DESIGN AND CHARACTERIZATION OF PHOTOPOLYMERIZABLE SEMI-
INTERPENETRATING NETWORKS FOR IN VITRO CHONDROGENESIS OF
HUMAN MESENCHYMAL STEM CELLS
by
AMANDA NICOLE BUXTON
Submitted in partial fulfillment of the requirements
For the degree of Doctor of Philosophy
Dissertation Adviser: Dr. Brian Johnstone
Department of Biomedical Engineering
CASE WESTERN RESERVE UNIVERSITY
August, 2007 CASE WESTERN RESERVE UNIVERSITY
SCHOOL OF GRADUATE STUDIES
We hereby approve the thesis/dissertation of
______Amanda N. Buxton______
candidate for the Doctor of Philosophy degree *.
(signed)_____Roger Marchant______(chair of the committee)
______Stuart Rowan______
______Steve Eppell______
______Brian Johnstone______
______Joseph Mansour______
______Lloyd A. Culp______
(date) ___April 3, 2007______
*We also certify that written approval has been obtained for any proprietary material contained therein.
TABLE OF CONTENTS
List of Tables iii
List of Figures iv
Acknowledgements vii
List of Abbreviations viii
Glossary ix
Abstract 1
Chapter One: Background & Significance 3
Structure & Function of Articular Cartilage 4
Formation of Cartilage 13
Articular Cartilage Pathology & Repair 17
Tissue Engineering Design Criteria 20
Cell Source 21
Scaffold Considerations 22
Addition of Bioactive Factors 25
Chapter Two: In Vitro Chondrogenesis in a Poly(ethylene glycol)
Diacrylate (PEGDA) Scaffold – Preliminary Experiments 30
In Vitro Chondrogenesis in Pellet Culture 30
MSC Encapsulation in 20% (w:v) PEGDA (6 kDa) Hydrogels 34
Examination of the Photoinitiator 36
Formation of the Semi-Interpenetrating Network 38
Conclusions 42
i Chapter Three: Design and Characterization of Photopolymerizable
Poly(ethylene glycol) Semi-Interpenetrating Networks for Chondrogenesis of Human Mesenchymal Stem Cells 44
Abstract 45
Introduction 46
Materials & Methods 49
Results 54
Discussion 67
Acknowledgements 72
Chapter Four: In Vitro Chondrogenesis of Human Mesenchymal Stem
Cells in Hydrogels is Affected by Temporal Exposure to Chondrogenic
Factors 73
Abstract 74
Introduction 75
Materials & Methods 76
Results 80
Discussion 89
Acknowledgements 94
Chapter Five: Conclusions & Future Work 95
Can Further Alterations Improve Matrix Elaboration? 104
What Type of Animal Model Should Be Explored? 110
Summary 111
Bibliography 112
ii LIST OF TABLES
Chapter Three
3-1. Experimentally determined swelling ratios, Q, for PEGDA scaffolds fabricated with and without PEG. 57
3-2. Molecular weight between crosslinks, MC, and mesh size, ξ, calculated for PEGDA hydrogels. 58
iii LIST OF FIGURES
Chapter One
1-1. Components of the extracellular matrix 7
1-2. Zonal architecture of adult articular cartilage 11
1-3. Endochondral ossification 15
Chapter Two
2-1. Toluidine blue stained histological sections and immunohistochemistry for collagen type II of human pellets after 1, 3, 6, 9, and 14 days of culture. 32
2-2. Immunohistochemistry for collagen types I, II, and X in day 1 and 14 pellets. 33
2-3. Chemical structure of PEGDA 35
2-4. Toluidine blue histological sections of 20% (w:v) PEGDA cell-seeded constructs. 37
2-5. Cellular viability of hMSCs subsequent to initiator and UV exposure, assessed through LDH activity. 39
2-6. Idealized and simplified depiction of an interpenetrating network. 40
2-7. Toluidine blue stained histological sections at 3 and 6 weeks. 43
Chapter Three
3-1. Extracellular matrix content of PEGDA (6 kDa) hydrogels polymerized with increasing concentrations of initiator. 56
3-2. Extracellular matrix content of constructs fabricated with the addition of PEG to PEGDA (6 kDa) at a PEGDA:PEG ratio of 2:1. 60
3-3. Extracellular matrix content of PEGDA (6 kDa) hydrogels
iv constructed at varying ratios of PEGDA:PEG. 62
3-4. Proteoglycan distribution in PEGDA (6 kDa) constructs fabricated with PEG at different ratios. 63
3-5. Extracellular matrix content of networks constructed with increased PEGDA molecular weight and PEG (88 kDa) at varying
PEGDA:PEG ratios. 64
3-6. Proteoglycan distribution in higher molecular weight PEGDA constructs fabricated with PEG (88 kDa). 66
Chapter Four
4-1. Comparison of extracellular matrix content at 6 weeks in constructs fabricated at different initial seeding densities. 81
4-2. Proteoglycan distribution of constructs made with different initial cell seeding density. 82
4-3. Extracellular matrix content of hydrogels fabricated with hMSCs pretreated in monolayer culture with defined chondrogenic medium. 84
4-4. Extracellular matrix content of hMSC-seeded scaffolds subjected to TGF-β1 withdrawal from defined chondrogenic medium. 86
4-5. Extracellular matrix content of hydrogels subjected to dexamethasone withdrawal from defined chondrogenic medium. 88
Chapter Five
5-1. Immunohistochemistry for collagen types II and X of hMSC-seeded
PEGDA:PEG networks cultured in defined chondrogenic medium that was subjected to removal of DEX or TGF-β1. 102
v 5-2. Immunohistochemistry for collagen types II and X of hMSC-seeded constructs cultured for 6 weeks in defined chondrogenic medium lacking or containing dexamethasone. 103
5-3. Extracellular matrix content of PEGDA scaffolds fabricated with or without tethered RGD. 106
5-4. Toluidine blue histological sections of hMSC-seeded constructs encapsulated in PEGDA polymer scaffolds with and without tethered RGD. 108
vi ACKNOWLEDGEMENTS
Any degree is an undertaking. While the efforts that merit the achievement must be those
of the student, they are not attained without support. The student, in fact, is supported by
an extensive social network – family, friends, peers, and faculty all aid in the endeavor.
This holds true in my experience and has been the case for everyone I have met in my
graduate work, without exception. To those who have invested countless hours teaching,
training, listening, encouraging, and loving me – I am eternally grateful. You know who
you are: my committee, my academic and research advisors, my current and previous lab- mates, my family and friends. Thank you so very much.
vii LIST OF ABBREVIATIONS
MSC: Mesenchymal Stem Cell
PEG: Poly(ethylene glycol)
PEGDA: Poly(ethylene glycol) Diacrylate
ECM: Extracellular Matrix
CS: Chondroitin Sulfate (chains)
KS: Keratan Sulfate (chains)
SLRPs: Small Leucine-Rich Proteoglycans
COMP: Cartilage Oligomeric Matrix Protein
OA: Osteoarthritis
DMEM: Dulbecco’s Modified Eagles Medium
FBS: Fetal Bovine Serum
PBS: Phosphate Buffered Saline
FGF: Fibroblast Growth Factor
TGF-β: Transforming Growth Factor-β
GC: Glucocorticosteroid
DEX: Dexamethasone
BMPs: Bone Morphogenic Proteins
LAP: Latency Associated Peptide
LTBP: Latent TGF-β Binding Proteins
LLC: Large Latency Complex
TβRI & II: TGF-β Receptors I & II
viii GLOSSARY
Hydrogels: insoluble polymer networks that are composed of water-soluble polymers, which are made insoluble by addition of a crosslinking component.
Semi-Interpenetrating Network: a hydrogel that employs two polymer components. Networks may be classified as covalent or non-covalent, depending on whether both polymers form covalent crosslinks. A non-covalent form is employed in this study, which uses the secondary (nonbinding) component to add stability, increase mass transport, and leave space for extracellular matrix elaboration.
Fibrillation: a process by which degradation of PGs and collagens occurs.
Osteochondritis Dessicans: a disorder leads to fracture through the subchondral bone and partial or complete separation of a fragment of the articular surface.
Osteoarthritis: the most common joint disease, caused by joint degeneration. This process involves the progressive loss of articular cartilage, followed by attempted repair of cartilage, remodeling and sclerosis of subchondral bone, and in many instances the formation of subchondral bone cysts and marginal osteophytes.
Crepitis: catching, grating, or grinding sensations during movement.
Endochondral Ossification: one of the two processes by which bone is formed. In this case, bone development occurs through a transitional cartilage element.
Chondrogenesis: the process by which cartilage is formed. Mesenchymal stem cells differentiate to become chondrocytes, which then synthesize all of the proteins and carbohydrates required to form the intricate extracellular matrix of which cartilage is comprised.
ix
Design and Characterization of Photopolymerizable Semi-Interpenetrating Networks for
In Vitro Chondrogenesis of Human Mesenchymal Stem Cells
ABSTRACT
by
AMANDA NICOLE BUXTON
Cartilage is formed through the process of chondrogenesis, in which mesenchymal stem
cells (MSCs) differentiate to become chondrocytes. Chondrocytes then synthesize all of
the components of the intricate extracellular matrix (ECM) of which cartilage is
comprised. Articular cartilage has a very poor reparative ability and extracellular matrix
degradation leads to the eventual destruction of the tissue. Tissue engineering presents a
possible avenue for its repair and regeneration. Because hydrogels are mimetic of the native, 3-dimensional, water-swollen environment in which chondrocytes exist, they have been the focus of much research in terms of cartilage tissue engineering, but success has been limited. This thesis describes the design and characterization of poly(ethylene diacrylate) (PEGDA) and poly(ethylene) (PEG)-based semi-interpenetrating networks that facilitated the formation of cartilage by human MSCs. Alterations in the molecular weight of the PEGDA crosslinker and the ratio of PEGDA:PEG, resulted in substantial changes in extracellular matrix elaboration. Proteoglycan content was significantly increased through lowering the PEGDA:PEG ratio at the lowest PEGDA molecular weight used (6 kDa). Furthermore, the profile of proteoglycan deposition within the scaffold was substantially altered with lower PEGDA:PEG ratio and with higher PEGDA
1 molecular weight. With higher PEGDA molecular weight, toluidine blue staining revealed intercellular/interterritorial deposition of proteoglycan, as compared with pericellular deposition exhibited by constructs fabricated at PEGDA:PEG of 2:1 with
PEGDA (6 kDa). Collagen content, however, was lower in constructs fabricated with higher PEGDA molecular weight at a PEGDA:PEG ratio of 1:2. The effects of culture conditions on subsequent matrix development in the optimized polymer hydrogel were also examined. Initial experiments defined the optimal cell seeding density that produced maximum matrix deposition throughout the construct at 25 x106 cells/ml. Pretreatment of monolayer cells with defined chondrogenic medium retarded matrix elaboration, while withdrawal of either TGF-β1, or dexamethasone (DEX) lowered collagen content per cell at all timepoints. These findings are informative for in vitro tissue engineering of cartilage, but may also aid in vivo implantation strategies for hydrogels that contain
MSCs rather than preformed cartilage.
2 CHAPTER ONE: BACKGROUND & SIGNIFICANCE
Synovial joints are formed from articular cartilage, subchondral and metaphyseal bone, synovium, ligaments, and joint capsules [1]. Articular cartilage is a highly specialized connective tissue covering the ends of long bones that serves as a low-friction surface which effectively distributes loads across synovial joints [1, 2]. Unique in that it contains no blood vessels, nerve fibers, or lymphatics, articular cartilage is composed of an extensive, hydrated extracellular matrix that is formed through the process of chondrogenesis—mesenchymal stem cells (MSCs) differentiate to become chondrocytes, which then synthesize all of the carbohydrates and proteins required to form the intricate extracellular matrix (ECM) of which cartilage is comprised. Chondrocytes account for less than 5% of the total volume of adult tissue. Predominantly lacking cell-cell contact, chondrocytes interact with the ECM for intercellular communication, delivery of nutrients, and removal of waste products. Even though tissue-embedded chondrocytes exhibit high metabolic activity, they do not divide after adolescence under normal conditions [2].
To better understand the challenges associated with tissue engineering cartilage constructs, we will first discuss the structure, function, and formation of the native tissue.
This will be followed by a treatise on pathology associated with disease and injury, and methods of repair. Finally, we will explore tissue engineering design criteria, with particular emphasis on a cartilaginous system.
3 Structure and Function of Articular Cartilage
The primary component of the ECM is water, which is approximately 70% of the weight
of adult cartilage [2]. The macromolecular components of the tissue that constitute the
collagens, proteoglycans, and noncollagenous proteins/glycoproteins bind water tightly
and prevent its escape (Figure 1-1). Chondrocytes synthesize and assemble all of the
components of the macromolecular framework, although it is not known whether these constituents are synthesized simultaneously or in regulated phases dependent on tissue
requirements. It is known, however, that macromolecules and their concentrations change
during different phases of development and aging in order to meet altered functional
needs.
Collagen represents more that 50% of the organic dry weight of cartilage, equivalent to
10 – 20% of the wet weight. In adult tissue approximately 90% of collagen is type II,
which belongs to the fibril-forming class of collagens (class 1). These fibrils form a
three-dimensional crosslinked network of insoluble collagen fibers, in which soluble
proteoglycans and glycoproteins are enmeshed, integrated, and/or chemically attached
[2]. The network of type II collagen fibrils provides for the tissue’s tensile strength and is
essential for maintaining cartilage volume and shape. Tensile strength increases with age,
due to the increase in covalent, intermolecular crosslinks between type II collagen
molecules which occurs with maturation.
Up to 10% of the total collagen in adult articular cartilage is made up collagen types VI,
IX, X, and XI, all of which contribute to the function of the tissue. Collagen type XI also
4 belongs to class 1 (fibril-forming) collagen, and research suggests that this collagen helps
organize and determine the final collagen type II fibril diameter. Collagen type IX
belongs to the class of short-helix molecules (class 3). One of the three chains comprising
collagen IX is glycosylated with a chondroitin sulfate chain, allowing collagen type IX to
also be classified as a proteoglycan. Type IX collagen is covalently crosslinked to type II
collagen, suggesting that it serves as an interfibrillar connector or bridging molecule.
Collagen type VI, also a member of the class of short-helix molecules, forms distinct
microfibrils. Type VI collagen is concentrated in the matrix surrounding chondrocytes
and in chondrons. Collagen type X also belongs to class 3 collagens. Synthesis of type X
collagen is restricted to the hypertrophic cells within the zone of calcified cartilage,
where articular cartilage attaches to underlying bone.
Proteoglycans, highly hydrophilic macromolecules comprised of heterogeneous glycoconjugates, are embedded in compressed (underhydrated) form within the collagen
fibrillar network (Figure 1-1). Articular cartilage’s ability to undergo reversible deformation is conferred by the proteoglycans; the most predominant member is aggrecan, whose concentration is 30 – 50 mg/ml within the tissue. Aggrecan is composed
of a core protein (Mr = 215 kDa), to which glycosaminoglycan side chains and N-linked
and O-linked oligosaccharides are covalently attached. The glycosaminoglycan chains,
consisting of keratan sulfate and chondroitin sulfate, comprise 90% of aggrecan’s mass.
The core protein contains three globular regions (G1, G2, and G3) and two extended,
interglobular domains. N-linked oligosaccharides are located within G1, the interglobular
domain between G1 and G2, and in the G3 regions. The interglobular domain between G2
5 and G3 contains O-linked oligosaccharides, more than 100 chondroitin sulfate chains, and
between 20 – 50 keratan sulfate chains. A moderately-sized aggrecan molecule is
approximately 80 - 100 nm in diameter, with a length that decreases with age. This
decrease is due to changes in the length and presence of chondroitin sulfate chains [3, 4].
With age, aggrecan becomes shorter and more enriched in keratan sulfate.
6
Figure 1-1. Components of the Extracellular Matrix. Chondrocytes are responsible for synthesis and organization of the extracellular matrix that comprises articular cartilage. Chondrocytes receive signals from growth factors, integrin binding, and from molecules within the network. The collagen-fibrillar network bounds the swelling pressure created by the proteoglycans, and the water that they attract, that reside within.
7
The G1 amino-terminal region of aggrecan also interacts noncovalently with hyaluronic
acid, which is comprised of an extracellular unsulfated high molecular weight
glycosaminoglycan (hyalorunan) consisting of many repeat units of N-acetyl
glucosamino-glucuronic acid. This interaction is stabilized by link protein, a small
glycoprotein that has homology to the G1 and G2 domains of aggrecan. Matrix assembly
and associations occur extracellularly, and as many as 200 aggrecan molecules
(monomers) can bind to a single hyaluronic acid molecule to form an aggregate [2].
Other proteoglycans, called small leucine-rich repeat proteoglycans (SLRPs), are present
in articular cartilage (Figure 1-1). The major SLRPs in cartilage are decorin, biglycan,
fibromodulin, and lumican [5]. Biglycan and decorin contain dermatan sulfate
glycosaminoglycans, while fibromodulin and lumican are rich in keratan sulfate
glycosaminoglycans. Biglycan and decorin are both found in low concentrations in the
tissue; however, each shows an age-related increase in concentration particularly in the
superficial layer of the tissue [2]. All SLRPs are capable of binding other ECM
components, including collagen types II and VI, fibronectin and elastin, and growth
factors such as TGF-β and EGF [5]. The function of SLRPs depends on their core protein
and their glycosaminoglycan chains. Their core proteins allow SLRPs to interact with
fibrillar collagen type II, regulating fibril diameter during its formation and possibly
fibril-fibril interaction in the ECM. Decorin, fibromodulin, and lumican all interact
directly with collagen fibrils, however, in vitro studies indicate that biglycan interaction depends on environmental conditions [5].
8 Articular cartilage contains other extracellular matrix proteins that are neither collagens
nor proteoglycans, including but not limited to: chondrocalcin, fibronectin, and cartilage
oligomeric matrix protein (COMP). Chondrocalcin is a 35 kDa protein composed of the
carboxy-terminal propeptide of type II collagen, which is released at sites of collagen
synthesis. Because it diffuses out of the tissue, chondrocalcin can be used as a marker of
collagen anabolism. Furthermore, chondrocalcin binds to apatite crystal with high
affinity, and is thought to be involved in calcification [2]. Fibronectin is synthesized
during early stages in cartilage development, and then downregulated. However,
digestion of the mature adult tissue reveals it presence throughout the ECM. The 36 kDa
cartilage matrix protein expresses a high binding affinity for chondrocytes, and may
mediate cellular interaction with the ECM during remodeling. COMP, a glycosylated
protein that contains some CS chains, is only detectable with high abundance in articular
cartilage. COMP seems to be localized in the chondrocyte territorial matrix, and is more enriched in the superficial layer of cartilage. It is currently thought to function in assembly and repair of the tissue.
As highly negatively-charged molecules, proteoglycans attract water and provide a
swelling pressure in the tissue. Upon loading, water is extruded from cartilage, and the
proteoglycans become more compressed. This compression increases the negative charge
density, which increases the resistance of water flow until equilibrium between loading
forces and swelling pressure is reached. Water returns to the tissue when the load is
removed, until the swelling pressure of the proteoglycans is again balanced by the
resistance of the collagen-fibrillar network. Because of their fixed charge density,
9 proteoglycans also act as ion exchangers. In this manner they may enhance the diffusion
of small molecules, influencing (even retarding) the diffusion of similar negatively
charged macromolecules while concentrating cationic molecules within the tissue. In general, mechanical loading of articular cartilage induces matrix and chondrocyte deformation, fluid flow, hydrostatic pressure gradients, streaming potentials, and other physiochemical alterations that regulate chondrocyte metabolism [2].
The mechanical properties of articular cartilage depend on the two main components of
the ECM: the solid macromolecular framework consisting of collagens and aggregating
proteoglycans, and water located within this framework [6]. Their structure-function
relationship is intimately wed; neither component acts alone. For simplicity, however,
one may assert that the interaction between water and proteoglycans confers the tissue’s
stiffness to compression, resilience, and durability – allowing for cartilage to undergo
reversible deformation. Concurrently, the collagen-fibrillar network provides for the
tissue’s tensile strength. Articular cartilage tolerates loading resulting from normal daily
activities so well that in many individuals it shows little or no evidence of degeneration
after 8 decades or more of daily use [6].
There are depth-related variations in the ECM and the chondrocytes that reside therein.
Chondrocytes in adult cartilage may be subdivided into three unmineralized zones, which are rather well-defined by the tidemark from the zone of calcified cartilage (Figure 1-2).
Zone I, called the superficial or tangential zone, has the highest cell density. Collagen fiber bundles in this zone are tangential to cells, which are relatively small, flat, and
10
Figure 1-2. Zonal architecture of adult articular cartilage. A. Zonal subpopulations in mature tissue. Chondrocytes in mature cartilage may be subdivided into three distinct unmineralized zones: the superficial or tangential (zone I), the transitional or intermediate (zone II), and the radial (zone III). Zones I-III are separated from the zone of calcified cartilage (zone IV) by the tidemark. Light photomicrographs of (B) a superficial zone chondrocyte, (C) a zone IIIa chondrocyte. ITM, interterritorial matrix; TM, territorial matrix; IF, intermediate filaments; G, glycogen, and (D) a zone IIIb chondrocyte near the tidemark (T). Adapted from Kuettner et al, 1986.
11 oriented with the long axis parallel to the surface of the tissue. Zone II, the transitional or intermediate zone, possesses larger and more round cells that are randomly dispersed in a matrix containing fibers extending in oblique directions. Cells increase further in size in zone III, and are arranged in columns with axes perpendicular to the surface. Cellular grouping in zone III reflects radial order in collagen fibers. The ECM becomes heavily mineralized in zone IV. Single cells or cell columns are buried within a layer of unmineralized matrix [7]. Zones I-III may only be viewed in adult animals – growing animals (and adults) lack a subdivision in zone II and III since the modeling of the epiphysis and shaping of the articular surface is incomplete. Cellular metabolic activity varies with zonal differences. Chondrocytes from the deep radial zone produce a well- organized matrix rich in PGs including KS-rich molecules, while cells from the superficial zone synthesize much less PG and only trace amounts of KS [2].
Chondrocytes in all zones exist predominantly without direct cell-cell contact within the
tissue. Each cell can be considered a functional unit of cartilage, responsible for ECM
maintenance in its immediate vicinity. The ECM surrounding a chondrocyte is compartmentalized – to a large extent based on the organization of the collagen-fibrillar
network [7]. The immediate extracellular vicinity surrounding the chondrocyte
membrane, called the pericellular or lacunar matrix, is rich in proteoglycans and
hyaluronic acid. Collagen is relatively absent from this domain. Adjacent to the
pericellular region is the territorial or capsular matrix, which is composed of a basket-like
network of crosslinked fibrillar collagen called a chondron. The chondron encapsulates
the individual chondrocyte and provides mechanical resistance for the cell; the majority
12 of collagen type VI is located in this region. Chondrocytes establish contact with the capsular matrix through numerous cytoplasmic processes rich in microfilaments (cilia), integrin-matrix molecule interactions, and via other extracellular matrix receptors. The largest and final compartment of the ECM is the interterritorial matrix. The farthest removed from the chondrocyte, the interterritorial matrix contains most of the collagen fibrils and proteoglycans. Chondrocytes in adult tissue exert active metabolic control over their territorial matrix; however, the ECM in the interterritorial matrix is less actively regulated, and seems to be metabolically more ‘inert’ [2]. Under physiologic conditions, chondrocytes regulate and balance synthesis of matrix molecules with degradation and loss from the ECM. This process helps maintain constant concentrations of ECM components, although they may be regulated on widely different timescales. For example the half-lives of hyaluronan and PGs are approximately 25 days, while the half-life of collagen type II is estimated to be many years [2].
Formation of Cartilage
In vivo formation of skeletal tissue occurs either through a transitional cartilage template,
or via direct differentiation into bone [8]. The former process, endochondral ossification,
is more prevalent in vertebrate development; however, the latter process
(intramembranous bone development) results in the formation of the flat cranial bones
and the clavicle [9, 10].
The process of condensation initiates and is required for successful chondrogenesis [11].
Undifferentiated mesenchymal cells migrate into the limb field, an area destined to
13 become either cartilage or bone in the developing embryo, and condense—that is, they increase cells/unit area of volume without proliferating (Figure 1-3). This condensation
step facilitates differentiation into chondrocytes. Undifferentiated mesenchymal cells
produce an extracellular matrix rich in collagen type I, hyaluronan, tenascin, and
fibronectin. Differentiation hallmarks the production of cartilage-specific collagen types
II, VI, IX, and XI, the proteoglycan aggrecan, and link protein (among other molecules)
[11, 12]. Chondrocytes become encased in this ECM, forming what is known as a
cartilage anlage [13]. The cartilage anlage is remodeled into bone over the course of 16-
25 years, the rate and duration of which is the largest determinant of the height of an
individual [10]. The cartilage anlage begins as hyaline cartilage. As the cartilage structure
grows, through chondrocyte proliferation and extracellular matrix production, two events
occur simultaneously: resident chondrocytes at the surface form a protective membrane,
called the perichondrium, while chondrocytes residing in the central region undergo
hypertrophy, apoptosis, and calcification [10, 14]. A collar of bone forms around this
central region, the function of which is to provide support for the calcifying cartilage
center and allow for blood vessel invasion. Subsequent vascularization of the shaft of
calcified cartilage allows for osteoblast recruitment. Osteoblasts align at the extremes of
the shaft (the diaphysis) and secrete new bone. The calcified cartilage in the central
region is degraded, adsorbed into the blood, and replaced with new bone. The two sites
where osteoblasts have aligned and begun secretion of new bone are called primary
centers of ossification. Bone growth is directed outward, towards the end of the
developing bone (epiphysis). Some of the degraded calcified cartilage is replaced by the
medullary, or marrow cavity, where bone marrow will reside.
14
Figure 1-3. Enchochondral ossification. (1) MSCs condense and undergo differentiation to chondrocytes. Cells at the border of the condensation form a membrane called the perichondrium. (2-3) Chondrocyte proliferation and matrix production enlarges the cartilaginous tissue. Chondrocytes in the central region cease proliferating, undergo hypertrophy, initiate mineralization of their surrounding matrix, promote vascularization through synthesis and secretion of growth and other factors, and direct adjacent perichondrial cells to become osteoblasts – which form a bone collar under the perichondrial membrane. The new composite structure is called the periosteum. (4-5) Blood vessel invasion from the periosteum brings bone-forming cells into the tissue. Osteoblasts align at the extremes of the cartilage shaft (diaphysis) and remodel calcified cartilage into bone – these two sites are known as the primary centers of ossification. Bone development and growth is directed outward, towards the ends of the bone (blue regions). (6-7) Vascularization of the epiphyses allows for the formation of secondary centers of ossification. (8) Cartilage located between the epiphyseal and diaphyseal centers of bone growth becomes known as the epiphyseal (growth) plate. (9-10) Longitudinal bone growth occurs through chondrocyte proliferation and matrix production at the growth plate, followed by a series of ossification steps. Growth ceases when the cartilage in the epiphyseal plate thins, allowing fusion of the two centers of ossification. Adapted from [15]
15 In the first few years after birth, blood vessel invasion of the epiphyses (ends) of bone allows for the formation of secondary centers of ossification [9, 10]. The cartilage located between the secondary (epiphyseal) and primary (diaphyseal) centers of ossification becomes the epiphyseal (growth) plate. Growth of this cartilage is directly responsible for lengthening of bone. Bone growth ceases when the cartilage in this plate thins, and is progressively replaced by bone – fusing the epiphyseal and diaphyseal centers of ossification.
The cartilaginous growth plate is organized into three functionally and structurally
distinct layers: the resting, the proliferative, and the hypertrophic zones. The resting zone
is comprised of irregularly arranged chondrocytes that rarely divide. Chondrocytes in the
proliferative zone appear flattened and are arranged in columns parallel to the long axis
of the bone. The proliferating chondrocytes farthest from the epiphysis stop replicating,
and enlarge to become hypertrophic chondrocytes. Apoptosis occurs next in these
terminally differentiated cells, which form the hypertrophic zone in a layer adjacent to the
metaphysis. Angiogenesis follows apoptosis, which allows the tissue to be invaded by
bone precursor cells that subsequently remodel cartilage into bone. During bone growth,
chondrogenesis and ossification are tightly coupled so that the width of the growth plate
remains relatively constant while new bone is formed at the junction of the growth plate
and the metaphyseal bone. Once bone reaches its ultimate length, the growth plate is
sealed – leaving permanent cartilage on the surfaces of articulating joints. This articular
cartilage is the only remnant of the cartilage anlage that initiated limb development.
16 Articular Cartilage Pathology & Repair
Damage to articular cartilage can be divided into two general categories: focal defects in
otherwise normal joints and degenerative defects [16]. Localized disruptions in articular
cartilage can extend to the underlying bone (osteochondral defects) or be limited to
articular cartilage. Osteochondral defects occur primarily in adolescents and young
adults. These may arise from a disorder called osteochondritis dessicans, which is the
result of joint loading that compromises the subchondral bone blood supply [16]. This
disorder leads to fracture through the subchondral bone and partial or complete separation
of a fragment of the articular surface. Impact or torsional loading of a joint may also
cause acute osteochondral fractures in these populations. Older individuals, young and
middle-aged adults, typically have a partial or complete separation of an articular
cartilage fragment (chondral defect) that leaves the subchondral bone intact after a similar joint loading injury. Differences in injury, which manifest with age, may be due to developmental changes in the articular surface. In particular, maturation and ossification of the subchondral bone plate, and the formation of the zone of calcified cartilage, reduce the chance that a load applied to the articular cartilage will cause an osteochondral defect
[16].
Focal defects possess well-defined boundaries; the defect is surrounded by clear regions
of structurally normal articular cartilage. Degenerative defects, however, lack this
definition. These lesions first appear as poorly demarcated regions of softening and
superficial fraying of the articular surface [16]. Although degenerative defects may remain unchanged for decades, they may also slowly extend to cover most if not all of an
17 articular surface. Furthermore, they may extend progressively into deeper layers of the
tissue. In most cases, these degenerations rarely develop before middle age. Degenerative defects occur in some autoimmune and metabolic diseases, and with osteoarthritis (OA).
OA, the most common joint disease, is caused by joint degeneration. This process involves the progressive loss of articular cartilage, followed by attempted repair of
cartilage, remodeling and sclerosis of subchondral bone, and in many instances the
formation of subchondral bone cysts and marginal osteophytes [6]. OA develops most
commonly in the absence of a known cause of joint degeneration; thus, the condition is
referred to as idiopathic or primary OA. Less frequently it develops as a results of injury
(posttraumatic OA), or a variety of hereditary, inflammatory, developmental, metabolic,
and neurologic disorders, all of which are referred to as secondary OA. Regardless of the
classification, the processes of joint degeneration in primary and secondary OA are
indistinguishable.
The structure of articular cartilage – lacking nerves, blood vessels, and lymphatics –
makes it difficult to track initial cellular changes in OA. The first signs therefore typically
present as a decrease in the freedom of movement in an active joint [6]. Diagnosis also
involves joint pain and crepitus (catching, grating, or grinding sensations during
movement). Radiographs confirm the diagnosis, with changes manifesting in three main
ways: narrowing of the cartilage space, increased density of subchondral bone, and the
presence of osteophytes. It should be noted, however, that radiographic changes may vary
considerably among patients; in some joints only one or two markers may be visible. As
OA progresses joint subluxation, deformity, and malalignment develop.
18 Tissue-associated changes with disease and injury have been examined in various animal models. Once the macromolecular organization of cartilage has been disturbed, by mechanical stress for example, fibrillation may result. During fibrillation, degradation of proteoglycans and collagens occurs. Once the collagen framework has been modified, proteoglycans embedded within the network may be able to attract more water, swell, and subsequently leak out of the tissue. Loss of proteoglycan leads to collapse of the collagen network, and aggregates of thick fibrils form. Reconstruction of functional tissue would require removal of the collapsed collagen and synthesis of the specific ECM constituents by the indigenous chondrocytes. Under normal circumstances, however, cellular attempts to repair the matrix result in production of fibrous cartilage with unsuitable biomechanical properties that lead to early failure of the tissue under normal physiologic load [17].
Given its avascular nature, articular cartilage itself has limited capacity for spontaneous self-regeneration; not surprisingly, most clinical treatments of injured articular cartilage target alleviation of pain and restoration of joint function rather than the regeneration of hyaline cartilage. Current therapies for damaged cartilage fall within the two categories of repair and regeneration. Repair refers to restoration of a damaged articular surface with new tissue that resembles but does not duplicate the structure, composition, and function of articular cartilage. Regeneration implies the formation of new tissue which is indistinguishable from that which it is replacing [6]. Several surgical repair techniques have been investigated including subchondral drilling, arthroscopic invasion, and microfracture technique. Alleviation of pain and improvement of joint movement,
19 however, are widely varied among patients [6]. Furthermore, none of these techniques
have successfully regenerated long-lasting hyaline cartilage tissue [18]. Allografts and
soft tissue replacements are also used as therapies for chondral defects. The main issue
with these treatments lies in acquisition of appropriate tissue – donor tissue availability.
Total joint arthroplasty is also used to repair damaged articular cartilage. Artificial joints
typically last 15 – 20 years, and come with encouraged modifications to the recipient’s activity level. However, surgery is invasive, and carries with it increased susceptibility to infection and future problems with implant fixation [19]. Furthermore, secondary surgeries (multiple replacements) on the same joint are not widely successful.
Tissue Engineering Design Criteria
Tissue engineering, combining of synthetic or natural biomaterials with cells to promote
tissue formation, has emerged as a field to develop functional substitutes for damaged
tissue—including cartilage. The ultimate goal of any tissue-engineered construct is the
restoration of the morphology and function of the tissue. Regeneration, however, is a
slow process that appears to recapitulate some of the key steps that occur in embryonic
development [20].
Tissue engineering strategies involve the singular or tandem use of cells, biomatrices or
scaffolds as delivery vehicles, and signaling molecules that provide biological cues for
cellular differentiation and/or functional modulation. The final construct formulation
requires careful consideration when selecting the cells to be utilized, the scaffold into
which they will be seeded (if one is to be used), and whether the environment wherein the
20 construct will be placed requires the use of bioactive factors, or the use of differential
adhesion to facilitate differentiation, matrix production, and/or cellular retention [20].
Cell Source
The source, number, and density of cells to be employed, as well as their age, phenotype, and developmental potential must be considered. Much of the previous research has studied the response of chondrocytes to various natural and synthetic scaffolds. Although these studies report that chondrocytes are viable and produce extracellular matrix, elaboration of matrix is limited at best. Problems with matrix production may be due to the decreased metabolic activity of mature chondrocytes, or difficulties in chondrocyte isolation and expansion. Acquisition of chondrocytes for culture and experimentation is time-consuming and difficult. Appropriate donor tissue must be found, a difficult task to achieve since the entire tissue is utilized in load-bearing joints. After the tissue is dissected, the extracellular matrix must be digested to release the encapsulated chondrocytes. Furthermore, once they are released from their 3-dimensional environment and placed in 2-dimensional (monolayer) culture, chondrocytes de-differentiate – they cease to produce mRNA and protein for the ECM molecules native and specific to cartilage.
Successful development of a functional tissue-engineered cartilage construct might
involve the recapitulation of developmental events, rather than the transplantation of
mature chondrocytes. The development of all musculoskeletal elements begins with
MSCs, and cells possessing this differentiation potential are found in adult tissues [21].
21 They are advantageous as a source of chondrogenic cells because they are more
accessible than native chondrocytes, yield higher numbers during in vitro expansion, and
are capable of differentiating along multiple lineages in vitro when cultured in defined
medium: adipocytes, chondrocytes, osteoblasts, muscle, and neuronal cell differentiation
have been reported [22-28]. A tissue engineering construct that promotes chondrogenesis
might stimulate not only a significant production of extracellular matrix, but also
facilitate the development of material properties that more closely mimic those of natural
articular cartilage. If such a construct were successful as an implant into a pathological
site, it could postpone or eliminate the need for joint replacement as a treatment for
degenerative joint diseases.
Scaffold Considerations
Both synthetic and natural scaffolds have been used in a wide variety of tissue
engineering approaches. Although the ideal physical, chemical, and mechanical
characteristics have not yet been elucidated for a cartilaginous system, several factors
should be considered when choosing an appropriate scaffold. These factors include, but
are not limited to the following: porosity, biodegradability, capability for adhesion and
integration into surrounding tissue, remodeling capacity, and architecture of the entity
and the surface. It may also be advantageous to employ a scaffold that may contain or
release bioactive factors – discrete, tailored components included to induce cellular
differentiation and/or proliferation. Finally, one must decide where to fabricate the
construct. Should it be designed and tested in vitro? Or should the device be implanted, optimized, and constructed in vivo, where it will ultimately function? There are no clear
22 answers to these questions – each case must be considered individually. There are no currently functional cartilage tissue-engineered constructs in routine surgical use.
Many scaffolds have been explored for their efficacy in cartilage tissue engineering, from porous solids and meshes to injectable gel networks with naturally-occurring and synthetic polymers. Poly(lactic acid) (PLA), poly(glycolic acid) (PGA), and their copolymers poly(lactic-co-glycolic acid) (PLGA) are some of the most commonly employed synthetic polymer scaffolds employed. Collagen matrices, gelatin sponges, and glycosaminoglycan-based materials are the more prevalent naturally-occurring porous systems utilized. Solid scaffolds require ex vivo cell seeding, through tissue culture or bioreactor culture, to enable cells to distribute evenly throughout the construct and to allow appropriate perfusion of media [29]. Porous structures seeded and cultured statically typically possess void spaces and poor cellular retention [30]. Several groups have used hyaluronic acid derivatives, bioreactors, and mechanical loading (singularly or in tandem) to increase matrix development and cellular retention [30-34]. Hydrogels have also been investigated as a means of augmenting cellular retention, with alginate the most commonly employed. Hydrogels are insoluble polymer networks that are composed of water-soluble (hydrophilic) polymers. These hydrophilic polymers are made insoluble by addition of a crosslinking component. Crosslinking may be facilitated through physical or covalent linkages—the former occurring as a response to altering temperature or ionic environment, and the latter through a chemical crosslinking mechanism. In the case of alginate, a naturally occurring polysaccharide, phase transition is triggered through the addition of calcium ions.
23
Hydrogels are attractive materials for cartilage tissue engineering because they provide a swollen, three-dimensional environment high in water content that resembles the environment of native tissue and allows chondrocytes to preserve their rounded cell morphology. Poly(ethylene glycol) (PEG) hydrogels have been extensively explored in tissue engineering applications due to their hydrophilicity, biocompatibility, and intrinsic resistance to protein adsorption and cell adhesion. Chondrocytes encapsulated in hydrogels proliferate and increase production of ECM macromolecules proteoglycan and collagen type II [19, 35-45]. Previous work has shown that aqueous solutions of acrylated
PEG derivatives can be safely photopolymerized in direct contact with cells and tissues
[19, 42-44, 46-52]. Photopolymerization is advantageous as a method for triggering phase change because it allows spatial and temporal control over the process of polymerization,
and can be carried out ex vivo or in situ.
Crosslink density closely controls both the physical and mechanical properties of the
resulting hydrogel [37-41, 51, 53-56]. Crosslink density and compressive modulus are
directly related. Increasing the crosslink density causes the equilibrium water content
within the hydrogel to decrease, and increases the compressive modulus. The crosslink
density may be tailored in several ways, including the following alterations: the
concentration and molecular weight of the crosslinker, the concentration of the initiator,
the duration of polymerization, the addition of other crosslinkers (that may or may not be
degradable), and the addition of non-binding elements. A semi-interpenetrating network
is a hydrogel that employs the latter, using two components to form a hydrogel. The first
24 element is the chemical crosslinker, and the second is a nonbinding component that may add stability, or it may serve as a spacer – providing space for nutrient delivery and waste removal [19, 57]. All of these mechanisms, in conjunction with the nature of the
hydrophilic polymer used as the core of the network, control the swelling ratio (i.e. water
content) of the resulting hydrogel.
Network crosslink density and compressive modulus, in particular, have been shown to
influence matrix production by chondrocytes encapsulated within PEG hydrogels.
Although a lower crosslink density alone has not been shown to increase the amount of
glycosaminoglycan content, localization of proteoglycan was more uniform in comparison to hydrogels constructed with a higher crosslink density [37-41, 58, 59].
Hydrogel systems that allow for gel degradation on a timescale similar to extracellular matrix production have also been constructed. When chondrocytes were seeded into this type of hydrogel with a higher initial crosslink density, one that would lower as matrix production progressed, researchers saw a significant increase in the production of ECM molecules. Because hydrogel properties influence extracellular matrix production, they must be optimized.
Addition of Bioactive Factors
Investigation of a scaffold for tissue engineering cartilage will mostly likely involve the
addition of bioactive factors to promote cellular retention, growth, or differentiation.
Members of the Transforming Growth Factor-β (TGF-β) superfamily are secreted
signaling molecules that regulate many aspects of growth and differentiation in a cell-
25 type-specific manner. The TGF-β superfamily includes 3 isoforms of TGF-β: 1) activins and inhibins, 2) growth and differentiation factors (GDF), and 3) the bone morphogenic proteins (BMPs) [60]. Based on sequence similarity and the signaling pathways utilized, the superfamily can be divided into 2 sub-families: 1) the TGF-β/activin/nodal subfamily, and 2) the BMP/GDF subfamily. TGF-β orthologs exist in all vertebrates and in genetically tractable invertebrates, and many of the functions of the superfamily are conserved.
TGF-β peptides, first synthesized as large precursor proteins containing a large
prodomain followed by a signal sequence, are secreted as latent (inactive) proteins, even
though the propeptide is cleaved in the cell. Their inactive state is due to a non-covalent
association with the N-terminal prodomain of the TGF-β precursor protein, also called
latency associated peptide (LAP). A family of latent TGF-β binding proteins (LTBP) also
associates with the inactive TGF-β peptide, forming what is called the large latency
complex (LLC). Both of these associations are thought to regulate TGF-β activity in vivo.
The mature, active TGF-β is a 25-kDa molecular weight dimer [60]. TGF-β1 is the most
abundant isoform largely found in platelets (20 mg/kg) and bone (200 μg/kg) [61].
Chondrogenesis and skeletal development, in particular, are closely regulated by TGFβ
[62, 63]. The TGF-β family is comprised of 3 closely related isoforms: TGF-β1, TGF-β2,
and TGF-β3. Despite sequence similarity, gene knockout studies in mice reveal striking
differences in bone development in vivo, providing evidence for isoform non-redundancy
[61]. Most chondrogenic systems involve defined medium containing TGF-β, and
26 systems involving human cells further require the glucocorticosteroid dexamethasone
(DEX). Previous work in our laboratory, and by others, has demonstrated in vitro
chondrogenesis through TGFβ induction of human MSCs, rabbit MSCs, goat MSCs,
periosteal cells, and adipose-derived stem cells in 3-dimensional culture [22, 25, 64-75].
Some tissue-engineered scaffolds also require the addition of bioactive factors to provide
a cellular ‘pavement’ which enhances adhesion or retention. The cell-binding sequences
from fibronectin and laminin have been employed to this effect in many 2-dimensional
systems. We will focus on the argument for fibronectin. Fibronectin’s usefulness in
chondrogenic systems is uncertain, although the rationale for its employment is clear.
Fibronectin has been implicated in cell-matrix interactions that play a role in mesenchymal cell condensation [76-82]. Fibronectin expression is increased in areas of cellular condensation and decreases as differentiation proceeds. Although the intact molecule’s role in cartilage development and maintenance is incompletely understood, previous in vitro experiments in a 3-dimensional aggregate system in our laboratory have illustrated the importance of fibronectin matrix assembly in forming cellular condensations, and subsequent chondrogenic differentiation of hMSCs (Pennington et al, reviewers abstract).
Cellular interactions with fibronectin are primarily mediated through the RGD tripeptide sequence contained within the tenth type III module, situated in the middle of the
molecule [77, 79, 83]. The RGD tripeptide sequence is found in many ECM proteins and
is the binding motif for cell-surface integrins, particularly the α5β1 integrin in
27 chondrocytes. Covalent binding of the RGD sequence to a substratum has been shown to
increase adhesion and/or spreading of many cell types, including fibroblasts, endothelial
cells, and smooth muscle cells [18, 46, 48, 84-95]. However, some studies have shown
that increased cell adhesion can result in decreased extracellular matrix protein
production by the cells [96]. However, it should be noted that a substantial portion of the
RGD-relevant literature has examined utility of the sequence in a 2-dimensional
culture/monolayer system.
In the next chapters I will discuss the design and use of a photopolymerizable hydrogel
for cartilage tissue engineering. Chapter two describes the initial hydrogel that was
explored, and the challenges that necessitated a re-design. Chapter three characterizes the
new scaffold – a semi-interpenetrating network that enables in vitro chondrogenic differentiation of human MSCs. Also discussed are the changes in extracellular matrix development that occur when multiple parameters of the network are altered: the concentration of photo-initiator utilized, the duration of UV exposure, the molecular weights of the crosslinking and physical elements and their ratio. Chapter four examines
the consequences of temporal exposure of cell-seeded constructs to growth factors which
are widely employed to induce in vitro chondrogenesis. This study is particularly relevant
since the in vivo climate – lacking nerves and blood vessels – is very different from that seen in vitro culture. Availability of growth and other factors may be limited in this environment and, thus, optimization of the concentrations and availability of growth factors may be critical to ensure the success of an implanted scaffold in which the cells are not pre-differentiated. Such work is also critical to the successful creation of
28 cartilaginous constructs created in vitro prior to implantation into the in vivo site. Chapter five provides conclusions for the body of work, and suggests avenues for future experimentation.
29 CHAPTER TWO:
IN VITRO CHONDROGENESIS IN A POLY(ETHYLENE GLYCOL)
DIACRYLATE (PEGDA) SCAFFOLD – PRELIMINARY EXPERIMENTS
Previous work in our laboratory successfully reported in vitro chondrogenesis of human
MSCs in aggregate culture [22, 25]. However, the pellets formed through this process
were small. Attempts to form larger aggregates – through manipulation of the volume and
number of cells – resulted in the formation of multiple smaller pellets. To increase the
size of the cartilage formed to that applicable in a clinically-relevant sized defect (with
diameter > 1 cm), we decided to employ a hydrogel scaffold. The experiments outlined
below describe the original pellet culture scaffold-free system, and subsequent attempts
to formalize a scaffold which would facilitate in vitro chondrogenesis of human MSCs
(hMSCs).
In Vitro Chondrogenesis in Pellet Culture
Methods
Human bone marrow was obtained from the iliac crests of consenting donors. Marrow aspirates were fractionated on a Percoll density gradient and plated in Dulbecco’s modified Eagle’s medium (DMEM) with 10% fetal bovine serum (FBS). Cells were cultured at 37° C, 5% CO2 with medium changes every three days. Once confluent, cells
were trypsinized, neutralized in DMEM + FBS, and re-suspended in individual 15 ml
conical tubes in defined medium at a concentration of 400,000 cells/ml. Conical tubes
were subsequently centrifuged for 5 minutes at 2000 rpm, and maintained in culture at
37°C, 5% CO2 for 14-21 days in a defined chondrogenic medium previously shown to
30 induce chondrogenic differentiation of these cells in aggregate culture [22, 25]. The
defined medium consisted of: high-glucose DMEM with ITS+ Premix (Collaborative
Biomedical Products: insulin (6.25 μg/ml), transferrin (6.25 μg/ml), selenous acid (6.25
μg/ml), and linoleic acid (5.35 μg/ml), with bovine serum albumin (1.25 μg/ml)). Sodium pyruvate (1 mM), ascorbate-2-phosphate (37.5 μg/ml), dexamethasone (10-7 M), TGF-β1
(10 ng/ml, recombinant human, Peprotech), and l-glutamine (4 mM) were also added.
Results
Pellets were harvested at various timepoints and sent for histological sectioning.
Representative histological sections were stained for toluidine blue, and immunohistochemistry was performed for collagen type II (Figure 2-1). Centrifuged cells form an aggregate in less than 24 hours. By day 3 of culture, collagen type I production has been downregulated, and remodeling of the existing collagen type I matrix has been initiated. The onset of collagen type II deposition begins on days 3-5, and extends throughout the pellet by 10-14 days of culture. Production of collagen type X (a marker of hypertrophy) is also found throughout the pellet by day 21.
Aggregates formed through this process have a diameter no greater than 1 mm. Attempts
to form larger pellets – by increasing cellular concentration or volume, resulted in the
formation of many smaller aggregates. Thus, there is a need for methods to scale-up these
constructs to the size of a clinically-relevant sized defect (> 1 cm). One method involves
the use of scaffolds that will hold higher cell numbers in a defined structure. A synthetic,
31
Figure 2-1. Toluidine blue stained histological sections and immunohistochemistry for collagen type II of human pellets after 1, 3, 6, 9, and 14 days of culture.
32
Figure 2-2. Immunohistochemistry for collagen types I (top), II (middle), and X (bottom) in day 1 (left column) and 14 (right column) pellets.
33 photopolymerizable polymer – poly(ethylene glycol) diacrylate (PEGDA) – was kindly
provided by Jennifer West (Rice University, TX) for use in our studies (Figure 2-3). In
her own work, Dr West had encapsulated rat smooth muscle cells in a 20% (w:v) PEGDA polymer [46-48]. Phase transition was accomplished with the photoinitiator
acetophenone, solvated in n-vinyl pyrrolidinone. Studies with this construct reported both
cell viability and extracellular matrix production.
MSC Encapsulation in 20% (w:v) PEGDA (6 kDa) Hydrogels
Methods
Our experimental protocol for pellet culture was modified to include hydrogel
encapsulation. Bone marrow preps were obtained and processed as before; however, once
confluence was reached cells were passaged with FGF-supplemented DMEM + FBS to maintain pluripotency. Previous research on encapsulated chondrocytes indicated the need for much higher cell-seeding densities, and 25-50 x 106 cell/ml were needed for
experimentation. [19, 37, 38, 40, 41, 45, 53, 59] Cells from passage 1 or 2 were used in
all experiments. Once the appropriate cell number was obtained, MSCs were trypsinized,
washed with DMEM + FBS, and then resuspended in defined chondrogenic medium.
This cell-solution was mixed 1:1 with PEGDA polymer solution, and 1.2% (w:v)
acetophenone initiator was added. The cell-polymer solution was mixed thoroughly, and
aliquoted into individual 80 μl wells of a custom-built fabrication chamber. The
fabrication plate was then exposed to long-wave, low-intensity UV light (λ = 365 nm) for
1 minute. Constructs were removed from the chamber, transferred to individual wells of a
34
Figure 2-3. Chemical structure of PEGDA. For the preliminary work, n = 136, which results in a molecular weight of 6,000 daltons (6 kDa).
35 sterile 12 well-plate, and cultured in vitro at 37°C, 5% CO2 in defined chondrogenic
medium for 6 weeks with medium changes every 4 days.
Results
After 6 weeks of culture, 2 hydrogels were sent for histological sectioning and staining
and the remaining were placed in 0.1 N NaOH overnight to break the covalent bonds of
the polymer network. The resulting polymer-cell solution was neutralized with 0.1 N
HCL, to a final pH of 6.0 in preparation for subsequent digestion overnight with papain at
60°C. Quantitative assays for DNA, collagen, and proteoglycan were then performed, facilitated by PicoGreen, hydroxyproline, and DMMB assays, respectively. Results for collagen and proteoglycan were below the level detectable by the assays. Although DNA content was indicative of cell survival, it was much lower than expected, given the initial seeding density. Examination of histological sections revealed the absence of extracellular matrix elaboration (Figure 2-4).
Examination of the Photoinitiator
A review of hydrogel tissue engineering literature indicated other choices of
photoinitiator, including hydroxycyclohexylphenylketone (HPK, Sigma-Aldrich 405612)
in 70% ethanol, and 1-[4-(2-hydroxyethoxy)-phenyl]-2-hydroxy-2-methyl-1-propane-1-
one (IrgacureTM or D2959, Ciba D2959, Switzerland) in water. A series of cytotoxicity
studies was performed on hMSCs in monolayer, assessed through lactose dehydrogenase
(LDH) activity, to determine whether there were significant differences in cellular viability with use of the three different photoinitiators.
36
Figure 2-4. Toluidine blue stained histological sections of 20% (w:v) PEGDA cell- seeded constructs.
37 The LDH assay was first optimized to determine the appropriate seeding density for
monolayer cells that would provide the maximum dynamic range. Based on the data from
live/dead positive and negative controls, a seeding density of 12 x 103 cells/well was
chosen. Human MSCs were plated at this initial seeding density, allowed to adhere
overnight, and then exposed to medium containing each of the three photoinitiators at
various concentrations based on the work of others [19, 97]: HPK and IrgacureTM were examined at 0.03, 0.06, and 0.12%, acetophenone at 0.3, 0.6, and 1.2%. Cells were exposed for 2, 4, 6, 8, 12, 16, 24, or 48 hours. Positive (live) controls were exposed to
DMEM + FBS, while all-dead controls were exposed to 1% Triton-X 100 for 2 hours
prior to examination. Results from all timepoints were similar to the shortest duration
examined (Figure 2-5). Cellular viability –determined by normalizing the absorbance of
the initiator in question to the all-live control – was strongly affected by the
photoinitiator. Irgacure and HPK exhibited similarly favorable cellular viabilities,
although HPK exhibited much higher standard deviation. Exposure to acetophenone,
however, significantly decreased MSC viability. Based on these results, Irgacure was
selected as the new photoinitiator.
Formation of the Semi-Interpenetrating Network
The review of the literature also revealed that greater matrix elaboration by chondrocytes had been reported with use of semi-interpenetrating networks [19, 57]. An interpenetrating network (IPN) is a hydrogel that contains two polymers. These may or may not be covalently linked during the polymerization process (Figure 2-6). A semi- interpenetrating network (semi-IPN) is one in which the second polymer either a) forms
38
1.2
1
0.8
0.6
0.4
0.2 Normalized Cellular Viability
0 D2959 HPK acetophenone all dead control Initiator at Lowest Concentration
Figure 2-5. Cellular viability of hMSCs subsequent to initiator and UV exposure, assessed through lactose dehydrogenase (LDH) activity.
39
Figure 2-6. Idealized and simplified depiction of an interpenetrating network (IPN). Adapted from Mauritz: http://www.psrc.usm.edu/mauritz/nano4.html
40
bonds at the same point as the first polymer, through the same reaction, or b) cannot form covalent linkages at all. The latter form of semi-IPN was employed in both chondrocyte studies.
The purpose of employing a non-covalent semi-IPN is to allow the second polymer to
serve as a physical spacer. This polymer cannot take part in a chemical reaction, but it
may serve as a bridge between chemical linkages of the first polymer. By filling up the
space between polymer chains, and potentially interfering with polymerization
(crosslinking) events, the nonbinding component may increase the distance between
crosslinks, and/or act as a temporary spacer – physically separating crosslinked chains.
Although some of this second non-covalent polymer may be physically entangled, it is
not chemically linked. It is therefore capable of extrusion (release) from the hydrogel
network. In this way, the second component increases nutrient delivery and provides
room for extracellular matrix elaboration.
Methods
Two changes were made to the original PEGDA hydrogel: use of the photo-sensitive
initiator IrgacureTM and addition of PEG-dimethyl ether (88 kDa) to the system to serve
as a non-covalent component. Both of these alterations necessitated changes to the
PEGDA hydrogel. We wanted to achieve polymerization within 10 minutes of UV
exposure (similar to previous studies), and set the initial PEGDA:PEG ratio at 2:1. The
formulation of the semi-IPN that allowed for both of these changes was 16% PEGDA (6
41 kDa), 8% PEG (88 kDa), with 0.06% IrgacureTM (all w:v).
Results
PEGDA semi-IPNs were seeded (1:1) with human MSCs at 25 x 106 cells/ml and
cultured in defined chondrogenic medium. Constructs were harvested at 3 and 6 weeks, fixed in 10% neutral buffered formalin, and paraffin-embedded. Constructs were
sectioned with 7 μm thickness and stained with toluidine blue to examine ECM
production by the encapsulated human MSCs. (Figure 2-7). Toluidine-blue stained
sections at both 3 (Figure 2-7 A,B) and 6 weeks (Figure 2-7 C,D) revealed production of
proteoglycan throughout the construct, with no evidence of preferential deposition or a
necrotic core. Immunohistochemistry for collagen type II, a marker of chondrogenesis,
revealed pericellular staining throughout the scaffold (Figure 2-7 E).
Conclusions
The modified IPN scaffold facilitated in vitro chondrogenesis of human MSCs. This
achievement allowed for the thorough investigation of how specific changes to the
network affected ECM production of differentiating hMSCs. The experiments and
calculations that describe systematic changes to the PEGDA semi-IPN are discussed in
the following chapter.
42
Figure 2-7. Toluidine blue stained histological sections at 3 weeks (A, B, at 20x magnification) and 6 weeks (C, D, at 4x magnification). Immunohistochemistry for collagen type II (E), and negative control (naïve serum – mouse IgG instead of primary antibody (Ab), F).
43 CHAPTER THREE:
Design and Characterization of Poly(ethylene glycol) Photopolymerizable Semi-
Interpenetrating Networks for Chondrogenesis of Human Mesenchymal Stem Cells
Reference:
Buxton AN, Zhu J, West JL, Marchant R, Yoo J, and Johnstone B (2007). Design and characterization of poly(ethylene glycol) photopolymerizable semi-interpenetrating networks for chondrogenesis of human mesenchymal stem cells. Tissue Eng. (in press)
44 ABSTRACT
Mesenchymal stem cells (MSCs) are used extensively in cartilage tissue engineering. We
have developed a photopolymerizable poly(ethylene glycol diacrylate) (PEGDA) and
poly(ethylene glycol) (PEG) semi-interpenetrating network that facilitates the in vitro
chondrogenesis of human MSCs (hMSCs). Network parameters were altered and tested
for their effects on subsequent matrix elaboration. The mesh size (ξ), calculated for each network based on equilibrium swelling ratios (Q), increased with lower PEGDA:PEG ratios and with higher PEGDA molecular weight. Increases in ξ correlated to changes in extracellular matrix content and deposition in hMSC-seeded networks cultured in vitro for 6 weeks in defined chondrogenic medium. Networks constructed with PEGDA (6 kDa) and PEG (88 kDa) at 1:2 displayed intercellular deposition of proteoglycan.
Furthermore, their proteoglycan contents were significantly elevated compared to
PEGDA (6 kDa) hydrogels constructed without the PEG component and those
constructed at a PEGDA:PEG ratio of 2:1, which both exhibited pericellular proteoglycan
deposition. However, networks constructed with PEGDA (12 and 20 kDa) and PEG (88
kDa) exhibited intercellular deposition of proteoglycan regardless of the ratio employed.
Collagen content was decreased in networks constructed with PEGDA (12 and 20 kDa)
and PEG (88 kDa) at ratio of 1:2, compared to those fabricated at the same PEGDA
molecular weights at ratio of 2:1. This study demonstrated that semi-interpenetrating
network parameters influence not only extracellular matrix content but also the
deposition of the matrix molecules by human mesenchymal stem cells undergoing
chondrogenesis. It is important that these parameters be considered carefully when
creating scaffolds for tissue-engineered cartilage.
45 INTRODUCTION
Articular cartilage is a highly specialized, avascular, connective tissue that plays an
important role in reducing friction and distributing shock in diarthrodial joints [2]. Adult
articular cartilage, however, has limited capacity to self-repair in response to trauma or
disease. Current clinical treatment methods frequently fail to restore normal function, and
there is a need for more effective therapies. With the advent of tissue engineering, there
has been a greatly increased effort to effect functional cartilage repair and/or regeneration.
Several scaffolds have been explored for use in cartilage tissue engineering, for both load
and non-load bearing applications, and many groups employ isolated chondrocytes in
these systems [19, 30, 37-41, 43-45, 53, 54, 59, 98-117]. Although these studies demonstrate feasibility and explore the relationships between the nature of the scaffold and cellular response, the isolation of autologous chondrocytes for human use is invasive, requiring biopsy of a non-loadbearing surface of a joint that may also be diseased.
Further, in vitro expansion of a clinically useful number of chondrocytes is hindered by de-differentiation caused by monolayer culture [104, 118, 119]. Bone marrow-derived mesenchymal stem cells (MSCs) have recently become a focus of cartilage research.
MSCs are a relatively unexplored source of chondrogenic cells. They are readily accessible and yield higher numbers during in vitro expansion than chondrocytes isolated from cartilage. Furthermore, when cultured under the appropriate conditions, MSCs can differentiate into chondrocytes.
46 In vitro chondrogenesis of isolated and expanded MSCs was first reported by our group
[22, 25]. The differentiation was accomplished in a scaffold-free cell culture system.
Since that time, tissue engineering cartilage using differentiation of human mesenchymal stem cells (hMSCs) has been reported with many types of scaffold including hydrogels, sponges and meshes. Hydrogels have advantages over porous scaffolds because they allow homogeneous cell-seeding during encapsulation, and extensive control over the process of phase transition. Hydrogels are water-insoluble networks that are capable of absorbing tissue-like quantities of aqueous solutions, creating a 3-dimensional environment that has efficient nutrient delivery and waste removal. One naturally occurring hydrogel is alginate, a linear polysaccharide soluble in aqueous or buffered salt solutions that can be crosslinked in the presence of calcium ions. Caterson et al.
successfully employed alginate to improve MSC loading and retention in a porous PLA
scaffold [120, 121] and Kavalkovich et al. examined the effects of seeding density on
glycosaminoglycan synthesis by MSCs [122]. These studies indicated that MSC differentiation in hydrogels was possible.
Hydrogels constructed from poly(ethylene glycol) (PEG) have been extensively explored
as cell scaffolds because of their hydrophilicity, biocompatibility, and intrinsic resistance
to protein adsorption and cell adhesion. Previous work has shown that aqueous solutions
of acrylated PEG derivatives can be safely photopolymerized in direct contact with cells
and tissues [19, 37-41, 43, 44, 46-48, 51, 53, 54, 58, 59, 85, 98]. Mann et al.
demonstrated both cellular viability and matrix production of smooth muscle cells
encapsulated in a photopolymerizable poly(ethylene glycol diacrylate) (PEGDA)
47 hydrogel [46-48]. Furthermore, Bryant et al. performed several studies that directly related chondrocyte morphology, metabolism, and matrix production to poly(ethylene oxide) (PEO) hydrogel parameters under both static and dynamically loaded conditions
[37-41, 53, 54, 58, 59]. In vitro chondrogenesis of MSCs in a photopolymerizable PEG hydrogel was first reported by Williams et al.[123] Overall, although these studies found expression of biochemical indicators or chondrogenesis, histological and immunohistochemical analysis revealed a limited production of extracellular matrix, indicating the need to understand what parameters of PEG hydrogels influence differentiation and matrix production.
PEG hydrogels can also be formed as semi-interpenetrating networks by using two components: the first is the element that provides crosslinks and the second is a nonbinding element that may add stability and may serve as a spacer – providing space for nutrient delivery and waste removal. Semi-interpenetrating networks have been produced in which chondrocytes elaborate extracellular matrix [19, 57], but the behavior
of hMSCs in this type of hydrogel has not been explored. The purpose of this study was
to examine the parameters of photopolymerizable PEG-based semi-interpenetrating
networks that affect the in vitro chondrogenesis of hMSCs. Multiple parameters of the
network were altered, and chondrogenic differentiation of hMSCs in defined
chondrogenic medium was explored for each variation. Specifically, the molecular
weight and availability of the PEG spacer, the molecular weight of the PEGDA
crosslinker, the PEGDA:PEG ratio, and the degree of polymerization were varied to
determine the properties that most affect matrix production by encapsulated cells. The
48 relationship between the scaffold and differentiating cells was characterized using
histological and immunohistochemical staining. Furthermore, the biochemical properties
of the resulting cell-seeded scaffolds were quantified using fluorimetric and
spectrophotometric assays.
MATERIALS & METHODS
Mesenchymal Stem Cell Isolation and Culture: MSCs were isolated and expanded from
human iliac crest bone marrow aspirates as previously described [22]. Briefly, human
bone marrow was obtained from the iliac crests of consenting donors. Marrow aspirates
were fractionated on a Percoll density gradient and plated in Dulbecco’s modified Eagle’s
medium (DMEM) with 10% fetal bovine serum (FBS). The adherent population of cells
was cultured at 37° C, 5% CO2 with medium changes every four days. Once these
primary cells were confluent, serum-containing DMEM was supplemented with
fibroblast growth factor (FGF-2, 10 ng/ml) to facilitate post-primary passage expansion
with retention of chondrogenic potential. Cells from either passage 1 or 2 were used for
all experiments.
Preparation of PEGDA: The PEGDA (6 kDa) polymer was produced as previously
described [47]. Briefly, 0.1 mM dry PEG was combined with 0.4 mM acryloyl chloride
and 0.2 mM triethylamine in anhydrous dichloromethane and stirred under argon
overnight. The mixture was then lyophilized and frozen. To fabricate PEGDA (12 kDa),
0.2 mmol PEG diol (12 kDa, Aldrich) was dissolved in a 100 ml flask containing 50 ml of anhydrous dichloromethane under nitrogen and magnetic stirring at room temperature.
49 After 10 min, 3 mmol acryloyl chloride and 1.6 mmol triethylamine were added. The
mixture was kept stirring under nitrogen overnight, followed by filtration to remove the
salts of triethylamine chloride. The resulting solution was precipitated in 400 ml of ether.
The final product was obtained by filtering the resulting precipitates, washing with ether
twice, and drying in a vacuum oven at room temperature. PEGDA (20 kDa) was
produced in similar fashion. To start, 0.15 mmol PEG diol (20 kDa, Aldrich) was
dissolved in a 100 ml flask containing 60 ml of anhydrous dichloromethane under
nitrogen and magnetic stirring at room temperature. After 10 min, 2.4 mmol acryloyl
chloride and 2.4 mmol triethylamine were added. Following stirring and filtration of the
above, the resulting solution was precipitated in 500 ml of ether. The final product was
obtained by filtering the resulting precipitates, washing with ether twice, and drying in a
vacuum oven at room temperature.
Cellular Encapsulation & Culture: PEGDA (0.16g; 6, 12, or 20 kDa) and (PEG)n-
dimethyl ether (17.6, 44, or 88 kDa, Polysciences, Warrington, PA) were combined in 0.5
ml sterile PBS at various ratios (2:1 or 1:2, w:w) and filter sterilized using 0.2 μm syringe
filters (Fisher Scientific, Santa Clara, CA). The resulting polymer solution was combined
(1:1, v/v) with expanded cells at 25 x 106 cells/ml. IrgacureTM initiator (Ciba, Tarrytown,
NY) was added (0.03%, 0.06%, or 0.12% w:v) and the subsequent cell-polymer solution was mixed thoroughly, aliquoted into 80 μl wells of a custom built fabrication chamber, and exposed to long-wave UV light for 4-10 minutes. After polymerization, the cylindrical cell-seeded constructs (7 mm diameter, 2 mm height) were transferred to individual wells of a sterile 12 well-plate and maintained in culture at 37°C, 5% CO2 for
50 six weeks in a defined chondrogenic medium previously shown to induce chondrogenic
differentiation of these cells in aggregate culture [22, 25]. The defined medium consisted
of high-glucose DMEM with ITS+ Premix (Collaborative Biomedical Products: insulin
(6.25 μg/ml), transferrin (6.25 μg/ml), selenous acid (6.25 μg/ml), and linoleic acid (5.35
μg/ml), with bovine serum albumin (1.25 μg/ml)). Sodium pyruvate (1 mM), ascorbate-
2-phosphate (37.5 μg/ml), dexamethasone (10-7 M), TGF-β1 (10 ng/ml, recombinant
human, Peprotech), and l-glutamine (4 mM) were also added.
Swelling Ratio & Mesh Size Calculations: Scaffolds were fabricated without cells for swelling ratio studies according to the above protocol for all PEGDA and PEG combinations and ratios. Constructs were placed in PBS at 37°C for 10 days. Once equilibrium had occurred, scaffolds were weighed to determine Weq. Constructs were
then dried in a vacuum under high heat for 3 hours and weighed to determine Wdry. The volumetric swelling ratio (Q) was determined using the following equation:
⎛ −WW ⎞ Q = ⎜ eq dry ⎟ (1) ⎜ ⎟ ⎝ Wdry ⎠
The molecular weight between crosslinks (MC) can be approximated using the Peppas-
Merrill model [124], which was modified from the Flory-Rehner model [125]:
ν [ )1ln( ++− μVVV 2 ] 21 V s ,2,2 s ,2 s −= 1 (2) 1 C MM n ⎡ ⎤ ⎛V ⎞ 3 1 ⎛V ⎞ V ⎢⎜ ,2 s ⎟ − ⎜ ,2 s ⎟⎥ ,2 r ⎢⎜ ⎟ ⎜ ⎟⎥ ⎝V ,2 r ⎠ 2 ⎝V ,2 r ⎠ ⎣⎢ ⎦⎥
Here Mn is the number average molecular weight of the polymer, ν is the specific volume
3 of bulk PEG in the amorphous state (0.893 cm /g), V1 is the molar volume of the solvent
51 (18 cm3/mol), μ is the Flory-Huggins polymer-solvent interaction parameter (0.426 for
PEG-water as determined by Merrill et al. [126]), and V2,r and V2,s are the volume fraction of the polymer at the relaxed state and swelling equilibrium, respectively. V2,r is equivalent to the volume concentration of the solution where crosslinking occurs
(14.3%). V2,s is equal to the reciprocal of the volumetric swelling ratio (v2,s = Vp/Veq =
1/Q; Vp = dry volume, Veq = volume in the equilibrium swollen state of the gel).
The mesh size, ξ , can be calculated on the basis of [127]:
1 1 − 3 2 2 =νξ ,2 s (r0 ) (3)
1 2 2 where ()r0 is the unperturbed mean-square end-to-end distance of the PEG:
1 1 12 2 2 ⎛ 2M C ⎞ ⎜ ⎟ 2 ()0 = lr ⎜ ⎟ Cn (4) ⎝ M r ⎠ and l is the average value of the bond length between C-C and C-O bonds in the PEG repeat unit [-O-CH2-CH2-] (1.46 A), Mr is the molecular mass of the PEG repeat unit (44 g/mol), and Cn is the characteristic ratio for PEG (4).
Histology and Immunohistochemistry: After 42 days in culture, constructs were fixed in
10% neutral buffered formalin (Fisher Scientific) before placing in Tissue-Loc
Histoscreen histocassettes (C-0250, Richard-Allan Scientific, Kalamazoo, MI). Samples were then processed and embedded in paraffin. Representative samples were selected, and 7 μm serial sections were cut and stained with toluidine blue for histological evaluation. For immunohistochemical analysis, unstained samples were deparaffinized
52 with xylene (Fisher Scientific) and blocked in 5% bovine serum albumin (BSA) in phosphate buffered saline (PBS) for 30 minutes before undergoing 30 minute digestion with pronase (1 mg/ml in PBS) or chondroitinase ABC (0.1 U/ml in Tris-Acetate, pH 8.0) to facilitate collagen and proteoglycan antibody access to the extracellular matrix, respectively. Proteoglycan and types I, II, and X collagen were immunolocalized with monoclonal antibodies at a 1:100 dilution in 1% BSA and visualized with a fluorescein isothiocyanate (FITC)-linked secondary antibody (55499, MP Biomedicals, Aurora, OH) at a dilution of 1:400 in 1% BSA. Specifically, anti-chondroitin sulfate (2B6) antibodies
(kindly provided by Bruce Caterson, U. Cardiff, Wales, UK), anti-collagen type I antibody (CP17L, EMD Biosciences, San Diego, CA), anti-collagen type II antibody (II-
II6B3, NIH Hybridoma Bank maintained at U. Iowa), and anti-collagen type X antibody
(kindly provided by Gary Gibson, Ph.D. Henry Ford Hospital & Medical Centers,
Detroit, MI) were used to characterize the extracellular matrix produced in the scaffolds.
Aggregates (14 days incubation) were used as positive controls, as previously described
[22, 25]. To assess the level of antibody specificity, two negative controls were also run: following digestion, sections were incubated either with mouse IgG, or without primary antibody before incubation with FITC-labeled secondary antibody.
Quantitative Assays: After 42 days in culture, constructs were removed from culture
media, rinsed with Tyrodes salt solution, and digested with NaOH (0.1 N) overnight at
60ºC. HCl (0.1 N) was added to neutralize and pH was adjusted to 6.0 [47]. Samples
were then digested for 18 h at 60ºC in 125 μg/ml papain in buffer (10 mM ETDA, 2 mM
cysteine, pH 6.0) (Sigma, St. Louis, MO). Sulfated glycosaminoglycan content of the
53 digested cell-polymer construct was assessed spectrophotometrically using the 1,9- dimethylmethylene blue (DMMB) dye assay (Polysciences, Warrington PA, pH 3.0)
[128]. Sample glycosaminoglycan content was compared to shark cartilage chondroitin
sulfate C standards (SIGMA-Aldrich, Oakville, Ontario, Canada). DNA content of the
digested, neutralized hydrogels was determined spectrofluorimetrically using the
PicoGreen fluorescent DNA binding dye assay according to manufacturer’s instructions
(P-7581, Molecular Probes). Sample fluorescence was compared to calf thymus DNA standards (Sigma, St. Louis, MO). Total collagen content was determined by oxidation of
OH-Pro residues in collagen with chloramine T trihydrate (ICN Biomedicals, Aurora,
OH) and developed with p-dimethylaminobenzaldehyde (Ehrlich’s reagent, ICN
Biomedicals) [129]. Sample concentrations were compared to hydroxyproline standard solutions made from trans-4-hydroxy-L-proline (56250, Fluka).
Statistical Analysis: Data for the biochemical composition of constructs were analyzed
by a one-way ANOVA using Tukey’s test for multiple comparisons. Each series of
experiments was run in triplicate with three separate human marrow preparations, with
n=5 samples for each biochemical analysis. Statistical significance was determined at a
value of p< 0.05 (Minitab®, Minitab Inc., State College, PA).
RESULTS
Degree of Network Polymerization
Preliminary experiments were conducted to determine the polymerization conditions that
could be used when forming the semi-interpenetrating networks. The duration of UV
54 exposure, and the concentration of Irgacure™ were varied to examine their effects on
matrix deposition. Neither proteoglycan, nor collagen content was significantly affected
by altering the duration of UV exposure from 4 to 10 minutes (data not shown).
Proteoglycan was unaffected by the concentration of Irgacure™ (p = 0.07); however, collagen content was significantly decreased at 0.03% (p = 0.001, Figure 3-1). Thus, the duration of UV exposure was set at 6 minutes, and Irgacure™ was maintained at 0.06%
(w:v) for all further experiments.
Swelling Ratios/Mesh Size
Volumetric swelling ratios (Q) for hydrogels using PEGDA (6 kDa) were determined
(Table 3-1). Altering PEG molecular weight (17.6, 44, or 88 kDa) did not affect Q at a
PEGDA:PEG ratio of 2:1; however, removal of the physical component significantly
decreased Q (p<0.001). Increasing the quantity of PEG, facilitated by decreasing the
PEGDA:PEG ratio to 1:2, significantly increased Q (p<0.001). Swelling ratios were also
calculated for PEGDA (12 kDa and 20 kDa) at 2:1 using PEG (88 kDa) as the physical
component (Table 3-1). In both cases, increasing the PEGDA molecular weight
significantly increased Q (p<0.001). Decreasing the ratio of PEGDA:PEG to 1:2 also
increased Q for all PEGDAs, (p<0.001).
The corresponding molecular weight between crosslinks (MC) and mesh size (ξ) were
calculated for PEGDA (6, 12, and 20 kDa) and PEG (88 kDa) with PEGDA:PEG ratios
of 2:1 and 1:2 (Table 3-2). Incorporation of the PEG spacer significantly increased ξ for
55
Figure 3-1. Extracellular matrix content of PEGDA (6 kDa) hydrogels polymerized with increasing concentration of initiator. DNA (A), proteoglycan (B), and collagen (C) content (mean +/- s.d.) of constructs cultured in defined medium for 6 weeks after initial hydrogel formation with 0.03, 0.06 or 0.12% (w/v) IrgacureTM concentrations. *p<0.001 compared with 0.03% (w/v) IrgacureTM.
56
Table 3-1. Experimentally determined swelling ratios for non-cell-seeded PEGDA hydrogels fabricated with and without PEG.
57
Table 3-2. Molecular weight between crosslinks, MC, and mesh size, ξ, calculated for PEGDA hydrogels.
58 all PEGDA molecular weights at both ratios (p<0.001). Further, ξ was statistically increased at a PEGDA:PEG ratio of 1:2 compared with 2:1, (p<0.001).
Contribution of the PEG Spacer with PEGDA (6 kDa)
A series of experiments was run to determine whether increases in the swelling ratio affected in vitro chondrogenesis of encapsulated hMSCs. Preliminary experiments were run using hydrogels constructed at a PEGDA:PEG of 2:1 with PEGDA (6 kDa) and PEG
(17.6, 44, or 88 kDa), since their swelling ratios were not statistically different. These were compared to hydrogels formed without PEG, which do exhibit a lower Q. All cell- seeded hydrogels were cultured for 6 weeks in vitro in defined medium. Human MSCs in gels constructed with PEG underwent chondrogenic differentiation and matrix production, and the matrix production was not statistically different between gels with the three different PEG molecular weights (Figure 3-2). However, hMSCs in hydrogels formed without PEG exhibited a significantly lower extracellular matrix production as evidenced by histological staining and quantitative assays for proteoglycan (p<0.001) and collagen (p=0.001). Since there was no statistically significant effect of the PEG molecular weight on matrix deposition, but its inclusion facilitated greater matrix production, PEG (88 kDa) was included for all further experiments.
Human MSC-seeded hydrogels were subsequently formed with PEGDA (6 kDa) and
PEG (88 kDa) at ratios of 2:1 and 1:2 to determine the effect on matrix deposition.
59
Figure 3-2. Extracellular matrix content of constructs fabricated with the addition of PEG to PEGDA (6 kDa) at a PEGDA:PEG ratio of 2:1. Proteoglycan (A) and collagen (B) content (mean +/- s.d.) of PEGDA (6 kDa) constructs fabricated at 16% (w:v) or with PEG (17.6, 44, or 88 kDa) at PEGDA:PEG of 2:1. *p<0.001 compared with PEGDA 6 kDa alone.
60 Quantitative assays indicated that hydrogels formed at a PEGDA:PEG ratio of 1:2 exhibited significantly higher proteoglycan content compared with hydrogels constructed at a PEGDA:PEG ratio of 2:1 (p=0.002, Figure 3-3). Total collagen content was unaffected. Histological sections stained for toluidine blue indicated marked differences in matrix production throughout the scaffold (Figure 3-4a). At 2:1 proteoglycan deposition was pericellular, but at 1:2 it was detected throughout the construct – although the strongest staining was in the pericellular region. Immunohistochemistry for chondroitin sulfate confirmed that the toluidine blue reflected proteoglycan deposition; staining extended beyond the pericellular region and into the intercellular space.
Immunohistochemistry for collagen types I, II, and X revealed similar patterns for all three collagen types (Figure 3-4b). Collagen type I was not detected. Collagen type II is a marker of chondrogenesis and exhibited a pericellular staining pattern throughout the scaffold. Evidence of hypertrophy was indicated by the presence of collagen type X in the pericellular region. Staining for collagen content of PEGDA (6 kDa) at 2:1 was very similar to 1:2, (not shown).
Hydrogels seeded with hMSCs were constructed with PEGDA (6, 12 kDa, or 20 kDa) at
PEGDA:PEG ratios of 2:1 and 1:2, using PEG (88 kDa). Altering PEGDA molecular weight with maintenance of the PEGDA:PEG ratio at 2:1 did not affect the proteoglycan content for any constructs compared with PEGDA (6 kDa) hydrogels (Figure 3-5A).
However, as seen in Figure 3-3, the proteoglycan content of PEGDA (6 kDa) hydrogels fabricated at 1:2 was significantly increased compared with PEGDA (6 kDa) constructs
61
Figure 3-3. Extracellular matrix content of PEGDA (6 kDa) hydrogels constructed at varying ratios of PEGDA:PEG. Proteoglycan (A) and collagen (B) content (mean +/- s.d.) of cultured PEGDA (6 kDa) constructs with PEGDA:PEG at 2:1 and 1:2. *p≤0.002 compared with PEGDA 6 kDa at 2:1.
62
Figure 3-4. Proteoglycan distribution in PEGDA (6 kDa) constructs fabricated with PEG at different ratios. (a) PEGDA:PEG of 2:1 (A,C,E) and 1:2 (B,D,F). Toluidine blue staining (A,B), immunohistochemical staining for chondroitin sulfate (C,D). Negative controls for chondroitin sulfate immunohistochemistry (E,F). (b) Immunohistochemistry for collagen types I (A), II (B) and X (C) with PEGDA(6 kDa):PEG at 1:2. Negative controls for collagen type I (D), II (E), and X (F). (Black scale bar = 200 μm, white scale bars = 100 μm.)
63
Figure 3-5. Extracellular matrix content of networks constructed with increased PEGDA molecular weight and PEG (88 kDa) at varying PEGDA:PEG ratios. Proteoglycan (A,B) and collagen (C,D) content (mean +/- s.d.) of cultured constructs fabricated with PEGDA (6, 12, or 20 kDa) and PEGDA:PEG of 2:1 (A,C) or 1:2 (B,D). *(B) p≤0.002 compared with PEGDA 6 kDa at 1:2; *(C) p≤0.002 compared with PEGDA 6 kDa at 2:1; *(D) p≤0.033 compared with PEGDA 6 kDa at 1:2.
64 made at 2:1 (Figure 3-5B). The use of different human cell preparations between experiments accounts for the discrepancy in magnitude of ECM content between the two figures, but the overall result is the same. Hydrogels constructed with higher PEGDA molecular weights (12 and 20 kDa) at PEGDA:PEG of 1:2 exhibited decreased proteoglycan content compared with PEGDA (6 kDa) constructs fabricated at the same ratio (p=0.002, Figure 5B). Furthermore, PEGDA (20 kDa) constructs made at 1:2 contained significantly less proteoglycan compared with those fabricated with the same
PEGDA at 2:1, (p=0.002). The collagen content of PEGDA (12 kDa) hydrogels constructed at PEGDA:PEG of 2:1 was elevated compared to all other formulations
(p=0.002, Figure 5C). However, PEGDA (20 kDa) networks fabricated at 1:2 exhibited decreased collagen content compared with those constructed with the same PEGDA at ratio of 2:1 (p=0.002), and with hydrogels constructed with PEGDA (6 or 12 kDa) at 1:2
(p=0.033).
Histological and immunohistochemical analyses revealed differences in matrix deposition
(Figure 3-6). With increased PEGDA molecular weight, proteoglycan distribution was altered from pericellular to intercellular, regardless of the ratio of PEGDA:PEG. The semi-interpenetrating network does not stain with toluidine blue application, and appears white during brightfield examination. Immunohistochemistry for proteoglycan confirmed intercellular deposition of sulfated-GAGs (data not shown). Quantitative assays revealed no change in DNA content of the hydrogels (data not shown). However, hydrogels constructed at 2:1 displayed a more uniform cellular distribution, and matrix deposition throughout the construct. Those constructed at a ratio of 1:2 exhibited intense
65
Figure 3-6. Proteoglycan distribution in higher molecular weight PEGDA constructs fabricated with PEG (88 kDa). Toluidine blue stained histological sections of hydrogels made with PEGDA 6 kDa (A,D), 12 kDa (B,E), 20 kDa (C,F) at PEGDA:PEG of 2:1 (A,B,C) and 1:2 (D,E,F). (Scale bar = 250 μm.)
66 metachromatic staining; however, cellular distribution appeared aggregated and was
localized to the center of the construct for PEGDA (12 and 20 kDa).
Immunohistochemistry for collagen types II and X revealed pericellular staining
throughout the scaffold at both PEGDA:PEG ratios for all PEGDA molecular weights
(data not shown). Collagen type I was not detected in constructs of any ratio.
DISCUSSION
The goal of this study was to investigate the effects of scaffold parameters on in vitro
chondrogenic differentiation of hMSCs encapsulated in a 3-dimensional hydrogel. A semi-interpenetrating network was created by incorporating high molecular weight PEG
into the PEGDA polymer solution, allowing for increased space for nutrient exchange
and matrix deposition through interruption of the crosslinking reaction. The addition of
PEG to PEGDA (6 kDa) hydrogels at PEGDA:PEG ratio of 2:1 significantly increased
proteoglycan content of hMSC-seeded constructs. Riley et al.[57] reported a similar
effect with bovine chondrocytes: an increase in both proteoglycan and collagen
production by cells in poly(ethylene glycol dimethacrylate) (PEGDM)/PEG constructs
over those in monomeric PEGDM constructs. They also found that incorporation of the
PEG component increased water uptake of chondrocyte-seeded constructs. Similarly,
addition of PEG to constructs in our system increased the swelling ratio and mesh size for
all hydrogels constructed with PEGDA (6, 12 or 20 kDa) compared with PEGDA (6 kDa)
gels without PEG. Altering the PEGDA:PEG ratio to 1:2 further increased both Q and ξ
for all PEGDA (6, 12, and 20 kDa), compared with hydrogels fabricated at 2:1.
67 Experimentally determined swelling ratios were used to calculate the corresponding MC and ξ of the resulting semi-interpenetrating networks, using previously established methods [124, 125, 127]. The Peppas-Merrill model applies to systems that have solvent present during monomer crosslinking [130-133]. Furthermore, in crosslinking swelling systems (v2,s < v2,r) with water-soluble monomers (μ < 0.5), the value calculated for MC is
always smaller than Mn/2 [132]. Similar to previous studies, the MC calculated for 16%
PEGDA (6 kDa) was 927 g/mol, far less than Mn of the starting material. Differences in
calculated MC values arise due to variability in the type and concentration of polymerization initiator, the duration of UV exposure, and the concentration of the monomer [130, 132, 133]. Although our analysis will only give an approximate result, calculations for MC were made to quantify the changes in the nature and order of ξ,
which arose as a result of changes to network parameters. The calculated values provide a
minimum value of the pore diameter.
Free radical polymerizations rarely go to completion. This is particularly true for our
system, which employs minimum levels of initiator and UV exposure. The calculations
for ξ thus provide a minimum value of the pore dimension that directly correlates with
changes to the network. All but the networks formed with the highest molecular weight
PEGDA possess minimum pore dimensions smaller than the diameter of nascent collagen
type II fibrils (10-20 nm) [134] and aggrecan (80-100 nm) [135]. However, pore sizes of
the network will not be uniform; rather, there will be a distribution of sizes. We may also
approximate ξ using the C-C and C-O bond lengths of the end groups and repeat unit of
the monomer. For example, for PEGDA (6 kDa) with a reaction at 100% completion and
68 perfect bond formation, the end-to-end distance of the monomer would be 60 nm.
Incorporation of the PEG spacer will certainly interrupt bond formation, such that this
second theoretical value must also be considered only a minimum value. Inclusion of the spacer, early termination of the reaction, side reactions, dangling ends, and chain entanglements will all result in increases to the pore diameter. For all of these reasons, the distribution of ξ for each network will range from ξ calculated, to a value greater than
the ξ approximated by bond lengths, which exceeds the dimensions of the extracellular
matrix proteins synthesized and secreted during chondrogenesis.
Increased PEGDA molecular weight did not affect proteoglycan content at a
PEGDA:PEG ratio of 2:1, but did affect the distribution. In contrast, at a ratio of 1:2,
proteoglycan content was significantly greater for PEGDA (6 kDa) hydrogels compared
with hydrogels fabricated with PEGDA (12 or 20 kDa). It was also greater than any
hydrogel made at a 2:1 ratio. Several groups have explored the effects of crosslinking
density and polymer chemistry on matrix production of chondrocytes encapsulated in
polymer networks [37, 40, 57-59]. Riley et al. [57] altered the mesh size of their semi- interpenetrating network through the addition of high molecular weight poly(ethylene oxide) (PEO) in differing ratios. They reported that constructs with 30% PEGDM (3.4 kDa), PEGDM:PEO of 30:70, contained more proteoglycan than those with 20%, 50%, or 100% PEGDM. Bryant et al.[37, 58, 59] altered the percentage of PEGDM (w:v) used to form the hydrogel, and copolymerized at different ratios with a second polymer possessing degradable linkages to affect hydrogel properties. They reported that proteoglycan content was elevated in a 50% degrading copolymer network, as compared
69 to 75% and 85% networks, and that it was homogeneously distributed throughout the
construct at 4 weeks in vitro. They hypothesized that the substantial increase in mesh size
facilitated intercellular deposition of larger proteoglycans. In contrast, although their 75%
and 85% gels exhibited GAG deposited in the intercellular space, void spaces were
visible in the cell-hydrogel constructs. They speculated that these voids were the result of
decreased mechanical integrity, which arose because of the high degree of degradation.
The results of these studies indicated that there may be an optimal mesh size for
proteoglycan production and deposition, as was also indicated for proteoglycan content in
our system. PEGDA (6 kDa) constructs made with decreased PEGDA:PEG ratio (at 1:2)
deposited proteoglycan in both the pericellular and the intercellular space—throughout
the entire scaffold. Constructs made with higher PEGDA molecular weight (12 or 20
kDa) also displayed intercellular deposition of proteoglycan, regardless of the ratio
employed. The quantifiable difference in ξ between all networks studied is <10 nm,
which is far less than that required to allow for diffusion of proteoglycans. However, as
discussed earlier, this value should be viewed as a minimum. That stated, this parameter
can still be used to design alterations to the semi-interpenetrating network that may allow for wider diffusion of proteoglycans and collagen.
Addition of PEG to PEGDA (6 kDa) hydrogels resulted in greater collagen content, in agreement with results found for bovine chondrocytes encapsulated in PEGDM:PEO semi-interpenetrating networks [57]. Thus, we anticipated that collagen content would also be greater if the ratio of PEGDA:PEG was decreased from 2:1 to 1:2; however, collagen content was unaffected for PEGDA (6 kDa) gels, and actually decreased for
70 PEGDA (12 and 20 kDa) constructs. Networks fabricated with PEGDA (12 and 20 kDa)
and PEG (88 kDa) exhibit significantly greater mesh sizes, ξ ≥ 10 nm, compared to those
values calculated for hydrogels formed with PEGDA (6 kDa) with or without PEG. Since
this study involves hMSCs undergoing chondrogenesis, the collagen fibrils may be
similar to those found in nascent tissues: embryonic cartilages possess very fine collagen type II fibrils of 10-20 nm in diameter, which differ from fibrils found in mature cartilage that range from 40 – 150 nm, depending on the zone in which they are located [134]. For
PEGDA (12 and 20 kDa), the minimum values for ξ based on bond lengths are 120 and
200 nm, respectively. It is thus possible that collagen may be lost from the constructs.
Medium was not assayed for the presence of collagen so this is conjecture and will be
examined in future work. Stem cells require cell-cell contact to undergo chondrogenesis
in vivo and in scaffold-free systems. For networks constructed with smaller molecular weight PEGDA, the close bond-formations surrounding the cell may compensate for direct cell-cell contact. It is possible that cells encapsulated within higher molecular
weight PEGDA constructs might receive feedback that induces a preferential reduction of
one or more types of collagen in the system, since their pore diameter may vary widely.
These changes will also be examined in future work on the influence of scaffolds on
extracellular matrix elaboration by differentiating hMSCs.
We have shown that a photopolymerizable PEG-based system can be used to fabricate hydrogels that facilitate chondrogenesis of hMSCs. As previously reported, hydrogel parameters greatly influence the matrix deposition of encapsulated cells. In our system,
proteoglycan deposition, in particular, is significantly impacted by increasing the
71 swelling ratio and mesh size of the resulting semi-interpenetrating networks. Increases in
ξ, even those on the order of 1 nm as quantified by volumetric swelling ratios, substantially alter proteoglycan content and deposition throughout the construct; collagen content, however, was less sensitive. Based on our current data, we would recommend construction and further exploration of networks that possess a minimum value of
ξ (derived from volumetric swelling ratios) in the range of 5-10 nm.
Because the gel system utilized here is photopolymerizable, it can be used to fabricate constructs in vitro and in vivo. This flexibility in polymerization allows the constructs to be formed at the exact location of damage and take on any desired shape. Sharma et al.
[136] recently used a similar PEG-based system to transdermally photopolymerize goat
MSCs subcutaneously implanted in nude mice, and examined subsequent cartilage formation in vivo. Since the parameters that constitute the semi-interpenetrating network have been systematically explored for their effects on chondrogenesis of hMSCs in vitro, future work would entail testing the scaffold for integration and repair in vivo in animal models.
ACKNOWLEDGEMENTS
The authors would like to thank Dr. Alex Jamieson for his assistance in calculating network parameters, and Teresa Pizzuto for her assistance in histology preparations.
Studies in this manuscript were supported in part by NIH RO1 AR048132 (BJ).
72 CHAPTER FOUR:
In Vitro Chondrogenesis of Human Mesenchymal Stem Cells in Hydrogels is
Affected by Temporal Exposure to Chondrogenic Factors
Expanded from:
Buxton AN, Yoo JU, and Johnstone B. (2007) In Vitro Chondrogenesis of Human
Mesenchymal Stem Cells is Affected by Temporal Exposure to Chondrogenic Factors.
Stem Cells (submitted)
73 ABSTRACT
Bone marrow-derived mesenchymal stem cells (MSCs) are advantageous in cartilage
tissue engineering because they yield high numbers during monolayer expansion, are
more accessible than native chondrocytes, and have been shown to differentiate along
multiple lineages including the chondrocyte pathway. This study examined the effects of
altering exposure to chondrogenic factors during in vitro culture on matrix elaboration of
human MSCs (hMSCs) seeded in a poly(ethylene glycol) (PEG)-based hydrogel. Initial
experiments performed to optimize cell seeding density discovered a direct correlation
between matrix production throughout the construct and seeding density. However,
matrix content normalized to DNA content revealed maximum collagen and proteoglycan
production at seeding densities of 10-25 x 106 cells/ml. Interestingly, constructs seeded
at the highest seeding density, 50 x 106 cells/ml, exhibited decreased matrix contribution
on a per cell basis. Withdrawal of transforming growth factor (TGF-β1) from defined
medium affected both cellular proliferation and collagen production. Dexamethasone
withdrawal decreased collagen production throughout the scaffold, and on a per-cell
basis. Monolayer hMSCs that were pre-treated with defined chondrogenic medium prior
to hydrogel encapsulation exhibited decreased extracellular matrix production that was
more pronounced with increased duration. These studies not only facilitate in vitro
engineering of cartilage, but also aid in implantation strategies for in vivo tissue engineered constructs.
74 INTRODUCTION
Bone marrow-derived mesenchymal stem cells (MSCs) have recently become a focus of
cartilage research. MSCs are advantageous as a source of chondrogenic cells because
they are more accessible than native chondrocytes, yield higher numbers during in vitro
expansion, and are capable of differentiating along multiple lineages in vitro when
cultured in defined medium [22-28]. In vitro chondrogenesis of isolated and expanded
bone marrow-derived MSCs was first reported by our group [22, 25]. The differentiation
was accomplished in a scaffold-free cell culture system that provided elements seen in
the pre-cartilage condensation phase of limb development. The defined medium that
facilitated differentiation included Transforming Growth Factor-β1 (TGF-β1) and
dexamethasone (DEX) [4]. MSCs can be driven to differentiate along the osteogenic
pathway with exposure to DEX [68-70, 137-142]. Exposure to DEX, without or with
other factors (ascorbic acid, vitamin D, growth factors, or other steroid hormones), can
also lead pluripotent mesenchymal cells or cell lines to differentiate into muscle cells,
adipocytes, and chondrocytes [22-26, 69]. Although DEX was sufficient to promote
chondrogenesis in some preparations of rabbit MSCs in the in vitro system, the addition
of TGF-β1 provided consistent differentiation of rabbit and human MSCs (hMSCs) [1, 4]
Since these initial findings, many groups have studied the combined effects of exposure
to DEX and TGF-β on both chondrocytes and MSCs derived from bone marrow, adipose,
and periosteum [22, 25, 64, 65, 72, 73, 75, 143-147].
The defined chondrogenic medium, or variations of it, has been used for tissue engineering of cartilage constructs by many groups [19, 22, 25, 33, 64, 65, 72, 73, 122,
75 123, 136, 144, 146-153]. We previously reported on its use with hMSCs in
photopolymerizable semi-interpenetrating network containing poly(ethylene glycol
diacrylate) (PEGDA) and poly(ethylene glycol) (PEG) [154]. The initial study concerned
the effects of various scaffold parameters for differentiation and matrix elaboration: the
presence of the non-binding PEG was important for matrix development, and its
importance increased when lower molecular weight PEGDA was used as the network
crosslinker. The purpose of the present study was to examine the effects of culture
conditions on subsequent matrix development in this polymer hydrogel. Initial
experiments defined the optimal cell seeding density. Subsequent experiments tested the
effects of altered exposure to the chondrogenic factors, including pretreatment with
defined chondrogenic medium of monolayer-expanded hMSCs prior to
photoencapsulation in the semi-interpenetrating network, and phased removal of TGF-β1
or DEX from the medium of cell-seeded constructs at various timepoints. Morphological
and immunohistochemical staining and quantification of the biochemical properties of the
resulting cell-seeded scaffolds were used to discern the effects of altering the various
culture parameters.
MATERIALS & METHODS
Mesenchymal Stem Cell Isolation and Culture: Mesenchymal stem cells (MSCs) were isolated and expanded from human iliac crest bone marrow aspirates as previously
described [22, 25]. Briefly, human bone marrow was obtained from the iliac crests of
consenting donors. Marrow aspirates were fractionated on a Percoll density gradient and
plated in Dulbecco’s modified Eagle’s medium (DMEM) with 10% fetal bovine serum
76 (FBS). Adherent cells were cultured at 37° C, 5% CO2 with medium changes every four
days. Once primary cells were confluent, serum-containing DMEM was supplemented
with fibroblast growth factor (FGF-2, 10 ng/ml) to facilitate post-primary passage
expansion with retention of chondrogenic potential. Cells from either passage 1 or 2 were
used for all experiments. For some experiments, once monolayer cells had reached
confluence, FBS-containing medium was removed and defined chondrogenic medium
was added to monolayer cells for 1, 3, or 6 days prior to trypsinization and
photoencapsulation. For comparison, control hydrogels were fabricated from hMSCs
from the same human prep. Control hMSCs were continuously exposed to DMEM + FBS
supplemented with FGF-2.
Preparation of PEGDA: The PEGDA (6 kDa) polymer was produced as previously
described [47]. Briefly, PEGDA was prepared by combining 0.1 mM dry PEG with 0.4
mM acryloyl chloride and 0.2 mM triethylamine in anhydrous dichloromethane and
stirred under argon overnight. The mixture was then lyophilized and frozen.
Cellular Encapsulation & Culture: PEGDA (0.16 g, 6 kDa) and (PEG)n-dimethyl ether
(Mw 88 kDa, Polysciences, Warrington, PA) were combined in 0.5 ml sterile PBS at 1:2
(w:w) and filter sterilized using 0.2 μm syringe filters (Fisher Scientific, Santa Clara,
CA). The resulting polymer solution was combined (1:1) with expanded cells with seeding densities ranging from 1 to 50 x 106 cells/ml. IrgacureTM initiator (Ciba,
Tarrytown, NY) was added (0.06%, w:v. The subsequent cell-polymer solution was
mixed thoroughly, aliquoted into 80 μl wells of a custom built fabrication chamber, and
77 exposed to long-wave UV light for 6 minutes. After polymerization, the cylindrical cell-
seeded constructs (7 mm diameter, 2 mm height) were transferred to individual wells of a sterile 12 well-plate and maintained in culture at 37°C, 5% CO2 for six weeks in a
defined chondrogenic medium [22, 25]. The defined medium consisted of high-glucose
DMEM with ITS+ Premix (Collaborative Biomedical Products: insulin (6.25 μg/ml), transferrin (6.25 μg/ml), selenous acid (6.25 μg/ml), and linoleic acid (5.35 μg/ml), with bovine serum albumin (BSA, 1.25 μg/ml)). Sodium pyruvate (1 mM), ascorbate-2- phosphate (37.5 μg/ml), dexamethasone (10-7 M), TGF-β1 (10 ng/ml, recombinant
human, Peprotech), and l-glutamine (4 mM) were also added. For some experiments,
either dexamethasone or TGF-β1 was withdrawn at 7, 14, 21, or 28 days in culture. For
comparison, hydrogels were fabricated from hMSCs from the same human prep, and
continuously exposed to either the complete chondrogenic medium or chondrogenic medium lacking TGF-β1.
Histology: After 42 days in culture, constructs were fixed in 10% neutral buffered
formalin (Fisher Scientific) before placing in Tissue-Loc Histoscreen histocassettes (C-
0250, Richard-Allan Scientific, Kalamazoo, MI). Samples were then processed and
embedded in paraffin. Representative samples were selected, and 7 μm serial sections
were cut and stained with toluidine blue for histological evaluation.
Quantitative Assays: After 42 days in culture, constructs were remo ved from culture
media, rinsed with Tyrodes salt solution, and digested with NaOH (0.1 N) overnight at
60ºC. The pH was adjusted to 6.0 with HCl (0.1 N) and the samples were then digested
78 for 18 h at 60ºC with 125 μg/ml papain in 10 mM EDTA, 2 mM cysteine, pH 6.0 (Sigma,
St. Louis, MO). [47]. Sulfated glycosaminoglycan content of the digested cell-polymer
construct was assessed spectrophotometrically using the 1,9-dimethylmethylene blue
(DMMB) dye assay (Polysciences, Warrington PA, pH 3.0) [128]. Sample
glycosaminoglycan content was compared to shark cartilage chondroitin sulfate C
standards (SIGMA-Aldrich, Oakville, Ontario, Canada). DNA content of the digested,
neutralized hydrogels was determined spectrofluorimetrically using the PicoGreen
fluorescent DNA binding dye assay according to manufacturer’s instructions (P-7581,
Molecular Probes). Sample fluorescence was compared to calf thymus DNA standards
(Sigma, St. Louis, MO). Total collagen content was determined by oxidation of
hydroxyproline residues in collagen with chloramine T trihydrate (ICN Biomedicals,
Aurora, OH), developed with p-dimethylaminobenzaldehyde (Ehrlich’s reagent, ICN
Biomedicals) [129]. Hydroxyproline is a marker for collagen synthesis [129]. Sample
concentrations were compared to hydroxyproline standard solutions made from trans-4-
hydroxy-L-proline (Fluka).
Statistical Analysis: Data for the biochemical composition of constructs were analyzed
by a one-way ANOVA using Tukey’s test for multiple comparisons. Each series of experiments was run using a minimum of three different marrow cell isolates, with 5
replicates for biochemical analysis for each condition. Statistical significance was
determined at a value of p< 0.05 (Minitab®, Minitab Inc., State College, PA).
79 RESULTS
Seeding Density
Preliminary experiments were conducted to determine the optimal seeding density of
hMSCs. Cells were encapsulated in hydrogels at 50, 25, 10, 5, 2.5, and 1.25 x 106 cells/ml and constructs were incubated for 6 weeks in defined chondrogenic medium. The results indicated that densities of 10-12.5 x 106 cells/ml facilitated the greatest matrix
deposition per cell (p<0.001, Figure 4-1A-C). Both proteoglycan and collagen production
per cell were significantly decreased at and below 5 x 106 cells/ml (p<0.001).
Furthermore, seeding at 50 x 106 cells/ml appeared to inhibit matrix production per cell.
The proteoglycan production per cell was not significantly affected at 25 x 106 cells/ml, but collagen production was decreased. The results on a per cell basis revealed a greater matrix deposition at 10-12.5 x 106 cells/ml, but seeding the hydrogels at or above 25 x
106 cells/ml provided the highest total proteoglycan and collagen deposition in the
constructs (p<0.001, Figure 4-1A-C). However, seeding at 50 x 106 cells/ml did not
provide greater total matrix elaboration than seeding at 25 x 106 cells/ml.
Toluidine blue stained sections confirmed differences in proteoglycan content and also indicated differences in distribution (Figure 4-2). At lower seeding densities, matrix deposition was predominantly pericellular, but at higher densities, more interterritorial matrix staining was seen. Furthermore, pericellular staining at lower seeding densities – 5
x 106 cells/ml and lower – appeared less intense than that exhibited at greater seeding
densities (Figure 4-2E-H). In all cases, proteoglycan deposition occurred through the
constructs, after 6 weeks of in vitro culture. Based on these results, an initial seeding
80 Figure 4-1. Comparison of extracellular matrix content at 6 weeks in constructs fabricated at different initial cell seeding densities. DNA (A), proteoglycan (B), and collagen (C) content (mean +/- s.d.) of constructs cultured in defined medium for 6 weeks after initial hydrogel formation with seeding densities of 50, 25, 12,5 10, 5,2.5, and 1.25 x 106 cells/ml. *p<0.001 compared with 50 x 106 cell/ml for the construct. +p<0.001 compared with 50 x 106 cells/ml production/DNA content.
81
Figure 4-2. Proteoglycan distribution of constructs made with different initial cell seeding densities. Toluidine blue staining of constructs seeded at 25 x 106 cell/ml (A,E), 10 x 106 cell/ml (B,F), 5 x 106 cell/ml (C,G), and 2.5 x 106 cells/ml. (A-C, scale bars = 500 μm.D-F, scale bars = 200 μm.)
82 density of 25 x 106 cells/ml was utilized in all subsequent experiments, with culture
duration of 6 weeks.
Pre-treatment with Defined Chondrogenic Medium during Monolayer Expansion
Cells were passaged in low-glucose DMEM containing 10% FBS supplemented with
FGF-2 to maintain pluripotency until confluence was reached. Once confluent, serum- containing medium was removed, and monolayer cells were exposed to defined
chondrogenic medium for 1-6 days before trypsinization and subsequent hydrogel
encapsulation. After 6 weeks of in vitro culture, hydrogels were harvested and the effects on matrix elaboration after subsequent encapsulation were determined. The results
indicated that pre-treatment did not enhance matrix elaboration in the hydrogels (Figure
4-3A-C). Proteoglycan production per cell was actually significantly lower with longer
durations of pretreatment (p<0.002). Collagen and proteoglycan content of the constructs
at 6 weeks of culture was significantly less with 1 day of monolayer pretreatment with
defined chondrogenic medium (p<0.001), and further affected by 3 or 6 days of
pretreatment (Figure 4-3A-C). The DNA content of pretreated constructs was also
significantly lower with pretreatment (p<0.001). Toluidine blue staining of histological
sections indicated a decline in proteoglycan deposition with pretreatment (Figure 4-3D-
F). In constructs fabricated with cells that had longer durations of pretreatment, pericellular
83
Figure 4-3. Extracellular matrix content of hydrogels fabricated with hMSCs pretreated in monolayer culture with defined chondrogenic medium. DNA (A), proteoglycan (B), and collagen content (C) (mean +/- s.d.) of constructs cultured without 0, 1, 3, or 6 days of monolayer pretreatment. *p<0.001 compared with no pretreatment for the construct. +p<0.002 compared with no pretreatment matrix production/DNA content. Toluidine blue staining of constructs cultured with 0 (D), 3 (E), or 6 (F) days of pretreatment. (Scale bars = 250 μm.)
84 deposition of proteoglycan was still seen but less intercellular deposition could be detected. Furthermore, hydrogels constructed with hMSCs pretreated for 6 days displayed limited proteoglycan staining even in the pericellular microenvironment.
TGF-β1 Withdrawal during Hydrogel Culture
A series of experiments was run to determine the relationship between exposure to TGF-
β1 and matrix deposition. TGF-β1 was withdrawn from chondrogenic feeding medium at day 7, 14, 21 or 28. Constructs had higher total DNA and proteoglycan content with
TGF-β1 withdrawal at all time points when compared with controls (p<0.001, Figure 4-
4). However, total collagen content of the constructs was lower with withdrawal at day 7 and day 14 (p<0.001). Proteoglycan content normalized to DNA was unchanged by TGF-
β1 withdrawal. However, collagen content normalized to DNA content was significantly decreased with any TGF-β1 withdrawal when compared with controls exposed to TGF-
β1 throughout the duration of culture (p<0.001). Constructs cultured without TGF-β1 did not undergo chondrogenesis (results not shown).
85
Figure 4-4. Extracellular matrix content of hMSC-seeded scaffolds subjected to TGF-β1 withdrawal from defined chondrogenic medium. DNA (A), proteoglycan (B), and collagen content (C) of constructs cultured with defined medium containing TGF-β1 for 7, 14, 21, 28 or 42 days. *p<0.001 compared with 42 days of exposure for the construct. +p<0.001 compared 42 days matrix production/DNA content.
86 Dexamethasone Withdrawal during Hydrogel Culture
DEX was withdrawn from defined chondrogenic medium at day 7, 14, 21 or 28. Total
DNA and proteoglycan content were unaltered (p=0.053), but collagen content of all constructs exposed to withdrawal was significantly decreased (p<0.001, Figure 4-5).
Collagen content normalized to DNA content of all groups was also lower compared with controls exposed to DEX throughout the duration of culture (p<0.001).
87
Figure 4-5. Extracellular matrix content of hydrogels subjected to DEX withdrawal from defined chondrogenic medium. DNA (A), proteoglycan (B), and collagen content (C) of constructs cultured in defined medium containing DEX for 42, 7, 14, 21, or 28 days. *p<0.001 compared with 42 days of exposure for the construct. +p<0.001 compared with 42 days matrix production/DNA content.
88 DISCUSSION
Scaffolds such as the semi-interpenetrating network used in this study are presently under
intense investigation for tissue engineering applications to repair and/or regenerate
cartilage defects. In addition to exploring the effects of alterations to the physical
parameters of the hydrogel on chondrogenesis, it is important to establish the biological
parameters that affect the differentiation and matrix production of hMSCs in this type of
hydrogel. We therefore examined the effects of cell seeding density, as well as bioactive
factor addition and withdrawal at different stages of the in vitro process of forming
cartilage.
We first established the optimal cell density that maximized extracellular matrix
deposition in the hydrogel. Networks seeded with 25-50 x 106 cells/ml contained the
greatest total proteoglycan and collagen content throughout the constructs after 6 weeks
of culture in chondrogenic medium. Interestingly, 50 x 106 cells/ml provided no
advantage over 25 x 106 cells/ml for either proteoglycan or collagen production.
Although cell-cell contact is advantageous for chondrogenic differentiation in scaffold-
free systems [22, 25], and is a prerequisite in vivo during limb development, it may have
to be considered differently in hydrogel scaffolds. Some of the signals for differentiation
that occur with high cell density, such as cell shape changes and cell-cell communicated
signals, may be replaced by the meshwork of the constraining hydrogel and the externally
added bioactive factors, respectively. Furthermore, the chemical linkages of the hydrogel
formed during phase transition/cellular encapsulation, may restrict the physical separation of cells that occurs with matrix production and deposition in vivo and in scaffold-free
89 systems. Thus, the cellular feedback may be different at the highest cell density explored
in this series (50 x 106 cells/ml), to the extent that matrix production is decreased by the
cell in response to sensing a higher surrounding matrix density.
Initial seeding densities of 10-25 x 106 cells/ml provided the highest matrix deposition
per cell, indicating that there is also a point at which the cell density is too low to provide
cell to cell paracrine communications that influences matrix production. This
phenomenon has been observed by others working with hydrogels: Kavalkovich et al.
found that differentiation in an alginate layer system was dependant on initial hMSC
seeding density, and that a seeding density of 25 x 106 cells/ml resulted in optimal
glycosaminoglycan synthesis per cell [122]. With cell densities lower than 25 x 106 cells/ml, matrix production per cell was reduced, with the lowest production per cell at the lowest cell density, as was found in our studies.
TGF-β plays an important role during chondrogenesis [62, 63], and we and others have previously established that in vitro chondrogenesis of bone marrow-derived MSCs in 3-D cell cultures can be achieved through TGF-β induction [22, 25, 64-66, 155, 156]. In tissue engineering experiments, several groups have demonstrated that exposure to TGF-
β1 increases matrix production of cultured chondrocytes encapsulated in 3-dimensional networks [19, 37-41, 44, 53, 54, 59, 98] and induces chondrogenesis in MSCs [17, 121-
123, 156-158]. However, in all these studies, TGF-β was added throughout the culture period. In contrast, O’Driscoll and co-workers have examined the effects of limiting chondroprogenitor cell exposure to TGF-β1 [72, 73]. They reported that enhancement of periosteal chondrogenesis was very similar with either 2 days or 14 days of TGF-β1
90 treatment. We tested the effects of altered temporal exposure to TGF-β1 on MSCs in the
hydrogel and found that withdrawal of TGF-β1 enhanced total proteoglycan content
measured at 6 weeks, regardless of whether withdrawal was done at 1, 2, 3 or 4 weeks.
However, DNA content was also significantly higher after withdrawal such that the
proteoglycan production per cell was not significantly different between withdrawal and
control conditions for any timepoint. Thus, one effect of TGF-β1 withdrawal appears to
be either greater cell proliferation and/or cell survival. However, withdrawal of TGF-β1
at day 7 or 14 reduced the total collagen content compared with sustained treatment of
TGF-β1 for all 42 days. Withdrawal at later times did not affect total collagen content;
however, for all timepoints, withdrawal significantly decreased the collagen production
per cell measured at 6 weeks. This is an important point for tissue engineering strategies
because it has been noted that collagen production in many MSC and chondrocyte
systems used to-date is inferior to that of native cartilage [61-63]. The collagen network
provides for the tensile strength of cartilage. A successful tissue engineered construct may require sufficient collagen content in order to mechanically function as well as the native tissue.
Alhadlaq et al. have experimented with exposure of TGF-β prior to seeding MSCs in scaffolds and subsequent defined chondrogenic conditions [149]. Scaffolds containing rat marrow-derived MSCs treated with TGF-β1 as a supplement to DMEM containing
FBS for 7 days prior to encapsulation in a PEG-based polymer had enhanced proteoglycan and collagen content. In the present study, cells were pretreated in defined chondrogenic medium for 1-6 days during monolayer culture just prior to encapsulation
91 in hydrogels. Pretreatment did not enhance matrix deposition of constructs: extracellular
matrix production was significantly decreased with 1 day of exposure, and further
inhibited with increased duration. The difference in results between the present study and
those of Alhadlaq et al. may be due to species-specific responses, but may also be due to
the use of serum-containing medium by Alhadlaq et al. [27-30]. Serum contains multiple
factors that are absent in the defined medium, and cells in culture are susceptible to
differences in serum production and quality. For example, re-differentiation of human-
expanded chondrocytes cultured in vitro was found to be highly dependent of the lot of
serum used [159], as is the proliferation and retention of differentiation capacity of
MSCs. The particular lot(s) of serum used by Alhadlaq et al. may have contained factors
appropriate for pre-differentiation of rat MSCs in 2-dimensional culture. It is possible
that the constituents of the defined medium that stimulate chondrogenesis in 3-
dimensional cultures lead to different responses of cells in 2-dimensional culture, altering
the cell phenotype such that they cannot be stimulated to undergo chondrogenesis upon
encapsulation in a 3-dimensional hydrogel.
The glucocorticosteroid (GC) dexamethasone (DEX) has been shown to induce cellular
differentiation [68-70, 137-142]. Exposure to DEX, in addition to other factors, can lead
pluripotent mesenchymal cells or cell lines to differentiate into various mesenchymal cell
phenotypes [22-25, 69]. Grigoriadis et al. [69] first reported the use of DEX to induce cartilage nodule formation, using the RCJ 3.1 cell line derived from fetal rat calvaria.
Previous work in our laboratory has examined the effects of DEX and TGF-β1 on
chondrogenesis in both rabbit and hMSC aggregate cultures [22, 25]. Although some
92 preparations of MSCs can be induced to undergo chondrogenesis with DEX but without the inclusion of TGF-β1, the addition of TGF-β1 promotes chondrogenesis in most preparations and stimulates a much greater extracellular matrix production. Since these initial findings, many groups have studied the combined effects of exposure to DEX and
TGF-β on chondrocytes, and on MSCs derived from bone marrow, adipose, and periosteum [22, 25, 64, 65, 72, 73, 75, 143-147]. Recently, Derfoul et al. [144] suggested that GC effects during in vitro chondrogenesis were mediated through the major active form of GC receptor, GRα, but the precise mechanism by which DEX promotes chondrogenesis is not known.
Although the effects of GCs on chondrogenesis have been examined, the duration of exposure necessary to affect differentiation and matrix development has not been determined. DEX withdrawal did not affect DNA or proteoglycan content in our system; however, it significantly decreased total collagen content and production per cell for all withdrawal timepoints analyzed. Previous studies have not included DEX withdrawal, rather they have studied the effects of addition of DEX. Grigoriadis et al. [68], reported that DEX treatment induced cartilage nodule formation of RCJ 3.1 cells. Furthermore, the effects of DEX exposure were not dependent on when addition occurred: exposure upon initial plating or at confluence produced equivalent results in their system. In contrast,
Fugita et al. [160] determined that the presence of DEX inhibited insulin-induced cellular condensation and subsequent cartilaginous nodule formation in the mouse ATDC5 cell line. Due to the differences between cell types utilized and culture conditions, it is
93 difficult to determine why the outcomes of DEX exposure are not equivalent in these
various systems.
We have developed a photopolymerizable semi-interpenetrating network that enables the
chondrogenic differentiation of hMSCs in vitro. Proteoglycan and collagen content
throughout the construct were dependent on initial seeding density, with 25 x 106 cells/ml resulting in maximal extracellular matrix elaboration. Pretreatment of monolayer cells with defined chondrogenic medium retarded matrix elaboration after encapsulation and subsequent in vitro culture. Limiting exposure to bioactive factors affected extracellular
matrix production of collagen. These findings are informative for in vitro tissue
engineering of cartilage, but may also aid in vivo implantation strategies for hydrogels that contain undifferentiated MSCs.
ACKNOWLEDGEMENTS
The authors would like to thank Teresa Pizzuto for her assistance in histology preparations, Dr. Jennifer West for supplying PEGDA and Dr Jamie Fitzgerald for his review of the manuscript. Studies in this manuscript were supported in part by NIH RO1
AR048132 (BJ).
94 CHAPTER FIVE: CONCLUSIONS & FUTURE WORK
The experiments outlined in this body of work describe the design and characterization of a photopolymerizable PEGDA:PEG semi-interpenetrating network that facilitates in vitro chondrogenesis of human mesenchymal stem cells. Prior to the onset of this research, no group had successfully demonstrated in vitro chondrogenesis of MSCs in a hydrogel.
Then in 2002, Kavalkovich et al. reported chondrogenesis of hMSCs in an alginate layer system. Williams et al. followed with reports of goat MSCs in a photopolymerizable
PEG-based scaffold in 2003. Although these reports, and those subsequent, have documented chondrogenic markers (mRNA expression of collagen type II and proteoglycan deposition), indicating feasibility, none have displayed substantial extracellular matrix elaboration within hydrogels. What has been missing from these studies is a careful examination of the network and biological parameters that affect chondrogenesis and extracellular matrix production.
In this thesis, multiple parameters of PEGDA:PEG semi-interpenetrating networks were explored for their effects on chondrogenesis and extracellular matrix production.
Alterations to the network were quantified by the mesh size, ξ, of the network – approximated by volumetric swelling ratios, Q, and the Peppas-Merrill model of calculating the average molecular weight between crosslinks, MC. Although this model is applicable for systems which are high swelling and have solvent present during monomer crosslinking, it does significantly underestimate the mesh size. Any hydrogel system will possess a distribution of mesh sizes. The ξ calculated by volumetric swelling ratios may be considered as the minimum pore size of the scaffold, keeping in mind that the actual
95 pore size can range to at least 1 order of magnitude greater. We come to this conclusion by considering a second minimum mesh size, which may be approximated by the end-to- end bond distances by the C-C and C-O bonds in the PEGDA polymer backbone.
Neglecting the presence of the PEG component, and assuming perfect bond formation and 100% completion in the reaction, the mesh size for the PEGDA (6 kDa) hydrogel would be 60 nm. Failure of the system to achieve 100% polymerization, dangling ends, and side reactions – all inherent in free radical polymerization reactions will increase the minimum mesh size for both scenarios, as will the presence of the PEG component and the cells.
With that in consideration, we may still employ the minimum mesh size calculated by volumetric swelling ratios to provide a measurable change in the pore diameter with alterations to the network. As such, changes in ξ on the order of 1 nm resulted in an expansion of the area in which proteoglycan was deposited throughout the construct.
Specifically, proteoglycan deposition shifted from pericellular to intercellular. As previously mentioned, the minimum mesh size (without PEG spacer) for PEGDA (6 kDa) constructs approximated by C-C and C-O bond distances is on the order of 60 nm – assuming perfect bond formation and 100% polymerization. Practical constraints lead us to believe that the presence of the PEG spacer, in particular at the decreased
PEGDA:PEG ratio, allows for a minimum mesh size in the range of 80-100 nm. A distribution of mesh sizes on this order would certainly facilitate deposition of aggrecan, which also possesses a solvated and chain-extended diameter in this range. This premise is supported by the fact that PEGDA (6 kDa) hydrogels were quantifiably impacted by
96 decreasing the ratio of PEGDA:PEG from 2:1 to 1:2. Constructs cultured at this decreased ratio exhibited statistically greater proteoglycan contents. However, hydrogels constructed with higher PEGDA molecular weights (12 and 20 kDa) at this same ratio did not maintain the increase in proteoglycan. This may be due to a dramatically increased mesh size. Bond length approximations provide 120 nm and 200 nm as the mesh size in these networks, respectively; effects of the addition of PEG and polymerization reaction constraints notwithstanding. Mesh size approximated by volumetric swelling ratios provides for a measurable ξ ≥ 10 nm in these constructs. Although they did not contain increased proteoglycan contents at either ratio, networks constructed with PEGDA (12
and 20 kDa) and PEG (88 kDa) exhibited interterritorial proteoglycan deposition
regardless of the PEGDA:PEG ratio.
Collagen production, however, did not follow a uniform trend. At the lowest PEGDA
molecular weight (6 kDa), collagen content was unaffected by decreasing the
PEGDA:PEG ratio. Networks constructed with higher PEGDA molecular weights (12 and 20 kDa) examined in this series exhibited a significant decrease in collagen content at
PEGDA:PEG ratio of 1:2, as compared with constructs fabricated with the same molecular weight PEGDA at a PEGDA:PEG ratio of 2:1. This decrease was also significant when compared with PEGDA:PEG of 1:2 using other PEGDAs (6 and 12 kDa). However, hydrogels constructed with PEGDA (12 kDa) exhibited a significant increase in collagen content at PEGDA:PEG of 2:1, compared to PEGDA (6 and 20 kDa) at the same ratio, and for all PEGDAs at 1:2.
97 Based on the results of this study, we would suggest maintaining the Peppas-Merrill
model of approximating ξ. Furthermore, it would be advantageous to determine whether
the non-binding secondary PEG polymer component is the largest contributor to the
significant changes in the profile proteoglycan deposition. In order to determine
conclusively that the high molecular weight PEG spacer is responsible for the increase in
proteoglycan and collagen, PEGDA constructs should be made with equivalent
volumetric swelling ratios and corresponding mesh sizes to those explored in this thesis
for all PEGDA molecular weights (6, 12, and 20 kDa). MSC-seeded constructs lacking
PEG should be cultured and examined for their effects on DNA, proteoglycan, and
collagen content. Chondrocytic differentiation of hMSCs and subsequent matrix
elaboration in these monomeric hydrogels should be compared with that seen in the
PEGDA:PEG semi-interpenetrating networks explored in this study. If hydrogels
constructed without PEG exhibit the same trend in matrix production, it is unlikely that a
semi-interpenetrating network is needed to facilitate chondrogenic differentiation and
matrix elaboration. However, particular attention should be paid to collagen content in
the resulting hydrogels. Only hMSC-seeded PEGDA (12 kDa) constructs fabricated with
PEG (88 kDa) at 2:1 exhibited an increase in collagen production in these studies.
Perhaps the minimum mesh size of ~10 nm, approximated by Q, in these constructs is optimal for collagen synthesis and/or deposition? To fully examine the collagen content, immunohistochemistry for collagen types I, II, and X should be performed to determine whether these monomeric constructs exhibit differences in their collagen profiles.
Quantitative data may also be obtained for the collagen contents. mRNA can be
quantified by real-time RT-PCR assays and quantitative ELISAs for collagen types I and
98 II are now commercially available. The ratios of different collagen types may be altered
by the mesh changes, even though total collagen content may not.
Biological parameters were also explored for their effects on differentiation and matrix
development. The initial seeding density was optimized at 25 x 106 for maximum
proteoglycan and collagen deposition throughout the construct. However, much lower
seeding densities (10-12.5 x 106 cells/ml) led to the highest collagen and proteoglycan production on a per-cell basis. Furthermore, constructs fabricated at the highest seeding
density explored in this study (50 x 106 cells/ml) exhibited significantly lower
proteoglycan and collagen production per cell. It is unclear, at this time, whether
constructs should be optimized on a per-cell or per-construct basis.
Human MSCs were also subjected to pretreatment with defined chondrogenic medium
during monolayer expansion, prior to encapsulation. Pretreatment, or any duration, with defined chondrogenic medium inhibited both proteoglycan and collagen production in constructs after 6 weeks of in vitro culture. It is possible that pre-treatment in monolayer
(2-dimensional) culture initiated differentiation along a different pathway, resulting in
cells that exhibited a fibroblastic phenotype. Perhaps, it is not possible to reverse these
effects, even upon transfer to a 3-dimensional system. Based on our studies, we conclude
that the in vitro, chondrogenic system we have developed requires a 3-dimensional
environment to function properly. Future experiments should include the pretreatment of
monolayer hMSCs prior to encapsulation with serum-containing low-glucose DMEM
supplemented with TGF-β1 for 1-7 days as done by Alhadlaq et al. [148-151, 161] and
99 the effects on subsequent matrix elaboration should be examined. This would provide a better comparison between the effects of pretreatment with the defined and serum- containing media.
Human MSC-seeded constructs were also subjected to temporal exposure (or withdrawal) of chondrogenic factors during the 6 week culture period. Additional TGF-β1 withdrawal studies should be run, with removal starting at earlier timepoints –as early as 24 hours after the onset of exposure, to determine the level of exposure necessary to replicate and/or exceed proteoglycan production with 6 weeks of culture. Proteoglycan content increased in cell-seeded constructs subjected to TGF-β1 withdrawal at various times of culture, but DNA content also increased such that proteoglycan production per cell was unaffected. More interesting was the fact that collagen production per cell was reduced with TGF-β1 withdrawal at any timepoint, and total collagen content was significantly lowered if withdrawal was done within the first two weeks of culture. In addition to re- examining the effects on proteoglycan production, the specific effects on collagen subpopulations should be determined. Quantitative assays for DNA, proteoglycan, and total collagen should be performed, as well as quantitative ELISAs and quantitative RT-
PCR for collagen types I, II, and X, and visualization by immunohistochemistry of cultured constructs. We feel it would be advantageous to focus on the collagen profile, because of the results of preliminary work with immunohistochemistry for collagen types
I, II, and X, performed on initial constructs subjected to TGF-β1 withdrawal. These early findings indicated a decrease in the production of collagen type X (Figure 5-1). Much work has yet to be completed that elucidates the exact timing for significant
100 chondrogenic events induced by TGF-β1, although it is known that expression is strictly
regulated. It’s difficult to say why a withdrawal in TGF-β1 would signal for a delay in
the onset of hypertrophy, evidenced by a reduction in the production of collagen type X;
however, a delay in the onset of hypertrophy could be a significant advantage to a
cartilaginous tissue engineered construct. In order to verify these findings, mRNA from
cultured constructs should be harvested, and examined for collagen type II and X
messages to determine the actual decrease (or elimination) of the mRNA for collagen
type X.
Preliminary findings on the effects of dexamethasone withdrawal did not indicate an
alteration in the production of collagen types II or X (Figure 5-1). This was intriguing
since withdrawal of DEX did not affect DNA or proteoglycan content, but was even more
detrimental to collagen production: withdrawal at any timepoint significantly lowered collagen production within the cell-seeded constructs. We expected more significant changes in matrix production, given that hMSCs in pellet culture require DEX to undergo chondrogenic differentiation. We ran a series of preliminary experiments on constructs cultured for 6 weeks in defined chondrogenic medium lacking DEX from the onset of culture. Immunohistochemistry for collagen type I, II, and X was performed on constructs subjected to DEX withdrawal. (Figure 5-2)
101
Figure 5-1. Immunohistochemistry for collagen types II and X on hMSC-seeded PEGDA:PEG networks cultured in defined chondrogenic medium that was subjected to removal of dexamethasone (DEX) or TGF-β1.
102
Figure 5-2. Immunohistochemistry for collagen types II and X of hMSC-seeded constructs cultured for 6 weeks in defined chondrogenic medium lacking (top) or containing (bottom) DEX.
103 Collagen type I was not produced in any constructs (cultured with or without DEX).
However, those cultured without DEX from the onset exhibited production of both
collagen types II and X. It is difficult to deduce whether collagen type X production is
augmented in constructs cultured without DEX from the onset of culture. The experiment
should be repeated, and should include quantitative ELISAs and quantitative RT-PCR for
collagen types I, II, and X to determine whether certain subpopulations are affected by
removal of DEX. If dexamethasone is not necessary to induce chondrogenesis in 3-
dimensional culture, it would be advantageous to eliminate from the defined
chondrogenic medium employed throughout these studies, particularly since withdrawal
of dexamethasone does seem (comparatively, qualitatively) to increase the intensity and
extent of collagen type X staining.
Can Further Alterations Improve Matrix Elaboration?
The concentration of proteoglycan in articular cartilage ranges from 30 – 50 mg/ml.
However, most constructs contain one tenth of this value, at most. Because proteoglycans
confer the tissue’s resistance to deformation, the properties of a tissue-engineered
construct must more closely mimic those of the native tissue. The inclusion of degradable
linkages into the network might increase proteoglycan deposition [37-41, 54, 59].
Degradable systems may take many forms. Some employ simple hydrolytic co-polymer
systems, while others incorporate cell-specific degradation sequences that are
hydrolytically cleaved upon encapsulated cellular activity. Mann et al. successfully
employed the latter technique with rat smooth muscle cells in a PEGDA hydrogel with
collagen type I collagenase-sensitive degradable linkages. The enzyme profile of our
104 differentiating MSCs is not known at this time. Future work may include determination
of MMPs, aggrecanases, and other enzymes produced by these cells as they differentiate.
Knowledge of this profile would be useful in tailoring a PEGDA network for degradation
in a cartilage-based system. A degradable system of this nature is advantageous because
it would facilitate removal of the hydrogel scaffold. Furthermore, it might also enhance
integration of the tissue engineered construct to the surrounding host cartilage. The
challenge with such degradable scaffolds is to design them such that the rate of
degradation matches the rate of matrix elaboration sufficiently to allow generation of
cartilage without too great a loss of matrix and cells.
It is also possible that other modifications to the gel structure could be advantageous. We
mentioned previously that the inclusion of sequences from bioactive factors, like the
RGD tripeptide from fibronectin, could potentially help facilitate cellular adhesion and
affect retention within the scaffold. However, some studies have shown that increased
cell adhesion can result in decreased extracellular matrix protein production by the cells
[96, 162]. Research in our own laboratory supports these findings. Quantitative data did
not reveal any changes in proteoglycan or collagen content with the addition of RGD to
the PEGDA polymer scaffold (Figure 5-3). It should be noted that two different methods
for tethering RGD to the polymer scaffold were employed in these studies. The first employed PEGDA (6 kDa) and an acrylated RGD tripeptide, which forms covalent linkages to the polymer backbone through a competing reaction for crosslinking sites
with PEGDA. The second method used PEGDA (6.8 kDa) constructed with RGD
105 Figure 5-3. Extracellular matrix content of PEGDA scaffolds fabricated with or without tethered RGD. covalently bonded in the middle of two PEG (3.4 kDa) repeat units, flanked by acrylate
end groups. A polymer of this nature leaves the two end-groups free to crosslink during
106 polymerization without competition from the tripeptide. In both cases, the addition of
RGD to the PEGDA backbone produced the same result. Toluidine blue stained sections from the RGD group were not discernibly different from those cultured without RGD
(Figure 5-4).
The inclusion of growth factors, including TGF-β, could prove useful in providing short- term exposure to cell-seeded constructs in vivo to initiate cellular differentiation.
Signaling factors, such as BMP-2, IGF, and PTHrP have been shown to enhance in vitro chondrogenesis [17, 68, 98, 145, 159, 163-166]. These factors could be added singularly or in combination to medium to determine the effects on subsequent extracellular matrix production of cell-seeded construct in vitro. Factors that enhance proteoglycan or collagen production could be further explored for their temporal affects (enhancing collagen production, integration, etc). The results may indicate improvements that could be made to media used for in vitro tissue engineering, but may also suggest factors that might be incorporated into cell-gel constructs that are set up in vivo in cartilage defects.
Exploration of gels modified by the inclusion of growth factors, perhaps in slow release vehicles, would be an interesting future objective.
The effects of mechanical loading on the metabolic activities of chondrocytes have been extensively studied [35, 87, 167-190]. Cellular loading in the form of hydrostatic, confined and/or unconfined compressive loading has been reported to increase
107
Figure 5-4. Toluidine blue stained histological sections of MSC-seeded constructs encapsulated in PEGDA polymer scaffolds without (A) and with (B) tethered RGD.
108 biosynthesis of macromolecules and subsequent matrix deposition of mature chondrocytes and bone-marrow derived MSCs in vitro [105, 170, 172, 187, 191-197].
Recently, Huang et al. reported that the dynamic compressive loading profile significantly impacts the resulting matrix deposition of mesenchymal progenitor cells
[191]. Further, they report that dynamic cyclic compressive loading is sufficient to induce chondrogenic differentiation of rabbit MSCs in vitro in non-permissive conditions
(defined medium lacking TGF-β1). Combinations of mechanical stimulation, and withdrawal or addition of growth factors might also substantially improve matrix elaboration. This is particularly important for collagen production because although there are many growth factors known to enhance proteoglycan production, enhancement of collagen production is more difficult. Given the extremely low levels of collagen production seen in hydrogels produced to-date, mechanical stimulation may be a very important addition to the in vitro tissue engineering of cartilage. It is for this reason that many groups are currently exploring bioreactors that incorporate mechanical loading [99-
101, 115-117, 177, 198-201].
Given the fact that physiologic loading is cyclic in nature – walking and running in particular occur with inherent periodicity – cyclic (not static) loading should be explored.
The nature and anatomy of cartilage provides some latitude in choosing whether this loading should take confined or unconfined form. In separate studies, the Johnstone laboratory has developed an unconfined system that may subject MSC-seeded constructs to cyclic physiologic loads. Cell-seeded constructs could be exposed to cyclic, unconfined compressive loading of various profiles (sinusoidal, square, ramp, etc. with
109 varying periodicity) for durations of 1 – 6 weeks to examine whether the properties change over the time course of loading, and whether mechanical loading enhances the chondrogenic differentiation and/or increases the rate of subsequent matrix production of
MSCs in vitro. In addition to the application of mechanical forces, mechanical studies should be also be performed to examine the initial properties of the unloaded semi- interpenetrating network, in addition to determining if (and how) the properties change over the duration of culture – both through the development of extracellular matrix and loss of the non-binding physical component (PEG). The biomechanical properties of the in vitro engineered cartilage constructs will need to be known and may guide the choice of conditions used for culturing hydrogels for implantation. Unfortunately, what is not known is what level of mechanical properties needs to be achieved prior to implantation.
If the biomechanical properties of the host tissue are sought and gained, these may not be optimal for integration since the tissue produced may behave like articular cartilage implants, which integrate very poorly [16].
What Type of Animal Model Should Be Explored?
Because these studies have clinical relevance to cartilage repair, construct biocompatibility and integration should be examined in an animal model. Smaller animal models, such as mouse or rabbit, do not provide as much insight to repair in human systems – predominantly due to the anatomy/size of the knee joint and differences in joint movement. It is more advantageous, then, to pursue an animal model of large-defect cartilage repair, such as goat or sheep, with quantitative biomechanical and biochemical analyses. Because the gel system utilized here is photopolymerizable, it can be used to
110 fabricate constructs ex vivo or in situ. This flexibility in polymerization allows the
constructs to be formed at the exact location of damage and take on any desired shape.
Constructs fabricated and cultured for 6 weeks in vitro (prior to in vivo implantation)
could be compared to cell-seeded constructs polymerized in situ. A delivery system for the required low-intensity, longwave UV light will have to be designed and constructed to fabricate constructs in situ. Furthermore, a sterile implantation protocol suitable for use in an operating room will have to be designed. A comparative study of in situ versus ex vivo formed cell-hydrogel implants could be valuable, with the time course of healing and resulting integration with the surrounding cartilage judged by systematic analyses of the morphologic, molecular, biochemical and biomechanical properties.
Summary
The semi-interpenetrating network that was developed and tested in this thesis work forms a strong platform from which many different aspects of scaffold design and MSC
differentiation can be explored. It is hoped that this will ultimately lead to the
development of clinically applicable implants for use in the repair and regeneration of
skeletal tissues, particularly, but not limited to, articular cartilage.
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