without channels..
IRCG100 constructs containing hMSCs within the guidance channels [Figure. 5-16 A:
100C_100S and 100C_50S] cultured for 21 days under differentiating conditions had significantly superior mechanical properties compared to corresponding IRCG100 constructs without channels [Figure 5-16A:control] ( p < 0.05). Similar results were shown for the compressive modulus of the tissue within these IRCG100 constructs
[Figure 5-16B]. The tissue compressive modulus was significantly higher for IRCG100 constructs containing guidance channels [Figure. 5-16 B: 100C_100S and 100C_50S]
150 when compared to the tissue compressive modulus of IRCG100 constructs without channels [Figure 5-16A: control].
There was no significant difference in the mechanical properties between constructs containing 50 µm spacing [Figure 5-16: 100C_50S] and constructs containing 100 µm spacing [Figure 5-16: 100C_100S]. However, the center of the constructs containing 100
µm spacing [Figure 5-16: 100C_100S Center] have significantly higher compressive modulus values than the edge regions [Figure 5-16: 100C_100S Edge], while constructs containing 50 µm spacing have no significant difference between the center and edge regions of the constructs [Figure 5-16: 100C_50S]. Therefore, constructs containing spacing of 50 µm [Figure 5-16: 100C_50S] result in more uniform compressive properties throughout the entire construct when compared to constructs with 100 µm spacing [Figure 5-16: 100C_100S]. This result suggests that IRCG100-100C_50S constructs have more evenly distributed tissue from the edge to the center than the
IRCG100-100C_100S constructs.
151
Figure 5-16. Effect of microscale guidance on mechanical properties of large scale IRCG100 constructs. IRCG100 constructs were subject to compressive testing. Mean values of the compressive modulus of IRCG100 constructs (A) and of tissue within the constructs (B) are shown.100C_100S: IRCG construct with a 100 μm thick scaffold containing 100 μm channels with a spacing of 100 μm; 100C_50S: IRCG construct with a 100 μm thick scaffold containing 100 μm channels with a spacing of 50 μm; Control: IRCG construct with a 100 μm thick scaffold without channels. *Statistically significant
152 difference ( p < 0.05). Each sample was compressed for 10 s at a strain rate of 10% s-1, and compression tests were performed on 6 samples per condition
5.3.4 Metabolism Analysis
Glucose consumption rate and lactate production rate for IRCG100 constructs during 21 days in chondrogenic culture were measured and presented in Figure 5-17. The per cell glucose consumption rate [Figure 5-17 (A)] significantly decreased (p < 0.05) from day 3 to day 7 for IRCG100 constructs containing 100 µm channels with 50 µm spacing (173.8
± 3.6 fmole/hr to 103.7 ± 3.6 fmole/hr) [Figure 5-17 (A): 100C_50S], IRCG100 constructs containing 100 µm channels with 100 µm spacing (170.8 ± 3.2 fmole/hr to
102.4 ± 8.5 fmole/hr) [Figure 5-17 (A): 100C_100S] and control constructs without channels (172.1 ± 3.0 fmole/hr to 123.2 ± 3.4 fmole/hr) [Figure 5-17 (A): Control].
Similar results were observed in lactate production rate. The per cell lactate production rate [Figure 5-17 (B)] for IRCG100 constructs significantly decreased (p < 0.05) from day 3 to day 7. The lactate production rate decreased from 259.2 ± 10.9 fmole/hr to 187.2
± 4.8 fmole/hr for the 100C_50S constructs, from 257.7 ± 4.7 fmole/hr to 189.3 ± 6.2 fmole/hr for the 100C_100S constructs, and from 254.5 ± 5.9 fmole/hr to 194.9 ± 1.9 fmole/hr for the control constructs. There was no significant change in the per cell glucose consumption rate and lactate production rate for all IRCG100 constructs from day 7 to day 21 (p > 0.05).
In addition, there was no significant difference for the per cell glucose consumption rate between constructs with channels (100C_50S constructs, 100C_100S constructs) and constructs without channel (control) at day 3, day 7, day 15, and day 21 [Figure 5-17 (A)].
153
Similar results were observed in lactate consumption rate except for day 21, at which the control was significantly higher than constructs with channels [Figure 5-17 (B)]
Figure 5-17. Glucose consumption rate (A) and lactate production rate (B) per cell for the IRCG100 constructs during 21 days in chondrogenic culture. Data represents the mean ±
154
SEM. *Statistically significant difference ( p < 0.05) compared to data with control construct at day 3. #Statistically significant difference ( p < 0.05) compared to data with constructs containing100 µm channels with 100 µm spacing (100C_100S) at day 3. +Statistically significant difference (p < 0.05) compared to data with constructs containing 100 µm channel and 50 µm spacing (100C_50S) at day 3.
5.4 Discussion
In chapter 4, we showed that microscale substrate features caused differentiating MSCs to preferentially arrange themselves and to deposit an oriented cartilage ECM. We demonstrated that smaller channels allowed us to control cellular orientation and ECM deposition, which resulted in enhanced mechanical properties in 2-dimensional tissue engineered constructs. Previous studies have shown that defects less than a critical size of
3mm diameter may lead to complete spontaneous repair of articular cartilage, while larger defects do not repair completely (Convery, Akeson et al. 1972; Shapiro, Koide et al. 1993; ButnariuEphrat, Robinson et al. 1996; Jackson, Lalor et al. 2001). Since current clinical methods fail to provide a long term solution for large defects, tissue engineering serves as a promising method to repair large cartilage defects. However, the production of a large scale tissue engineered construct remains a challenge, as current constructs lack the necessary mechanical properties to be used in the clinc. . In this study, we developed a new method that utilized the microscale contact guidance of MSCs we investigated in our previous study (Chou, Rivera et al. 2012) and fabricated clinically useful, large-scale
3-dimesional constructs with an approximate size of1.8 cm (length) x 3.5 cm (diameter).
Two different techniques were used in this study to fabricate these large-scale constructs: an autonomous roll-up procedure and an induced roll-up procedure.
155
In the autonomous roll-up procedure, MSCs produced contractile forces during chondrogenesis to roll-up the 2-dimensional construct into a 3-dimensional cylinder.
Cells bind to substrates using integrins (DeMali, Wennerberg et al. 2003). When cells bind to the substrate, they routinely pull on the matrix and on other cells near where they are attached using their contractile cytoskeleton and produce a contraction force within the cell.Other groups have reported that cell contraction occurred in collagen matrices
(Mochitate, Pawelek et al. 1991; Schiro, Chan et al. 1991; Tomasek, Haaksma et al.
1992). Autonomous rolled-up constructs seeded with hMSCs (ARCG) started folding on day 4 and became half rolled constructs by day 7. Previous studies have shown matrix contraction in MSC-seeded collagen scaffolds (Moutos, Estes et al.; Nehrer, Breinan et al.
1997; Noeth, Rackwitz et al. 2007). Nehrer et al. found that the diameter of their chondrocyte-seeded collagen type I matrices decreased by day 7 due to cell-generated contractile forces (Nehrer, Breinan et al. 1997). Other studies have shown that chondrocytes were able to contract type I and type II collagen-GAG matrices using cell- generated contractile forces regardless of pore diameter (25–250 mm) or compressive stiffness (145–690 Pa) (Lee, Breinan et al. 2000). In addition, it has been reported that chondrocytes were able to bend collagen fibrils via cell motility activity and further manipulate the ECM to a specific architecture (Ohsawa, Yasui et al. 1982; Lee and
Loeser 1999). Therefore, we propose that these cellular contractile forces that occur during chondrogenesis caused our autonomously rolled –up constructs to roll-up on their own.
By means of these cell-generated contractile forces, autonomous rolled-up constructs
(ARCGs) were fabricated using CG membrane scaffolds of different thicknesses. Dense
156
GAG and type II collagen, as visualized by toluidine blue staining and type II collagen immunohistochemistry, were deposited in ARCG constructs [Figure 5-5 (A)-(E), Figure
5-7 (A)-(B)]. ARCG constructs with thinner membranes (ARCG250) yield better tissue- scaffold integration when compared to constructs with thicker membranes (ACRG 1000)
[Figure 5-5 (B)-(C), Figure 5-7 (A)-(B)]. We attribute this enhanced tissue-scaffold integration to the ability of the thinner membrane (ACRG250) to more easily roll-up, enhancing the contact between the tissue and scaffold during the roll-up, and also the ability of the thinner membrane to enhance nutrient transfer due to the lower scaffold thickness when compared to the ACRG1000 constructs. Comparing autonomous rolled- up (ARCGs) and induced rolled-up constructs (IRCGs) with the same scaffold thickness, much more ECM was deposited in the autonomous ARCG250 constructs [Figure 5-7:
(A)-(B)], when compared to the induced IRCG250 constructs [Figure 5-8: (B)-(C)]. The autonomous roll-up process gradually folded the membranes, which allowed the cells to synthesize ECM gradually over time during the roll-up process without blocking the nutrient transport by thick scaffold that occurs during the induced roll-up procedure.
However, the size and shape of ACRG constructs were limited and not controllable, and multi-tissue layers (more than 2) were not formed in the ACRG constructs (1-2 spirals in the construct).Therefore, these autonomous rolled-up constructs cannot be tailor-made to meet specific defect size and geometric requirements, which limits their ability to be used in a clinical setting.
Although we encountered challenges when fabricating induced rolled-up constructs
(IRCGs) initially [Figure 5-8: (A)-(B)] due to limited nutrient diffusion, by reducing the thickness of the CG membrane to 100 µm, multi-layer hMSC seeded CG constructs were
157 successfully fabricated [Figure 5-9: (D)]. ECM within the constructs containing microchannels (IRCG100-100C_50S, IRCG100-100C_100S) were more uniformly distributed than IRCG constructs without channels (IRCG Control) [Figure 5-10] which has been shown via both toluidine blue stained cross sections [Figure 5-10: (a)-(c)] and longitudinal sections [Figure 5-10: (A)-(C)]. The hMSCs in constructs without small channels (IRCG Control) tended to aggregate into clumps in certain regions, which we have shown in chapter 4 with the 2-dimensional constructs [Figure 4-7]. We believe that the uneven distribution of ECM within the control rolled-up constructs was caused by this tendency of the hMSCs to aggregate on scaffolds without channels. From these results, we hypothesize that the channels allow for more uniform cellular seeding and also help anchor the cells in place, therefore, preventing aggregation. When compared to the induced CG membrane-hMSC constructs, the induced rolled-up CG sponge-hMSC constructs showed inferior tissue-scaffold integration and only formed 1-2 tissue layers.
These CG sponge based constructs were more difficult to roll-up due to their lack of rigidity when compared to the CG membrane based constructs. We believe this technical difficulty in rolling-up the CG sponge constructs led to less tissue-scaffold contact and poor integration. Due to these advantages of the CG membrane and the superior mechanical properties of the CG membrane, we chose to use the CG membrane rather than the CG sponge for these studies.
In this study, we employed different materials (CG sponge and CG membrane),, techniques (autonomous roll-up and induced roll-up) and thicknesses (100 to 1000 µm) for fabricating large-scale rolled-up constructs. Based on our ECM deposition results, the optimal construct design is the IRCG100 (100 µm thickness) construct containing the100
158
µm microchannels. We further investigated the ECM alignment, mechanical properties, and metabolism for the IRCG 100 constructs.
First, we used SHG to visualize the collagen fibril alignment within the constructs. SHG is a powerful tool for imaging collagen fibrils in various tissues (Chen, Nadiarynkh et al.).
In this chapter, through SHG imaging, we further confirmed that oriented cells aligned along the length of the small guidance channels (100 µm in this study) produced aligned collagen fibrils that were also oriented in the direction of channel length [Figure 5-14:
(A)]. In contrast, the collagen in randomly-seeded constructs without channels showed no preferential alignment [Figure 5-14: (B)]. To further study collagen fibrils at the tissue level for the IRCG constructs, polarized light microscopy images revealed cell produced collagenous matrix in the IRCG constructs and collagen fibrils aligned along the channel length [Figure 5-15: (A)-(B)]. These results further confirm our hypothesis in 3- dimensional constructs that microscale substrate features cause differentiating MSCs to preferentially arrange themselves and to deposit an oriented cartilage ECM. We have shown that MSCs align and elongate along the length of the small guidance channels
[Figure 4-7], and we further demonstrated that collagenous matrix produced by moving cells in 3-dimensional IRCG100 constructs is aligned with respect to the cell movements
[Figure 5-15: (A)-(B)]. Similar studies have indicated that similarly elongated cells exhibited contractile forces along the direction of elongation and led to the alignment of collagen fibers (Hata, Hori et al. 1984; Wang, Grood et al. 2000; Wang, Jia et al. 2003).
Cells are able to manipulate the ECM around them into a specific architecture via cell motility activity and cell produced contraction force (Ohsawa, Yasui et al. 1982; Lee and
Loeser 1999).
159
Further, we wanted to investigate if the collagen fibril alignment within the IRCG100 constructs with guidance channels led to enhanced mechanical properties. Adult human hyaline cartilage shows depth dependent mechanical properties and a compressive elasticity modulus that varies from 1.16±0.20 MPa (0–125 µm from the joint surface; superficial zone) to 7.75±1.45 MPa (1250–1500 µm from the joint surface; deep zone)
(Chen, Falcovitz et al. 2001). The tissue compressive elasticity modulus for the IRCG100 constructs containing guidance channels was approximately 1.16 to 2.07 MPa [Figure 5-
16: (B)], which is comparable to the scale of native articular cartilage found in the literature (Setton, Elliott et al. 1999; Chen, Falcovitz et al. 2001). IRCG100 constructs containing microscalee guidance channels (1.16 to 2.07 MPa) displayed superior mechanical properties when compared to constructs without guidance channels (0.35 to
0.39 MPa), which demonstrated that microscale guidance has an effect on the mechanical function of the resulting tissue within 3-dimensional construct. The compressive elasticity modulus for the center region of the IRCG100 construct was higher than the compressive elasticity modulus for the edge region [Figure 5-16], which may be with a result of the hypoxic environment within the center of the constructs. It has been reported that MSC-based chondrogenesis is enhanced under hypoxic conditions and hypothesized that the low oxygen environment may be an essential element for in vivo chondrogenesis.
(Lennon, Edmison et al. 2001; Grayson, Zhao et al. 2006; Pfander and Gelse 2007;
Kanichai, Ferguson et al. 2008). In particular, Kanichai et al. reported that MSC culture under hypoxia displayed a significant increase in proteoglycan deposition and collagen type II expression after 3 weeks in culture (Kanichai, Ferguson et al. 2008). Since the center region of the construct is directly surrounded by less chondrogenic medium when
160 compared to the edges, these diffusion limitations create a more hypoxic environment within the center region of the construct. Based on the previously mentioned hypoxic studies and the hypoxic nature of native cartilage tissue, we hypothesize that the hypoxic conditions within the center region of the constructs lead to enhanced mechanical properties. Current 3-dimensional tissue engineered cartilage,fabricated using stem cells and various types of scaffolds including poly(e-caprolactone), poly(L-lactic acid), collagen type I, hyaluronic acid hydrogels, gelatin, alginate, and agarose, typically displays a compressive elasticity modulus between 10-15 KPa (Awad, Wickham et al.
2004; Bian, Zhai et al. 2011; Huang, Farrell et al. 2011). Our IRCG100 constructs containing guidance channels displayed a compressive modulus elasticity modulus of
1.16 to 2.07 MPa and therefore, displayed relatively superior mechanical properties when compared to most current TE constructs.
Next, we monitored the cell metabolism during chondrogenesis. Glucose metabolism has been reported an important factor contributing to ECM synthesis in cartilage tissue engineering (Nettles, Chilkoti et al. 2009). Glucose consumption and lactate production rates in cartilage have been regarded as indicators for monitoring cell metabolism and matrix synthesis during chondrogenesis (Lane, Brighton et al. 1977; Otte 1991; Lee and
Urban 1997; Lee, Wilkins et al. 2002; Nettles, Chilkoti et al. 2009).Other studies have shown that oxygen consumption of cells in articular cartilage is only 2–5% of the consumption of kidney or liver cells (Lee and Urban 1997) and that anaerobic glycolysis is considered as the principal mechanism in which glucose is metabolized to lactate in cartilage (Lee and Urban 1997). Per cell glucose consumption for the ICRG100 constructs showed a significant decrease from day 1 to day 7 (p<0.05) and no further
161 changes from day 7 to day 21 (p >0.05). There were no significant differences in per cell glucose consumption [Figure 5-17 (A)] and lactate production rates [Figure 5-17 (B) ] between the control ICRG 100 constructs without channels and the ICRG 100 constructs containing 100 µm channels, which suggests that the channels have no influence on cell metabolism during the 21 days the cells undergo chondrogenesis [Figure 5-17]. The average metabolic values were 108.7 fmole/hr for glucose consumption and 172.0 fmole/hr for lactate production from day 7 to day 21. These values are approximately half of the values from a previous study for chondrogenesis of hMSC pellets between day 7 and day 21 (Pattappa, Heywood et al.). The difference in metabolic values may due to the different systems (scaffold free pellet culture and CG based constructs) since cellular biosynthetic activity is dependent on the biomaterial used. The ratio of lactate production to glucose consumption is ideally 2 (1 mole of glucose is metabolized to 2 moles of lactate) for anaerobic glycolysis. This ratio for the IRCG100 constructs is approximately
1.58, which indicates glucose metabolism occurs through the anaerobic pathway and that only 60% of the glucose was converted to lactate, which suggests that part of the glucose may be used for ECM synthesis. Teixeira et al. reported a ratio of 2 was observed for single chondrocytes cultured in hydrogels and that the ratio dropped to 1 in aggregate culture (Teixeira, Leijten et al.). They proposed that higher cell-cell contact within the aggregate may stimulate cell metabolism and convert part of glucose for biosynthesis instead of energy consumption (Teixeira, Leijten et al.), which helps explain our result.
We feel that the hMSCs within these rolled-up scaffolds utilize the glucose to produce
GAG and other ECM components during chondrogenesis.
162
Our study demonstrated that microscale guidance has an effect on hMSC differentiation and ECM deposition within these large-scale constructs. While we imposed some organization on the newly synthesized ECM and successfully enhanced the mechanical function of the resulting tissue in the large-scale IRCG100 construct, there are still certain limitations. First, while native cartilage possesses a compressive elasticity modulus that varies from 1.16 to 7.75 MPa, the IRCG constructs only reached the initial scale of native cartilage (Chen, Falcovitz et al. 2001). This limitation of our constructs may be due to the hole along the center of the construct and local areas of poor integration between the scaffold and tissue. In future studies, we will investigate methods to eliminate the central cylindrical hole and to enhance integration between the scaffold and tissue.
5.5 Conclusion
In this work, we extended the previous 2-dimensional construct findings (Chou, Rivera et al. 2012) to a 3-dimensional construct for preclinical experiments. We successfully created 3-dimensional constructs for in vivo applications by rolling up 2-dimensional hMSC-seeded CG based scaffold and formed 3-dimensional rolled-up cartilage constructs with microscale architecture to guide the differentiating hMSCs. Different materials (CG sponge and CG membrane), techniques (autonomous roll-up and induced roll-up), and thicknesses (100 to 1000µm) were used in this study for fabricating large- scale rolled-up constructs. IRCG100 constructs containing microchannles displayed superior mechanical and tissue properties among all constructs. Dense ECM rich in both
GAG and type II collagen throughout the outer 3 layers of the construct were observed in the IRCG100 constructs. Importantly, we demonstrated that microscale guidance channels incorporated within the 3-dimensional IRCG100 cartilage constructs led to the
163 production of aligned cell-produced collagenous matrix and enhanced mechanical function.
5.6 Reference
Adelola O. Oseni, C. C., Maria Z. Boland, and P. E. B. a. A. M. Seifalian (2011). "Cartilage Tissue Engineering: the Application of Nanomaterials and Stem Cell Technology." Tissue Engineering for Tissue and Organ Regeneration
Awad, H. A., M. Q. Wickham, et al. (2004). "Chondrogenic differentiation of adipose- derived adult stem cells in agarose, alginate, and gelatin scaffolds." Biomaterials 25(16): 3211-3222.
Bian, L., D. Y. Zhai, et al. (2011). "Enhanced MSC chondrogenesis following delivery of TGF-beta 3 from alginate microspheres within hyaluronic acid hydrogels in vitro and in vivo." Biomaterials 32(27): 6425-6434.
ButnariuEphrat, M., D. Robinson, et al. (1996). "Resurfacing of goat articular cartilage by chondrocytes derived from bone marrow." Clinical orthopaedics and related research(330): 234-243. Chen, S. S., Y. H. Falcovitz, et al. (2001). "Depth-dependent compressive properties of normal aged human femoral head articular cartilage: relationship to fixed charge density." Osteoarthritis and Cartilage 9(6): 561-569.
Chen, X., O. Nadiarynkh, et al. "Second harmonic generation microscopy for quantitative analysis of collagen fibrillar structure." Nature Protocols 7(4): 654-669.
Chou, C.-L., A. L. Rivera, et al. (2012). “Micrometer scale guidance of mesenchymal stem cells to form structurally oriented cartilage extracellular matrix. Tissue Engineering Part A, 19(9-10): 1081-1090.
Chung, C. and J. A. Burdick (2009). "Influence of three-dimensional hyaluronic acid microenvironments on mesenchymal stem cell chondrogenesis." Tissue Engineering Part A 15(2): 243-254.
Convery, F. R., W. H. Akeson, et al. (1972). "The repair of large osteochondral defects. An experimental study in horses." Clinical orthopaedics and related research 82: 253-62.
Delon, I. and N. H. Brown (2007). "Integrins and the actin cytoskeleton." Current Opinion in Cell Biology 19(1): 43-50.
DeMali, K. A., K. Wennerberg, et al. (2003). "Integrin signaling to the actin cytoskeleton." Current Opinion in Cell Biology 15(5): 572-582.
164
Dickhut, A., E. Gottwald, et al. (2008). "Chondrogenesis of mesenchymal stem cells in gel-like biomaterials in vitro and in vivo." Frontiers in Bioscience 13: 4517-4528.
Djouad, F., D. Mrugala, et al. (2006). "Engineered mesenchymal stem cells for cartilage repair." Regenerative Medicine 1(4): 529-537.
Gong, Y. Y., J. X. Xue, et al. (2011). "A sandwich model for engineering cartilage with acellular cartilage sheets and chondrocytes." Biomaterials 32(9): 2265-2273.
Grayson, W. L., F. Zhao, et al. (2006). "Effects of hypoxia on human mesenchymal stem cell expansion and plasticity in 3D constructs." Journal of Cellular Physiology 207(2): 331-339.
Hata, R. I., H. Hori, et al. (1984). "Selective-inhibition of type-I collagen-synthesis in osteoblastic cells by epidermal growth-factor." Endocrinology 115(3): 867-876.
Huang, A. H., M. J. Farrell, et al. (2011). "Mechanics and mechanobiology of mesenchymal stem cell-based engineered cartilage." Journal of Biomechanics 43(1): 128- 136.
Hunziker, E. B. (2002). "Articular cartilage repair: basic science and clinical progress. A review of the current status and prospects." Osteoarthritis and Cartilage 10(6): 432-463.
Huttenlocher, A. and A. R. Horwitz "Integrins in cell migration." Cold Spring Harbor perspectives in biology 3(9): a005074-a005074.
Jackson, D. W., P. A. Lalor, et al. (2001). "Spontaneous repair of full-thickness defects of articular cartilage in a goat model - A preliminary study." Journal of Bone and Joint Surgery-American Volume 83A(1): 53-64.
Janakiraman, V., B. L. Kienitz, et al. (2007). "Lithography technique for topographical micropatterning of collagen-glycosaminoglycan membranes for tissue engineering applications." J Med Device 1(3): 233-237.
Jakobsen, R. B., A. Shandadfar, et al. (2010). "Chondrogenesis in a hyaluronic acid scaffold: comparison between chondrocytes and MSC from bone marrow and adipose tissue." Knee Surgery Sports Traumatology Arthroscopy 18(10): 1407-1416.
Janjanin, S., W.-J. Li, et al. (2008). "Mold-shaped, nanofiber scaffold-based cartilage engineering using human mesenchymal stem cells and bioreactor." Journal of Surgical Research 149(1): 47-56.
Kanichai, M., D. Ferguson, et al. (2008). "Hypoxia promotes chondrogenesis in rat mesenchymal stem cells: A role for AKT and hypoxia-inducible factor (HIF)-1 alpha." Journal of Cellular Physiology 216(3): 708-715.
165
Lane, J. M., C. T. Brighton, et al. (1977). "Anaerobic and aerobic metabolism in articular-cartilage." Journal of Rheumatology 4(4): 334-342.
Lee, C. R., H. A. Breinan, et al. (2000). "Articular cartilage chondrocytes in type I and type II collagen-GAG matrices exhibit contractile behavior in vitro." Tissue Engineering 6(5): 555-565.
Lee, G. M. and R. F. Loeser (1999). "Cell surface receptors transmit sufficient force to bend collagen fibrils." Experimental Cell Research 248(1): 294-305.
Lee, R. B. and J. P. G. Urban (1997). "Evidence for a negative Pasteur effect in articular cartilage." Biochemical Journal 321: 95-102.
Lee, R. B., R. J. Wilkins, et al. (2002). "The effect of mechanical stress on cartilage energy metabolism." Biorheology 39(1-2): 133-143.
Lennon, D. P., J. M. Edmison, et al. (2001). "Cultivation of rat marrow-derived mesenchymal stem cells in reduced oxygen tension: Effects on in vitro and in vivo osteochondrogenesis." Journal of Cellular Physiology 187(3): 345-355.
Li, W. J., R. Tuli, et al. (2005). "A three-dimensional nanofibrous scaffold for cartilage tissue engineering using human mesenchymal stem cells." Biomaterials 26(6): 599-609.
Liang, W.-H., B. L. Kienitz, et al. (2010). "Concentrated collagen-chondroitin sulfate scaffolds for tissue engineering applications." Journal of Biomedical Materials Research Part A 94A(4): 1050-1060.
Mochitate, K., P. Pawelek, et al. (1991). "Stress-relaxation of contracted collagen gels - disruption of actin filament bundles, release of cell-surface fibronectin, and down- regulation of dna and protein-synthesis." Experimental Cell Research 193(1): 198-207.
Moutos, F. T., B. T. Estes, et al. "Multifunctional hybrid three-dimensionally woven scaffolds for cartilage tissue engineering." Macromolecular Bioscience 10(11): 1355- 1364.
Nehrer, S., H. A. Breinan, et al. (1997). "Canine chondrocytes seeded in type I and type II collagen implants investigated in vitro." Journal of Biomedical Materials Research 38(2): 95-104.
Nettles, D. L., A. Chilkoti, et al. (2009). "Early metabolite levels predict long-term matrix accumulation for chondrocytes in elastin-like polypeptide biopolymer scaffolds." Tissue Engineering Part A 15(8): 2113-2121.
Noeth, U., L. Rackwitz, et al. (2007). "Chondrogenic differentiation of human mesenchymal stem cells in collagen type I hydrogels." Journal of Biomedical Materials Research Part A 83A(3): 626-635.
166
Ohsawa, S., N. Yasui, et al. (1982). "Contraction of collagen gel by the dedifferentiated chondrocytes." Cell Biology International Reports 6(8): 767-774.
Otte, P. (1991). "Basic cell-metabolism of articular-cartilage - manometric studies." Zeitschrift Fur Rheumatologie 50(5): 304-312.
Ousema, P. H., F. T. Moutos, et al. (2012). "The inhibition by interleukin 1 of MSC chondrogenesis and the development of biomechanical properties in biomimetic 3D woven PCL scaffolds." Biomaterials 33(35): 8967-8974.
Pattappa, G., H. K. Heywood, et al. "The metabolism of human mesenchymal stem cells during proliferation and differentiation." Journal of Cellular Physiology 226(10): 2562- 2570.
Pfander, D. and K. Gelse (2007). "Hypoxia and osteoarthritis: how chondrocytes survive hypoxic environments." Current Opinion in Rheumatology 19(5): 457-462.
Schiro, J. A., B. M. C. Chan, et al. (1991). "Integrin-alpha-2-beta-1 (vla-2) mediates reorganization and contraction of collagen matrices by human-cells." Cell 67(2): 403-410.
Setton, L. A., D. M. Elliott, et al. (1999). "Altered mechanics of cartilage with osteoarthritis: human osteoarthritis and an experimental model of joint degeneration." Osteoarthritis and Cartilage 7(1): 2-14.
Shapiro, F., S. Koide, et al. (1993). "Cell origin and differentiation in the repair of full- thickness defects of articular-cartilage." Journal of Bone and Joint Surgery-American Volume 75A(4): 532-553.
Shirazi, R., A. Shirazi-Adl, et al. (2008). "Role of cartilage collagen fibrils networks in knee joint biomechanics under compression." Journal of Biomechanics 41(16): 3340- 3348.
Spadaccio, C., A. Rainer, et al. (2009). "Poly-l-lactic acid/hydroxyapatite electrospun nanocomposites induce chondrogenic differentiation of human MSC." Annals of Biomedical Engineering 37(7): 1376-1389.
Teixeira, L. S. M., J. C. H. Leijten, et al. "High throughput generated micro-aggregates of chondrocytes stimulate cartilage formation in vitro and in vivo." European Cells & Materials 23: 387-399.
Tomasek, J. J., C. J. Haaksma, et al. (1992). "Fibroblast contraction occurs on release of tension in attached collagen lattices - dependency on an organized actin cytoskeleton and serum." Anatomical Record 232(3): 359-368.
167
Wang, J. H. C., E. S. Grood, et al. (2000). "Alignment and proliferation of MC3T3-E1 osteoblasts in microgrooved silicone substrata subjected to cyclic stretching." Journal of Biomechanics 33(6): 729-735.
Wang, J. H. C., F. Y. Jia, et al. (2003). "Cell orientation determines the alignment of cell- produced collagenous matrix." Journal of Biomechanics 36(1): 97-102.
Wang, Y. Z., D. J. Blasioli, et al. (2006). "Cartilage tissue engineering with silk scaffolds and human articular chondrocytes." Biomaterials 27(25): 4434-4442.
Wang, Y. Z., U. J. Kim, et al. (2005). "In vitro cartilage tissue engineering with 3D porous aqueous-derived silk scaffolds and mesenchymal stem cells." Biomaterials 26(34): 7082-7094.
Wilson, W., J. M. Huyghe, et al. (2007). "Depth-dependent compressive equilibrium properties of articular cartilage explained by its composition." Biomechanics and Modeling in Mechanobiology 6(1-2): 43-53.
Woo, S. L. Y., W. H. Akeson, et al. (1976). "Measurements of nonhomogeneous, directional mechanical properties of articular cartilage in tension." Journal of Biomechanics 9(12): 785-791.
Xue, J. X., Y. Y. Gong, et al. (2012). "Chondrogenic differentiation of bone marrow- derived mesenchymal stem cells induced by acellular cartilage sheets." Biomaterials 33(24): 5832-5840.
Yang, Q., J. Peng, et al. (2008). "A cartilage ECM-derived 3-D porous acellular matrix scaffold for in vivo cartilage tissue engineering with PKH26-labeled chondrogenic bone marrow-derived mesenchymal stem cells." Biomaterials 29(15): 2378-2387.
Yannas, I. V., J. F. Burke, et al. (1980). "Design of an artificial skin .2. Control of chemical-composition." Journal of Biomedical Materials Research 14(2): 107-132.
Zwingmann, J., A. T. Mehlhorn, et al. (2007). "Chondrogenic differentiation of human articular chondrocytes differs in biodegradable PGA/PLA scaffolds." Tissue Engineering 13(9): 2335-2343.
168
Chapter 6
CONCLUSIONS
6.1 Conclusions
Current clinical treatments for articular cartilage injuries usually lead to the formation of fibrocartilage tissue which is biomechanically and structurally inferior to the hyaline cartilage. Tissue engineering offers the potential to provide a long-term solution for cartilage defects; however, current tissue engineered constructs lack the microscale structural organization of native cartilage. Therefore, current tissue engineered constructs have suboptimal mechanical properties that prevent them from being used in the clinic.
We hypothesized that the suboptimal mechanical properties of cartilage constructs obtained via these approaches were due to their inability to reproduce the organized ultrastructure present in native cartilage tissue.
In this study, we addressed the major challenge of current cartilage tissue engineering: the inferior mechanical properties. The overall goal of this dissertation was to investigate the influence of the microscale guidance features on the architecture of the differentiating tissue and further develop clinically useful human mesenchymal stem cell (hMSC) based constructs with enhanced mechanical properties. Accordingly, the research objectives were divided into the following aims: (i) Control of hMSC selective adhesion on the scaffolds, (ii) Investigation of the effect of microscale guidance on MSC-based condrogenesis, and (iii) Development of large-scale 3-dimensional clinically relevant
169 cartilage constructs with an organized microscale structure. The findings of this project, limitations, and possible improvements are summarized in this section.
(i) Control of hMSC selective adhesion on collagen glycosaminoglycan (GG) and polydimethylsiloxane (PDMS) Surface
To study the effects of the guidance channels on cellular orientation and function for hMSC-based chondrogenesis, as the first step, the hMSCs must be localized within the channels. Therefore, the goal of this study was to develop a technique that allows for control of hMSC adhesion and spatial organization on CG membranes and PDMS membranes. In this study, we improved the collagen soft- lithography technique developed earlier (Janakiraman, Kienitz et al. 2007) to micropattern the collagen membrane for use with differentiating MSCs and developed a method of tuning surface chemistry and topography to direct cell selectively and adhere MSCs to the defined areas.
Through microfluid techniques, CG membrane soft lithography, and F108 modification, selective attachment and spreading of hMSCs within the channels was ensured. We demonstrate selective seeding of viable MSCs within the channels, and the chondrogenic potential of MSCs seeded in these channels were investigated. This study served as a first step towards the creation of tissue engineered constructs with built-in cellar architecture.
(ii) Investigate the effect of microscale guidance on MSC-based chondrogenesis
Next, the effect of microscale guidance on cell alignment and ECM function in an hMSC-based chondrogenesis model was investigated. In this work, we tested the hypothesis that microscale substrate features would cause differentiating MSCs to preferentially arrange themselves and to deposit an oriented cartilage ECM similar to native articular cartilage. MSCs aligned and produced mature collagen fibrils along the
170 length of the channel in smaller linear channels of widths from 25–100 µm compared to larger linear channels of widths 500–1000 µm. Further, substrates with smaller microchannels that led to cell alignment also led to superior mechanical properties compared to constructs with randomly seeded cells or selectively seeded cells in larger channels. The ultimate stress and modulus of elasticity of constructs with cells seeded in smaller channels increased by as much as four folds. In this study, we demonstrate that microfabricated matrices can induce differentiating hMSCs to form oriented collagen microstructures and proved that an increase in cellular and collagen type II alignment within the smaller channels led to enhanced mechanical strength of the resultant ECM.
These findings provided new parameters which help optimize the design of functional hMSC-based tissue-engineered cartilage using collagen-based substrates and can be used to fabricate large clinically useful hMSC-based cartilage constructs with superior mechanical properties.
(iii) Design of large-scale 3-dimensional cartilage constructs with microchannels for preclinical studies
The 2-dimensional constructs do not provide a sufficient solution for a 3-dimensional cartilage defect, and therefore, lack clinical relevance. Lastly, we extended the 2- dimensional construct findings to a 3-dimensional construct for preclinical studies. Based on the parameters from the 2-dimensional studies (Chou, Rivera et al. 2012), we introduce guidance channels into collagen-based scaffolds, optimized the scaffold design, and successfully created 3-dimensional constructs for in vivo applications. By rolling up
2-dimensional hMSC-seeded CG based scaffolds, 3-dimensional rolled-up large-scale
(3.5 mm in diameter × 18 mm in length) cartilage constructs with organized microscale
171 architecture were fabricated. Extensive GAG and collagen type II were present in the
ECM of the 3-dimensional constructs, which are both indicative of chondrogenesis. We demonstrated that the microscale guidance channels incorporated within the 3- demensional cartilage constructs led to the production of aligned cell-produced collagenous matrix and enhanced mechanical function. The tissue modulus of elasticity of 3-demensional cartilage constructs containing guidance channels increased by as much as six folds compared to constructs without channels, which is comparable to the scale of the native cartilage and higher than most current tissue-engineered constructs. We demonstrated that the incorporation of our microscale guidance features into 3- dimensional hMSC-seeded collagen-based constructs led to the formation of a superior architecture, homogenous cartilaginous tissue, and enhanced mechanical properties.
In conclusion, these findings provide us with the fundamental knowledge of how microscale guidance channels regulate matrix deposition for long term cartilage construct development and therefore, provide us the information on design parameters for rational tissue engineering of cartilage constructs. The design parameters for rational tissue engineering in this work can be utilized for designing next-generation scaffolds for structured chondrogenesis of cartilage constructs as well as used for other stem cell-based tissue engineering research where spatial cellular architecture plays a key role such as nerves and blood vessels tissue.
172
6.2 Future Directions
In this project, we developed large-scale 3-dimensional cartilage constructs that exhibited an organized microscale structure and enhanced mechanical properties compared to most current tissue engineered constructs. We utilized microscale guidance channels to guide differentiating hMSCs to align and produce oriented ECM that displayed enhanced mechanical properties. However, the roll-up technique for developing large scale cartilage constructs has its limitations. One current challenge for our large-scale constructs is the integration of rolled-up cell layers withthe CG scaffold layers. In many regions of the construct, excellent integration was observed; however, in some regions, the integration was suboptimal. To improve the integration, we need to optimize the culture conditions by testing different culture variables including by increasing the cell attachment time before roll-up, reducing the time for the roll-up procedure, and minimizing the scaffold to tissue ratio. Another issue is the uneven tissue formation in large-scale constructs and the lack of tissue in the middle of the construct. We will use bioreactor culture to overcome the issue of uneven tissue formation. Also, we plan to inject cells into the middle of the construct to fill the tissue-free hole that exists in the middle of the cylindrical construct. In addition, we will investigate different methods of rolling up the construct to form a more compact structure that lacks a central hole within the 3-dimensional cylinder.
In addition to the above challenges, nutrient transfer through these 3-dimensional constructs also remains a challenge, which we plan to investigate and optimize via mathematical modeling. Mathematical models will be developed for both static and
173 bioreactor culture to calculate the mass transfer rates through the membrane layers and tissue layers. In this work, experimental data and theoretical calculations will be used to create a model that can predict the influence of the transport of glucose, oxygen, and growth factors for culturing large-scale constructs based on different values for the construct parameters., Using this model, we will be able to optimize the construct parameters such as the number of tissue layers and the scaffold thickness to obtain optimal nutrient transport throughout the construct.
In addition to rolling-up these 2-dimensional hMSC-CG scaffold constructs to form 3- dimensional constructs, future studies will also investigate stacking multiple 2- dimensional constructs to form stacked 3-dimensional cartilage constructs also containing these guidance channels. For these stacked constructs, the guidance channels will be changed to align the MSCs and ECM perpendicular to the membrane rather than parallel, which is the case of the rectangular guidance channels used in this project. We will change the guidance channels to deeper circular wells of small diameter. When designing these circular wells, we will minimize the diameter to depth ratio so that the cells align along the depth of the circular wells and therefore, align and produce ECM perpendicular to the membrane surface. Using this method, we will create 3-dimensional stacked cartilage constructs with ECM that emulates the organization of collagen fibrils in the deep zone of native cartilage (fibrils are aligned perpendicular to the subchondral bone and parallel to the direction of compression), which are the regions in native cartilage that are responsible for its excellent compressive mechanical properties. We will need to optimize culture conditions for these stacked scaffolds including cell attachment time, cell density, scaffold thickness, and number of constructs to stack. We will also study the
174 integration between layers by shear testing. Future directions of this project will focus on improving the current roll-up procedure of these 2-dimensional constructs with guidance channels and on developing this stacking method.
6.3 References
Chou, C.-L., A. L. Rivera, et al. (2012). “Micrometer scale guidance of mesenchymal stem cells to form structurally oriented cartilage extracellular matrix. Tissue Engineering Part A, 19(9-10): 1081-1090.
Janakiraman, V., B. L. Kienitz, et al. (2007). "Lithography technique for topographical micropatterning of collagen-glycosaminoglycan membranes for tissue engineering applications." J Med Device 1(3): 233-237.
175
BIBLIOGRAPHY
Adams, M. E., M. H. Atkinson, et al. (1995). "The role of viscosupplenentation with hylan G-F 20 (Synvisc(R)) in the treatment of osteoarthritis of the knee: A Canadian multicenter trial comparing hylan G-F 20 alone, hylan G-F 20 with non-steroidal anti- inflammatory drugs (NSAIDs) and NSAIDs alone." Osteoarthritis and Cartilage 3(4): 213-225.
Adelola O. Oseni, C. C., Maria Z. Boland, and P. E. B. a. A. M. Seifalian (2011). "Cartilage Tissue Engineering: the Application of Nanomaterials and Stem Cell Technology." Tissue Engineering for Tissue and Organ RegenerationU.K.
Ahmed, T. A. E. and M. T. Hincke (2010). "Strategies for Articular Cartilage Lesion Repair and Functional Restoration." Tissue Engineering Part B-Reviews 16(3): 305-329.
Alaerts, J. A., V. M. De Cupere, et al. (2001). "Surface characterization of poly(methyl methacrylate) microgrooved for contact guidance of mammalian cells. Biomaterials 22(12): 1635-1642.
Almarza, A. J. and K. A. Athanasiou (2004). "Design characteristics for the tissue engineering of cartilaginous tissues." Annals of Biomedical Engineering 32(1): 2-17.
Archer, C. W., S. Redman, et al. (2006). "Enhancing tissue integration in cartilage repair procedures." Journal of Anatomy 209(4): 481-493.
Aubin, H., J. W. Nichol, et al. (2010). "Directed 3D cell alignment and elongation in microengineered hydrogels." Biomaterials 31(27): 6941-6951.
Awad, H. A., M. Q. Wickham, et al. (2004). "Chondrogenic differentiation of adipose- derived adult stem cells in agarose, alginate, and gelatin scaffolds." Biomaterials 25(16): 3211-3222.
Banu, N., Y. Banu, et al. (2005). "Biodegradable polymers in chondrogenesis of human articular chondrocytes." Journal of artificial organs : the official journal of the Japanese Society for Artificial Organs 8(3): 184-91.
Benninghoff, A. (1925). "Form un Bau der Gelenkknorpel in ihren Beziehungen zur Funktion." Zeitschrift für Zellforschung und Mikroskopische Anatomie 2(5): 783-862.
Bentley, G. (2004). "A prospective, randomised comparison of autologous chondrocyte implantation versus mosaicplasty for osteochondral defects in the knee - Reply." Journal of Bone and Joint Surgery-British Volume 86B(4): 619-620.
176
Bentley, G., L. C. Biant, et al. (2003). "A prospective, randomised comparison of autologous chondrocyte implantation versus mosaicplasty for osteochondral defects in the knee." Journal of Bone and Joint Surgery-British Volume 85B(2): 223-230.
Bhosale, A. M. and J. B. Richardson (2008). "Articular cartilage: structure, injuries and review of management." British Medical Bulletin 87(1): 77-95.
Bian, L., D. Y. Zhai, et al. (2011). "Enhanced MSC chondrogenesis following delivery of TGF-beta 3 from alginate microspheres within hyaluronic acid hydrogels in vitro and in vivo." Biomaterials 32(27): 6425-6434.
Birk, D. E. and R. L. Trelstad (1986). "Extracellular compartments in tendon morphogenesis - collagen fibril, bundle, and macroaggregate formation." Journal of Cell Biology 103(1): 231-240.
Brittberg, M., A. Lindahl, et al. (1994). "Treatment of deep cartilage defects in the knee with autologous chondrocyte transplantation." New England Journal of Medicine 331(14): 889-895.
Brittberg M, P. L., Sjogren-Jansson E, Tallheden T, Lindahl A (2003). "Articular cartilage engineering with autologous chondrocytes transplantation. A review of recent developments." J Bone Joint Surg Am 85-A (3): 109–115.
Buckwalter, J. A. (1998). "Articular cartilage: Injuries and potential for healing." Journal of Orthopaedic & Sports Physical Therapy 28(4): 192-202.
Buckwalter, J. A. (2002). "Articular cartilage injuries." Clinical Orthopaedics and Related Research(402): 21-37.
Buckwalter, J. A. and H. J. Mankin (1998). "Articular cartilage: tissue design and chondrocyte-matrix interactions." Instructional course lectures 47: 477-86.
Buckwalter, J. A., L. C. Rosenberg, et al. (1990). Articular-cartilage - composition, structure, response to injury, and methods of facilitating repair. Articular Cartilage and Knee Joint Function: Basic Science and Arthroscopy: 19-56.
Bullough, P. and J. Goodfellow (1968). "The significance of the fine structure of articular cartilage." The Journal of bone and joint surgery. British volume 50(4): 852-7.
Bullough, P. G. (1981). "The geometry of diarthrodial joints, its physiologic maintenance, and the possible significance of age-related-changes in geometry-to-load distribution and the development of osteo-arthritis." Clinical Orthopaedics and Related Research(156): 61-66.
177
ButnariuEphrat, M., D. Robinson, et al. (1996). "Resurfacing of goat articular cartilage by chondrocytes derived from bone marrow." Clinical orthopaedics and related research(330): 234-243.
Butt, A. M., M. C. I. M. Amin, et al. "In vitro characterization of pluronic f127 and d- alpha-tocopheryl polyethylene glycol 1000 succinate mixed micelles as nanocarriers for targeted anticancer-drug delivery." Journal of Nanomaterials.
C. M. Jack, S. S. Rajaratnam, et al. (2012). "The modified tibial tubercle osteotomy for anterior knee pain due to chondromalacia patellae in adults.A five-year prospective study." Bone Joint Res 1(8): 167-173.
Cahill (1995). "Osteochondritis Dissecans of the Knee: Treatment of Juvenile and Adult Forms." J Am Acad Orthop Surg 3(4): 237-247.
Cancedda, R., B. Dozin, et al. (2003). "Tissue engineering and cell therapy of cartilage and bone." Matrix Biology 22(1): 81-91.
Caplan, A. I., M. Elyaderani, et al. (1997). "Principles of cartilage repair and regeneration." Clinical Orthopaedics and Related Research(342): 254-269.
Caplan, A. I. and V. M. Goldberg (1999). "Orthopaedic tissue engineering - Editorial comment." Clinical Orthopaedics and Related Research(367): S2-S4.
Chanlalit, C., C. Kasemkijwattana, et al. (2007). "Autologous chondrocyte implantation for traumatic large cartilage defect." Journal of the Medical Association of Thailand 90(7): 1435-1442.
Chapekar, M. S. (2000). "Tissue engineering: Challenges and opportunities." Journal of Biomedical Materials Research 53(6): 617-620.
Charriere, G., M. Bejot, et al. (1989). "Reactions to a bovine collagen implant - clinical and immunological study in 705 patients." Journal of the American Academy of Dermatology 21(6): 1203-1208.
Chen, A. C., W. C. Bae, et al. (2001). "Depth- and strain-dependent mechanical and electromechanical properties of full-thickness bovine articular cartilage in confined compression." Journal of Biomechanics 34(1): 1-12.
Chen, C. S., M. Mrksich, et al. (1998). "Micropatterned surfaces for control of cell shape, position, and function." Biotechnology Progress 14(3): 356-363.
Chen, F. H., K. T. Rousche, et al. (2006). "Technology Insight: adult stem cells in cartilage regeneration and tissue engineering." Nat Clin Pract Rheum 2(7): 373-382.
178
Chen, S. S., Y. H. Falcovitz, et al. (2001). "Depth-dependent compressive properties of normal aged human femoral head articular cartilage: relationship to fixed charge density." Osteoarthritis and Cartilage 9(6): 561-569.
Chen, X., O. Nadiarynkh, et al. "Second harmonic generation microscopy for quantitative analysis of collagen fibrillar structure." Nature Protocols 7(4): 654-669.
Chou, C.-L., A. L. Rivera, et al. (2012). “Micrometer scale guidance of mesenchymal stem cells to form structurally oriented cartilage extracellular matrix. Tissue Engineering Part A, 19(9-10): 1081-1090.
Chung, C. and J. A. Burdick (2009). "Influence of three-dimensional hyaluronic acid microenvironments on mesenchymal stem cell chondrogenesis." Tissue Eng Part A 15(2): 243-54.
Chung, C. and J. A. Burdick (2009). "Influence of three-dimensional hyaluronic acid microenvironments on mesenchymal stem cell chondrogenesis." Tissue Engineering Part A 15(2): 243-254.
Clar, C., E. Cummins, et al. (2005). "Clinical and cost-effectiveness of autologous chondrocyte implantation for cartilage defects in knee joints: systematic review and economic evaluation." Health Technology Assessment 9(47): III-+.
Clark, J. M. (1991). "Variation of collagen fiber alignment in a joint surface - a scanning electron-microscope study of the tibial plateau in dog, rabbit, and man." Journal of Orthopaedic Research 9(2): 246-257.
Clarke, I. C. (1971). "Articular cartilage a review and scanning electron microscope study part 1 the inter territorial fibrillar architecture." Journal of Bone and Joint Surgery British Volume 53(4): 732-750.
Cohen, N. P., R. J. Foster, et al. (1998). "Composition and dynamics of articular cartilage: Structure, function, and maintaining healthy state." Journal of Orthopaedic & Sports Physical Therapy 28(4): 203-215.
Cole, A. G. (2011). "A review of diversity in the evolution and development of cartilage: the search for the origin of the chondrocyte." European Cells & Materials 21: 122-129.
Cole, A. G. and B. K. Hall (2004). "Cartilage is a metazoan tissue; integrating data from nonvertebrate sources." Acta Zoologica 85(2): 69-80.
Convery, F. R., W. H. Akeson, et al. (1972). "The repair of large osteochondral defects. An experimental study in horses." Clinical orthopaedics and related research 82: 253-62.
179
Csucs, G., R. Michel, et al. (2003). "Microcontact printing of novel co-polymers in combination with proteins for cell-biological applications." Biomaterials 24(10): 1713- 1720.
Curl, W. W., J. Krome, et al. (1997). "Cartilage injuries: A review of 31,516 knee arthroscopies." Arthroscopy 13(4): 456-460.
Delon, I. and N. H. Brown (2007). "Integrins and the actin cytoskeleton." Current Opinion in Cell Biology 19(1): 43-50.
DeMali, K. A., K. Wennerberg, et al. (2003). "Integrin signaling to the actin cytoskeleton." Current Opinion in Cell Biology 15(5): 572-582.
Detterline, A. J., S. Goldberg, et al. (2005). "Treatment options for articular cartilage defects of the knee." Orthopaedic Nursing 24(5): 361-366.
Dickhut, A., E. Gottwald, et al. (2008). "Chondrogenesis of mesenchymal stem cells in gel-like biomaterials in vitro and in vivo." Frontiers in Bioscience 13: 4517-4528.
Dickhut, A., E. Gottwald, et al. (2008). "Chondrogenesis of mesenchymal stem cells in gel-like biomaterials in vitro and in vivo." Front Biosci 13: 4517-28.
Dickinson, R. B., S. Guido, et al. (1994). "Biased cell-migration of fibroblasts exhibiting contact guidance in oriented collagen gels." Annals of Biomedical Engineering 22(4): 342-356.
Djouad, F., D. Mrugala, et al. (2006). "Engineered mesenchymal stem cells for cartilage repair." Regenerative Medicine 1(4): 529-537.
Djouad, F., D. Mrugala, et al. (2006). "Engineered mesenchymal stem cells for cartilage repair." Regen Med 1(4): 529-37.
Dunn, G. A. and A. F. Brown (1986). "Alignment of fibroblasts on grooved surfaces described by a simple geometric transformatioN." Journal of Cell Science 83: 313-340.
Eaglstein, W. H., O. M. Alvarez, et al. (1999). "Acute excisional wounds treated with a tissue-engineered skin (Apligraf)." Dermatologic Surgery 25(3): 195-201.
Eastwood, M., V. C. Mudera, et al. (1998). "Effect of precise mechanical loading on fibroblast populated collagen lattices: Morphological changes." Cell Motility and the Cytoskeleton 40(1): 13-21.
Erickson, I. E., A. H. Huang, et al. (2009). "Macromer density influences mesenchymal stem cell chondrogenesis and maturation in photocrosslinked hyaluronic acid hydrogels." Osteoarthritis and Cartilage 17(12): 1639-1648.
180
Erickson, I. E., S. R. Kestle, et al. (2012). "High mesenchyrnal stem cell seeding densities in hyaluronic acid hydrogels produce engineered cartilage with native tissue properties." Acta Biomaterialia 8(8): 3027-3034.
Fedewa, M. M., T. R. Oegema, et al. (1998). "Chondrocytes in culture produce a mechanically functional tissue." Journal of Orthopaedic Research 16(2): 227-236.
Ficat, R. P., C. Ficat, et al. (1979). "Spongialization - new treatment for diseased patellae." Clinical Orthopaedics and Related Research(144): 74-83.
Francioli, S. E., C. Candrian, et al. (2010). "Effect of three-dimensional expansion and cell seeding density on the cartilage-forming capacity of human articular chondrocytes in type II collagen sponges." Journal of Biomedical Materials Research Part A 95A(3): 924- 931.
Freed, L. E., G. Vunjaknovakovic, et al. (1994). "Biodegradable polymer scaffolds for tissue engineering." Bio-Technology 12(7): 689-693.
Frenkel, S. R., B. Toolan, et al. (1997). "Chondrocyte transplantation using a collagen bilayer matrix for cartilage repair." Journal of Bone and Joint Surgery-British Volume 79B(5): 831-836.
Friedenstein, A. J. (1995). "Marrow stromal fibroblasts." Calcified Tissue International 56: S17-S17.
Gemmiti, C. V. and R. E. Guldberg (2006). "Fluid flow increases type II collagen deposition and tensile mechanical properties in bioreactor-grown tissue-engineered cartilage." Tissue Engineering 12(3): 469-479.
Gendler, E., S. Gendler, et al. (1984). "Toxic reactions evoked by glutaraldehyde-fixed pericardium and cardiac-valve tissue bioprosthesis." Journal of Biomedical Materials Research 18(7): 727-736.
Gendler, E., S. Gendler, et al. (1984). "Toxic reactions evoked by glutaraldehyde-fixed pericardium and cardiac valve tissue bioprosthesis." J Biomed Mater Res 18(7): 727-36.
Gilbert, J. E. (1998). "Current treatment options for the restoration of articular cartilage." Am J Knee Surg 11(1): 42-6.
Goldring, M. B. (2000). "The role of the chondrocyte in osteoarthritis." Arthritis and Rheumatism 43(9): 1916-1926.
Gong, Y. Y., J. X. Xue, et al. (2011). "A sandwich model for engineering cartilage with acellular cartilage sheets and chondrocytes." Biomaterials 32(9): 2265-2273.
181
Gooding, C. R., W. Bartlett, et al. (2006). "A prospective, ranomised study comparing two techniques of autologous chondrocyte implantation for osteochondral defects in the knee: Periosteum covered versus type I/III collagen covered." Knee 13(3): 203-210.
Gratz, K. R., V. W. Wong, et al. (2006). "Biomechanical assessment of tissue retrieved after in vivo cartilage defect repair: tensile modulus of repair tissue and integration with host cartilage." Journal of Biomechanics 39(1): 138-146.
Grayson, W. L., F. Zhao, et al. (2006). "Effects of hypoxia on human mesenchymal stem cell expansion and plasticity in 3D constructs." Journal of Cellular Physiology 207(2): 331-339.
Hangody, L., P. Feczko, et al. (2001). "Mosaicplasty for the treatment of articular defects of the knee and ankle." Clinical Orthopaedics and Related Research(391): S328-S336.
Hangody, L., G. Vasarhelyi, et al. (2008). "Autologous osteochondral grafting-technique and long-term results." Injury-International Journal of the Care of the Injured 39: S32- S39.
Hardingham, T. E. and A. J. Fosang (1992). "Proteoglycans - many forms and many functions." Faseb Journal 6(3): 861-870.
Harley, B. A., A. K. Lynn, et al. (2010). "Design of a multiphase osteochondral scaffold. II. Fabrication of a mineralized collagen-glycosaminoglycan scaffold." Journal of Biomedical Materials Research Part A 92A(3): 1066-1077.
Hasler, E. M., W. Herzog, et al. (1999). "Articular cartilage biomechanics: Theoretical models, material properties, and biosynthetic response." Critical Reviews in Biomedical Engineering 27(6): 415-488.
Hata, R. I., H. Hori, et al. (1984). "Selective-inhibition of type-i collagen-synthesis in osteoblastic cells by epidermal growth-factor." Endocrinology 115(3): 867-876.
Helmick, C. G., D. T. Felson, et al. (2008). "Estimates of the prevalence of arthritis and other rheumatic conditions in the United States." Arthritis and Rheumatism 58(1): 15-25.
Heng, B. C., T. Cao, et al. (2004). "Directing stem cell differentiation into the chondrogenic lineage in vitro." Stem Cells 22(7): 1152-1167.
Hice, G., D. Freedman, et al. (1990). "Scanning and light microscopic study of irrigated and nonirrigated joints following burr surgery performed through a small incision." Journal of Foot Surgery 29(4): 337-344.
Hirono, T., K. Torimitsu, et al. (1988). "Recognition of artificial microstructures by sensory nerve-fibers in culture." Brain Research 446(1): 189-194.
182
Hoffman-Kim, D., J. A. Mitchel, et al. (2010). Topography, Cell Response, and Nerve Regeneration. Annual Review of Biomedical Engineering, Vol 12. 12: 203-231.
Huang, A. H., B. M. Baker, et al. (2012). "Sliding contact loading enhances the tensile properties of mesenchymal stem cell-seeded hydrogels." European Cells & Materials 24: 29-45.
Huang, A. H., M. J. Farrell, et al. (2010). "Long-term dynamic loading improves the mechanical properties of chondrogenic mesenchymal stem cell-laden hydrogels." European Cells & Materials 19: 72-85.
Huang, A. H., M. J. Farrell, et al. (2011). "Mechanics and mechanobiology of mesenchymal stem cell-based engineered cartilage." Journal of Biomechanics 43(1): 128- 136.
Huang, C. Y., A. Stankiewicz, et al. (2005). "Anisotropy, inhomogeneity, and tension- compression nonlinearity of human glenohumeral cartilage in finite deformation." Journal of Biomechanics 38(4): 799-809.
Huang, N. F., R. J. Lee, et al. (2010). "Engineering of aligned skeletal muscle by micropatterning." American journal of translational research 2(1): 43-55.
Hunziker, E. B. (1999). "Articular cartilage repair: are the intrinsic biological constraints undermining this process insuperable?" Osteoarthritis and Cartilage 7(1): 15-28.
Hunziker, E. B. (2002). "Articular cartilage repair: basic science and clinical progress. A review of the current status and prospects." Osteoarthritis and Cartilage 10(6): 432-463.
Hunziker, E. B., M. Michel, et al. (1997). "Ultrastructure of adult human articular cartilage matrix after cryotechnical processing." Microsc Res Tech 37(4): 271-84.
Hutmacher, D. W. (2000). "Scaffolds in tissue engineering bone and cartilage." Biomaterials 21(24): 2529-2543.
Huttenlocher, A. and A. R. Horwitz "Integrins in cell migration." Cold Spring Harbor perspectives in biology 3(9): a005074-a005074.
Hwang, C. M., Y. Park, et al. (2009). "Controlled cellular orientation on PLGA microfibers with defined diameters." Biomedical Microdevices 11(4): 739-746.
Im, G. I., D. Y. Kim, et al. (2001). "Repair of cartilage defect in the rabbit with cultured mesenchymal stem cells from bone marrow." Journal of Bone and Joint Surgery-British Volume 83B(2): 289-294.
183
Insall, J. N. (1967). "Intra-articular surgery for degenerative arthritis of the knee. A report of the work of the late K. H. Pridie." The Journal of bone and joint surgery. British volume 49(2): 211-28.
Jackson, D. W., P. A. Lalor, et al. (2001). "Spontaneous repair of full-thickness defects of articular cartilage in a goat model - A preliminary study." Journal of Bone and Joint Surgery-American Volume 83A(1): 53-64.
Jackson, R. W. and C. Dietrichs (2003). "The results of arthroscopic lavage and debridement of osteoarthritic knees based on the severity of degeneration: A 4-to 6-year symptomatic follow-up." Arthroscopy-the Journal of Arthroscopic and Related Surgery 19(1): 13-20.
Jakobsen, R. B., A. Shahdadfar, et al. (2010). "Chondrogenesis in a hyaluronic acid scaffold: comparison between chondrocytes and MSC from bone marrow and adipose tissue." Knee Surg Sports Traumatol Arthrosc 18(10): 1407-16.
Jakobsen, R. B., A. Shandadfar, et al. (2010). "Chondrogenesis in a hyaluronic acid scaffold: comparison between chondrocytes and MSC from bone marrow and adipose tissue." Knee Surgery Sports Traumatology Arthroscopy 18(10): 1407-1416.
Janakiraman, V., B. L. Kienitz, et al. (2007). "Lithography technique for topographical micropatterning of collagen-glycosaminoglycan membranes for tissue engineering applications." J Med Device 1(3): 233-237.
Janjanin, S., W.-J. Li, et al. (2008). "Mold-shaped, nanofiber scaffold-based cartilage engineering using human mesenchymal stem cells and bioreactor." Journal of Surgical Research 149(1): 47-56.
Jason R. Fritz, D. P. a. H. S. C. (2009). "Current Challenges in Cartilage Tissue Engineering: A Review of Current Cellular-Based Therapies." Current Rheumatology Reviews 5(1): 8-14 (7)
Johnstone, B. and J. U. Yoo (1999). "Autologous mesenchymal progenitor cells in articular cartilage repair." Clinical Orthopaedics and Related Research(367): S156-S162.
Kane, R. S., S. Takayama, et al. (1999). "Patterning proteins and cells using soft lithography." Biomaterials 20(23-24): 2363-2376.
Kanichai, M., D. Ferguson, et al. (2008). "Hypoxia promotes chondrogenesis in rat mesenchymal stem cells: A role for AKT and hypoxia-inducible factor (HIF)-1 alpha." Journal of Cellular Physiology 216(3): 708-715.
Kapoor, A., E. H. G. Caporali, et al. (2010). "Microtopographically patterned surfaces promote the alignment of tenocytes and extracellular collagen." Acta Biomaterialia 6(7): 2580-2589.
184
Kasemkijwattana, C., S. Kesprayura, et al. (2009). "Autologous Chondrocytes Implantation with Three-Dimensional Collagen Scaffold." Journal of the Medical Association of Thailand 92(10): 1282-1286.
Kempson, G. E., M. A. R. Freeman, et al. (1968). "Tensile properties of articular cartilage." Nature 220(5172): 1127-&.
Khan, I. M., S. J. Gilbert, et al. (2008). "Cartilage integration: evaluation of the reasons for failure of integration during cartilage repair. A review." European Cells & Materials 16: 26-39.
Kingshott, P. and H. J. Griesser (1999). "Surfaces that resist bioadhesion." Current Opinion in Solid State & Materials Science 4(4): 403-412.
Kuo, C. K., W. J. Li, et al. (2006). "Cartilage tissue engineering: its potential and uses." Current Opinion in Rheumatology 18(1): 64-73.
Lai, W. M., J. S. Hou, et al. (1991). "A triphasic theory for the swelling and deformation behaviors of articular-cartilage." Journal of Biomechanical Engineering-Transactions of the Asme 113(3): 245-258.
Lane, J. G., J. B. Massie, et al. (2004). "Follow-up of osteochondral plug transfers in a goat model - A 6-month study." American Journal of Sports Medicine 32(6): 1440-1450.
Lane, J. M., C. T. Brighton, et al. (1977). "Anaerobic and aerobic metabolism in articular-cartilage." Journal of Rheumatology 4(4): 334-342.
Langer, R. and J. P. Vacanti (1993). "Tissue engineeringI." Science 260(5110): 920-926.
Lee, C. H., A. Singla, et al. (2001). "Biomedical applications of collagen." International Journal of Pharmaceutics 221(1-2): 1-22.
Lee, C. R., H. A. Breinan, et al. (2000). "Articular cartilage chondrocytes in type I and type II collagen-GAG matrices exhibit contractile behavior in vitro." Tissue Engineering 6(5): 555-565.
Lee, E. H. and J. H. P. Hui (2006). "The potential of stem cells in orthopaedic surgery." Journal of Bone and Joint Surgery-British Volume 88B(7): 841-851.
Lee, G. M. and R. F. Loeser (1999). "Cell surface receptors transmit sufficient force to bend collagen fibrils." Experimental Cell Research 248(1): 294-305.
Lee, R. B. and J. P. G. Urban (1997). "Evidence for a negative Pasteur effect in articular cartilage." Biochemical Journal 321: 95-102.
185
Lee, R. B., R. J. Wilkins, et al. (2002). "The effect of mechanical stress on cartilage energy metabolism." Biorheology 39(1-2): 133-143.
Lennon, D. P., J. M. Edmison, et al. (2001). "Cultivation of rat marrow-derived mesenchymal stem cells in reduced oxygen tension: Effects on in vitro and in vivo osteochondrogenesis." Journal of Cellular Physiology 187(3): 345-355.
Li, J. T. and K. D. Caldwell (1996). "Plasma protein interactions with Pluronic(TM)- treated colloids." Colloids and Surfaces B-Biointerfaces 7(1-2): 9-22.
Li, W. J., R. Tuli, et al. (2005). "A three-dimensional nanofibrous scaffold for cartilage tissue engineering using human mesenchymal stem cells." Biomaterials 26(6): 599-609.
Liang, W.-H., B. L. Kienitz, et al. (2010). "Concentrated collagen-chondroitin sulfate scaffolds for tissue engineering applications." Journal of Biomedical Materials Research Part A 94A(4): 1050-1060.
Liang, W. H., B. L. Kienitz, et al. (2010). "Concentrated collagen-chondroitin sulfate scaffolds for tissue engineering applications." J Biomed Mater Res A 94(4): 1050-60.
Linn, F. C. and L. Sokoloff (1965). "Movement and composition of interstitial fluid of cartilage." Arthritis and Rheumatism 8(4): 481-&.
Liu, V. A., W. E. Jastromb, et al. (2002). "Engineering protein and cell adhesivity using PEO-terminated triblock polymers." Journal of Biomedical Materials Research 60(1): 126-134.
Lu, Q. J., A. Simionescu, et al. (2005). "Novel capillary channel fiber scaffolds for guided tissue engineering." Acta Biomaterialia 1(6): 607-614.
Lu, X. L. and V. C. Mow (2008). "Biomechanics of articular cartilage and determination of material properties." Medicine and Science in Sports and Exercise 40(2): 193-199.
Magne, D., C. Vinatier, et al. (2005). "Mesenchymal stem cell therapy to rebuild cartilage." Trends in Molecular Medicine 11(11): 519-526.
Mankin, H. J. (1974). "Reaction of articular-cartilage to injury and osteoarthritis .1." New England Journal of Medicine 291(24): 1285-1292.
Mankin, H. J. and A. Z. Thrasher (1975). "Water-content and binding in normal and osteoarthritic human cartilage." Journal of Bone and Joint Surgery-American Volume A 57(1): 76-80.
Manwaring, M. E., J. F. Walsh, et al. (2004). "Contact guidance induced organization of extracellular matrix." Biomaterials 25(17): 3631-3638.
186
Maroudas, A. (1979). "Physico-chemical properties of articular cartilage." In: Freeman M.A.R., editor. Adult Articular Cartilage. Tunbridge Wells, England: Pitman Medical: 215-290.
McLaren, A. C., C. P. Blokker, et al. (1991). "Arthroscopic debridement of the knee for osteoarthrosis." Canadian Journal of Surgery 34(6): 595-598.
Meyle, J., K. Gultig, et al. (1995). "Variation in contact guidance by human-cells on a microstructured surface (VOL 29, PG 81, 1995)." Journal of Biomedical Materials Research 29(7): 905-905.
Michaud, C. M., M. T. McKenna, et al. (2006). "The burden of disease and injury in the United States 1996." Population health metrics 4: 11-11.
Minas, T. (2001). "Autologous chondrocyte implantation for focal chondral defects of the knee." Clinical Orthopaedics and Related Research(391): S349-S361.
MMWR (2001). CDC. Prevalence of disabilities and associated health conditions among adults-United States, 1999. Morbidity and Mortality Weekly Report. 50: 120-125.
MMWR (2007). "National and state medical expenditures and lost earnings attributable to arthritis and other rheumatic conditions--United States, 2003." MMWR. Morbidity and mortality weekly report 56(1): 4-7.
MMWR (2010). "Prevalence of doctor-diagnosed arthritis and arthritis-attributable activity limitation --- United States, 2007-2009." MMWR. Morbidity and mortality weekly report 59(39): 1261-5.
Mochitate, K., P. Pawelek, et al. (1991). "Stress-relaxation of contracted collagen gels - disruption of actin filament bundles, release of cell-surface fibronectin, and down- regulation of dna and protein-synthesis." Experimental Cell Research 193(1): 198-207.
Moutos, F. T., B. T. Estes, et al. "Multifunctional Hybrid Three-dimensionally Woven Scaffolds for Cartilage Tissue Engineering." Macromolecular Bioscience 10(11): 1355- 1364.
Mrksich, M., L. E. Dike, et al. (1997). "Using microcontact printing to pattern the attachment of mammalian cells to self-assembled monolayers of alkanethiolates on transparent films of gold and silver." Experimental Cell Research 235(2): 305-313.
Nayyer, L., K. H. Patel, et al. "Tissue engineering: revolution and challenge in auricular cartilage reconstruction." Plastic and Reconstructive Surgery 129(5): 1123-1137.
187
Nehrer, S., H. A. Breinan, et al. (1997). "Canine chondrocytes seeded in type I and type II collagen implants investigated in vitro." Journal of Biomedical Materials Research 38(2): 95-104.
Nehrer, S., M. Spector, et al. (1999). "Histologic analysis of tissue after failed cartilage repair procedures." Clinical Orthopaedics and Related Research(365): 149-162.
Nerurkar, N. L., W. Han, et al. (2011). "Homologous structure-function relationships between native fibrocartilage and tissue engineered from MSC-seeded nanofibrous scaffolds." Biomaterials 32(2): 461-468.
Nettles, D. L., A. Chilkoti, et al. (2009). "Early metabolite levels predict long-term matrix accumulation for chondrocytes in elastin-like polypeptide biopolymer scaffolds." Tissue Engineering Part A 15(8): 2113-2121.
Noeth, U., L. Rackwitz, et al. (2007). "Chondrogenic differentiation of human mesenchymal stem cells in collagen type I hydrogels." Journal of Biomedical Materials Research Part A 83A(3): 626-635.
O'Driscoll, S. W. (1998). "The healing and regeneration of articular cartilage." Journal of Bone and Joint Surgery-American Volume 80A(12): 1795-1812.
Ohsawa, S., N. Yasui, et al. (1982). "Contraction of collagen gel by the dedifferentiated chondrocyteS." Cell Biology International Reports 6(8): 767-774.
Otte, P. (1991). "Basic cell-metabolism of articular-cartilage - manometric studies." Zeitschrift Fur Rheumatologie 50(5): 304-312.
Ousema, P. H., F. T. Moutos, et al. (2012). "The inhibition by interleukin 1 of MSC chondrogenesis and the development of biomechanical properties in biomimetic 3D woven PCL scaffolds." Biomaterials 33(35): 8967-8974.
Owen, M. and A. J. Friedenstein (1988). "Stromal stem-cells - marrow-derived osteogenic precursors." Ciba Foundation Symposia 136: 42-60.
Palecek, S. P., J. C. Loftus, et al. (1997). "Integrin-ligand binding properties govern cell migration speed through cell-substratum adhesiveness." Nature 385(6616): 537-540.
Parenteau-Bareil, R., R. Gauvin, et al. (2010). "Collagen-based biomaterials for tissue engineering applications." Materials 3(3): 1863-1887.
Patel, N., R. Padera, et al. (1998). "Spatially controlled cell engineering on biodegradable polymer surfaces." Faseb Journal 12(14): 1447-1454.
188
Pattappa, G., H. K. Heywood, et al. "The metabolism of human mesenchymal stem cells during proliferation and differentiation." Journal of Cellular Physiology 226(10): 2562- 2570.
Pelaez, D., C.-Y. C. Huang, et al. (2009). "Cyclic compression maintains viability and induces chondrogenesis of human mesenchymal stem cells in fibrin gel scaffolds." Stem Cells and Development 18(1): 93-102.
Peloso, P. M. and J. M. Scheiman (2001). "The economic implications of cyclooxygenase-2-specific inhibitors." American Journal of Medicine 110(Supplement 3A): 50S-54S.
Peterson, L., T. Minas, et al. (2000). "Two- to 9-year outcome after autologous chondrocyte transplantation of the knee." Clinical Orthopaedics and Related Research(374): 212-234.
Pfander, D. and K. Gelse (2007). "Hypoxia and osteoarthritis: how chondrocytes survive hypoxic environments." Current Opinion in Rheumatology 19(5): 457-462.
Pieter K Bos , M. L. v. M., G.J.v. Osch (2010). "Articular cartilage repair and the evolving role of regenerative medicine " Open Access Surgery 3(1): 109-122.
Pittenger, M. F., A. M. Mackay, et al. (1999). "Multilineage potential of adult human mesenchymal stem cells." Science 284(5411): 143-147.
Ponticiello, M. S., R. M. Schinagl, et al. (2000). "Gelatin-based resorbable sponge as a carrier matrix for human mesenchymal stem cells in cartilage regeneration therapy." Journal of Biomedical Materials Research 52(2): 246-255.
Redman, S. N., S. F. Oldfield, et al. (2005). "Current strategies for articular cartilage repair." European Cells & Materials 9: 23-32; discussion 23-32.
Risbud, M. V. and M. Sittinger (2002). "Tissue engineering: advances in in vitro cartilage generation." Trends in Biotechnology 20(8): 351-356.
Roach, P., T. Parker, et al. (2010). "Surface strategies for control of neuronal cell adhesion: A review." Surface Science Reports 65(6): 145-173.
Ruoslahti, E. (1996). "RGD and other recognition sequences for integrins." Annual Review of Cell and Developmental Biology 12: 697-715.
Salinas, C. N. and K. S. Anseth (2009). "Mesenchymal stem cells for craniofacial tissue regeneration: designing hydrogel delivery vehicles." J Dent Res 88(8): 681-92.
189
Schinagl, R. M., D. Gurskis, et al. (1997). "Depth-dependent confined compression modulus of full-thickness bovine articular cartilage." Journal of Orthopaedic Research 15(4): 499-506.
Schiro, J. A., B. M. C. Chan, et al. (1991). "Integrin-alpha-2-beta-1 (vla-2) mediates reorganization and contraction of collagen matrices by human-cells." Cell 67(2): 403-410.
Schmalenberg, K. E. and K. E. Uhrich (2005). "Micropatterned polymer substrates control alignment of proliferating Schwann cells to direct neuronal regeneration." Biomaterials 26(12): 1423-1430.
Schmidt, C. E. and J. M. Baier (2000). "Acellular vascular tissues: natural biomaterials for tissue repair and tissue engineering." Biomaterials 21(22): 2215-2231.
Schmidt, H., K. J. Schulze, et al. (1988). "Results in the treatment of cartilage lesions by drilling according to the pridie method." Beitraege zur Orthopaedie und Traumatologie 35(3): 117-122.
Schnabel, M., S. Marlovits, et al. (2002). "Dedifferentiation-associated changes in morphology and gene expression in primary human articular chondrocytes in cell culture." Osteoarthritis and Cartilage 10(1): 62-70.
Setton, L. (2008). "Polymer therapeutics - Reservoir drugs." Nature Materials 7(3): 172- 174.
Setton, L. A., D. M. Elliott, et al. (1999). "Altered mechanics of cartilage with osteoarthritis: human osteoarthritis and an experimental model of joint degeneration." Osteoarthritis and Cartilage 7(1): 2-14.
Shapiro, F., S. Koide, et al. (1993). "Cell origin and differentiation in the repair of full- thickness defects of articular-cartilage." Journal of Bone and Joint Surgery-American Volume 75A(4): 532-553.
Sharma, C. G., Sneh; Dinda, Amit K.; Mishra, Narayan C (2011). "Cartilage tissue engineering: current scenario and challenges." Advanced Materials Letters 2(2): 90.
Shimizu, C., M. Yoshioka, et al. (1998). "Long-term effects of hyaluronan on experimental osteoarthritis in the rabbit knee." Osteoarthritis and Cartilage 6(1): 1-9.
Shirazi, R., A. Shirazi-Adl, et al. (2008). "Role of cartilage collagen fibrils networks in knee joint biomechanics under compression." Journal of Biomechanics 41(16): 3340- 3348.
Simon, L. S. (1999). "Osteoarthritis: a review." Clinical cornerstone 2(2): 26-37.
190
Smith, G. D., J. B. Richardson, et al. (2003). "Autologous chondrocyte implantation and osteochondral cylinder transplantation in cartilage repair of the knee joint." Journal of Bone and Joint Surgery-American Volume 85A(12): 2487-2488.
Solchaga, L. A., J. F. Welter, et al. (2004). "Generation of pluripotent stem cells and their differentiation to the chondrocytic phenotype." Cartilage and Osteoarthritis, Vol 1: Cellular and Molecular Tools 100: 53-67.
Song, L., D. Baksh, et al. (2004). "Mesenchymal stem cell-based cartilage tissue engineering: cells, scaffold and biology." Cytotherapy 6(6): 596-601.
Sorensen, A., T. Alekseeva, et al. (2007). "Long-term neurite orientation on astrocyte monolayers. aligned by microtopography." Biomaterials 28(36): 5498-5508.
Spadaccio, C., A. Rainer, et al. (2009). "Poly-L-lactic acid/hydroxyapatite electrospun nanocomposites induce chondrogenic differentiation of human MSC." Ann Biomed Eng 37(7): 1376-89.
Speer, D. P., M. Chvapil, et al. (1979). "Enhancement of healing in osteochondral defects by collagen sponge implants." Clinical Orthopaedics and Related Research(144): 326- 335.
Steadman, J. R., K. K. Briggs, et al. (2003). "Outcomes of microfracture for traumatic chondral defects of the knee: Average 11-year follow-up." Arthroscopy: The Journal of Arthroscopic & Related Surgery 19(5): 477-484.
Stockwell, R. A. "Biological structure and function", Vol. 7. Biology of Cartilage Cells. Viii+329p. Cambridge University Press: New York, N.Y., USA; Cambridge, England. Illus
Tan, J. L., W. Liu, et al. (2004). "Simple approach to micropattern cells on common culture substrates by tuning substrate wettability." Tissue Engineering 10(5-6): 865-872.
Tat, S. K., J.-P. Pelletier, et al. (2009). "Treatment with ephrin B2 positively impacts the abnormal metabolism of human osteoarthritic chondrocytes." Arthritis Research & Therapy 11(4).
Teixeira, L. S. M., J. C. H. Leijten, et al. "High throughput generated micro-aggregates of chondrocytes stimulate cartilage formation in vitro and in vivo." European Cells & Materials 23: 387-399.
Temenoff, J. S. and A. G. Mikos (2000). "Review: tissue engineering for regeneration of articular cartilage." Biomaterials 21(5): 431-440.
Tessier-Lavigne, M. and C. S. Goodman (1996). "The molecular biology of axon guidance." Science 274(5290): 1123-33.
191
Tharmalingam, T., H. Ghebeh, et al. (2008). "Pluronic enhances the robustness and reduces the cell attachment of mammalian cells." Molecular Biotechnology 39(2): 167- 177.
Thomson, J. A., J. Itskovitz-Eldor, et al. (1998). "Embryonic stem cell lines derived from human blastocysts." Science 282(5391): 1145-1147.
Tognana, E., A. Borrione, et al. (2007). "Hyalograft (R) C: Hyaluronan-based scaffolds in tissue-engineered cartilage." Cells Tissues Organs 186(2): 97-103.
Toh, W. S., E. H. Lee, et al. (2011). "Potential of Human Embryonic Stem Cells in Cartilage Tissue Engineering and Regenerative Medicine." Stem Cell Reviews and Reports 7(3): 544-559.
Tomasek, J. J., C. J. Haaksma, et al. (1992). "Fibroblast contraction occurs on release of tension in attached collagen lattices - dependency on an organized actin cytoskeleton and serum." Anatomical Record 232(3): 359-368.
Tuan, R. S., G. Boland, et al. (2003). "Adult mesenchymal stem cells and cell-based tissue engineering." Arthritis Research & Therapy 5(1): 32-45.
Van Susante, J. L. C., J. Pieper, et al. (2001). "Linkage of chondroitin-sulfate to type I collagen scaffolds stimulates the bioactivity of seeded chondrocytes in vitro." Biomaterials 22(17): 2359-2369.
Wakitani, S., T. Goto, et al. (1994). "Mesenchymal cell-based repair of large, full- thickness defects of articular-cartilage." Journal of Bone and Joint Surgery-American Volume 76A(4): 579-592.
Wang, J. H. C., E. S. Grood, et al. (2000). "Alignment and proliferation of MC3T3-E1 osteoblasts in microgrooved silicone substrata subjected to cyclic stretching." Journal of Biomechanics 33(6): 729-735.
Wang, J. H. C., F. Y. Jia, et al. (2003). "Cell orientation determines the alignment of cell- produced collagenous matrix." Journal of Biomechanics 36(1): 97-102.
Wang, Y., U. J. Kim, et al. (2005). "In vitro cartilage tissue engineering with 3D porous aqueous-derived silk scaffolds and mesenchymal stem cells." Biomaterials 26(34): 7082- 94.
Wang, Y. C. and C. C. Ho (2004). "Micropatterning of proteins and mammalian cells on biomaterials." FASEB Journal 18(1): 525-+.
Wang, Y. Z., D. J. Blasioli, et al. (2006). "Cartilage tissue engineering with silk scaffolds and human articular chondrocytes." Biomaterials 27(25): 4434-4442.
192
Wang, Y. Z., U. J. Kim, et al. (2005). "In vitro cartilage tissue engineering with 3D porous aqueous-derived silk scaffolds and mesenchymal stem cells." Biomaterials 26(34): 7082-7094.
Wei, Y., W. Zeng, et al. (2012). "Chondrogenic differentiation of induced pluripotent stem cells from osteoarthritic chondrocytes in alginate matrix." European Cells & Materials 23: 1-12.
Weiss, P. (1934). "In vitro experiments on the factors determining the course of the outgrowing nerve fiber." Journal of Experimental Zoology 68(3): 393-448.
Weiss, P. (1945). "Experiments on cell and axon orientation in vitro; the role of colloidal exudates in tissue organization." The Journal of experimental zoology 100: 353-86.
Welter, J. F., L. A. Solchaga, et al. (2007). "Simplification of aggregate culture of human mesenchymal stem cells as a chondrogenic screening assay." Biotechniques 42(6): 732, 734-7.
Whitesides, G. M., E. Ostuni, et al. (2001). "Soft lithography in biology and biochemistry." Annu Rev Biomed Eng 3: 335-73.
Williams, G. M., E. F. Chan, et al. (2010). "Shape, loading, and motion in the bioengineering design, fabrication, and testing of personalized synovial joints." Journal of Biomechanics 43(1): 156-165.
Williams, G. M., T. J. Klein, et al. (2005). "Cell density alters matrix accumulation in two distinct fractions and the mechanical integrity of alginate-chondrocyte constructs." Acta Biomaterialia 1(6): 625-633.
Williams, R. J. and S. C. Gamradt (2008). "Articular cartilage repair using a resorbable matrix scaffold." Instructional course lectures 57: 563-71.
Wilson, W., J. M. Huyghe, et al. (2007). "Depth-dependent compressive equilibrium properties of articular cartilage explained by its composition." Biomechanics and Modeling in Mechanobiology 6(1-2): 43-53.
Wilson, W., C. C. van Donkelaar, et al. (2005). "A fibril-reinforced poroviscoelastic swelling model for articular cartilage." Journal of Biomechanics 38(6): 1195-1204.
Wilson, W., C. C. van Donkelaar, et al. (2004). "Stresses in the local collagen network of articular cartilage: a poroviscoelastic fibril-reinforced finite element study." Journal of Biomechanics 37(3): 357-366.
Wilson, W., C. C. van Donkelaar, et al. (2005). "Stresses in the local collagen network of articular cartilage: a poroviscoelastic fibril-reinforced finite element study (vol 37, pg 357, 2004)." Journal of Biomechanics 38(10): 2138-2140.
193
Wong, M., P. Wuethrich, et al. (1996). "Zone-specific cell biosynthetic activity in mature bovine articular cartilage: A new method using confocal microscopic stereology and quantitative autoradiography." Journal of Orthopaedic Research 14(3): 424-432.
Woo, S. L. Y., W. H. Akeson, et al. (1976). "Measurements of nonhomogeneous, directional mechanical properties of articular cartilage in tension." Journal of Biomechanics 9(12): 785-791.
Wynn, J. J. and C. E. Alexander "Increasing organ donation and transplantation: the US experience over the past decade." Transplant International 24(4): 324-332.
Xue, J. X., Y. Y. Gong, et al. (2012). "Chondrogenic differentiation of bone marrow- derived mesenchymal stem cells induced by acellular cartilage sheets." Biomaterials 33(24): 5832-40.
Yahyouche, A., X. Zhidao, et al. (2011). "Macrophage-mediated degradation of crosslinked collagen scaffolds." Acta Biomater 7(1): 278-86.
Yang, Q., J. Peng, et al. (2008). "A cartilage ECM-derived 3-D porous acellular matrix scaffold for in vivo cartilage tissue engineering with PKH26-labeled chondrogenic bone marrow-derived mesenchymal stem cells." Biomaterials 29(15): 2378-2387.
Yannas, I. V., J. F. Burke, et al. (1980). "Design of an artificial skin .2. Control of chemical-composition." Journal of Biomedical Materials Research 14(2): 107-132.
Zwingmann, J., A. T. Mehlhorn, et al. (2007). "Chondrogenic differentiation of human articular chondrocytes differs in biodegradable PGA/PLA scaffolds." Tissue Engineering 13(9): 2335-2343.
194