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Substrate stiffness influences osteogenic differentiation of induced pluripotent stem cell-derived mesenchymal stem cells

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Substrate stiffness influences osteogenic differentiation of induced pluripotent stem cell-derived mesenchymal stem cells
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Treadwell, Karl Alexander ( author )
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English
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Stem cells -- Biotechnology -- Cytology ( lcsh )
Multipotent stem cells ( lcsh )
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bibliography ( marcgt )
theses ( marcgt )
non-fiction ( marcgt )

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Stem cells are able to differentiate into a variety of tissue-specific cells, demonstrating a unique promise for tissue engineering and orthopaedic regenerative therapies. Routine culture techniques utilize soluble factors that chemically influence differentiation, though the mechanical properties of the surrounding environment are proving to significantly impact cell differentiation as well. Mesenchymal stem cells (MSCs), a widely studied cell source for bone regeneration, have repeatedly demonstrated these mechanotransductive properties in vitro, favoring relatively rigid substrates under osteogenic differentiation conditions. The effect of substrate stiffness on osteogenic differentiation of mesenchymal progenitors derived from induced pluripotent stem cells (iPSC-MPs), a novel cell source for bone regeneration, has not been widely studied. By culturing iPSC-MPs on Polydimethylsiloxane (PDMS) substrates with varying stiffness, we tested the hypothesis that comparatively rigid substrates could promote more efficient osteogenic differentiation than softer substrates. The stiffness of the substrates, characterized by Young’s modulus, ranged from 0.5 to 2.5 MPa. Cells cultured on these substrates were run against controls grown on tissue culture plastic (TCP). Effects were observed for cultures grown in both complete culture media (CCM) and osteogenic differentiation media (ODM). Gene expression at 3, 7, 14 and 21 days, quantified by RT-qPCR, shows significant upregulation of osteogenic markers, including runt-related transcription factor 2 (RUNX2), alkaline phosphatase (ALP), and osteocalcin (OCN) for cells cultured on 2.245 MPa substrates in ODM. In comparison to cells cultured on softer substrates and especially TCP, these cells also displayed an increase in ALP protein activity and calcium deposition as determined by alizarin red s staining and quantification. Cells cultured in CCM showed trends for osteogenic differentiation, suggesting that mechanical properties of the substrate may affect cell fate without chemical differentiation signaling. With recent studies showing the efficacy of iPSCs as a novel stem cell source for regenerative medicine, it is becoming ever more necessary to establish a fast and efficient method for differentiating these cells for orthopaedic applications. This study shows how substrate stiffness can be used to promote faster, more efficient osteogenic differentiation in iPSC-MPs. This information could be used to design scaffolds that would promote bone formation in vivo.
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Thesis (M.S.) - University of Colorado Denver
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Includes bibliographic references.
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Department of Bioengineering
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by Karl Alexander Treadwell.

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Full Text
SUBSTRATE STIFFNESS INFLUENCES OSTEOGENIC DIFFERENTIATION OF
INDUCED PLURIPOTENT STEM CELL-DERIVED MESENCHYMAL STEM
CELLS
by
KARL ALEXANDER TREADWELL
B.S., University of Arizona, 2011
A thesis submitted to the
Faculty of the Graduate School of the
University of Colorado in partial fulfillment
Of the requirements for the degree of
Master of Science
Bioengineering
2015
1


This thesis for the Master of Science degree by
Karl Alexander Treadwell
has been approved for the
Bioengineering Program
by
Dae Won Park, Chair
Karin Payne, Advisor
Michael Yeager
November 19, 2015
11


Treadwell, Karl Alexander (M.S., Bioengineering)
Substrate Stiffness Influences Osteogenic Differentiation of Induced Pluripotent Stem
Cell-Derived Mesenchymal Stem Cells
Thesis directed by Assistant Professor Karin Payne.
ABSTRACT
Stem cells are able to differentiate into a variety of tissue-specific cells,
demonstrating a unique promise for tissue engineering and orthopaedic regenerative
therapies. Routine culture techniques utilize soluble factors that chemically influence
differentiation, though the mechanical properties of the surrounding environment are
proving to significantly impact cell differentiation as well. Mesenchymal stem cells
(MSCs), a widely studied cell source for bone regeneration, have repeatedly
demonstrated these mechanotransductive properties in vitro, favoring relatively rigid
substrates under osteogenic differentiation conditions. The effect of substrate stiffness on
osteogenic differentiation of mesenchymal progenitors derived from induced pluripotent
stem cells (iPSC-MPs), a novel cell source for bone regeneration, has not been widely
studied. By culturing iPSC-MPs on Polydimethylsiloxane (PDMS) substrates with
varying stiffness, we tested the hypothesis that comparatively rigid substrates could
promote more efficient osteogenic differentiation than softer substrates. The stiffness of
the substrates, characterized by Youngs modulus, ranged from 0.5 to 2.5 MPa. Cells
cultured on these substrates were run against controls grown on tissue culture plastic
(TCP). Effects were observed for cultures grown in both complete culture media (CCM)
and osteogenic differentiation media (ODM). Gene expression at 3, 7, 14 and 21 days,
quantified by RT-qPCR, shows significant upregulation of osteogenic markers, including
runt-related transcription factor 2 (RUNX2), alkaline phosphatase (.ALP), and osteocalcin


('OCN) for cells cultured on 2.245 MPa substrates in ODM. In comparison to cells
cultured on softer substrates and especially TCP, these cells also displayed an increase in
ALP protein activity and calcium deposition as determined by alizarin red s staining and
quantification. Cells cultured in CCM showed trends for osteogenic differentiation,
suggesting that mechanical properties of the substrate may affect cell fate without
chemical differentiation signaling. With recent studies showing the efficacy of iPSCs as
a novel stem cell source for regenerative medicine, it is becoming ever more necessary to
establish a fast and efficient method for differentiating these cells for orthopaedic
applications. This study shows how substrate stiffness can be used to promote faster,
more efficient osteogenic differentiation in iPSC-MPs. This information could be used to
design scaffolds that would promote bone formation in vivo.
The form and content of this abstract are approved. I recommend its publication.
Approved: Karin Payne
IV


TABLE OF CONTENTS
CHAPTER
I. INTRODUCTION AND GOALS...............................................01
Stem Cells..........................................................02
Bone................................................................07
Mechanosensing and Mechanotransduction..............................09
The Cellular Environment............................................13
Goals and Hypothesis................................................14
II. MATERIALS AM) METHODS................................................15
Cell Culture........................................................15
Substrate Material Synthesis........................................16
Material Properties.................................................17
Cell Plating and Culture............................................18
Cell Proliferation..................................................18
Actin Staining......................................................19
RT-qPCR.............................................................20
Pico Green & ALP Activity...........................................22
Alizarin Red S......................................................24
Statistical Analysis................................................26
III. RESULTS..............................................................27
Substrate Stiffness.................................................27
Cell Spreading and Proliferation....................................28
Gene Expression.....................................................31
Alkaline Phosphatase Activity.......................................36
Calcium Deposition..................................................38
IV. DISCUSSION...........................................................41
Cell Proliferation..................................................41
Gene Expression.....................................................42
ALP Activity........................................................45
Calcium Deposition..................................................46
Mechanical Influence in CCM Cultures................................48
Future Directions...................................................48
Implications for Scaffold Seeding and Bioreactors...................50
V. CONCLUSIONS..........................................................51
WORKS CITED................................................................52
v


CHAPTERI
INTRODUCTION AND GOALS
According to the United States Bone and Joint Initiative (USBJI), there are
approximately 223.6 million cases each year in the United States that involve diseases,
disorders, and injuries related to bones, joints, and muscles [1], This results in an
estimated annual medical care cost of $212.7 billion [1], Bone injuries are a common
occurrence in all age groups, but especially in individuals over the age of 50, where 1 in 2
women and 1 in 4 men will have an osteoporosis-related fracture [1], With an aging
population and prolonged life expectancy, the occurrence of bone injuries is only
expected to increase. Despite advances in orthopaedic medicine, some fractures remain
difficult to treat or do not heal well, with 5-10% resulting in non-unions and severe
functional impairment [2], Stimulation of bone formation is of major clinical
significance in orthopaedic procedures related to non-unions and can also be effective in
cases of spinal fusion, joint fusion and for the repair of segmental bone defects. Implants
and bone allografts have been widely used in orthopaedics to bridge the gap between the
injured bones, but they lack a biological component that will allow successful integration
into the surrounding native tissue. Thus, regenerative medicine techniques to promote
bone stimulation have gained significant interest.
Regenerative medicine focuses on developing therapies that will repair, replace,
or promote the regeneration of damaged or diseased tissue. One key component of
regenerative medicine is the use of stem cells, which have the potential to become any of
the tissue-specific cells in the body. Some of the current applications of stem cells
include therapeutic treatments of cancer, neurodegeneration, third degree bums, and loss
of vision due to chemical destruction of the cornea [3], The regenerative capabilities of
1


stem cells have also shown promise for repairing cartilage, treating spinal cord injuries,
and reversing damage caused by critical limb ischemia [4], However, there are many
different kinds of stem cells that can be obtained from different sources and can vary in
their regenerative ability.
Stem Cells
Stem cells are the foundation for every organ and tissue in the body. They are
characterized by the ability to self-renew, creating identical daughter cells, and to
differentiate into specific tissues or organs. Though there are many different kinds of
stem cells, they are classified based on their differentiation potential as either totipotent,
pluripotent, or multipotent.
Totipotent Stem Cells
At the outset of embryonic development, two gametes (i.e. ovum and sperm)
combine to form a zygote which contains all of the genetic information necessary for the
embryo to develop. These are the first stem cells of the new organism and they are
totipotent, meaning they have the potential to form any cell in the developing embryo, as
well as the placenta. Only these early stem cells are totipotent, with truly universal
differentiation potential. As the embryo develops, other kinds of more specialized stem
cells form, which are defined by their commitment to given cell lineages.
Pluripotent Stem Cells
Pluripotent stem cells differ from totipotent cells only in that they cannot form
placenta. They retain the ability to form any one of the three germ layers, differentiating
into any specialized cell of a specific tissue or organ. They also maintain the capacity to
proliferate indefinitely in a series of self-renewals. Embryonic stem cells (ESCs),
2


embryonic germ cells (EGCs), and induced pluripotent stem cells (iPSCs) all exhibit
behaviors of pluripotency [3], ESCs were first derived from the inner cell mass of a
mouse blastocyst in 1981, and subsequently from a human blastocyst in 1998 [5-7],
Formed in the early development of mammals, the inner cell mass of the blastocyst is the
structure that subsequently forms an embryo, which is the inherent source of controversy
for ESCs, as the harvested blastocyst has the potential to grow into a full human being
[3], The embryonic germ cell serves as the progenitor of adult gametes and are found
during late embryonic to early fetal development [8], IPSCs are special in that they are
not derived from embryonic development and will be discussed in more detail on page 5.
One potential consequence with the use of pluripotent stem cells is the risk of
teratoma formation. A teratoma is a nonmalignant tumor composed of cells derived from
all three embryonic germ layers [9], With the inherent potential to form this disorganized
mixture of tissue, there is a strong motivation for understanding the factors that control
stem cell differentiation. Utilizing monoclonal antibodies, small molecules, anti-
angiogenic agents, suicide genes, and pharmacological agents, significant progress has
been made towards inhibiting uncontrolled pluripotency [10-12], Further work that
develops protocols for the safe and effective differentiation of these cells will greatly
facilitate the advancement of stem cell-based therapies.
Multipotent Stem Cells
Multipotent stem cells are more specialized than pluripotent stem cells and
include adult stem cells that replenish dying cells or repair damaged tissue. Multipotent
cells can be harvested from a variety of sources, but have limited proliferative ability.
Depending on the type of multipotent cell, they are able only to form the differentiated
3


cell types of a specific tissue. These include neural stem cells (which form neurons,
astrocytes, and oligodendrocytes), endothelial stem cells (blood vessels), hematopoietic
stem cells (blood cells), mesenchymal stem cells (bone, cartilage, muscle, and fat), and
many others. In the interest of orthopaedic research, discussions will be limited to
mesenchymal stem cells.
Mesenchymal Stem Cells
Mesenchymal Stem Cells (MSCs) are a type of multipotent stem cell that can
differentiate into a variety of cell types and maintains the ability to self-renew, however
at a limited capacity. Human MSCs are defined as being plastic adherent, expressing the
surface molecules CD105, CD73, and CD90, and showing multilineage differentiation in
vitro towards the osteogenic (bone), adipogenic (fat), and chondrogenic (cartilage)
lineages, though they are not limited to these cell lines [13], MSC therapies are among
the most common of stem cell applications, as they are free of ethical harvesting
concerns, have numerous sources, show little risk of producing an immunogenic
response, and virtually no risk of teratoma formation (unlike pluripotent cells) [14],
They are being investigated for treating a variety of diseases including myocardial
infarction, liver cirrhosis, diabetes, spinal cord injuries, osteoarthritis, and many more
[14].
MSCs derived from harvested bone marrow (BM-MSCs) are the most commonly
studied stem cells for orthopaedic regenerative medicine. In cases of trauma, abnormal
development, and congenital malformations, MSCs have shown particular promise for
scaffold seeding and inciting bone and cartilage regeneration. Scaffold implants provide
support in bone and cartilage defects, while serving as a template for seeded MSCs to
4


encourage growth and regeneration of the damaged tissue [15], They have been shown to
enhance bone healing in both small and large animal models, whether seeded on scaffolds
or implanted directly [16], They have also shown some success clinically, when used for
the treatment of non-unions in distal tibial fractures, demonstrating no adverse effects and
reducing union time by half [17],
The clinical application of stem cell therapies appears limitless, but there are a
number of issues surrounding the harvest and use of stem cells. For example, the
proportion of BM-MSCs obtained from bone marrow aspiration is less than 0.01% [16,
18], Translation of BM-MSC technology could is also limited by the fact that human
MSCs cannot be expanded indefinitely in culture, and their differentiation capacity has
been reported to decrease with aging and aging-related diseases [19-21], This can limit
their therapeutic potential in bone regenerative medicine, especially in older patients.
Thus, there is an emerging interest in the identification of alternative cell sources for
MSCs.
Induced Pluripotent Stem Cells
In 2006, Kazutoshi Takahashi and Shinya Yamanaka demonstrated that by
introducing four factors (Oct3/4, Sox2, c-Myc, and Klf4) to adult mouse fibroblasts
cultured under ESC culture conditions, they could induce pluripotency in the fibroblasts
[22], The reprogramming of fibroblasts into what has been designated as induced
pluripotent stem cells (iPSCs) was a scientific breakthrough that introduced a novel stem
cell source that could potentially revolutionize regenerative medicine. iPSCs are
advantageous to current stem cell sources because they can be generated directly from the
cells of the patient requiring therapy, whereby they are customized to match the patients
5


immune system, making the use of immunosuppressants for preventing rejection
unnecessary. This inherently means that they can serve as an autologous stem cell
source, while negating the moral ambiguity with using ESCs.
Beyond what Takahashi and Yamanaka achieved in 2006, researchers have since
improved reprogramming techniques and shown that iPSCs can be reliably differentiated
down a multitude of cell lineages. In 2007, Wemig et al. significantly improved the
techniques that were developed a year earlier, and showed that DNA methylation, gene
expression and chromatin state of the iPSCs were similar to those of ESCs [23], In
subsequent years, Takahashi furthered his research, demonstrating similar pluripotency
induction in adult human fibroblasts and in 2010 Warren et al. devised a safer, more
effective reprogramming method based on mRNA administration [24, 25], Moreover,
cellular reprogramming of skin fibroblasts from older individuals has been reported,
suggesting that iPSCs may provide a source of rejuvenated adult stem cells for patients
that may not have optimal BM-MSCs [26], These findings could prove to be very
advantageous for bone repair.
IPSCs could inherently become any cell in the body but although this is exciting
for regenerative medicine, this pluripotency comes with the risk of forming teratomas
upon implantation [27], Thus it is important to ensure complete differentiation of the
iPSCs towards the desired lineage in vitro before implantation in vivo. The generation of
functional iPS-derived MSCs has been demonstrated, as well as further differentiation
towards bone and cartilage both in vitro and in vivo [28, 29], Furthermore, differentiation
of iPSCs towards a neuronal lineage has also been studied extensively, with even more
specific generation of both Purkinje neurons and motor neurons [30-32],
6


The scope of this study is to observe osteogenic differentiation, pushing iPSC
derivatives to generate bone, so it is important to understand the structure of bone and the
different cell types that make up osseous tissue.
Bone
Bone is a rigid organ of the vertebral skeleton that supports and protects various
other organs of the body. Osteoblasts, osteoclasts, and osteocytes are the three types of
cells that make up bone, forming either cortical or cancellous (spongy) tissue. Cortical
bone forms the outer shell of most long bones and is much denser than cancellous bone,
serving as the primary supportive structure of the organ. The foundational structure of
cortical bone is the osteon, which consists of layers of mesenchymal-derived osteoblasts
that are responsible for synthesizing new bone. Osteoblasts bud vesicles containing
hydroxyapatite, a calcium phosphate, to be deposited between collagen fibrils of the
extracellular matrix, which leads to the formation of new bone [33], Once these cells
become trapped by the bone matrix that they deposit, they differentiate into osteocytes
that reside in little voids known as lacunae, and are responsible for directing routine bone
turnover. Osteocytes communicate through canals known as canaliculi that connect the
lacunae, directing osteoblasts to lay down new bone and osteoclasts to resorb old bone.
Unlike osteoblasts and osteocytes, osteoclasts are derived from monocytes. The blood
supply for the bone is transported through osteonic canals known as Haversian and
Volkmanns canals. The interior of the bone consists of cancellous tissue, made up of a
porous network of thin osteoblast formations, resembling the structure of a sponge.
Within the spaces of the cancellous bone resides the bone marrow and hematopoietic
7


stem cells that give rise to platelets and red and white blood cells. A diagram of long
bone structure can be seen in Figure 1.1.
Compact Bone & Spongy (Cancellous Bone)
Lacunae containing osteocytes Osteon of compact bone
Figure 1.1 Structural anatomy of a long bone.
Regenerative orthopaedic therapies aim to utilize MSCs in order to generate
viable osteoblasts to enhance bone formation in cases of breaks and malunions. In spite
of this potential, current culturing techniques are rather expensive, time-consuming, and
can yield non-homogenous cultures under differentiation. In regards to scaffold seeding,
it has been shown that scaffold design and material choice can have varied effects on
stem cell differentiation alluding to an environmental influence on cell behavior [34-36],
For these reasons and more, it is becoming more crucial to understand what factors are
involved in the growth, proliferation, and differentiation of bone cells and what can be
done to improve culturing techniques. The next section will provide an overview of the
effects the mechanical environment can have on stem cell behavior.
8


Mechanosensing and Mechanotransduction
One growing area of stem cell research is investigating how the cellular
microenvironment affects stem cell activity. Researchers have documented the effects of
surface micro-topography, biochemical environment, substrate material, three-
dimensional scaffold constructs, and mechanical vibration on the growth and
differentiation of stem cells [37-40], In particular, the mechanical properties of the
surrounding environment have been found to affect many cellular processes in a variety
of cell types. It has been demonstrated that naive MSCs show extreme sensitivity to
tissue-level matrix elasticity, becoming neurogenic on soft substrates, myogenic on stiffer
substrates, and osteogenic on a comparatively rigid substrate [41], MSCs, in particular,
have been shown to become either osteogenic, chondrogenic, myogenic, or adipogenic
due to matrix elasticity specification, and have even shown memory response to
mechanical dosing influence [42-46], Substrate influence has also been observed in
mesodermal differentiation of human embryonic stem cells and subsequent terminal
differentiation for both chondrogenic and osteogenic lineages [47-49], It has shown
regulatory influences on kidney cell morphology and locomotion, as well as fibroblast
polarization [50, 51], Even iPSCs have shown mechanotransductive properties,
demonstrating neural and cardiac differentiation when plated on a soft matrix (0.6 kPa)
[52], This inherent ability of cells to recognize variations in matrix stiffness is clearly an
important factor in the regulation of stem cell fate. The mechanism by which cells can
feel their surrounding environment and convert this stimulus into an electrochemical
response is referred to as mechanotransduction.
9


Extracellular Matrix-Integrin-Cytoskeleton Mechanosensing Pathway
The Extracellular Matrix-Integrin-Cytoskeleton (EIC) mechanosensing pathway
allows for mechanical stresses or vibrations to be rapidly transferred from cell surface
receptors to distinct structures in the cell and nucleus, resulting in regulation of cellular
functions such as cell attachment, proliferation, migration and differentiation [53], This
mechanosensing occurs as cellular integrins bind to the extracellular matrix. As the cell
contracts and pulls on the surface, the cytoskeletal filaments reorient themselves,
resulting in nucleus distortion and redistribution of the nucleolus (Figure 1.2) [54], This
t CSK Tension I ECM Rigidity
Figure 1. 2 Mechanotransduction. The cell attaches to the extracellular matrix (ECM)
through attachment points called integrins. If the cell is attached to a relatively rigid
substrate (left), then as the cell contracts, cytoskeletal (CKS) tension goes up resulting in
nuclear redistribution. If the ECM is flexible (right) the cell can more readily pidl on the
substrate, resulting in a different cellular reaction. Image reprintedfrom Cellular
mechanotransduction: putting all the pieces together again, by Donald E. Ingber, The
FASEB Journal, vol. 20. no. 7. Pgs. 811-827. 2006.
pathway can be activated not only through cell-cell and cell-matrix interactions, but also
by gravitational forces. This effect is evident when astronauts develop osteopenia (a
decrease in bone density) as a result of prolonged space flights [55], Studies have shown
how mechanical stimulation can influence cellular activity in a variety of cells including
dermal and cardiac fibroblasts, cardiac myocytes, endothelial cells, and bone and
cartilage cells [56], Most studies culture cells on flexible substrates, or apply direct force
to adhesion molecules to observe cellular responses, though others have used
10


centrifugation to simulate gravitational changes [57, 58], The sections below will briefly
discuss the roles of each component in the EIC mechanosensing pathway.
Extracellular Matrix
The extracellular matrix (ECM) provides primary structural and biochemical
support for the cell, but the stiffness of the ECM can have an influence on cell behavior
as well. Focal adhesions bind to the ECM and as the cell contracts, the ECM deforms by
an amount relative to its mechanical properties. This provides a certain amount of give
that the cell is capable of detecting. In many cases, cells respond by remodeling the ECM
through newly activated/deactivated gene expression, or by direct mechanical
manipulation of the ECM fibrils [59], The newly remodeled matrix is most often more
resistant to the applied forces.
Integrins
Integrins are transmembrane molecular structures that bind cells to the
extracellular matrix, and have been shown to be an integral part of cellular
mechanosensing. They serve as the structural mediators between the ECM and the
cytoskeleton of the cell. When introduced to a mechanical stress, integrins can either
transduce the force into a chemical response or transmit the force directly to the
cytoskeleton. This stimulates downstream reactions but the specific processes and targets
of each of these stimulations are largely unknown [53, 55], It is hypothesized that the
mechanical signals can be used to induce conformational changes in the integrins
themselves and several studies have shown force application to cause the attached focal
adhesions to stabilize and increase in size and strength [58, 59],
11


Cytoskeleton
The cytoskeleton is made up of a network of microfilaments and serves as the
supportive structure of the cell, both generating force and bearing elastic deformation
[55], Just as muscles pull on specific parts of bones, physical manipulation can influence
the cytoskeleton to mechanically distort specific parts of the nucleus, inducing changes in
cellular organization and gene expression [54], Additionally, the simulated effects of
gravity can be studied with centrifugation. Li et al. showed that osteoblasts exposed to
mild centrifugal force exhibited temporary and reversible changes in gene transcription
which helped to verify the role of the cytoskeleton in mechanosensing [57],
Cellular and Physiological Responses
Though the mechanism for how mechanical energy is transduced into chemical
changes in the cell is not fully understood, new studies are emerging that are beginning to
map the chemical pathways of mechanotransduction. For example, the application of
mechanical stress has been shown to increase focal adhesion kinase phosphorylation and
specifically, Rho kinase is hypothesized to be affected by stress as it is heavily involved
in the generation of cytoskeletal tension [58], Still other studies have hypothesized that
some mechanisms may influence cellular structures directly (i.e. ion channels, nuclear
pores, chromosomes, individual genes, etc.), remaining independent of chemical
signaling mechanisms [54],
As investigations of the EIC mechanosensing pathway continue, the relevance of
its application becomes more apparent. Cardiac fibroblasts have been shown to increase
ECM gene expression and growth factor activation when exposed to mechanical stress in
vitro, and cyclic loading has been shown to induce membrane matrix metalloproteinase
production similar to that found in ischemic hearts [60], ECM restructuring, influenced
12


by cyclic stretch, is a vital process in the strengthening of vascular smooth muscle cells,
which would otherwise result in an aneurysm [59], Others have hypothesized that
mechanosensing can be relevant in cases of hypertrophic scarring as well as tendon and
ligament regeneration [55], Further studies into how cells transduce mechanical signals
into biochemical changes would progress the understanding of mechanosensing, giving a
better visualization of a cells working environment and ultimately providing greater
knowledge for the culture and control of cellular activity.
The Cellular Environment
Cells ability to sense the mechanical properties of their environment presents a
wide variety of stimuli that can affect how cells grow, proliferate, and differentiate.
Research with regards to the microenvironment includes surface micro-nanotopography,
soluble factors, extracellular matrix composition and distribution, and mechanical
stress/strain conditions [61], Each has been shown to influence the efficacy of cell
culture, which has major implications for tissue engineering and regenerative medicine.
Schulze et al. demonstrated how primary mouse motor neurons were successfully guided
and directed towards growth of axons into grid-like neurite networks when cultured on
microtopographical substrates coated with rolled-up Si0/Si02 [62], Similarly, human
MSCs cultured on 10 pm micropost-textured PDMS demonstrated higher gene
expression of osteoblast-specific markers accompanied by substantial bone matrix
formation and mineralization when compared to smooth surface controls [37], These
results were amplified when biochemical soluble factors intended for osteogenic
differentiation were added to the culture media [37], Soluble factors have also been
shown to improve cell survival and proliferation as well as induce cellular
vascularization, regenerate neurons, and even maintain a stem cell phenotype [61],
13


Finally, matrix composition variability can have significant effects on the differentiation
of stem cells and can modulate the phenotype of both endothelial and mesangial cell
populations in culture [39, 63], The compilation of all this data can be utilized to design
bioreactors which mimic the mechanical microenvironment for functional engineered
tissues that undergo mechanical loading under in vivo conditions (i.e. cartilage, bone,
tendons, heart valves, etc.), which is becoming the standard for large scale cell culture
[61, 64],
Goals and Hypotheses
IPSCs, like ESCs, show particular potential for regenerative medicine due to their
ability to proliferate and differentiate into cells of all three germ layers, yet they are more
attractive due to their lack of controversy and virtually limitless availability. In 2014,
Tang et al. demonstrated the promise for iPSCs to promote bone regeneration through
cell seeding of calcium phosphate cement scaffolds and subsequent osteogenesis [15],
Procedures such as this require a homogeneous bank of mesenchymal stem cells derived
from iPSCs, referred to as mesenchymal progenitors (iPSC-MPs), demonstrating the need
for efficient methods of iPSC differentiation. This, in conjunction with the influence of
mechanotransduction and the cellular environment, presents a motivation for studying the
effects of substrate stiffness on iPSC-MP differentiation and osteogenesis. The goal of
this study is to observe the effects of culturing iPSC-MPs on polydimethylsiloxane
(PDMS) substrates of varying stiffness and culturing in either control MSC medium or
osteogenic differentiation medium, with the hypothesis that relatively stiffer substrates
will influence faster, more homogenous osteogenic differentiation, especially compared
to current culture methods.
14


CHAPTER II
MATERIALS AND METHODS
Cell Culture
Induced pluripotent stem cells (iPSCs) were generated from human fibroblasts
collected from the skin biopsy of a 50 year old female. The fibroblasts were
reprogrammed using an optimized mRNA-based approach previously described by
Warren et al. [25], Briefly, mRNA molecules encoding the reprogramming factors Oct4,
Sox2, Klf4, and c-Myc were introduced into the fibroblasts by direct transfection.
Generated iPSCs were positive for the expression of endogenous pluripotency markers
(iOCT4, NANOG, TRA-1-81) by immunofluorescence and were shown to display a
normal karyotype.
iPSCs were then differentiated into mesenchymal progenitors (iPSC-MPs) by a
method previously described by Marolt et al. [65], In brief, iPSCs were cultured in
induction medium (Knockout DMEM (Life Technologies, Carlsbad, CA, USA)
supplemented with 20% FBS (Atlanta biologicals, Lawrenceville, GA, USA), 2 mM
glutagro supplement (Coming, Manassas, VA, USA), 0.1 mM non-essential amino acids
(Corning, Manassas, VA, USA), 0.1 mM 2-Mercaptoethanol (Fisher Scientific, Fair
Lawn, NJ, USA), and 1% penicillin streptomycin (Thermo Scientific, Logan, UT, USA)),
hereby referred to as MSC differentiation media, for one week and passaged until they
achieved a homogenous fibroblast-like morphology under microscopic examination.
Mesenchymal differentiation was confirmed by flow cytometry for mesenchymal
markers CD90, CD73, and CD105 (BD Biosciences, San Jose, CA, USA) when
compared to bone marrow mesenchymal stem cells (BM-MSCs) [65, 66], Cells were
then collected, frozen down as cell stocks, and placed in liquid nitrogen storage.
15


2
Cells were thawed after being removed from storage and plated in a 225cm tissue
culture treated flask with 30 mL of MSC differentiation media. Cells were incubated at
37C, 5% CO2, expanded, and plated for differentiation studies at passage 9.
PDMS culture trials were conducted three times, using frozen cell stocks from the
same set of iPSC-MPs.
Substrate Material Synthesis
PDMS substrates were fabricated using a two part silicone elastomer kit
(SYLGARD 184, Dow Corning, Midland, MI, USA), which was chosen for its
biocompatibility and regular use in mechanotransductive differentiation studies [37, 44,
47, 67], The stiffness of each substrate was varied by altering the base to curing agent
ratio. Five substrates were characterized by the percentage of curing agent that made up
the final solution (4%, 8%, 12%, 16%, and 20%). Each substrate was mixed and
degassed, and then 2 mL of the final solution was transferred in triplicate to 6-well plates
and allowed to cure at room temperature for at least 60 hours. Substrates were then
rinsed in 70% ethanol and allowed to air dry.
Through a method previously described by Pelham and Wang, Type I collagen
was covalently crosslinked to the PDMS surface with N-Sulfosuccinimidyl-6-(4'-azido-
2'nitrophenylamino) hexanoate (Sulfo- SANPAH, CovaChem, Loves Park, IL, USA)
[50], A 200 mg/mL solution of Sulfo-SANPAH was created by dissolving in
dimethylsulfoxide (DMSO, Sigma Aldrich, St. Louis, MO, USA), which was further
diluted with 50 mM 4-(2-hydroxyethyl)-l-piperzaineethanesulfonic acid (HEPES, GE
Healthcare Life Sciences, Logan, UT, USA), pH 8.5, to a final concentration of 0.5
mg/mL. The Sulfo-SANPAH solution was used to cover the PDMS substrates, which
were then exposed to UV light from a transilluminator for 30 minutes in a cell culture
16


hood (Model: 1385, Thermo Fisher Scientific, Marietta, OH, USA). Excess Sulfo-
SANPAH solution was aspirated and plates were exposed to UV light for an additional
30 minutes. A 0.05 mg/mL collagen concentration was made by diluting Collagen Type
I, Rat Tail (Coming Life Sciences, Tewksbury, MA, USA) in 0.02 N acetic acid. Wells
were washed three times in sterile phosphate buffered saline (PBS) and 1 mL of the
collagen solution was added to each well and allowed to incubate overnight at 4C.
Collagen solution was aspirated and wells were washed twice with sterile PBS. Plates
were wrapped in Parafilm M (Bemis NA, Neenah, WI, USA) and stored at 4C until
needed.
PDMS substrates were fabricated separately for each of three trials to account for
variability between batches.
Material Properties
Material properties of PDMS samples were measured by displacement controlled
nanoindentation using a HYSITRON TI 950 Tribolndenter (Hysitron, Minneapolis, MN,
USA). A 250 pm radius sapphire spherical tip (Ti-0185, Hysitron) was used to indent to
a depth of 4000 nm at a rate of 800 nm/s where it was held for 30 seconds before
retracting at the same rate. Force measurements were read in a 5x5 point matrix with an
even separation of 200 pm. The Hertz model with a Poisson ratio of 0.5 was used to
calculate Youngs modulus, E, based on the retracting force curves with the assumption
that the sphere had an infinitely larger E than the PDMS samples.
Testing was conducted on nine samples from each condition (three trials of
triplicate samples), and averages of the samples are reported along with standard
deviations.
17


Cell Plating and Culture
iPSC-MPs were plated from a single cell suspension at a density of 30,000
cells/well on 6-well PDMS-coated plates and were run against controls plated at the same
density on tissue culture plastic (TCP). Cells were cultured in either Complete Culture
Media (CCM, MEM-alpha (Life Technologies, Carlsbad, CA, USA) supplemented with
16.5% FBS (Atlanta biologicals, Lawrenceville, GA, USA), 2 mM L-glutamine (Corning,
Manassas, VA, USA), and 1% penicillin streptomycin (Thermo Scientific, Logan, UT,
USA) or Osteogenic Differentiation Media (ODM, CCM supplemented with 10 nM
dexamethasone (Sigma, St. Louis, MO, USA), 20 mM P-glycerophosphate (Sigma, St.
Louis, MO, USA), and 50 pM L-ascorbic acid 2-phosphate (Fisher Scientific, Fair Lawn,
NJ, USA)). Cells were incubated at 37C, 5% CO2, and media was changed three times
per week (Mondays, Wednesdays, and Fridays). Cells were collected at 3, 7, 14, and 21
day time points.
Culture trials were conducted three times in triplicate.
Cell Proliferation
The alamarBlue assay (Thermo Scientific, Logan, UT, USA) incorporates a
fluorometric oxidation-reduction indicator that reacts in response to the chemical
reduction of growth medium indicating the metabolic activity of cellular growth. A
repeated measure of this assay was conducted at 3, 7, 14, and 21 day time points. A
100% reduced positive control was made by autoclaving 5 mL of CCM with 500 pL of
alamarBlue reagent. One milliliter of this solution was added to one well of a 6-well
plate with 100 pL of ultrapure water. Negative control was made of 1 mL of CCM only.
The respective media for each experimental sample (either CCM or ODM) was aspirated
and replaced with 1 mL of fresh media plus 100 pL of the alamarBlue reagent. Samples,
18


including controls, were incubated at 37C, 5% CO2 for 2.5 hours, then 100 pL of each
sample was plated in triplicate on a 96-well plate and fluorescence measurements (530-
560 nm excitation wavelength and 590 nm emission wavelength) were read using a
Glomax Multi Detection System plate reader (Promega, Madison, WI, USA). After
reading, 6-well sample plates were washed once with PBS and returned to culture with 2
mL of their respective media for future readings.
To calculate the percent reduction of the alamarBlue reagent, relative
fluorescence units (RFU) for each sample were averaged and used in the following
equation:
Experimental RFU value-Negative control RFU value ^ 100 1
100% reduced positive control RFU value-Negative control RFU value ^ *
AlamarBlue analysis was conducted at four time points over the course of one
culture trial, though proliferation was monitored regularly with microscopy over three
culture trials.
Actin Staining
After 3 days of culture, cells were washed twice with PBS and fixed in 10%
neutral buffered formalin for 10 minutes at room temperature. Wells were washed two
more times with PBS and plates were wrapped in Parafilm M and stored at 4C in PBS.
Phalloidin (F-actin) staining was performed by applying ActinGreen 488 ReadyProbes
reagent (Life Technologies, Eugene, OR, USA) to fixed cells using 2 drops/mL of PBS
and incubated for 30 minutes at room temperature. Wells were washed with PBS and
imaged with an Eclipse TE2000-S (Nikon, Melville, NY, USA).
Phalloidin staining was performed on one set of samples from a single culture
trial.
19


RT-qPCR
RT-qPCR strategies were conducted in adherence to the guidelines for the
Minimum Information for Publication of Quantitative Real-Time PCR Experiments
(MIQE) [68],
Cells were collected for reverse transcription polymerase chain reaction (RT-
qPCR) at 3, 7, 14, and 21 day time points. RNA was isolated following manufacturers
instructions for the RNeasy Plus Mini Kit (Qiagen, Valencia, CA, USA). Isolated RNA
was quantified using an Epoch Microplate Spectrophotometer and a Take3 plate (BioTek,
Winooski, VT, USA) and diluted with sterile DNase- and RNase-free water (Fisher
Scientific, Fair Lawn, NJ, USA) to a final concentration of no more than 50 ng/pL.
Isolated RNA samples were stored at -80C until cDNA reverse transcription.
cDNA reverse transcription was carried out following manufacturers instructions
for the High-Capacity cDNA Reverse Transcription Kit (Life Technologies, Carlsbad,
CA, USA), including RNase inhibitor. Samples were once again quantified with the
Epoch Microplate Spectrophotometer and Take3 plate and diluted to a concentration of
50 ng/pL. cDNA samples were stored at -80C until RT-qPCR analysis.
Following manufacturers instructions for the SsoAdvanced SYBR Green
Supermix (Bio-Rad Laboratories, Inc., Hercules, CA, USA), RT-qPCR was carried out
using primers at a final concentration of 500 nM and 100 ng of the DNA template per
reaction. Target genes were selected as markers of osteogenesis and include alkaline
phosphatase (ALP), TWIST' runt-related transcription factor 2 (RUNX2), and osteocalcin
('OCN). Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as a
housekeeping gene for normalizing expression and has been used in this capacity
previously during osteogenic differentiation experiments [49], ALP is typically used as
20


an early marker of osteoblastic differentiation whose expression has been shown to
increase over time in confluent monolayer bone-derived cell cultures [69], ALP also
plays a direct role in mineralization, hydrolyzing several phosphates in order to prepare
ions for deposition [33], RUNX2 is another osteogenic-related marker shown to regulate
the non-collagenous, late stage, osteoblastic marker, OCN [70, 71], Finally, TWIST is a
basic helix-loop-helix transcription factor that, as its expression decreases, has been
shown to increase ALP and RUNX2 expression, and was chosen for its use in previous
osteogenic differentiation studies [49, 72-74], Efficiencies for GAPDH, ALP, OCN,
RUNX2, and TWIST were 112%, 101.5%, 114%, 109%, and 103% respectively.
Temperature cycling was conducted using a CFX Connect Real-Time System
(Bio-Rad Laboratories, Inc., Hercules, CA, USA) and cycling conditions for all genes can
be seen in Table 2.1.
Table 2.1 Cycling Conditions for cDNA
Cycling Step Temperature Time # Cycles
Enzyme activation/Initial DNA denaturation 95C 30 sec 1
Denaturation 95C 5 sec 40
Annealing/Extension 60C 30 sec 40
Melt Curve 65-95C (in 0.5C increments) 5 sec/step 1
Results were analyzed using the Bio-Rad CFX Manager software, which utilized the
mathematical model for relative quantification described by Pfaffl, where the efficiency
of a primer, E, was related to percent efficiency in the following way [75]:
E = (% Efficiency x 0.01) + 1 OR% Efficiency = (E 1) x 100 (2)
21


In this way, an E of 2 was equal to 100% efficiency and represented perfect doubling
with every cycle. Using a Day 0 sample as control, the relative quantity (RQ) of any
experimental sample for a given gene of interest (GOI) was calculated as:
ry/~% _ (control) Cq (sample)
kQgOI V-GOI U
where
Cq (control) = Average Cq for the control sample
Cq (sample) = Average Cq for experimental samples
The relative quantities of the samples were normalized to GAPDH expression with the
following equation:
*r i j r* KQsample (GOI)
Normalized Expressionsample (G0/) = ------------- (4
m£sample(gapdh)
Results were compared using one-way ANOVA analysis with a post-hoc Tukey
test and differences were considered significant when p < 0.05.
RT-qPCR was conducted on triplicate samples over three culture trials. Results
are displayed as the representative average of the triplicate of one trial with error bars
representing standard error of the mean.
Pico Green & ALP Activity
Samples were lysed at 14 and 21 days and alkaline phosphatase (ALP) activity
was quantified in an enzyme immunoassay. Triton X-100 (Sigma, St. Louis, MO, USA)
was diluted to a concentration of 0.1% in distilled water to create the cell lysis buffer.
Samples were first washed with PBS, then 250 pL of the Triton X-100 buffer was added
to each well and samples were incubated for 30 minutes at 4C. Plates were then
wrapped in Parafilm M and stored at -20C overnight.
22


Preparing the SigmaFast p-Nitrophenyl Phosphate (pNPP, Sigma, St. Louis, MO,
USA) solution required vortexing 1 pNPP tablet with 1 Tris tablet in 20 mL of dH20.
Experimental samples were plated in triplicate in 96-well plates with 100 pL of pNPP
solution, 90 pL dH20, and 10 pL of lysed sample. One triplicate of lysis buffer was run
alone to control for background value. Plates were wrapped in aluminum foil to protect
from light and incubated for 30 minutes at room temperature before measuring
absorbance in the Epoch Microplate Spectrophotometer at 405 nm.
ALP activity for each sample was normalized to its DNA content by first
quantifying DNA concentration using the Quant-iT PicoGreen dsDNA Assay (Life
Technologies, Carlsbad, CA, USA) as summarized below. To create a standard curve, a
serial dilution of the provided DNA standard was made with concentrations increasing
two-fold from 15.625 ng/mL to 1000 ng/mL, and included a null sample. Samples were
diluted 1:10 in 0.01% Triton X-100. 100 pL of the standards and samples were added, in
duplicate, to 100 pL of PicoGreen solution (0.05% PicoGreen in IX TE Buffer -
provided in kit) in a black 96-well plate. As a reference blank, a duplicate of 1:1
PicoGreen Solution: Cell Lysis Buffer was plated alongside the samples. After 5 minutes
of incubation at room temperature wrapped in aluminum foil, fluorescence intensity in
the blue spectrum (480 nm excitation/520 nm emission) was measured with the Promega
Glomax Multi Detection System.
After duplicates were averaged, ALP activity [ALPact, pmol pNPP/mL] was
determined from the equation:
_ (Optical Density-Cell Lysis Buff er Blank) x (total volume) x (dilution)
act 18.45 x (sample volume) ^ ^ ^
and normalized [ALPn0rm, nmol pNPP/pg DNA] to DNA content using the equation:
23


ALP
norm ~
ALP net AA nmol
---- x 1000---------
DNAqUclnt [ITnol
(6)
where DNAquant was determined from the PicoGreen fluorescence readings. From the standard
curve, a linear relationship between fluorescence and DNA quantity was determined in the form:
(I" Innv/>^r/>nr/> hi
Fluorescence = a x DNAquant + b OR DNAquant = --------------------------------- (7)
where a and b are constants determined by the standard curve.
ALP activity and PicoGreen analysis was for one culture trial and compared
against RT-qPCR results. Averages of the triplicates and standard deviations are
presented.
Alizarin Red S
Alizarin red s (ARS, Sigma, St. Louis, MO, USA) was used to stain calcium
deposits which certain osteogenic cell lines, specifically osteoblasts, form in culture.
Samples were fixed and stained at 7, 14, and 21 day time points. Media was aspirated
from wells, which were then rinsed with 2 mL of PBS before incubating for 60 minutes at
room temperature in 2 mL of 10% Buffered Formalin (Sigma, St. Louis, MO, USA).
Formalin was aspirated and wells were rinsed with dLLO before incubating for 20
minutes at room temperature in 2 mL of ARS, and finally rinsed 3-4 times with dLLO.
Wells were imaged with an AMG EVOS XL Core Microscope (Fisher Scientific, Fair
Lawn, NJ, USA) and plates were scanned with a LaserJet Pro 500 M570dn (Hewlett-
Packard, Palo Alto, CA, USA). Wells were covered in dH20 and plates were wrapped in
Parafilm M for storage at 4C until quantitative destaining.
For quantitative destaining, a protocol was followed that was originally developed
by Gregory et al [76], Water was aspirated from ARS stained wells and 800 pL of 10%
acetic acid was added to each well before 30 minutes of incubation at room temperature
with gentle shaking. The cell layer was detached with a cell scraper and transferred along
24


with the acetic acid to a centrifuge tube where it was vortexed, heated to 85C for 10
minutes, and put on ice for 5 minutes. The samples were then centrifuged at 20,000 xg
for 15 minutes and 500 pL of the supernatant was neutralized to a pH between 4.1 and
4.5 with 200 pL of 10% ammonium hydroxide. 150 pL of each sample was plated in
triplicate to a 96-well plate and read at 405 nm with the Epoch Microplate.
Spectrophotometer along with the alizarin red standards listed in Table 2.2.
Table 2.2 Alizarin Red Serial Dilutions. ARS solution is originally 29.2141 mM
concentration and is diluted with buffer made from 7.5 mL of 10% acetic acid and 3 mL
of 10% ammonium hydroxide. A 2 mM working stock solution was made by adding 342
piL ARS to 5 mL of dilution buffer which was then diluted two-fold in series for high
range standards. 15 uk of the 2 mM working stock was added to 985 uk of dilution
buffer to create a 30 uM concentration which was then diluted two fold in series for low
range standards.____________________________________________________________________
Range Concentrations
High 2 mM 1 mM 500 pM 250 pM 125 pM 62.5 pM 31.3 pM 0
Low 30 pM 15 pM 7.5 pM 3.75 pM 1.88 pM 0.94 pM 0.47 pM 0
The standard curve generated a linear relationship between absorbance, A, and
quantification, ARqUant. After subtracting the blank value, the following equation was
applied to determine quantity of ARS stain based on absorbance:
A 3. X ARqUant T b OR ARqUant (8)
Where a and b are constants determined by the standard curve. Equation 8 was used to
determine quantification of ARS.
ARS staining was conducted for all three culture trials at the 21 day time point
and once after 14 days of culture, each time in triplicate. Results were consistent across
all trials and representative images are presented. Quantitative destaining was conducted
on one set of triplicate samples and displayed as an average of the three with error bars
representing the standard deviation.
25


Statistical Analysis
To compare groups, assumptions of parametric data were tested using Shapiro-
Wilk test for normality of data distribution and Levenes test for homogeneity of
variance. For parametric data, one-way ANOVA and Tukey post hoc analysis was used.
For non-parametric data, a Kruskal-Wallis test was used with a post hoc pairwise Mann-
Whitney U with a Bonferroni correction. All analyses will be performed with SigmaPlot
v. 11.2. Significant differences are reported with a p-value < 0.05.
26


CHAPTER HI
RESULTS
Substrate Stiffness
Substrates of PDMS were fabricated by mixing different percent concentrations of
the crosslinking component of the two-part elastomer. For the rest of this work, samples
will be referred to by the percent concentration of the crosslinker in the substrate on
which the cells were plated (4%, 8%, 12%, 16%, 20%, and TCP for reference).
Nanoindentation results showed elastic modulus to be 0.6790.422, 1.1820.251,
1.7130.328, 2.2450.309, and 2.4470.459 MPa for 4%, 8%, 12%, 16%, and 20%
substrates, respectively (Figure 3.1). A one-way ANOVA statistical analysis showed all
samples to be significantly different from each other (p<0.05) except for 16% and 20%.
3.5
4% 8% 12% 16% 20%
Figure 3.1 -Material Properties for PDMS Samples. Samples were measured by
displacement controlled nanoindentation as an average of 25 points over an area of 0.64
mm2. Three samples were fabricated at each of three time points to account for
variability between batches.
27


Cell Spreading and Proliferation
Cells were plated at a seeding density of 30,000 cells per well of 6-well tissue
culture plates (growth area: 9.5 cm ). Wells were imaged after 24 hours (Figure 3.2a) to
ensure cellular attachment, which appeared consistent among all substrates. Wells were
repeatedly imaged over the time course of the experiment at regular intervals and showed
similar growth and proliferation until cells reached confluency at about Day 7.
Phalloidin staining after 3 days of culture (Figure 3.2b) showed early cellular
spreading. PDMS substrates hindered imaging by reducing brightness and resolution, but
post-imaging contrast enhancement improved visibility. Though F-actin concentration on
TCP appeared to be more spread out than that on stiffer substrates (especially 12% and
16%), no significant differences could be observed among conditions for either cells
grown in CCM or ODM.
Percent oxidation-reduction indicated by alamarBlue showed similar proliferation
rates among the different substrates. Figure 3.3a shows that proliferation activity
continued to increase over the three week incubation period and held no significant
difference between the different substrates when cultured in CCM. When cultured in
ODM, however, stiffer substrates (16% and 20%) showed significantly reduced
proliferation at Day 21 in comparison to the CCM counterparts. These samples even
reduced in comparison to their Day 14 ODM proliferation rate. Day 7 for ODM also
showed a significant decrease in proliferation rate, compared to samples cultured in
CCM. This decrease in proliferation may be indicative of higher differentiation rates at
this time point. Figure 3.3b is a representative image of several conditions where over-
confluency resulted in peeling of the cellular monolayer, which led to three dimensional
28


cultures of cells on the stiffer PDMS substrates (particularly 16% and 20%) and
significantly reduced confluency in the peeled areas.
CCM W ODM
4% 8% 4% 8%
12% 16% 12% 16%

20% TCP 20% - fCP 1 *
CCM 0 b) ODM
4% 8% 4% 8%
12% l6% 12% 16%
20% TCP 20% TCP
Figure 3.2 Cellular Attachment and Spreading, (a) Microscopic images of wells, taken
after one day of culture on PDMS substrates and Tissue Culture Plastic (TCP). Images
were taken at 100X magnification, (b) After three days of culture in either CCM or
ODM, cells were fixed, stained, and imaged to show cell spreading. Cells were imaged
at 200X magnification and contrast was enhanced post-imaging to increase stain
visibility.


(a)
CC\1 ODM
Figure 3.3 Cellular Proliferation Rates. AlamarBlue measurements were repeated on
the same set ofplates at 3, 7, 14, and 21 days of culture, (a) Graphs represent cidtures
grown in Complete Culture Media (CCM) and Osteogenic Differentiation Media (ODM)
over the time course of incubation, (b) Representative image at Day 21 of cellular
peeling and detachment of cells cultured on 20% in ODM. Image taken at 40X
magnification.
30


Gene Expression
Gene expression was quantified by RT-qPCR, presented as a fold increase of Day
0 gene expression, and normalized to the housekeeper GAPDH. Four genes were chosen
for representation of osteogenic differentiation (ALP, TWIST, RUNX2, and OCN). Figure
3.4 displays relative expression of these genes at 3, 7, 14, and 21 days of incubation for
both CCM and ODM cultures. An asterisk (*) is used to denote statistically significant
differences (p<0.05) between a given substrates expression level in comparison to TCP
in order to emphasize the effect of specialized substrate fabrication compared to
traditional cell culture methods. Significant differences among varying PDMS substrates
are also noted.
ALP expression in CCM shows no significant difference between substrates with
the exception of an increase in 12% compared to TCP at Day 14. However, it is
important to note the trend for the middle PDMS substrates (12% and 16%) to express
slightly greater amounts of ALP compared to both the softer (4% and 8%) and stiffer
(20%) substrates. Additionally, it can be noted that TCP regularly expressed less ALP
compared to all other substrates. Cells cultured in ODM show similar trends in ALP
expression, with significant differences in 8% and 12% at different time points.
Upregulation of ALP began early in both culture media, though more significantly in
ODM, and continued to be highly expressed at the later time points.
Cells cultured in both CCM and ODM showed trends for TWIST expression to
decrease to almost Day 0 expression levels at the 7 and 14 day time points. This was
reversed at the final time point, where expression of TWIST returned to levels similar to
Day 3. Significant differences again favored moderately stiff samples (8%, 12%, and
16%), though changes in expression levels are greater in ODM samples than in CCM.
31


It
ODM CCM
Relative Normalized Expression
h- K> UJ 4- KS*
0 o o o o
o o o o o o
1 1 1 I I
K>
o
H
9
Relative Normalized Expression
i-> u> ji. a\ ~j yo
U*
o
o
o
o
KJ
o
o
UJ 4- ^
o o o
o o o
4^
00
-V

K>
0s
'
to
o
^
9
j
K)
o
&
cn

00


ee
ODM CCM
(q)


K
ODM CCM
Relative Normalized Expression
Relative Normalized Expression
(0)


S£
ODM
Relative Normalized Expression
O O o u*
O O O
CCM
a
p
C/5
K)
u
03
'C
C/)
(P)


Figure 3.4 Gene Expression Data. Time-course relative expression of osteogenic
markers. Markers for ALP (a), TWIST (b), RUNX2 (c), and OCN (d) were analyzed.
Expression of all genes were normalized to the housekeeper GAPDH and expressed as a
relative fold increase on Day 0 iPSC-MPs. *p<0.05 for material compared to TCP
Expression of RUNX2 showed no significant changes in either media for the first
week of culture. At the 14 and 21 day time points, marked increases in expression could
first be noticed in ODM and CCM respectively. Day 21 ODM samples saw the greatest
expression of RUNX2 with 4% expressing significantly more over 12%, 16%, 20%, and
TCP samples.
OCN expression saw no significant increase until Day 14 for ODM cultures.
Here, again, the trend for 12% and 16% substrates to be upregulating slightly more,
though no statistically significant difference could be determined. The increase in
expression of OCN at Day 21 in CCM is noteworthy given the lack of osteogenic signals
in the media. All samples cultured on PDMS substrates show osteogenic trends,
especially on 16% substrates, and the upregulation of osteogenic markers is notable when
comparing to samples cultured on TCP.
Alkaline Phosphatase Activity
ALP activity [nmol] was measured at 14 and 21 day time points and normalized
to the amount of DNA content [pig]. Figure 3.5a shows that in the CCM condition 12%
had significantly greater ALP activity than both the softest substrate (4%) and the control
(TCP). After 21 days, 8% and 16% also showed significantly greater activity in
comparison with the control. It is only the softest substrate (4%), in fact, which showed
no significant difference when compared to TCP. Interestingly, ALP activity increased
between Day 14 and Day 21 across all conditions, again alluding to a possibility for
substrate stiffness to affect differentiation events without chemical signaling.
36


When cultured in ODM (Figure 3.5b), cells plated on TCP showed the least
amount of ALP activity at both time points, even compared to 4%. Additionally, the
stiffest substrate (20%) became statistically different compared to TCP in this condition,
and even showed the greatest amount of ALP activity at the end of the three week period.
It can also be seen that these activity levels mirror gene expression trends discussed in the
previous section.
80
70 -
60 -
Iso-
c:
; 40 -
i 30 -
; 20 -
10 -
0 --
14 Days
p < 0.05
* *
III!
4% 8% 12% 16% 20%
Figure 3.5 ALP Activity. Cells were lysed after 14 and 21 days of culture in either
CCM (a) or ODM (b), and ALP activity was measured and normalized to DNA content
determined by PicoGreen fluorenscence. *p<0.05 for material compared to TCP
37


Calcium Deposition
Alizarin red s was used to stain
for calcium deposition after 7, 14, and
21 days of culture in CCM and ODM.
Samples cultured in CCM showed no
indication of calcium deposition in any
condition (data not shown).
Representative images of calcium
deposition by cells cultured in ODM
can be seen in Figure 3.6.
Samples fixed and stained
after one week showed no signs of
calcium mineralization when cultured
in ODM, indicating that iPSC-MPs
had not deposited any matrix.
However, within the next 7 days,
evidence of calcium could be seen
from the staining. Qualitatively, it is
apparent that 16% substrates resulted
in the greatest amount of calcium
deposits, while 12% and 20% had a
comparable follow-up. Cells cultured
Figure 3.6 Calcium Deposition Data. Alizarin
red s was used to stain calcium deposition at 7,
14, and 21 day time points for cells cultured in
CCM and ODM. Plates were scanned and
representative images are displayed. Cells
cultured in CCM showed no indication of calcium
deposition at any time point (data not shown).
38


on TCP, however, showed no indication of calcium deposition after 14 days of
incubation. This is notable as all PDMS substrates showed at least some level of
mineralization at this time point.
(a) (b)
4% 8% 4% 8%
12% 16% 12% 16%
20% ^7 20% Z* TCP
Figure 3.7 Microscopy. Microscopic images of cells cultured in osteogenic media after
21 days at 40Xmagnification. Pictures were taken before (a) and after (b) Alizarin red s
staining.
After 21 days of culture, calcium deposition was clearly increased on all
substrates, with 16% producing the largest amount of the mineral. Figure 3.7 shows the
same wells at 40X magnification both before (a) and after (b) ARS staining. The images
in Figure 3.7b more clearly show cell morphology independent of mineral staining. It is
especially noteworthy on the 20% PDMS, where cells had peeled away from the
substrate, that the PDMS itself had no effect on the outcome of the stain.
Quantification of the ARS confirmed qualitative evidence of reduced mineral
deposition on TCP when compared to PDMS. Graphical comparisons of ARS
quantification can be seen in Figure 3.8. After 14 days, 12% and 16% substrates resulted
39


in significantly greater amounts of calcium compared to TCP. 16% also showed
significantly more deposition than softer PDMS substrates, and continued to mineralize
more calcium than TCP after 21 days. After 21 days, all PDMS substrates (except 4%)
had resulted in significantly greater calcium deposition than TCP.
4o 8o 12o 16o 20 o TCP 4o 8o 12o 16b 20o TCP
Figure 3.8 Quantifying Alizarin red s. Quantitative destaining of ARS was performed
after plates were imaged at 14 (a) and 21 (b) day time points for ODM cultures. Plates
were readfor absorbance at 405 nm and run against a standard curve generatedfrom 2-
fold serial dilutions of ARS. *p 0.05 for material compared to TCP
40


CHAPTER IV
DISCUSSION
Cell Proliferation
Microscopic imaging of plates after 24 hours shows cell attachment after seeding
at a density of 30,000 cells per well (Figure 3.2a). At this point it appears as though cell
attachment was slightly decreased on TCP in both media conditions, though due to visual
interference by the PDMS substrate, it is difficult to form any definitive conclusions. By
Day 3, actin spreading in ODM samples appears to be slightly greater than in CCM
cultures, as indicated by less localized fluorescence of the ActinGreen reagent (Figure
3.2b). This is in line with previous investigations that showed that osteogenic
differentiation increases with an enhanced degree of spreading [77, 78], Within the
ODM culture, 12% and 16% appear to have less localized actin fibers than other PDMS
substrates, though cells cultured on TCP seem to present the greatest amount of spreading
at this time point.
The effect of substrate stiffness on proliferation was further analyzed by
quantitative measure over the 21 day incubation period. AlamarBlue data shows
consistent proliferation rates among different substrates over the time course of
incubation when cultured in CCM. All samples show significantly increased
proliferation as incubation time increases. In ODM cultures, the reduction in
proliferation by all samples at Day 7, when compared to CCM, suggests differentiation
induced by the osteogenic media [79], At this point, the osteogenic development
sequence would predict a significant increase in alkaline phosphatase activity, followed
by an upregulation of osteocalcin expression, and subsequent mineral deposition, all of
which will be discussed later [79], The decrease in proliferation by the stiffer substrates
41


at Day 14 and Day 21 is more likely a result of cellular detachment rather than an
indicator for further differentiation events. This was a common occurrence for the stiffer
substrates (especially 20%) cultured in ODM to become over confluent and begin
receding into a three dimensional mass. With the exception of these samples, the mean
proliferation of ODM specimens at Day 14 is notably higher than those cultured in CCM.
Perhaps this reduction in proliferation of the CCM samples is indicative of a delayed
differentiation event caused by mechanical stimuli without the presence of chemical
signals. Further evidence for this hypothesis will be discussed.
Gene Expression
The upregulation of ALP is a distinct indicator of early osteogenic differentiation
events that resemble previous investigations of osteogenesis [69, 80], It is evident as
early as Day 7 that the stiffness of the substrate has a significant effect on the cells
expression of the gene. By Day 14, cells cultured on the 12% sample are expressing
significantly more ALP than TCP cultures, not only in ODM but also notably in CCM.
This early upregulation of ALP is correlated with the down regulation of the TWIST gene.
TWIST serves as an early mediator of the mesoderm, which subsequently forms
mesenchyme and, as noted from previous investigations, was expected to decrease during
osteogenic differentiation to allow for RUNX2 and ALP upregulation [72-74], The high
expression of the gene at the early stages is indicative of the mesenchymal cell type and
subsequent downregulation, notable at Day 7, substantiates previous data for the
influence of TWIST on ALP [72],
Additionally, the downregulation of TWIST in osteoblast precursors coincides
with the upregulation of RUNX2, which conforms to results of previous investigations
[73, 74], Though few significant differences can be noted, the data indicates trends for
42


moderately stiff substrates (primarily 12%) to have regularly higher expression levels of
osteogenic markers in both CCM and ODM. For RUNX2, it is evident by Day 14 that the
gene was being upregulated in cells cultured in ODM, which is generally accepted as a
significant measure of osteoblastic differentiation [70], It has also been shown that the
transcription factor of RUNX2 is a direct regulator of ALP, contributing to its significant
increase in expression in the later time points [81],
The final genetic indicator of osteogenesis, OCN, is considered a late stage
marker and thought to be specifically expressed in mature osteoblasts [71], There is an
apparent upregulation of OCN at the earliest RT-qPCR time point in ODM, which is in
line with previous investigations, but more significant expression is not noticed until the
14 and 21 day time points [71], Contrary to its effects on ALP and RUNX2, the
downregulation of TTFZSThas been shown to correlate with a downregulation of OCN
[72], This may explain the correspondingly reduced expression of OCN at Day 7 and
increased expression at Day 14, but this is an enigmatic correlation that may require
further investigation. Regardless, the notable increase in OCN expression at the later
time points is indicative of terminal osteoblast differentiation.
Though no definitive indication of osteoblastic differentiation occurred in cells
cultured in CCM, Figure 4.1 illustrates similar gene expression profiles for CCM at Day
21 and ODM at Day 14. The consistent under regulation of osteogenic markers by cells
cultured on TCP alludes to a possibility for substrate stiffness to influence differentiation
43


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Figure 4.1 Similarities between CCM and ODM cultures. Gene expression results for
cells cultured in CCM at Day 21 (top) and ODM at Day 14(bottom). Correlations in
expression profiles can be drawn between the two conditions at the different time points
and trends indicate favor for PDMS substrates of moderate stiffness. *p<0.05 for
material compared to TCP
44


without the presence of osteogenic chemical signals, however at a much slower rate. In
order to further study these effects, future investigations should draw out incubation
periods for at least four weeks.
Many of the recurring trends in the RT-qPCR data indicate osteogenic
differentiation, though no definitive conclusions can be reached from this data alone. It is
important to note the tendency for moderately stiff substrates (particularly 12% and 16%)
to regularly show higher expression of osteogenic markers than other PDMS substrates
not only in ODM cultures, but also in CCM. Additionally, TCP regularly produced the
lowest expression of osteogenic markers when cultured in ODM as well as in the
previously stated CCM cultures, which supports the hypothesis that TCP may not be the
best culture substrate for osteogenic differentiation.
ALP Activity
The previous section summarized cellular activity at the gene level alluding favor
for 12% and 16% substrates in osteogenic differentiation, especially compared to TCP.
So for a higher order of confirmation, an assay at the protein level was conducted to
confirm results. Figure 4.2 illustrates the qualitative similarities between gene expression
results for ALP, and protein ALP activity. There is a notable correlation between ALP
gene expression and ALP protein activity, which can confirm reliability of RT-qPCR
results.
As an osteogenic marker in and of itself, robust activity of ALP is considered a
measurable marker of successful differentiation as it contributes greatly to mineralization
and is shown to be one of the first functional genes to be expressed in the process of
calcification [82],
45


14 Days CCM 21Days-CCM
4% 8% 12% 16% 20% TCP 4% 8% 12% 16% 20% TCP
Figure 4.2 Comparison of gene and protein alkaline phosphatase (ALP) activity.
Qualitative comparison among common conditions shows very similar relative
expression of ALP. Common trends among conditions show PDMS substrates of
moderate stiffness (8%, 12%, and 16%) regularly expressing greater amounts of
activity, while the control condition (plated on TCP) commonly showed the least amount
of ALP.
Calcium Deposition
Results of alizarin red s staining provide additional support for the data resulting
from RT-qPCR and ALP activity. Qualitatively, it is apparent that moderate to stiff
substrates (namely 12%, 16%, and 20%) produce the greatest amount of calcium
deposition, a definitive marker of terminal osteoblast differentiation. Quantitative
destaining showed that after 21 days of culture in ODM, cells cultured on all but the
softest substrate (4%) produced significantly more calcium than did the TCP control.
Similar results can be observed for ALP activity under the same conditions. Even greater
46


evidence for the effectiveness of material stiffness on the osteogenic differentiation of
iPSC-MPs is at the 14 day time point, where all PDMS substrates deposited a noticeable
amount of calcium while TCP did not (Figure 3.6). Again, this data is supportive of that
provided by RT-qPCR and ALP activity at this time point and indicates a strong
correlation between the 12% and 16% substrates and efficient osteogenesis. Even the
stiffest PDMS substrate (20%), which was common to experience over confluency and
peeling (Figure 3.7), showed greater amounts of calcium deposition than the control,
which is also supported by ALP activity and RT-qPCR results.
Should a strong correlation between ALP activity and mineralization be assumed,
as has been shown in previous investigations, the threshold of ALP activity to produce
calcium deposition could be determined from this data [82, 83], Referencing Figure 3.6,
at Day 14 all PDMS substrates have noteworthy amounts of calcium deposited onto the
substrate, while TCP shows negligible deposition. Figure 3.5b (Day 14) shows that ALP
activity for TCP is below a concentration of 10 nmol ALP/pg DNA, while all PDMS
substrates have a concentration higher than 10. By 21 days, TCP passes this threshold
and begins to show a significant amount of calcium deposits. Noting CCM ALP
concentration levels after 21 days, even the substrate with the highest amount of activity
(12%) never crossed the 8 nmol ALP/pg DNA mark, nor showed signs of significant
amounts of calcium deposits. Based on this data, the threshold of ALP activity for active
calcium deposition could fall between 7 and 12 nmol ALP/pg DNA (highest ALP
concentration without calcium deposition and lowest ALP concentration with calcium
deposition Figures 3.5b and 3.6), but the hypothesis requires further investigation.
47


Mechanical Influence in CCM Cultures
An interesting trend can be observed for cells cultured in CCM that seems to
follow the osteoblast development sequence, however at a slightly slower pace than the
ODM cultures [79], The alamarBlue results show a slight drop in proliferation,
compared to ODM, at Day 14. This reduction in proliferation could be indicative of the
beginning of differentiation events, which were evident at Day 7 for the ODM cultures.
Figure 4.1 illustrates similarities between the expression of genes for CCM at Day 21 and
ODM at Day 14. Though ALP expression is lower in CCM, the expression profile across
the different substrates is very similar and most other conditions show comparable gene
expression levels. ALP protein activity in CCM cultures did not quite reach the levels
observed in ODM, yet, as discussed in the previous section, the activity was approaching
a level that may be inducive to mineral deposition. There is enough evidence to support a
hypothesis for mechanical stimuli to induce osteogenic differentiation in iPSC-MPs
without the influence of chemical signaling, though cells may require culture for two to
three weeks longer than what was tested in this study.
Future Directions
Results indicate that iPSC-MPs plated on moderately stiff substrates (12% and
16%) display significantly greater osteogenic differentiation at faster rates than both
softer and stiffer substrates especially compared to cells cultured on TCP. These
conditions correspond to a plating stiffness of about 2.245 MPa. These results are
slightly lower, though comparable to those from previous investigations where Evans et
al. showed that 2.7 MPa substrates encouraged terminal osteogenic differentiation of
embryonic stem cells and Wang et al. demonstrated enhanced osteogenic differentiation
of MSCs on substrates near 3.0 MPa [44, 49], Based on these studies, 20% PDMS
48


substrates (2.4470.459 MPa) should have induced greater osteogenic differentiation
than softer substrates, however over-confluence and cellular peeling was common and
may have affected results of the assays. In contrast to CCM cultures that should be
drawn out, it would be beneficial to reduce the incubation period of ODM cultures and
increase the frequency of conducted assays. Similarly, in cultures of peeled cells (Figure
3.3b) it would be beneficial to compare osteogenic markers for cells that collected into 3-
D culture to the cells that remained in monolayer. This could be conducted by scraping
the peeled cells and analyzing them separately from the unpeeled cells, or staining the
entire well with fluorescent antibodies for osteogenic markers and making visual
comparisons. Assays such as these could indicate whether or not peeling was beneficial
for osteogenesis and confirm the effectiveness of the substrate to induce differentiation.
As per previously established protocols, collagen type I was crosslinked with the
PDMS substrates and TCP for consistency, in order to promote cellular attachment to the
elastomer [50], Collagen is a major component of the extra-cellular matrix for bone
formation and could have influenced cellular differentiation. Confirmation of successful
crosslinking of collagen across all substrates should be conducted in future studies so as
to ensure that this did not vary across the different substrates, whereby affecting the
outcome of the experiment. Similarly, cellular attachment without collagen crosslinking
could also be observed to determine the necessity for the use of this protein.
With substrate stiffness optimized for osteogenic differentiation of iPSC-MPs,
further studies are required in order to best replicate the microenvironment conducive for
bone modelling. The microenvironment that can affect cellular behavior is composed of
the extracellular matrix, the surrounding cells, signals from autocrine, endocrine, and
49


paracrine signaling, nanotopography, and extracellular mechanical forces such as shear
stress caused by blood flow [61], Studies have observed many of these effects on
osteogenesis, and results from these experiments in conjunction with what is known
about bone anatomy and modelling can be used to generate a matrix of culture conditions
to test the efficacy of combining all of these stimuli in order to create an environment
optimized for osteogenic culturing [37, 39, 40, 61, 84],
Implications for Scaffold Seeding & Bioreactors
Bioreactors, designed to mimic the mechanical microenvironment for functional
engineered tissues, can use the data from this study in order to recreate plating surfaces
with material properties similar to that of the 12% and 16% substrates that produced the
greatest results for osteogenic differentiation. The resulting cell cultures could then
provide a virtually unlimited source of viable, homogenous cells that could be utilized for
a variety of emerging stem cell therapies [2, 3, 14, 16], Methods developed here could
also be applied to priming cells for scaffold seeding which has been shown to improve
bone regenerative capabilities of iPSC-MPs on bone scaffold implants [15],
Furthermore, advances in biomaterial development allow researchers to more tightly
control the mechanical properties of the scaffolds themselves [35, 36], Data obtained
from this study can provide information that could guide the design of scaffolds to
promote the osteogenic differentiation of iPSC-MPs and eventually lead to better bone
formation in vivo.
50


CHAPTER V
CONCLUSIONS
From plating and early spreading, through proliferation and gene expression, to
protein activity and mineral deposition, it is clear that osteogenic differentiation of iPSC-
MPs favors stiffer substrates with an elastic modulus about 2.2450.3 MPa. This is
especially evident for cells cultured in osteogenic media, which showed significantly
increased levels of calcium deposition in addition to an upregulation of osteogenic gene
expression and protein activity. When cultured in CCM, cells displayed trends for
osteogenic differentiation without the presence of soluble differentiation factors, however
at a slower pace compared to ODM cultures. Further investigation is required in order to
verify these effects on CCM cultures.
The results from this study clearly indicate a cellular preference for substrates
with particular mechanical properties that allude to methods for faster, more efficient cell
differentiation techniques. The results from this study can be used for optimizing iPSC-
MP culture techniques in order to provide viable, homogenous cultures of safe and
effective stem cells for orthopaedic tissue engineering.
51


WORKS CITED
1. Weinstein, S.I., Yelin, Edward EL, Watkins-Castillo, Sylvia I. The Burden of
Musculoskeletal Diseases in the United States. Prevalence, Societal and
Economi cCost2014 [cited 2015 10/13],
2. Fox, J.M., Genever, Paul G., Use of Adult Stem Cells for Orthopedic Regenerative
Medicine Applications. Cell & Tissue Transplantation & Therapy, 2014. 6: p. 19-
25.
3. Watt, F.M. and R.R. Driskell, The therapeutic potential of stem cells. Philos Trans
R Soc Lond B Biol Sci, 2010. 365(1537): p. 155-63.
4. Rao, M., Stem cells and regenerative medicine. Stem Cell Res Ther, 2012. 3(4): p.
27.
5. Evans, M.J. and M.H. Kaufman, Establishment in culture of pluripotential cells
from mouse embryos. Nature, 1981. 292(5819): p. 154-6.
6. Martin, G.R., Isolation of a pluripotent cell line from early mouse embryos
cultured in medium conditioned by teratocarcinoma stem cells. Proc Natl Acad
Sci USA, 1981. 78(12): p. 7634-8.
7. Thomson, J.A., et al., Embryonic stem cell lines derivedfrom human blastocysts.
Science, 1998. 282(5391): p. 1145-7.
8. Kerr, C.L., et al., Embryonic germ cells: when germ cells become stem cells.
Semin Reprod Med, 2006. 24(5): p. 304-13.
9. Wesselschmidt, R.L., The teratoma assay: an in vivo assessment of pluripotency.
Methods Mol Biol, 2011. 767: p. 231-41.
10. Lee, M.O., et al., Inhibition ofpluripotent stem cell-derived teratoma formation
by small molecules. Proc Natl Acad Sci USA, 2013. 110(35): p. E3281-90.
11. Mohseni, R., Hamidieh, Amir Ali, Verdi, Javad, Shoae-Hassani, Alireza, Safe
Transplantation of Pluripotent Stem Cell by Preventing Teratoma Formation.
Stem Cell Research and Therapy, 2014. 4: p. 212-218.
12. Kimbrel, E. A. and R. Lanza, Hope for regenerative treatments: toward safe
transplantation of human pluripotent stem-cell-based therapies. Reeen Med,
2015. 10(2): p. 99-102.
13. Fernandez Vallone, V.B., et al., Mesenchymal stem cells and their use in therapy:
what has been achieved? Differentiation, 2013. 85(1-2): p. 1-10.
14. Wei, X., et al., Mesenchymal stem cells: a new trendfor cell therapy. Acta
Pharmacol Sin, 2013. 34(6): p. 747-54.
52


15. Tang, M., et al., Human inducedpluripotent stem cell-derived mesenchymal stem
cell seeding on calcium phosphate scaffoldfor bone regeneration. Tissue Eng
Part A, 2014. 20(7-8): p. 1295-305.
16. Bajada, S., et al., Updates on stem cells and their applications in regenerative
medicine. J Tissue Eng Regen Med, 2008. 2(4): p. 169-83.
17. Liebergall, M., et al., Stem cell-based therapy for prevention of delayedfracture
union: a randomized and prospective preliminary study. Mol Ther, 2013. 21(8):
p. 1631-8.
18. Caplan, A.I., The mesengenicprocess. Clin Plast Surg, 1994. 21(3): p. 429-35.
19. Payne, K.A., D.M. Didiano, and C.R. Chu, Donor sex and age influence the
chondrogenic potential of human femoral bone marrow stem cells. Osteoarthritis
Cartilage, 2010. 18(5): p. 705-13.
20. D'lppolito, G., et al., Age-related osteogenic potential of mesenchymal stromal
stem cells from human vertebral bone marrow. J Bone Miner Res, 1999. 14(7): p.
1115-22.
21. Muschler, G.F., et al., Age- and gender-related changes in the cellularity of
human bone marrow and the prevalence of osteoblastic progenitors. J Orthop
Res, 2001. 19(1): p. 117-25.
22. Takahashi, K. and S. Yamanaka, Induction ofpluripotent stem cells from mouse
embryonic and adult fibroblast cultures by definedfactors. Cell, 2006. 126(4): p.
663-76.
23. Wemig, M., et al., In vitro reprogramming offibroblasts into a pluripotent ES-
cell-like state. Nature, 2007. 448(7151): p. 318-24.
24. Takahashi, K., et al., Induction ofpluripotent stem cells from adult human
fibroblasts by definedfactors. Cell, 2007. 131(5): p. 861-72.
25. Warren, L., et al., Highly efficient reprogramming topluripotency and directed
differentiation of human cells with synthetic modified mRNA. Cell Stem Cell,
2010. 7(5): p. 618-30.
26. Lapasset, L., et al., Rejuvenating senescent and centenarian human cells by
reprogramming through the pluripotent state. Genes Dev, 2011. 25(21): p. 2248-
53.
27. Hipp, J. and A. Atala, Sources of stem cells for regenerative medicine. Stem Cell
Rev, 2008. 4(1): p. 3-11.
28. Hynes, K., et al., Generation offunctional mesenchymal stem cells from different
induced pluripotent stem cell lines. Stem Cells Dev, 2014. 23(10): p. 1084-96.
53


29. Phillips, M.D., et al., Directed differentiation of human induced pluripotent stem
cells toward bone and cartilage: in vitro versus in vivo assays. Stem Cells Transl
Med, 2014. 3(7): p. 867-78.
30. Denham, M. and M. Dottori, Neural differentiation of induced pluripotent stem
cells. Methods Mol Biol, 2011. 793: p. 99-110.
31. Wang, S., et al., Differentiation of human induced pluripotent stem cells to mature
functionalPurkinje neurons. Sci Rep, 2015. 5: p. 9232.
32. Karumbayaram, S., et al., Directed differentiation of human-induced pluripotent
stem cells generates active motor neurons. Stem Cells, 2009. 27(4): p. 806-11.
33. Orimo, H., The mechanism of mineralization and the role of alkaline phosphatase
in health and disease. J Nippon Med Sch, 2010. 77(1): p. 4-12.
34. Xin, X., M. Hussain, and J.J. Mao, Continuing differentiation of human
mesenchymal stem cells and induced chondrogenic and osteogenic lineages in
electrospun PLGA nanofiber scaffold. Biomaterials, 2007. 28(2): p. 316-25.
35. Meinel, L., et al., Bone tissue engineering using human mesenchymal stem cells:
effects of scaffold material and medium flow. Ann Biomed Eng, 2004. 32(1): p.
112-22.
36. Wang, Y., et al., In vitro cartilage tissue engineering with 3D porous aqueous-
derived silk scaffolds and mesenchymal stem cells. Biomaterials, 2005. 26(34): p.
7082-94.
37. Song, S., et al., The synergistic effect of micro-topography and biochemical
culture environment to promote angiogenesis and osteogenic differentiation of
human mesenchymal stem cells. Acta Biomater, 2015. 18: p. 100-11.
38. Kawase, E., Efficient Expansion of Dissociated Human Pluripotent Stem Cells
Using a Synthetic Substrate. Methods Mol Biol, 2016. 1307: p. 61-9.
39. Battista, S., et al., The effect of matrix composition of 3D constructs on embryonic
stem cell differentiation. Biomaterials, 2005. 26(31): p. 6194-207.
40. Edwards, J.H. and G.C. Reilly, Vibration stimuli and the differentiation of
musculoskeletal progenitor cells: Review of results in vitro and in vivo. World J
Stem Cells, 2015. 7(3): p. 568-82.
41. Engler, A. J., et al., Matrix elasticity directs stem cell lineage specification. Cell,
2006. 126(4): p. 677-89.
42. Huang, C., J. Dai, and X.A. Zhang, Environmental physical cues determine the
lineage specification of mesenchymal stem cells. Biochim Biophys Acta, 2015.
1850(6): p. 1261-6.
54


43. Rowlands, A.S., P.A. George, and J.J. Cooper-White, Directing osteogenic and
myogenic differentiation o/MSCs: interplay of stiffness and adhesive ligand
presentation. Am J Physiol Cell Physiol, 2008. 295(4): p. C1037-44.
44. Wang, P. Y., W.B. Tsai, and N.H. Voelcker, Screening of rat mesenchymal stem
cell behaviour onpolydimethylsiloxane stiffness gradients. Acta Biomater, 2012.
8(2): p. 519-30.
45. Steward, A.J. and D.J. Kelly, Mechanical regulation of mesenchymal stem cell
differentiation. J Anat, 2014.
46. Yang, C., et al., Mechanical memory and dosing influence stem cell fate. Nat
Mater, 2014. 13(6): p. 645-52.
47. Eroshenko, N., et al., Effect of substrate stiffness on early human embryonic stem
cell differentiation. J Biol Eng, 2013. 7(1): p. 7.
48. Allen, J.L., M.E. Cooke, and T. Alliston, ECM stiffness primes the TGFbeta
pathway to promote chondrocyte differentiation. Mol Biol Cell, 2012. 23(18): p.
3731-42.
49. Evans, N.D., et al., Substrate Stiffness Affects Early Differentiation Events in
Embryonic Stem Cells. European Cells & Materials, 2009. 18: p. 1-14.
50. Pelham, R.J., Jr. and Y. Wang, Cell locomotion andfocal adhesions are regulated
by substrate flexibility. ProcNatl Acad Sci USA, 1997. 94(25): p. 13661-5.
51. Prager-Khoutorsky, M., et al., Fibroblast polarization is a matrix-rigidity-
dependent process controlled by focal adhesion mechanosensing. Nat Cell Biol,
2011. 13(12): p. 1457-65.
52. Macri-Pellizzeri, L., et al., Substrate stiffness and composition specifically direct
differentiation of induced pluripotent stem cells. Tissue Eng Part A, 2015. 21(9-
10): p. 1633-41.
53. Sebastine, I.M. and D.J. Williams, The role of mechanical stimulation in
engineering of extracellular matrix (ECM). Conf Proc IEEE Eng Med Biol Soc,
2006. 1: p. 3648-51.
54. Maniotis, A.J., C.S. Chen, and D.E. Ingber, Demonstration of mechanical
connections between integrins, cytoskeletalfilaments, and nucleoplasm that
stabilize nuclear structure. Proc Natl Acad Sci USA, 1997. 94(3): p. 849-54.
55. Sarasa-Renedo, A. and M. Chiquet, Mechanical signals regulating extracellular
matrix gene expression in fibroblasts. Scand J Med Sci Sports, 2005. 15(4): p.
223-30.
55


56. Wang, J.H. and B.P. Thampatty, An introductory review of cell mechanobiology.
Biomech Model Mechanobiol, 2006. 5(1): p. 1-16.
57. Li, J., et al., The role of extracellular matrix, integrins, and cytoskeleton in
mechanotransduction of centrifugal loading. Mol Cell Biochem, 2008. 309(1-2):
p. 41-8.
58. Chen, C.S., J. Tan, and J. Tien, Mechanotransduction at cell-matrix and cell-cell
contacts. Annu Rev Biomed Eng, 2004. 6: p. 275-302.
59. Schwartz, M. A., Integrins and extracellular matrix in mechanotransduction. Cold
Spring Harb Perspect Biol, 2010. 2(12): p. a005066.
60. MacKenna, D., S.R. Summerour, and F.J. Villarreal, Role of mechanical factors
in modulating cardiac fibroblast function and extracellular matrix synthesis.
Cardiovasc Res, 2000. 46(2): p. 257-63.
61. Barthes, J., etal., Cell microenvironment engineering and monitoring for tissue
engineering and regenerative medicine: the recent advances. Biomed Res Int,
2014. 2014: p. 921905.
62. Sabine Schulze, G.H., Matthias Krause, Deborah Aubyn, Vladimir A. Bolanos
Quinones, Christine K. Schmidt, Yongfeng Mei, and Oliver G. Schmidt,
Morphological Differentiation of Neurons on Microtopographic Substrates
Fabricated by Rolled-Up Nanotechnology. Advanced Biomaterials, 2010. 12(9):
p. 558-564.
63. Madri, J.A. and M. Marx, Matrix composition, organization and soluble factors:
modulators of microvascular cell differentiation in vitro. Kidney Int, 1992. 41(3):
p. 560-5.
64. Chu, L. and D.K. Robinson, Industrial choices for protein production by large-
scale cell culture. Curr Opin Biotechnol, 2001. 12(2): p. 180-7.
65. Marolt, D., et al., Engineering bone tissue from human embryonic stem cells. Proc
Natl Acad Sci USA, 2012. 109(22): p. 8705-9.
66. de Peppo, G.M., et al., Human embryonic mesodermal progenitors highly
resemble human mesenchymal stem cells and display high potential for tissue
engineering applications. Tissue Eng Part A, 2010. 16(7): p. 2161-82.
67. Trappmann, B., et al., Extracellular-matrix tethering regulates stem-cell fate. Nat
Mater, 2012. 11(7): p. 642-9.
68. Bustin, S. A., et al., The MIQE guidelines: minimum information for publication of
quantitative real-time PCR experiments. Clin Chem, 2009. 55(4): p. 611-22.
56


69. Hoemann, C.D., H. El-Gabalawy, and M.D. McKee, In vitro osteogenesis assays:
influence of the primary cell source on alkaline phosphatase activity and
mineralization. Pathol Biol (Paris), 2009. 57(4): p. 318-23.
70. Bruderer, M., et al., Role and regulation ofRUNX2 in osteogenesis. Eur Cell
Mater, 2014. 28: p. 269-86.
71. Born, A.K., S. Lischer, andK. Maniura-Weber, Watching osteogenesis: life
monitoring of osteogenic differentiation using an osteocalcin reporter. J Cell
Biochem, 2012. 113(1): p. 313-21.
72. Yousfi, M., et al., Increased bone formation and decreased osteocalcin
expression induced by reduced Twist dosage in Saethre-Chotzen syndrome. J Clin
Invest, 2001. 107(9): p. 1153-61.
73. Bialek, P., et al., A twist code determines the onset of osteoblast differentiation.
Dev Cell, 2004. 6(3): p. 423-35.
74. Kronenberg, H.M., Twist genes regulate Runx2 and bone formation. Dev Cell,
2004. 6(3): p. 317-8.
75. Pfaffl, M.W., A new mathematical model for relative quantification in real-time
RT-PCR. Nucleic Acids Res, 2001. 29(9): p. e45.
76. Gregory, C. A., et al., An Alizarin red-based assay of mineralization by adherent
cells in culture: comparison with cetylpyridinium chloride extraction. Anal
Biochem, 2004. 329(1): p. 77-84.
77. Song, W., Kawazoe, Naoki, Chen, Guoping, Dependence of Spreading and
Differentiation of Mesenchymal Stem Cells on Micropatterned Surface Area.
Journal of Nanomaterials, 2011. 2011.
78. McBeath, R., et al., Cell shape, cytoskeletal tension, andRhoA regulate stem cell
lineage commitment. Dev Cell, 2004. 6(4): p. 483-95.
79. Owen, T.A., et al., Progressive development of the rat osteoblast phenotype in
vitro: reciprocal relationships in expression of genes associated with osteoblast
proliferation and differentiation during formation of the bone extracellular
matrix. J Cell Physiol, 1990. 143(3): p. 420-30.
80. Phillips, J.E., et al., Glucocorticoid-induced osteogenesis is negatively regulated
by Runx2/Cbfal serine phosphorylation. J Cell Sci, 2006. 119(Pt 3): p. 581-91.
81. Weng, J. J. and Y. Su, Nuclear matrix-targeting of the osteogenic factor Runx2 is
essential for its recognition and activation of the alkaline phosphatase gene.
Biochim Biophys Acta, 2013. 1830(3): p. 2839-52.
57


82. Golub, E.E., Boesze-Battaglia, Kathleen, The role of alkaline phosphatase in
mineralization. Current Opinion in Orthopaedics, 2007. 18(5): p. 5.
83. Leboy, P.S., et al., Ascorbic acid induces alkaline phosphatase, type X collagen,
and calcium deposition in cultured chick chondrocytes. J Biol Chem, 1989.
264(29): p. 17281-6.
84. Dorst, K., Derek Rammelkamp, Michael Hadjiargyrou, and Yizhi Meng, The
Effect of Exogenous Zinc Concentration on the Responsiveness ofMC3T3-El
Pre-Osteoblasts to Surface Microtopography: Part II (Differentiation). Materials,
2014. 7(2): p. 1097-1112.
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i SUBSTRATE STIFFNESS INFLUENCES OSTEOGENIC DIFFERENTIATION OF INDUCED PLURIPOTENT STEM CELL DERIVED MESENCHYMAL STEM CELLS by KARL ALEXANDER TREADWELL B.S., University of Arizona, 2011 A thesis submitted to the Faculty of the Graduate Schoo l of the University of Colorado in partial fulfillment Of the requirements for the degree of Master of Science Bioengineering 2015

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ii This thesis for the Master of Science degree by Karl Alexander Treadwell has been approved for the Bioengineering Pro gram by Dae Won Park, Chair Karin Payne, Advisor Michael Yeager November 19, 2015

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iii Treadwell, Karl Alexander (M.S., Bioengineering) S ubstrate S tiffness I nfluences O steogenic D ifferentiation of I nduced P luripotent S tem C ell D erived M esenchymal S tem C ells Thesis directed by Assistant Professor Karin Payne. ABSTRACT Stem cells are able to differentiate into a variety of tissue specific cells, demonstrating a unique promise for tissue engineering and orthopaedic regenerative therapies. Routine culture tech niques utilize soluble factors that chemically influence differentiation, though the mechanical properties of the surrounding environment are proving to significantly impact cell differentiation as well Mesenchymal stem cells (MSCs), a widely studied cel l source for bone regeneration, have repeatedly demonstrated these mechanotransductive properties in vitro favoring relatively rigid substrates under osteogenic differentiation conditions. The effect of substrate stiffness on osteogenic differentiation o f mesenchymal progenitors derived from induced pluripotent stem cells (iPSC MPs), a novel cell source for bone regeneration, has not been widely studied. By culturing iPSC MPs on Polydimethylsiloxane (PDMS) sub strates with varying stiffness, we tested the hypothesis that comparatively rigid substrates could promote more efficient osteogenic differentiation than softer substrates. The stiffness of cultured on these substra tes were run against controls grown on tissue culture plastic (TCP) Effects were observed for cultures grown in both complete culture media (CCM) and osteogenic differentiation media (ODM) Gene expression at 3, 7, 14 and 21 days, quantified by RT qPCR, s hows significant upregulation of osteogenic markers, including r unt related transcription factor 2 ( RUNX2 ), alkaline p hosphatase ( ALP ), and o steocalcin

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iv ( OCN ) for cells cultured on 2.245 MPa substrates in ODM In comparison to cells cultured on softer su bstrates and especially TCP, these cells also displayed an increase in ALP protein activity and calcium deposition as determined by a lizarin r ed s staining and quantification. Cells cultured in CCM showed trends for osteogenic differentiation, suggesting that mechanical properties of the substrate may affect cell fate without chemical differentiation signaling. With recent studies showing the efficacy of iPSCs as a novel stem cell source for regenerative medicine, it is becoming ever more necessary to est ablish a fast and efficient method for differentiating these cells for orthopaedic applications. This study shows how substrate stiffness can be used to promote faster, more efficient osteogenic differentiation in iPSC MPs. This information could be used to design scaffolds that would promote bone formation in vivo The form and content of this abstract are approved. I recommend its publication. Approved: Karin Payne

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v TABLE OF CONTENTS CHAPTER I. Stem Cells ..02 .09 .13 Goals and Hypothesis II. Cell Culture Substrate Material Synthesis Material Properties Cell Plating Cell Proliferation Ac RT q PCR Pico Green & ALP Activity Alizarin Red III. Cell Spreading and Proliferation Gene Expression Calcium Deposition 38 IV. Mechanical Influence in CCM C V.

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1 C HAPTER I INTRODUCTION AND GOA LS According to the United States Bone and Joint Initiative (USBJI), there are approximately 223.6 million cases each year in the United States that involve diseases, disorders, and injuries related to bones, joints, and muscles [1] This result s in an estimated annual medical care cost of $212.7 billion [1] Bone injuries are a common occurrence in all age groups, but especially in individuals over the age of 50, where 1 in 2 women and 1 in 4 men will have an osteoporosis related fracture [1] With an aging population and prolonged life expectancy, the occurrence of bone injuries is only expected to increase. Despite advances in orthopaedic medicine, some f r actures remain difficult to t reat or do not heal well with 5 10% resulting in non unions and severe functional impairment [2] Stimulation of bone formation is of major clinical significance in orthopaedic procedures related to non unions and can also be effective in cases of spinal fusion joint fusion and for the repair of segmental bone defects. Implants and bone allografts have been widely used in orthopaedics to bridge the gap between the injured bones, but they lack a biological component that will allow successful integration into the surrounding native tissue. Thus, regenerative medicine techniques to promote bone stimulation have gained significant interest. Regenerative medicine focuses on developing therapies that will repair, replac e, or promote the regeneration of damaged or diseased tissue. One key component of regenerative medicine is the use of stem cells, which have the potential to become any of the tissue specific cells in the body Some of the current applications of stem c ells include therapeutic treatments of cancer, neurodegeneration, third degree burns, and loss of vision due to chemical destruction of the cornea [3] T he regenerative capabilities of

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2 stem cells have also shown promise for repairing cartilage, treating spinal cord injuries, and reversing damage caused by critical limb ischemia [4] However, there are many different kinds of s tem cells that can be obtained from different source s and can vary in their regenerative ability. Stem Cells Stem cells are the foundation for every organ and tissue in the body. They are characterized by the ability to self renew, creating identical daughter cells, and to differentiate into specific tis sues or organs. Though there are many different kinds of stem cells, they are classified based on their differentiation potential as either totipotent, pluripotent, or multipotent. Totipotent Stem Cells At the outset of embryonic development, two gametes (i.e. ovum and sperm) combine to form a zygote which contains all of the genetic information necessary for the embryo to develop. These are the first stem cells of the new organism and they are totipotent, meaning they have the potential to form any cell in the developing embryo, as well as the placenta. Only these early stem cells are totipotent, with truly un iversal differentiation potential. As the embryo develops, other kinds of more specialized stem cells form, which are defined by their commitment to given cell lineage s Pluripotent Stem Cells Pluripotent stem cells differ from totipotent cells only in that they can not form placenta. They retain the ability to form any one of the three germ layers, differentiating into any specialized cell of a s pecific tissue or organ They also maintain the capacity to proliferate indefinitely in a series of self renewals Embryonic stem cells (ESCs),

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3 embryonic germ cells (EGCs), and induced pluripot ent stem cells (iPSCs) all exhibit behavior s of pluripotency [3] ESCs were first derived from the inner cell mass of a mouse blastocyst in 1981 and subsequently from a human blastocy st in 1998 [5 7] Formed in the early development of mammals, the inner cell mass of the blastocyst is the structure that subsequently forms an embryo, which is the inherent source of controversy for ESCs, as the harvested blastocyst has the potential to grow into a full human being [3] T he embryonic germ cell serves as the progenito r of adult gametes and are found during late embryonic to early fetal development [8] I PSC s are special in that they are not derived from embryonic development and will be discussed in more detail on page 5 One potential consequence with the use of pl uripotent stem cells is the risk of teratoma formation. A teratoma is a nonmalignant tumor composed of cells derived from all three embryonic germ layers [9] With the inherent potential to form this disorganized mixture of tissue, there is a strong motivation for understanding the factors that control stem cell differentiation. U tilizing monoclonal antibodies, small molecules, anti angiogenic agents, suicide genes, and pharmacological agents significant progress has been made towards inhibiting unc ontrolled pluripotency [10 12] Further work that develop s protocols for the safe and effective differentiation of these cells will greatly facilitate the advance ment of stem cell base d therapies Multipotent Stem Cells Multipotent stem cells are more specialized than pluripotent stem cells and include adult stem cells that replenish dying cells or repair damaged tissue Multipotent cells can be harvested from a variety of sources, but have limi ted proliferative ability Depending on the type of multipotent cell, t hey are able only to form the differentiated

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4 cell types of a specific tissue. These include neural stem cells ( which form neurons, astrocytes, and oligodendrocytes), endothelial stem cells (blood vessel s), hematopoietic stem cells (blood cells), mesenchymal stem cells (bone, cartilage, muscle, and fat) and many others In the interest of orthopaedic research, discussions will be limited to mesenchymal stem cells Mesenchymal Stem Cells Mesenchymal Ste m Cells (MSCs) are a type of multipotent stem cell that can differentiate into a variety of cell types and maintains the ability to self renew, however at a limited capacity. Human MSCs are defined as being plastic adherent, expressing the surface molecul es CD105, CD73, and CD90 and showing multilineage differentiation in vitro towards the osteogenic (bone), adipo genic (fat), and chondro genic (cartilage) lineages though they are not limited to these cell lines [13] MSC therapies are among the most common of stem cell applications, as they are free of ethical harvesting concerns have numerous sources, show little risk of producing an immunogenic response, and virtually n o risk of teratoma formation (unlike pluripotent cells) [14] They are being investigated for treating a varie ty of diseases including myocardial infarction, liver cirrhosis, diabetes, spinal cord injuries, osteoarthritis, and many more [14] MSCs derived from harvested bone marrow (BM MSCs) are the most commonly studied stem cells for orthopaedic regenerative medicine. In cases of trauma, abnormal development, and congenital malformations, MSCs have shown particular pr omise for scaffold seeding and inciting bone and cartilage regeneration. Scaffold implants provide support in bone and cartilage defects, while serving as a template for seeded MSCs to

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5 encourage growth and regeneration of the damaged tissue [15] They have been shown to enhance bone healing in both small and large animal models whether seeded on scaffolds or implanted directly [16] They have also shown some success clinically, when used for t he treatment of non unions in distal tibial fractures, demonstrating no adverse effects and reducing union time by half [17] The clinical application of stem cell therapies appears limitless, but there are a num ber of issues surrounding the harvest and use of stem cells. For example, the proportion of BM MSCs obtained from bone marrow aspiration is less than 0.01% [16, 18] Translation of BM MSC technology could is also limited by the fact that human MSCs cannot be expanded indefinitely in culture, and their differentiation capacity has been reported to decrease with aging and aging related diseases [19 21] This can limit their therapeutic potential in bone regenerative medicine, especially in older patients. Thus, t here is an emerging interest in the identification of alternative cell sources for MSCs. Induced Pluripotent Stem Cells In 2006, Kazutoshi Takahashi and Shinya Yaman aka demonstrated that by introducing four factors (Oct3/4, Sox2, c Myc, and Klf4) to adult mouse fibroblasts cultured under ESC culture conditions, they could induce pluripotency in the fibroblasts [22] The repro gramming of fibroblasts into what has been designated as induced pluripotent stem cells (iPSCs) was a scientific breakthrough that introduced a novel stem cell source that could potentially revolutionize regenerative medicine. iPSCs are advantageous to cu rrent stem cell sources because they can be generated directly from the

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6 immune system, making the use of immunosuppressants for preventing rejection unnecessary. Th is inherently means that they can serve as an autologous stem cell source, while negating the moral ambiguity with using ESCs. Beyond what Takahashi and Yamanaka achieved in 2006, researchers have since improved reprogramming techniques and shown that iP SCs can be reliably differentiated down a multitude of cell lineages. In 2007, Wernig et al. significantly improved the techniques that were developed a year earlier, and showed that DNA methylation, gene expression and chromatin state of the iPSCs we re s imilar to those of ES Cs [23] In subsequent years, Takahashi furthered his research, demonstrating similar pluripotency induction in adult human fibroblasts and in 2010 Warren et al. devised a safer, more effectiv e reprogramming method based on mRNA administration [24, 25] Moreover, cellular reprogramming of skin fibroblasts from older individuals has been reported, suggesting that iPSCs may provide a source of rejuvenate d adult stem cells for patients that may not have optimal BM MSCs [26] These findings could prove to be very advantageous for bone repair. IPSCs could inherently become any cell in the body but although this is e xciting for regenerative medicine, this pluripotency comes with the risk of forming teratomas upon implantation [27] Thus it is important to ensure complete differentiation of the iPSCs towards the desired lineage in vitro before implantation in vivo T he generation of functional iPS derived MSC s has been demonstrated as well as furth er differentiation towards bone and cartilage both in vitro and in vivo [28, 29] Furthermore, d ifferentiation of iPSCs towards a neuronal lineage has also been studied extensively, with even more specific generat ion of both Purkinje neurons and motor neurons [30 32]

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7 The scope of this study is to observe osteogenic differentiation, pushing iPSC derivatives to generate bone, so it is important to understand the structure of bone and the different cell types that make up osseous tissue. Bone Bone is a rigid organ of the vertebral skeleton that supports and protects various other organs of the body. Osteoblasts osteoclasts, and osteocytes are the three typ es of cells that make up bone, forming either cortical or cancellous (spongy) tissue. Cortical bone forms the outer shell of most long bones and is much denser than cancellous bone serving as the primary supportive structure of the organ The foundational structure of cortical bone is the osteon, which consists of layers of mesenchymal derived osteoblasts that are responsible for synthesizing new bone. Osteoblasts bud vesicles containing hydroxyapatite, a calcium phosphate, to be deposited between collagen fibrils of t he extracellular matrix, which leads to the formation of new bone [33] Once these cells b ecome trapped by the bone matrix that they deposit, they differentiate into osteocytes that reside in little voids known as lacunae and are responsible for directing routine bone turnover. Osteocytes communicate through canals known as canaliculi that co nnect the lacunae, directing osteoblasts to lay down new bone and osteoclasts to resorb old bone. Unlike osteoblasts and osteocytes, osteoclasts are derived from monocytes. The blood supply for the bone is transported through osteonic canals known as Hav ersian and Vo l s of cancellous tissue, made up of a porous network of thin osteoblast formations resembling the structure of a sponge Within the spaces of the cancellous bone resides the bone marrow and he matopoietic

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8 stem cells that give rise to platelets and red and white blood cells. A diagram of long bone structure can be seen in Figure 1.1 Figure 1. 1 Structural anatomy of a long bone. Regenerative orthopaedic therapie s aim to utilize MSCs in order to generate viable osteoblasts to enhance bone formation in cases of breaks and malunions. In spite of this potential, current culturing techniques are rather expensive, time consuming, and can yield non homogenous cultures under differentiation. In regards to scaffold seeding, it has been shown that scaffold design and material choice can have varied effects on stem cell differentiation alluding to an environmental influence on cell behavior [34 36] For these reasons and more, it is becoming more crucial to understand what factors are involved in the growth, proliferation, and differentiation of bone cells and what can be done to improve culturing techniques. The next section will provide an overview of the effects the mechanical environment can have on stem cell behavior.

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9 Mechanosensing and Mechanotransduction One growing area of stem cell research is investigating how the cellular microenvironment affects stem cell activity. Res earchers have documented the effects of surface micro topography, biochemical environment, substrate material, three dimensional scaffold constructs, and mechanical vibration on the growth and differentiation of stem cells [37 40] In particular, the mechanical properties of the surrounding environment have been foun d to affect many cellular processes in a variety of cell types. It has been demonstrated that nave MSCs show extreme sensitivity to tissue level matr ix elasticity, becoming neurogenic on soft substrates, myogenic on stiffer substrates, and osteogenic on a comparatively rigid substrate [41] MSCs, in particular, have been shown to become either osteogenic chon drogenic, myogenic, or adipogenic due to matrix elasticity specification, and have even shown memory response to mechanical dosing influence [42 46] Substrate influence has also been observed in mesodermal differ entiatio n of human embryonic stem cells and subsequent terminal differentiation for both chondrogen ic and osteogen ic lineages [47 49] It has shown regulatory influences on kidney cell morphology and locomotion, a s well as fibroblast polarization [50, 51] Even iPSCs have shown mechanotransductive properties, demonstrating neural and cardiac differentiation when plated on a soft matrix (0.6 kPa) [52] This inherent ability of cells to recognize variations in matrix stiffness is clearly an important factor in the regulation of stem cell fate. The mechanism by which cells can feel their surrounding environment and convert this stimulus into an electrochemical response is referred to as m echanotransduction.

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10 Extracellular Matrix Integrin Cytoskeleton Mechanosensing Pathway The Extracellular Matrix Integrin Cytoskeleton (EIC) mechanosensing pathway allows for mechanical stresses or vibrations to be rapidly transferred from cell surface re ceptors to distinct structures in the cell and nucleus, resulting in regulation of cellular functions such as cell attachment, proliferation, migration and differentiation [53] This mechanosensing occurs as cellu lar integrins bind to the extracellu lar matrix A s the cell contracts and pulls on the surface, the cytoskeletal filaments reorient themselves, resulting in nucleus distortion and redistribution of the nucleolus (Figure 1. 2 ) [54] This Figure 1. 2 Mechanotransduction. The cell attaches to the extracellular matrix (ECM) through attachment points called integrins. If the cell is attached to a relatively rigid substrate (left), then as the cell contracts, cytoskeletal (CKS) tension goes up resulting in nuclear redistribution. If the ECM is flexible (right) the cell can more readily pull on the Cellular mech anotransduction: putting all the pieces together again The FASEB Journal vol. 20. no. 7. Pgs. 811 827. 2006. pathway can be activated not only through cell cell and cell matrix interactions, but also by gravit ational forces. This e ffect is evident when astronauts develop osteopenia (a decrease in bone density) as a result of prolonged space flights [55] Studies have shown how mechanical stimulation can influence cellular activity in a variety of cells including dermal and cardiac fibroblasts, cardiac myocytes, endothelial cells, and bone and cartilage cells [56] Most studies culture cells on flexible substrates, or apply direct force to adhesion molecules to observe cellular responses, though others have used

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11 centrifugation to simulate gravitational changes [57, 58] The sections below will briefly discuss the roles of each component in the EIC mechanosensing pathway. Extracellular Matrix The extracellular matrix (ECM) provides primary structural and biochemical support for the cell, but the stiffness of the ECM can have an influence on cell behavior as well. Focal adhesions bind to the ECM and as the cell contracts, the ECM deforms by an amount relative to i ts mechanical properties This provides a certain amount of give that the cell is capable of detecting. In many cases, cells respond by remodeling the ECM through newly activated/deactivated gene expression, or by direct mechanical manipulation of the ECM fibrils [59] The newly remodeled matrix is most often more resistant to the applied forces. Integrins Integrins are transmembrane molecular structures that bind cells to the extrace llular matrix and have been shown to be an integral part of cellular mechanosensing. They serve as the structural mediators between the ECM and the cytoskeleton of the cell. When introduced to a mechanical stress integrins can either transduce the fo rce into a chemical response or transmit the force directly to the cytoskeleton This stimulates downstream reactions but the specific processes and targets of each of these stimulation s are largely unknown [53, 55 ] It is hypothesized that the mechanical signals can be used to induce conformational changes in the integrins themselves and several studies have shown force application to cause the a ttached focal adhesions to stabilize and increase in size and streng th [58, 59]

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12 Cytoskeleton The cytoskeleton is made up of a network of microfilaments and serves as the supportive structure of the cell, both generating force and bearing elastic deformation [55] Just as muscles pull on specific parts of bones, physical manipulation can influence the cytoskeleton to mechanically distort specific parts of the nucleus, inducing changes in cellular organization and gene expression [54] Additionally the simulated effects of gravity can be studied with centrifugation. Li et al. showed that osteoblasts exposed to mild centrifugal force exhibited temporary and reversible changes in gene transcription which helped to verif y the role of the cytoskeleton in mechanosensing [57] Cellular and Physiological Responses Though th e mechanism for how mechanical energy is transduced into chemical changes in the cell is not fully understood, new studies are emerging that are beginning to map the chemical pathways of mechanotransduction. For example, the application of mechanical stre ss has been shown to increase f ocal a dhesion k inase phosphorylation and specifically, Rho kinase is hypothesized to be affected by stress as it is heavily involved in the generation of cytoskeletal tension [58] Still o ther studies have hypothesized that some mechanisms may inf luence cellular structures directly (i.e. ion channels, nuclear pores, chromosomes, individual genes, etc.) remaining independent of chemical signaling mechanisms [54] As investigations of the EIC mechanosensing pathway continue, the relevance of its application becomes more apparent. Cardiac fibroblasts have been shown to increase ECM gene expression and growth factor activation when exposed to mechanical stress in vitro and cyclic loading has been shown to ind uce me mbrane matrix metalloproteinase production similar to that found in ischemic hearts [60] ECM restructuring, influenced

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13 by cyclic stretch, is a vital process in the strengthening of vascular smooth muscle cells, which would otherwi se result in an aneurysm [59] O thers have hypothesized that mechanose nsing can be relevant in cases of hypertrophic scarring as well as tendon and ligament regeneration [55] Further studies into how cells transduce mechanical signals into biochemical changes would progress the understanding of mechanosensing, giving a better visualization ing greater knowledge for the culture and control of cel lular activity. The Cellular Environment ability to sense the mechanical properties of their environment presents a wide variety of stimuli that can affect how cells grow, proliferate, and differentiate R esearch with regards to the microenv ironme nt includes surface micro nanotopography, soluble factors, extracellular matrix composition a nd distribution and mechanical stress/strain conditions [61] Each has been shown to influence the efficacy of cell cul ture, which has major implications for tissue engineering and regenerative medicine. Schulze et al. demonstrated how primary mouse motor neurons were successfully guided and directed towards growth of axons into grid like neurite networks when cultured on microtopographical substrates coated with rolled up SiO/SiO 2 [62] Similarly, human MSCs cultured on 10 m micropost textured PDMS demonstrated higher gene expression of osteoblast specific markers accompanied by substantial bone matrix formation and mineralization when compared to smooth surface controls [37] T he se results were amplified when biochemical soluble factors intended for osteogenic differentiation were added to the culture media [37] Soluble factors have also been shown to improve cell survival and proliferation as well as induce cellular vascularization, regenerate neurons, and even maintain a stem cell phenotype [61]

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14 Final ly, matrix composition variability can have significant effects on the differentiation of stem cells and can modulate the phenotype of both endothelial and mesangial cell populations in culture [39, 63] The compi lation of all this data can be utilized to design bioreactors which mimic the mechanical microenvironment for functional engineered tissues that undergo mechanical loading under in vivo conditions (i.e. cartilage, bone, tendons, heart valves, etc.) which is becoming the standard for large scale cell culture [61, 64] Goals and Hypotheses IPSCs like ESCs show particular potential for regenerative medicine due to their ability to proliferate and differentiate i nto cells of all three germ layers, yet they are more attractive due to their lack of controversy and virtually limitless availability In 2014, Tang et al. demonstrated the promise for iPSCs to promote bone regeneration through cell seeding of calcium ph osphate cement scaffolds and subsequent osteogenesis [15] Procedures such as this require a homogeneous bank of mesenchymal stem cells derived from iPSCs, referred to as mesenchymal progenitors (iPSC MPs), demons trating the need for efficient methods of iPSC differentiation. This, in conjunction with the influence of mechanotransduction and the cellular environment presents a motivation for studying the effects of substrate stiffness on iPSC MP differentiation a nd osteogenesis. The goal of this study is to observe the effects of culturing iPSC MPs on p olydimethylsiloxane (PDMS) substrates of varying stiffness and cultur ing in either control MSC medium or osteogenic differentiation medium with the hypothesis tha t relatively stiffer substrates will influence faster, more homogenous osteogenic differentiation especially compared to current culture methods

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15 CHAPTER II MATERIALS AND METHODS Cell Culture Induced pluripotent stem cells (iPSCs) were generated from hum an f ibrobla s ts collected from the skin biopsy of a 50 year old female The fibroblasts were reprogrammed using an optimized mRNA based approach previously described by Warren et al. [25] Briefly, mRNA molecules encoding the reprogramming factors Oct4, Sox2, Klf4, and c Myc were introduced into the fibroblasts by direct transfection Generated iPSCs were positive for the expression of endogenous p l uripotency markers ( OCT4 NANOG TRA 1 81 ) by immunofluorescence and were shown to display a normal karyotype iPSCs were then differentiated into mesenchymal progenitors (iPSC MPs) by a method previously described by Marolt et al. [65] In brief, iPSCs were cultured in inducti on medium (Knockout DMEM (Life Technologies, Carlsbad, CA, USA) supplemented with 20% FBS (Atlanta biologicals, Lawrenceville, GA, USA), 2 mM glutagro supplement (Corning, Manassas, VA, USA), 0.1 mM non essential amino acids (Corning, Manassas, VA, USA), 0 .1 mM 2 Mercaptoethanol (Fisher Scientific, Fair Lawn, NJ, USA), and 1% penicillin streptomycin (Thermo Scientific, Logan, UT, USA)), hereby referred to as MSC differentiation media for one we ek and passaged until they achieved a homogenous fibroblast lik e morphology under microscopic examination. Mesenchymal differentiation was confirmed by flow cytometry for mesenchymal markers CD90, CD73, and CD105 (BD Biosciences, San Jose, CA, USA) when compared to bone marrow mesenchymal stem cells (BM MSCs) [65, 66] Cells were then collected, frozen down as cell stocks, and placed in liquid nitrogen storage.

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16 Cells were thawed after being removed from storage and plated in a 225cm 2 tissue culture treated flask with 30 mL of MSC differentiation media Cells were incubated at 37C, 5% CO 2 expanded and plated for differentiation studies at passage 9 PDMS culture trials were conducted three times, using frozen cell stocks from the same set of iPS C MPs. Substrate Material S yn thesis PDMS substrates were fabricated using a two part silicone elastomer kit (SYLGARD 184, Dow Corning Midland, MI, USA) which was chosen for its biocompatibility and regular use in mechanotransductive differentiation studies [37, 44, 47, 67] The stiffness of each substrate was varied by altering the base to curing agent ratio. Five s ubstrates were characterized by the percentage of curing agent that made up the final solution (4%, 8%, 12%, 16%, and 20%) Ea ch substrate was mixed and degassed and then 2 mL of the final solution was transferred in triplicate to 6 well plate s and allowed to cure at room temperature for at least 60 hours. Substrates were then rinsed in 70% ethanol and allowed to air dry. Thr ough a method previously described by Pelham and Wang T ype I collagen was covalently crosslinked to the PDMS surface with N Sulfosuccinimidyl 6 (4' azido 2'nitrophenylamino) hexanoate (Sulfo SANPAH, CovaChem Loves Park, IL, USA) [50] A 200 mg/mL solution of Sulfo SANPAH was created by dissolving in dimethylsulfoxide (DMSO, Sigma Aldrich, St. Louis, MO, USA) which was further diluted with 50 mM 4 (2 hydroxyethyl) 1 piperzaineethanesulfonic acid ( HEPES, GE Healt hcare Life Sciences, Logan, UT, USA) pH 8.5, to a final concentration of 0.5 mg/mL. The Sulfo SANPAH solution was used to cover the PDMS substrates, which were then exposed to UV light from a transilluminator for 30 minutes in a cell culture

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17 hood ( Model : 1385 Thermo Fisher Scientific, Marietta, OH, USA). Excess Sulfo SANPAH solution was aspirated and plates were exposed to UV light for an additional 30 minutes. A 0.05 mg/mL collagen concentration was made by diluting Collagen Type I, Rat Tail ( Corning Life Sciences Tewksbury MA, USA) in 0.02 N acetic acid. Wells were washed three times in sterile phosphate buffered saline (PBS) and 1 mL of the collagen solution was added to each well and allowed to incubate overnight at 4C. Collagen solution was a spirated and wells were washed twice with sterile PBS. Plates were wrapped in P arafilm M (Bemis NA, Neenah, WI, USA) and stored at 4C until needed. PDMS substrates were fabricated separately for each of three trials to account for variability between bat ches Material Properties Material properties of PDMS samples were measured by displacement controlled nanoindentation using a HYSITRON TI 950 TriboIndenter (Hysitron, Minneapolis, MN, USA). A 250 m radius sapphire spherical tip ( Ti 0185, Hysitron ) was u sed to indent to a depth of 4000 nm at a rate of 800 nm/s where it was held for 30 seconds before retracting at the same rate. Force measurements were read in a 5 5 point matrix with an even separation of 200 m. The Hertz model with a Poisson ratio of 0 .5 was used to calculate based on the retracting force curves with the assumption that the sphere had an infinitely larger E than the PDMS samples. Testing was conducted on nine samples from each condition (three trials of triplicate s amples), and averages of the samples are reported along with standard deviations.

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18 Cell Plating and Culture iPSC MP s were plated from a single cell suspension at a density of 30,000 cells/well on 6 well PDMS coated plates and were run against controls pla ted at the same density on tissue culture plastic (TCP) Cells were cultured in either Complete Culture Media (CCM, MEM alpha (Life Technologies, Carlsbad, CA, USA) supplemented with 16.5% FBS (Atlanta biologicals, Lawrenceville, GA, USA) 2 mM L glutamin e (Corning, Manassas, VA, USA) and 1% penicillin streptomycin (Thermo Scientific, Logan, UT, USA ) or Osteogenic Differentiation Media (ODM, CCM supplemented with 10 nM dexamethasone (Sigma, St. Louis, MO, USA) glycero phosphate (Sigma, St. Louis, MO, USA) ascorbic acid 2 phosphate (Fisher Scientific, Fair Lawn, NJ, USA) ). Cells were incubated at 37C, 5% CO 2 and media was changed three times per week (M ondays W ednesdays and F ridays ). Cells were collected at 3 7 14, and 21 day ti me points Culture trials were cond ucted three times in triplicate. Cell Proliferation The alamarBlue assay (Thermo Scientific, Logan, UT, USA) incorporates a fluorometric oxidation reduction indicator that reacts in response to the chemical reduction of growth medium indicating the metabolic activity of cellular growth. A repeated measure of this assay was conducted at 3, 7 14 and 21 day time points. A 100% reduced positive control was made by autoclaving 5 mL of CCM with 500 L of alamarBlue reagent One milliliter of this solution was added to one well of a 6 well plate with 100 L of ultrapure water. Negative control was made of 1 mL of CCM only. The respective media for each experimental sample (either CCM or ODM) was aspirated and replaced with 1 mL of fresh media plus 100 L of the alamarBlue reagent. S amples

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19 including controls, were incubated at 37C, 5% CO 2 for 2.5 hours, then 100 L of each sample was plat ed in triplicate on a 96 well plate and fluorescence measurements ( 530 560 nm excitat ion wavelength and 590 nm emission wavelength ) were read using a Glomax Multi Detection System plate reader (Promega, Madison, WI, USA). After reading, 6 well sample plates were washed once with PBS and returned to culture with 2 mL of their respective me dia for future readings. To calculate the percent r eduction of the alamarBlue r eagent r elative f luorescence u nits (RFU) for each sample were averaged and used in the following equation: ( 1 ) A lamarBlue analysis was conducted at four time points over the course of one culture trial, though proliferation was monitored regularly with microscopy over three culture trials Actin Staining After 3 days of culture, cells were washed twice with PBS and fixed in 10% neutral b uffered f ormalin for 10 minutes at room temperature. Wells were washed two more times with PBS and plates were wrapped in Parafilm M and stored at 4C in PBS. Phalloidin (F actin) staining was performed by applying ActinGre en 488 ReadyProbes reagent (Life Technologies, Eugene, OR, USA) to fixed cells using 2 drops / mL of PBS and incubated for 30 minutes at room temperature. Wells were washed with PBS and imaged with an Eclipse TE2000 S (Nikon, Melville, NY, USA). Phalloidin staining was performed on one set of samples from a single culture trial.

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20 RT qPCR RT qPCR strategies were conducted in adherence to the guidelines for the Minimum Information for Publication of Quantitative Real Time PCR Experiments (MIQE) [68] Cells were collected for reverse transcription polymerase chain reaction (RT qPCR) at 3, 7, 14, and 21 day instructions for the RNeasy Plus Mini Kit (Q iagen Valencia CA, USA). Isolated RNA was quantified using an Epoch Microplate Spectrophotometer and a Take3 plate (BioTek, Winooski, VT, USA) and diluted with sterile DNase and RNase free water (Fisher Scientific, Fair Lawn, NJ, USA) to a final concentration of no m Isolated RNA samples were stored at 80C until cDNA reverse transcription. for the High Capacity cDNA Reverse Transcription Kit ( Life Technologies, Carl sbad, CA, USA ), including RNase inhibitor. Samples were once again quantified with the Epoch Microplate Spectrophotometer and Take3 plate and diluted to a concentration of 80C until RT qPCR analysis. Following manu the SsoAdvanced SYBR Green Supermix (Bio Rad Laboratories, Inc., Hercules, CA, USA), RT qPCR was carried out using primers at a final concentration of 500 nM and 100 ng of the DNA template per reaction. Target genes were select ed as markers of osteogenesis and include alkaline phosphatase ( ALP ), TWIST runt related transcription factor 2 ( RUNX2 ), and osteocalcin ( OCN ). Glyceraldehyde 3 phosphate dehydrogenase ( GAPDH ) was used as a housekeeping gene for normalizing expression an d has been used in this capacity previously during osteogenic differentiation experiments [49] ALP is typically used as

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21 an early marker of osteoblastic differentiation whose expression has been shown to increase over time in confluent monolayer bone derived cell cultures [69] ALP also plays a direct role in mineralization, hydrolyzing several phosphates in order to prepare ions for deposition [33] RUNX2 is another osteogenic related marker shown to regulate the non collagenous, late stage osteoblastic marke r OCN [70, 71] Finally, TWIST is a basic helix loop helix transcription factor that, as its expression decreases, has been shown to increase ALP and RUNX2 expression, and was chosen for its use in previous osteo genic differentiation studies [49, 72 74] Efficiencies for GAPDH ALP OCN RUNX2 and TWIST were 112%, 101.5%, 114%, 109%, and 103% respectively. Temperature cycling was conducted using a CFX Connect Real Time System (Bio Rad Laboratories, Inc., Hercules, CA, USA) and cycling conditions for all genes can be seen in Table 2 .1. Table 2 .1 Cycling Conditions for cDNA Cycling Step Temperature Time # Cycles Enzyme activation/Initial DNA denaturation 95C 30 sec 1 Denaturation 95C 5 sec 40 Annealing/Extension 60C 30 sec 40 Melt Curve 65 95C (in 0.5C increments) 5 sec/step 1 Results were analyzed using the Bio Rad CFX Manager software, which utilized the mathematical model for relative quantification describ ed by Pfaffl, where the efficiency of a primer, E, was related to percent efficiency in the following way [75] : ( 2 )

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22 In this way, an E of 2 was equal to 100% efficiency and represented perfect doubling with every cycle. Using a Day 0 sample as control, the relative quantity (RQ) of any experimental sample for a given gene of interes t (GOI) was calculated as: ( 3 ) where C q (control) = Average C q for the control sample C q (sample) = Average C q for experimental sample s The relative quantities of the samples were normalized to GAPDH expression with the following equation: ( 4 ) Results were compared using one way ANOVA analysis with a post hoc Tukey test and differences were considered significant when p < 0.05. RT qPCR was conducted on triplicate samples over three culture trials. Results are displayed as the representative average of the triplicate of one trial with error bars representing standard error of the mean Pico Green & ALP Activity Samples were lysed at 14 and 21 days and alkaline phosphatase (ALP) activity was quantified in an enz yme immunoassay Triton X 100 (Sigma, St. Louis, MO, USA) was diluted to a concentration of 0.1% in distilled water to create the cell lysis buffer Samples were first washed with PBS, then 250 L of the Triton X 100 buffer was added to each well and sam ples were incubated for 30 minutes at 4C. Plates were then wrapped in Parafilm M and stored at 20C over night.

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23 Preparing the SigmaFast p Nitrophenyl Phosphate (pNPP, Sigma, St. Louis, MO, USA) solution require d vortexing 1 pNPP tablet with 1 Tris tabl et in 20 mL of dH 2 O. Experimental samples were plated in triplicate in 96 well plates with 100 L of pNPP solution, 90 L dH 2 O, and 10 L of lysed sample. O ne triplicate of lysis buffer was run alone to control for background value. P lates were wrapped in aluminum foil to protect from light and incubate d for 30 minutes at room temperature before measuring absorbance in the Epoch Microplate Spectrophotometer at 405 nm. ALP activity for each sample was normalized to its DNA content by first quantifying D NA concentration using the Quant iT PicoGreen dsDNA Assay (Life Technologies, Carlsbad, CA, USA) a s summarized below. To create a standard curve, a serial dilution of the provided DNA standard was made with concentrations increasing two fold from 15.625 n g/mL to 1000 ng/mL, and included a null sample. Samples were diluted 1:10 in 0.01% Triton X 100 100 L of the standards and samples were added in duplicate, to 100 L of PicoGreen solution (0.05% PicoGreen in 1X TE Buffer provided in kit) in a black 96 well plate. As a reference blank, a duplicate of 1:1 PicoGreen Solution: Cell Lysis Buffer was plated alongside the samples. After 5 minutes of incubation at room temperature wrapped in aluminum foil, fluorescence intensity i n the blue spectrum (480 n m excitation/520 nm emission) was measured with the Promega Glomax Multi Detection System. After duplicates were averaged, ALP activity [ ALP act determined from the equation: ( 5 ) and normalized [ALP norm nmol pNPP/g DNA] to DNA content using the equation:

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24 ( 6 ) where DNA quant was determined from the PicoGreen fluore scence readings. From the standard curve, a linear relationship between fluorescence and DNA quantity was determined in the form: OR ( 7 ) where a and b are constants determined by the standard curve. ALP activity and PicoGreen analysis was for one culture trial and compared against RT qPCR results. Average s of the triplicates and standard deviation s are presented. A lizarin Red S Alizarin r ed s ( ARS, Sigma, St. Louis, MO, USA) was used to stain calcium deposits which certain osteogenic cell lines, specifically osteoblasts, form in culture Samples were fixed and stained at 7, 14 and 21 day time points. Media was as pirated from wells, which were then rinsed with 2 mL of PBS before incubating for 60 minutes at room temperature in 2 mL of 10% Buffered Formalin (Sigma, St. Louis, MO, USA). Formalin was aspirated and wells were rinsed with dH 2 O before incubating for 20 minutes at room temperature in 2 mL of ARS and finally rinsed 3 4 times with dH 2 O Wells were imaged with an AMG EVOS XL Core Microscope ( Fisher Scientific, Fair Lawn, NJ, USA ) and plates were scanned with a LaserJet Pro 500 M570dn (Hewlett Packard, Palo Alto, CA, USA). Wells were covered in dH 2 O and p lates were wrapped in Parafilm M for storage at 4C until quantitative destaining. For quantitative destaining, a protocol was followed that was originally developed by Gregory et al [76] W ater was aspirated from ARS stained wells and 800 L of 10% acetic acid was added t o each well before 30 minutes of incubation at room temperature with gentle shaking. The c ell layer was detached with a cell scraper and transferred along

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25 with the acetic acid to a centrifuge tube where it was vortexed, heated to 85C for 10 minutes, and put on ice for 5 minutes. The samples were then centrifuged at 20,000 xg for 15 minutes and 500 L of the supernatant was neutralized to a pH between 4.1 and 4.5 with 200 L of 10% ammonium hydroxide. 150 L of each sample was plated in triplicate to a 9 6 well plate and read at 405 nm with the Epoch Microplate Spectrophotometer along with the a lizarin r ed s tandards listed in Table 2 .2 Table 2 .2 Alizarin Red Serial Dilutions. ARS solution is originally 29.2141 mM concentration and is diluted with buf fer made from 7.5 mL of 10% acetic acid and 3 mL of 10% ammonium hydroxide. A 2 mM working stock solution was made by adding 342 L ARS to 5 mL of dilution buffer which was then diluted two fold in series for high range standards. 15 L of the 2 mM worki ng stock was added to 985 L of dilution buffer to create a 30 M concentration which was then diluted two fold in series for low range standards. Range Concentrations High 2 mM 1 mM 500 M 250 M 125 M 62.5 M 31.3 M 0 Low 30 M 15 M 7.5 M 3.75 M 1 .88 M 0.94 M 0.47 M 0 The standard curve generated a linear relationship between absorbance A, and quantification, AR quant After subtracting the blank value, the following equation was applied to determine quantity of ARS stain based on absorbance : OR ( 8 ) Where a and b are constants determined by the standard curve Equation 8 was used to determine qua n tification of ARS ARS staining was conducted for all three culture trials at the 21 day time point and once after 14 days of culture, each time in triplicate. Results were consistent across all trials and representative images are presented. Quantitative destaining was conducted on one set of triplicate samples and displayed as an average of the three with error bars representing the standard deviation.

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26 Statistical Analysis To compare groups, assumptions of parametric data were tested using Shapiro r homogeneity of variance. For parametric data, one way ANOVA and Tukey p ost hoc analysis was used. For non parametric data, a Kruskal Wallis test was used with a post hoc pairwise Mann Whitney U with a Bonferroni correction. All analyses will be perfor med w ith SigmaPlot v.11.2 Significant differences are reported with a p value < 0.05

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27 CHAPTER III RESULTS Substrate Stiffness Substrates of PDMS were fabricated by mixing different percent concentrations of the crosslinking component of the two part e lastomer. For the rest of this work samples will be referred to by the percent concentration of the crosslinker in the substrate on which the cells were plated (4%, 8%, 12%, 16%, 20%, and TCP for reference). Nanoindentation results showed elastic m odulu s to be 0.679 0.42 2 1.1 82 0.2 51 1.7 1 30.3 28 2. 245 0.3 09 and 2. 447 0. 459 MPa for 4%, 8%, 12%, 16%, and 20% substrates, respectively (Figure 3.1) A one way ANOVA statistical analysis showed all samples to be significantly different from each other (p<0 .05) except for 16% and 20% Figure 3. 1 Material Properties for PDMS Samples. Samples were measured by displacement controlled nanoindentation as an average of 25 points over an area of 0.64 mm 2 Three samples were fabricate d at each of three time points to account for variability between batches. 0 0.5 1 1.5 2 2.5 3 3.5 4% 8% 12% 16% 20% Elastic Modulus (MPa)

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28 Cell Spreading and Proliferation Cells were plated at a seeding density of 30,000 cells per well of 6 well tissue culture plates (growth area: 9.5 cm 2 ). Wells were imaged after 24 hours (Figure 3. 2 a) to ensure cellular attachment, which appeared consistent among all substrates. Wells were repeatedly imaged over the time course of the experiment at regular intervals and show ed similar growth and proliferation until cells reached confluency at about Day 7 Phalloidin staining after 3 days of culture (Figure 3. 2 b) showed early cellular spreading. PDMS substrates hindered imaging by reducing brightness and resolution, but post imaging contrast enhancement improved visibility. Th ough F actin concentration on TCP appeared to be more spread out than that on stiffer substrates (especially 12% and 16%), no significant differences could be observed among conditions for either cells grown in CCM or ODM. Percent oxidation reduction indic ated by alamarBlue showed similar proliferation rates among the different substrates. Figure 3. 3 a shows that proliferation activity continued to increase over the three week incubation period and held no significa nt difference between the different substrates when cultured in CCM. When cultured in ODM, however, stiffer substrates (16% and 20%) showed significantly reduced proliferation at Day 21 in comparison to the CCM counterparts. These samples even reduced in comparison to their Day 14 ODM proliferation rate. Day 7 for ODM also showed a significant decrease in proliferation rate, compared to samples cultured in CCM. This decrease in proliferation may be indicative of higher differentiation rates at this time point. Figure 3. 3 b is a representative image of several conditions where over confluency resulted in peeling of the cellular monolayer, which led to three dimensional

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29 (a ) CCM ODM cultures of cells on the stiffer PDMS substrates (particularly 16% and 20%) and sign ificantly reduced confluency in the peeled areas. CCM ODM (b) Figure 3. 2 Cellular Attachment and Spreading. (a) Microscopic images of wells, taken after one day of culture on PDMS substrates and Tissue Culture Plastic (TCP). Images were taken at 100X magnification. (b) After three days of culture in either CCM or ODM, cells were fixed, stained, and imaged to show cell spreading. Cells were imaged at 200X ma gnification and contrast was enhanced post imaging to increase stain visibility.

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30 Figure 3. 3 Cellular Proliferation Rates AlamarBlue measurements were repeated on the same set of plates at 3, 7, 14, and 21 days of culture. (a) Graphs represent cultures grown in Complete Cultur e Media (CCM) and Osteogenic Differentiation Media (ODM) over the time course of incubation (b) Representative image at Day 21 of cellular peeling and detachment of cells cultured on 20% in ODM. Image taken at 40X magnification. (a) (b)

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31 Gene Expression Gene ex pression was quantified by RT qPCR, presented as a fold increase of Day 0 gene expression, and normalized to the housekeeper GAPDH Four genes were chosen for representation of osteogenic differentiation ( ALP TWIST RUNX2 and OCN ) Figure 3. 4 displays relative expression of these genes at 3, 7, 14, and 21 day s of incubation for both CCM and ODM cultures. An asterisk (*) is used to denote statistically significant differences (p<0.05) in order to emphasize the effect of specialized substrate fabrication compared to traditional cell culture methods. Significant differences among varying PDMS substrates are also noted ALP expression in CCM shows no significant difference between substrate s with the exception of an increase in 12% compared to TCP at D ay 14. However, it is important to note the trend for the middle PDMS substrates (12% and 16%) to express slightly greater amounts of ALP compared to both the softer (4% and 8%) and stiffer (2 0%) substrates. Additionally, it can be noted that TCP regularly expressed less ALP compared to all other substrates. Cells cultured in ODM show similar trends in ALP expression, with significant differences in 8% and 12% at different time points. Upreg ulation of ALP began early in both culture media, though more significantly in ODM, and continued to be highly expressed at the later time points. Cells cultured in both CCM and ODM showed trends for TWIST expression to decrease to almost Day 0 expressio n levels at the 7 and 14 day time points. This was reversed at the final time point, where expression of TWIST returned to levels similar to Day 3. Significant differences again favored moderately stiff samples (8%, 12%, and 16%) though changes in expre ssion levels are greater in ODM samples than in CCM.

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32 (a)

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33 (b)

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34 (c)

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35 (d)

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36 Figure 3.4 Gene Expression Data. Time course relative expression of osteogenic markers. Markers for ALP (a), TWIST (b) RUNX2 (c) and OCN (d ) were analyzed. Expression of all genes were normalized to the housekeeper GAPDH and expressed as a relative fold increase on Day 0 iPSC MPs. *p<0.05 for material compared to TCP Expression of RUNX2 showed no significant changes in either media for the first week of culture. At th e 14 and 21 day time points, marked increases in expression could first be noticed in ODM and CCM respectively. Day 21 ODM samples saw the greatest expression of RUNX2 with 4% expressing significantly more over 12%, 16%, 20%, and TCP samples OCN expressi on saw no significant increase until D ay 14 for ODM cultures. Here, again the trend for 12% and 16% substrates to be upregulating slightly more, though no statistically significant difference could be determined. The increase in expression of OCN at Day 21 in CCM is noteworthy given the lack of osteogenic signals in the media A ll samples cultured on PDMS substrates show osteogenic trends, especially on 16% substrates, and the upregulation of osteogenic markers is notable when comparing to samples cultur ed on TCP A lkaline Phosphatase Activity ALP activity [nmol] was measured at 14 and 21 day time points and normalized to the amount of DNA content [g]. Figure 3. 5 a shows that in the CCM condition 12% had significantly greater ALP activity than both the softest substrate (4%) and the control (TCP). After 21 days, 8% and 16% also showed significantly greater activity in comparison with the control. It is only the softest substrate (4%), in fact, which showed no significant difference when compared to TCP Interestingly, ALP activity increased between Day 14 and Day 21 across all conditions, again alluding to a possibility for substrate stiffness to affect differentiation events without chemical signaling.

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37 When cultured in ODM (Figure 3. 5 b), cells plate d on TCP showed the least amount of ALP activity at both time points, even compared to 4%. Additionally, the stiffest substrate (20%) became statistically different compared to TCP in this condition, and even showed the greatest amount of ALP activity at the end of the three week period. It can also be seen that these activity levels mirror gene expression trends discussed in the previous section. Figure 3. 5 ALP Activity. Cells were lysed after 14 and 21 days of culture in either CCM (a) or ODM (b) and ALP activity was measured and normalized to DNA content determined by PicoGreen fluorenscence. *p<0.05 for material compared to TCP (a) ( b )

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38 Calcium Deposition Alizarin r ed s was used to stain for calcium deposition after 7, 14, and 21 days of culture in C CM and ODM. Samples cultured in CCM showed no indication of calcium deposition in any condition (data not shown). Representative images of calcium deposition by cells cultured in ODM can be seen in Figure 3. 6 Samples fixed and stained after one week s howed no signs of calcium mineralization when cultured in ODM, indicating that iPSC MPs had not deposited any matrix However, within the next 7 days evidence of calcium could be seen from the staining. Qualitatively it is apparent that 16% substrates resulted in the greatest amount of calcium deposits, while 12% and 20% had a comparable follow up. Cells cultured Figure 3. 6 Calcium Deposition Data. Alizarin red s was used to stain calcium deposition at 7, 14, and 21 day time points for cells cultured in CCM and ODM. Plates were scanned and representative images are displayed. Cells cultured in CCM showed no indication of calcium deposition at any time point (data not shown).

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39 Figure 3. 7 Microscopy. Microscopic images of cells cultur ed in osteogenic media after 21 days at 40X magnification. Pictures were taken before (a) and after (b) Alizarin r ed s staining. on TCP however, showed no indication of calcium deposition after 14 days of incubation. This is notable as all PDMS substrates showed at le ast some level of mineralization at this time point (a) (b) A fter 21 days of culture, calcium deposition was clearly increased on all subs trates, with 16% producing the largest amount of the mineral. Figure 3. 7 shows the same wells at 40X magnification both before (a) and after (b) ARS staining. The images in Figure 3. 7 b more clearly show cell morphology independent of mineral staining. It is especially noteworthy on the 20% PDMS wher e cells had peeled away from the substrate, that the PDMS itself had no effect on the outcome of the stain. Quantification of the ARS confirmed qualitative evidence of reduced mineral deposition on TCP when compared to PDMS Graphical comparisons of ARS q uantification can be seen in Figure 3. 8 After 14 days, 12% and 16% substrates resulted

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40 in significantly greater amounts of calcium compared to TCP. 16% also showed significantly more deposition than softer PDMS substrates and continued to mineralize mo re calcium than TCP after 21 days After 21 days all PDMS substrates (except 4%) had resulted in significantly greater calcium deposition than TCP. Figure 3. 8 Quantifying Alizarin r ed s Quantitative destaining of ARS was performed after plates were imaged at 14 (a) and 21 (b) day time points for ODM cultures Plates were read for absorbance at 405 nm and run against a standard curve generated from 2 fol d serial dilutions of ARS *p<0.05 for material compared to TCP

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41 CHAPTER I V DISCUSSION Cell Pro liferation Microscopic imaging of plates after 24 hours shows cell attachment after seeding at a density of 30,000 cells per well (Figure 3. 2 a). At this point it appears as though cell attachment was slightly decreased on TCP in both media conditions, tho ugh due to visual interference by the PDMS substrate, it is difficult to form any definitive conclusions. By Day 3, actin spreading in ODM samples appears to be slightly greater than in CCM cultures, as indicated by less localized fluorescence of the Acti nGreen reagent (Figure 3. 2 b). This is in line with previous investigations that showed that osteogenic differentiation increases with an enhanced degree of spreading [77, 78] Within the ODM culture, 12% and 16% appear to have less localized actin fibers than other PDMS substrates, though cells cultured on TCP seem to present the greatest amount of spreading at this time point. The effect of substrate stiffness on proliferation was further analyzed by quantitati ve measure over the 21 day incubation period. AlamarBlue data shows consistent proliferation rates among different substrates over the time course of incubation when cultured in CCM. All samples show significantly increased proliferation as incubation ti me increases. In ODM cultures the reduction in proliferation by all samples at Day 7 when compared to CCM, suggest s differentiation induced by the osteogenic media [79] At this point the osteogenic developmen t sequence would predict a significant increase in alkaline phosphatase activity, followed by an upregulation of osteocalcin expression, and subsequent mineral deposition, all of which will be discussed later [79] The decrease in proliferation by the stiffer substrates

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42 at Day 14 and Day 21 is more likely a result of cellular detachment rather than an indicator for further differentiation events. This was a common occurrence for the stiffer substrates (especially 20%) cultured in ODM to become over confluent and begin receding into a three dimensional mass. With the exception of these samples, the mean proliferation of ODM specimens at Day 14 is notably higher than those cultured in CCM. Perhaps this reduction in proliferation of the CCM samples is indicative of a delayed differentiation event caused by mechanical stimuli without the presence of chemical signals. Further evidence for this hypothesis will be discussed. Gene Expression The upregulation of ALP is a distinct indicator of early osteogenic differentiation events that resemble previous investigations of osteogenesis [69, 80] I t is evident as early as D ay 7 that the stiffness of the substrate has a significant e expression of the gene. By D ay 14 cells cultured on the 12% sample are expressing significantly more ALP than TCP cultures not only in ODM but also notably in CCM. This early upregulation of ALP is correlated with the down regulatio n of the TWIST gene. TWIST serves as an early mediator of the mesoderm, which subsequently forms mesenchyme and, as noted from previous investigations, was expected to decrease during osteogenic differentiation to allow for RUNX2 and ALP upregulation [72 74] The high expression of the gene at the early stages is indicative of the mesenchymal cell type and subsequent downregulation, notable at D ay 7, substantiates previous data for the influence of TWIST on ALP [72] Additionally, the downregulation of TWIST in osteoblast precursors coincides with the upregulation of RUNX2 which conforms to results of previous investigations [73, 74] Though few significant differences can be noted, the data indicates trends for

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43 moderately stiff substrates (primarily 12%) to have regularly higher expression levels of osteogenic markers in both CCM and ODM For RUNX2 it is evident by Day 14 that the gene was being upregulated in cells cultured in ODM, which is generally accepted as a significant measure of osteoblastic differentiation [70] It has also been shown that the transcription factor of RUNX2 is a direct regulator of ALP contributing to its significant i ncrease in expression in the later time points [81] The final genetic indicator of osteogenesis, OCN is considered a late stage marker and thought to be specifically expressed in mature osteoblasts [71] There is an apparent upregulation of OCN at the earliest RT qPCR time point in ODM, which is in line with previous investigations, but more significant expression is not noticed until the 14 and 21 day time points [71] Contrary to its effects on ALP and RUNX2 the downregulation of TWIST has been shown to correlate with a downregulation of OCN [72] This may explain the correspondingly reduced expression of OCN at Day 7 and increased expression at Day 14, but this is an enigmatic correlation that may require further investigation. Regardless, the notable increase in OCN expression at the later time points is indicative of terminal osteo blast differentiation. Though no definitive indication of osteoblastic differentiation occurr ed in cells cultured in CCM, Figure 4.1 illustrates similar gene expression profiles for CCM at Day 21 and ODM at Day 14. The consistent under regulation of osteo genic markers by cells cultured on TCP alludes to a possibility for substrate stiffness to influence differentiation

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44 Figure 4.1 Similarities between CCM and ODM cultures. Gene expression results for cells cultured in CCM at Day 21 (top ) and ODM at Day 14 (bottom) Correlations in expression profiles can be drawn between the two conditions at the different time points and trends indicate favor for PDMS substrates of moderate stiffness. *p<0.05 for material compared to TCP

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45 without the presence of osteoge nic chemical signals, however at a much slower rate. In order to further study these effects, f uture investigations should draw out incubation periods for at least four weeks. Many of the recurring trends in the RT qPCR data indicate osteogenic differenti ation, though no definitive conclusions can be reached from this data alone. It is important to note the tendency for moderately stiff substrates (particularly 12% and 16%) to regularly show higher expression of osteogenic markers than other PDMS substrat es not only in ODM cultures, but also in CCM. Additionally, TCP regularly produced the lowest expression of osteogenic markers when cultured in ODM as well as in the previously stated CCM cultures, which supports the hypothesis that TCP may not be the bes t culture substrate for osteogenic differentiation. ALP Activity The previous section summarized cellular activity at the gene level alluding favor for 12% and 16% substrates in osteogenic differentiation, especially compared to TCP. S o for a higher ord er of confirmation, an assay at the protein level was conducted to confirm results. Figure 4.2 illustrates the qualitative similarities between gene expression results for ALP and protein ALP activity. There is a notable correlation between ALP gene exp ression and ALP protein activity, which can confirm reliability of RT qPCR results. As an osteogenic marker in and of itself, robust activity of ALP is considered a measurable marker of successful differentiation as it contributes greatly to mineralization and is shown to be one of the first functional genes to be expressed in the process of calcification [82]

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46 C alcium Deposition Results of a lizarin r ed s staining provide additional support for the data resulting from RT qPCR and ALP activity. Qualitatively, it is apparent that moderate to stiff substrates (namely 12%, 16%, and 20%) produce the greatest amount of calcium deposition, a definitive marker of terminal osteoblast differentiation. Quantitative destaini ng showed that a fter 21 days of culture in ODM cells cultured on all but the softest substrate (4%) produced significantly more calcium than did the TCP control Similar results can be observed for ALP activity under the same conditions. Even greater Figure 4.2 Comparison of gene and protein alkaline phosphatase (ALP) activity. Qualitative comparison among common condi tions shows very similar relative expression of ALP. Common trends among conditions show PDMS substrates of moderate stiffness (8%, 12%, and 16%) regularly expressing greater amounts of activity, while the control condition (plated on TCP) commonly showed the least amount of ALP. D14 ODM

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47 ev idence for the effectiveness of material stiffness on the osteogenic differentiation of iPSC MPs is at the 14 day time point, where all PDMS substrates deposited a noticeable amount of calcium while TCP did not ( Figure 3. 6 ). Again, this data is supportive of that provided by RT qPCR and ALP activity at this time point and indicates a strong correlation between the 12% and 16% substrates and efficient osteogenesis. Even the stiffest PDMS substrate (20%) which was common to experience over confluency and p eeling (Figure 3. 7 ), showed greater amounts of calcium deposition than the control, which is also supported by ALP activity and RT qPCR results. Should a strong correlation between ALP activity and mineralization be assumed, as has been shown in previous i nvestigations, the threshold of ALP activity to produce calcium deposition could be determined from this data [82, 83] Referencing Figure 3. 6 at Day 14 a ll PDMS substrates have noteworthy amounts of calcium depo sited onto the substrate, while TCP shows negligible deposition. Figure 3. 5 b (Day 14) shows that ALP activity for TCP is below a concentration of 10 nmol ALP/g DNA, while all PDMS substrates have a concentration higher than 10. By 21 days TCP passes th is threshold and begins to show a significant amount of calcium deposits. Noting CCM ALP concentration levels after 21 days even the substrate with the highest amount of activity (12%) never crossed the 8 nmol ALP/g DNA mark nor showed signs of signifi cant amount s of calcium deposit s Based on this data, the threshold of ALP activity for active calcium deposition could fall between 7 and 12 nmol ALP/g DNA (highest ALP concentration without calcium deposition and lowest ALP concentration with calcium d eposition Figures 3. 5 b and 3. 6 ), but the hypothesis requires further investigation.

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48 Mechanical Influence in CCM Cultures An inter esting trend can be observed for cells cultured in CCM that seem s to follow the osteoblast development sequence, however at a slightly slower pace than the ODM cultures [79] The alamarBlue results show a slight drop in proliferation, compared to ODM, at D ay 14. This reduction in proliferation could be indicative of the beginning o f differentiation events, which were evident at Day 7 for the ODM cultures. Figure 4.1 illustrates similarities between the expression of genes for CCM at Day 21 and ODM at Day 14. Though ALP expression is lower in CCM, the expression profile across the different substrates is very similar and most other conditions show comparable gene expression levels. ALP protein activity in CCM cultures did not quite reach the levels observed in ODM, yet, as discussed in the previous section, the activity was approac hing a level that may be inducive to mineral deposition. There is enough evidence to support a hypothesis for mechanical stimuli to induce osteogenic differentiation in iPSC M P s without the influence of chemical signaling, though cells may require culture for two to three weeks longer than what was tested in this study Future Directions Results indicate that iPSC MPs plated on moderately stiff substrates (12% and 16%) display significantly greater osteogenic differentiation at faster rates than both softe r and stiffer substrates especially compared to cells cultured on TCP. These conditions correspond to a plating stiffness of about 2.245 MPa. These results are slightly lower, though comparable to those from previous investigations where Evans et al. sho wed that 2.7 MPa substrates encouraged terminal osteogenic differentiation of embryonic stem cells and Wang et al. demonstrated enhanced osteogenic differentiation of MSCs on substrates near 3.0 MPa [44, 49] Base d on these studies, 20% PDMS

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49 substrates (2.4470.459 MPa) should have induced greater osteogenic differentiation than softer substrates, however over confluence and cellular peeling was common and may have affected results of the assays. In contrast to CC M cultures that should be drawn out, it would be beneficial to reduce the incubation period of ODM cultures and increase the frequency of conducted assays. Similarly, in cultures of peeled cells (Figure 3.3b) it would be beneficial to compare osteogenic m arkers for cells that collected into 3 D culture to the cells that remained in monolayer. This could be conducted by scraping the peeled cells and analyzing them separately from the unpeeled cells, or staining the entire well with fluorescent antibodies f or osteogenic markers and making visual comparisons Assays such as these could indicate whether or not peeling was beneficial for osteogenesis and confirm the effectiveness of the substrate to induce differentiation As per previously established protoco ls, collagen type I was crosslinked with the PDMS substrates and TCP for consistency, in order to promote cellular attachment to the elastomer [50] Collagen is a major component of the extra cellular matrix for b one formation and could have influenced cellular differentiation. Confirmation of successful crosslinking of collagen across all substrates should be conducted in future studies so as to ensure that this did not vary across the different substrates, where by affecting the outcome of the experiment. Similarly, cellular attachment without collagen crosslinking could also be observed to determine the necessity for the use of this protein. With substrate stiffness optimized for osteogenic differentiation of iP SC MPs, further studies are required in order to best replicate the microenvironment conducive for bone modelling. The microenvironment that can affect cellular behavior is composed of the extracellular matrix, the surrounding cells, signals from autocrin e, endocrine, and

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50 paracrine signaling, nanotopography, and extracellular mechanical forces such as shear stress caused by blood flow [61] Studies have observed many of these effects on osteogenesis, and results f rom these experiments in conjunction with what is known about bone anatomy and modelling can be used to generate a matrix of culture conditions to test the efficacy of combining all of these stimuli in order to create an environment optimized for osteogeni c culturing [37, 39, 40, 61, 84] Im plications for Scaffold Seeding & Bioreacto r s Bioreactors, designed to mimic the mechanical microenvironment for function al engineered tissues, can use the data from this study in order to recreate plating surfaces with material properties similar to that of the 12% and 16% substrates that produced the greatest results for osteogenic differentiation. The resulting cell cultures could then provide a virtually unlimited source of viable homogenous cells that could be utilized for a variety of emerging stem cell therapies [2, 3, 14, 16] M ethods developed here could also be applied to priming cells for scaffold seeding which has been show n to improve bone regenerative capabilities of iPS C M P s on bone scaffold implants [15] Furthermore, advances in biomaterial development allow researchers to more tightly control the mechanical properties of the s caffolds themselves [35, 36] Data obtained from this study can provide information that could guide the design of scaffolds to promote the osteogenic differentiation of iPSC MPs and eventually lead to better bone formation in vivo

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51 CHAPTER V CONCLUSIONS From plating and early spreading, through proliferation and gene expression, to protein activity and mineral deposition, it is clear that osteogenic differentiation of iPSC MPs favors stiffer substrates with a n elastic modulus about 2. 245 0.3 MPa This is especially evident for cells cultured in osteogenic media, which showed significantly increased levels of calcium deposition in addition to an upregulation of osteogenic gene expression and protein activity When cultured in CCM, cells displayed trends for osteogenic differentiation without the presence of soluble differentiation factors however at a slower pace compared to ODM cultures F urther investigation is required in order to verify the se effects on CCM cultures The results from this study clearly indicate a cellular preference for substrates with particular mechanical properties that allude to methods for faster, more efficient cell differentiation techniques. The results from this study can be u sed for optimizing iPSC MP culture techniques in order to provide viable, homogenous cultures of safe and effective stem cells for orthopaedic tissue engineering

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52 WORKS CITED 1. Weinstein, S.I., Yelin, Edward H., Watkins Castillo, Sylvia I. The Burden of Musculoskeletal Diseases in the United States Prevalence, Societal and Economic Cost 2014 [cited 2015 10/13]. 2. Fox, J.M., Genever, Paul G., Use of Adult Stem Cells for Orthopedic Regenerative Medicine Applications. Cell & Tissu e Transplantation & Therapy, 2014. 6 : p. 19 25. 3. Watt, F.M. and R.R. Driskell, The therapeutic potential of stem cells. Philos Trans R Soc Lond B Biol Sci, 2010. 365 (1537): p. 155 63. 4. Rao, M., Stem cells and regenerative medicine. Stem Cell Res Ther, 2012. 3 (4): p. 27. 5. Evans, M.J. and M.H. Kaufman, Establishment in culture of pluripotential cells from mouse embryos. Nature, 1981. 292 (5819): p. 154 6. 6. Martin, G.R., Isolation of a pluripotent cell line from early mouse embryos cultured in medium co nditioned by teratocarcinoma stem cells. Proc Natl Acad Sci U S A, 1981. 78 (12): p. 7634 8. 7. Thomson, J.A., et al., Embryonic stem cell lines derived from human blastocysts. Science, 1998. 282 (5391): p. 1145 7. 8. Kerr, C.L., et al., Embryonic germ cells : when germ cells become stem cells. Semin Reprod Med, 2006. 24 (5): p. 304 13. 9. Wesselschmidt, R.L., The teratoma assay: an in vivo assessment of pluripotency. Methods Mol Biol, 2011. 767 : p. 231 41. 10. Lee, M.O., et al., Inhibition of pluripotent stem cell derived teratoma formation by small molecules. Proc Natl Acad Sci U S A, 2013. 110 (35): p. E3281 90. 11. Mohseni, R., Hamidieh, Amir Ali, Verdi, Javad, Shoae Hassani, Alireza, Safe Transplantation of Pluripotent Stem Cell by Preventing Teratoma Format ion. Stem Cell Research and Therapy, 2014. 4 : p. 212 218. 12. Kimbrel, E.A. and R. Lanza, Hope for regenerative treatments: toward safe transplantation of human pluripotent stem cell based therapies. Regen Med, 2015. 10 (2): p. 99 102. 13. Fernandez Vallone V.B., et al., Mesenchymal stem cells and their use in therapy: what has been achieved? Differentiation, 2013. 85 (1 2): p. 1 10. 14. Wei, X., et al., Mesenchymal stem cells: a new trend for cell therapy. Acta Pharmacol Sin, 2013. 34 (6): p. 747 54.

PAGE 58

53 15. Tan g, M., et al., Human induced pluripotent stem cell derived mesenchymal stem cell seeding on calcium phosphate scaffold for bone regeneration. Tissue Eng Part A, 2014. 20 (7 8): p. 1295 305. 16. Bajada, S., et al., Updates on stem cells and their application s in regenerative medicine. J Tissue Eng Regen Med, 2008. 2 (4): p. 169 83. 17. Liebergall, M., et al., Stem cell based therapy for prevention of delayed fracture union: a randomized and prospective preliminary study. Mol Ther, 2013. 21 (8): p. 1631 8. 18. C aplan, A.I., The mesengenic process. Clin Plast Surg, 1994. 21 (3): p. 429 35. 19. Payne, K.A., D.M. Didiano, and C.R. Chu, Donor sex and age influence the chondrogenic potential of human femoral bone marrow stem cells. Osteoarthritis Cartilage, 2010. 18 (5) : p. 705 13. 20. D'Ippolito, G., et al., Age related osteogenic potential of mesenchymal stromal stem cells from human vertebral bone marrow. J Bone Miner Res, 1999. 14 (7): p. 1115 22. 21. Muschler, G.F., et al., Age and gender related changes in the cell ularity of human bone marrow and the prevalence of osteoblastic progenitors. J Orthop Res, 2001. 19 (1): p. 117 25. 22. Takahashi, K. and S. Yamanaka, Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell, 2006. 126 (4): p. 663 76. 23. Wernig, M., et al., In vitro reprogramming of fibroblasts into a pluripotent ES cell like state. Nature, 2007. 448 (7151): p. 318 24. 24. Takahashi, K., et al., Induction of pluripotent stem cells from adult human fibrobl asts by defined factors. Cell, 2007. 131 (5): p. 861 72. 25. Warren, L., et al., Highly efficient reprogramming to pluripotency and directed differentiation of human cells with synthetic modified mRNA. Cell Stem Cell, 2010. 7 (5): p. 618 30. 26. Lapasset, L. et al., Rejuvenating senescent and centenarian human cells by reprogramming through the pluripotent state. Genes Dev, 2011. 25 (21): p. 2248 53. 27. Hipp, J. and A. Atala, Sources of stem cells for regenerative medicine. Stem Cell Rev, 2008. 4 (1): p. 3 11 28. Hynes, K., et al., Generation of functional mesenchymal stem cells from different induced pluripotent stem cell lines. Stem Cells Dev, 2014. 23 (10): p. 1084 96.

PAGE 59

54 29. Phillips, M.D., et al., Directed differentiation of human induced pluripotent stem ce lls toward bone and cartilage: in vitro versus in vivo assays. Stem Cells Transl Med, 2014. 3 (7): p. 867 78. 30. Denham, M. and M. Dottori, Neural differentiation of induced pluripotent stem cells. Methods Mol Biol, 2011. 793 : p. 99 110. 31. Wang, S., et a l., Differentiation of human induced pluripotent stem cells to mature functional Purkinje neurons. Sci Rep, 2015. 5 : p. 9232. 32. Karumbayaram, S., et al., Directed differentiation of human induced pluripotent stem cells generates active motor neurons. Ste m Cells, 2009. 27 (4): p. 806 11. 33. Orimo, H., The mechanism of mineralization and the role of alkaline phosphatase in health and disease. J Nippon Med Sch, 2010. 77 (1): p. 4 12. 34. Xin, X., M. Hussain, and J.J. Mao, Continuing differentiation of human m esenchymal stem cells and induced chondrogenic and osteogenic lineages in electrospun PLGA nanofiber scaffold. Biomaterials, 2007. 28 (2): p. 316 25. 35. Meinel, L., et al., Bone tissue engineering using human mesenchymal stem cells: effects of scaffold mat erial and medium flow. Ann Biomed Eng, 2004. 32 (1): p. 112 22. 36. Wang, Y., et al., In vitro cartilage tissue engineering with 3D porous aqueous derived silk scaffolds and mesenchymal stem cells. Biomaterials, 2005. 26 (34): p. 7082 94. 37. Song, S., et al ., The synergistic effect of micro topography and biochemical culture environment to promote angiogenesis and osteogenic differentiation of human mesenchymal stem cells. Acta Biomater, 2015. 18 : p. 100 11. 38. Kawase, E., Efficient Expansion of Dissociated Human Pluripotent Stem Cells Using a Synthetic Substrate. Methods Mol Biol, 2016. 1307 : p. 61 9. 39. Battista, S., et al., The effect of matrix composition of 3D constructs on embryonic stem cell differentiation. Biomaterials, 2005. 26 (31): p. 6194 207. 4 0. Edwards, J.H. and G.C. Reilly, Vibration stimuli and the differentiation of musculoskeletal progenitor cells: Review of results in vitro and in vivo. World J Stem Cells, 2015. 7 (3): p. 568 82. 41. Engler, A.J., et al., Matrix elasticity directs stem cel l lineage specification. Cell, 2006. 126 (4): p. 677 89. 42. Huang, C., J. Dai, and X.A. Zhang, Environmental physical cues determine the lineage specification of mesenchymal stem cells. Biochim Biophys Acta, 2015. 1850 (6): p. 1261 6.

PAGE 60

55 43. Rowlands, A.S., P. A. George, and J.J. Cooper White, Directing osteogenic and myogenic differentiation of MSCs: interplay of stiffness and adhesive ligand presentation. Am J Physiol Cell Physiol, 2008. 295 (4): p. C1037 44. 44. Wang, P.Y., W.B. Tsai, and N.H. Voelcker, Screen ing of rat mesenchymal stem cell behaviour on polydimethylsiloxane stiffness gradients. Acta Biomater, 2012. 8 (2): p. 519 30. 45. Steward, A.J. and D.J. Kelly, Mechanical regulation of mesenchymal stem cell differentiation. J Anat, 2014. 46. Yang, C., et a l., Mechanical memory and dosing influence stem cell fate. Nat Mater, 2014. 13 (6): p. 645 52. 47. Eroshenko, N., et al., Effect of substrate stiffness on early human embryonic stem cell differentiation. J Biol Eng, 2013. 7 (1): p. 7. 48. Allen, J.L., M.E. C ooke, and T. Alliston, ECM stiffness primes the TGFbeta pathway to promote chondrocyte differentiation. Mol Biol Cell, 2012. 23 (18): p. 3731 42. 49. Evans, N.D., et al., Substrate Stiffness Affects Early Differentiation Events in Embryonic Stem Cells. Euro pean Cells & Materials, 2009. 18 : p. 1 14. 50. Pelham, R.J., Jr. and Y. Wang, Cell locomotion and focal adhesions are regulated by substrate flexibility. Proc Natl Acad Sci U S A, 1997. 94 (25): p. 13661 5. 51. Prager Khoutorsky, M., et al., Fibroblast pola rization is a matrix rigidity dependent process controlled by focal adhesion mechanosensing. Nat Cell Biol, 2011. 13 (12): p. 1457 65. 52. Macri Pellizzeri, L., et al., Substrate stiffness and composition specifically direct differentiation of induced pluri potent stem cells. Tissue Eng Part A, 2015. 21 (9 10): p. 1633 41. 53. Sebastine, I.M. and D.J. Williams, The role of mechanical stimulation in engineering of extracellular matrix (ECM). Conf Proc IEEE Eng Med Biol Soc, 2006. 1 : p. 3648 51. 54. Maniotis, A. J., C.S. Chen, and D.E. Ingber, Demonstration of mechanical connections between integrins, cytoskeletal filaments, and nucleoplasm that stabilize nuclear structure. Proc Natl Acad Sci U S A, 1997. 94 (3): p. 849 54. 55. Sarasa Renedo, A. and M. Chiquet, Mec hanical signals regulating extracellular matrix gene expression in fibroblasts. Scand J Med Sci Sports, 2005. 15 (4): p. 223 30.

PAGE 61

56 56. Wang, J.H. and B.P. Thampatty, An introductory review of cell mechanobiology. Biomech Model Mechanobiol, 2006. 5 (1): p. 1 16 57. Li, J., et al., The role of extracellular matrix, integrins, and cytoskeleton in mechanotransduction of centrifugal loading. Mol Cell Biochem, 2008. 309 (1 2): p. 41 8. 58. Chen, C.S., J. Tan, and J. Tien, Mechanotransduction at cell matrix and cell c ell contacts. Annu Rev Biomed Eng, 2004. 6 : p. 275 302. 59. Schwartz, M.A., Integrins and extracellular matrix in mechanotransduction. Cold Spring Harb Perspect Biol, 2010. 2 (12): p. a005066. 60. MacKenna, D., S.R. Summerour, and F.J. Villarreal, Role of m echanical factors in modulating cardiac fibroblast function and extracellular matrix synthesis. Cardiovasc Res, 2000. 46 (2): p. 257 63. 61. Barthes, J., et al., Cell microenvironment engineering and monitoring for tissue engineering and regenerative medici ne: the recent advances. Biomed Res Int, 2014. 2014 : p. 921905. 62. Sabine Schulze, G.H., Matthias Krause, Deborah Aubyn, Vladimir A. Bolanos Quinones, Christine K. Schmidt, Yongfeng Mei, and Oliver G. Schmidt, Morphological Differentiation of Neurons on M icrotopographic Substrates Fabricated by Rolled Up Nanotechnology. Advanced Biomaterials, 2010. 12 (9): p. 558 564. 63. Madri, J.A. and M. Marx, Matrix composition, organization and soluble factors: modulators of microvascular cell differentiation in vitro. Kidney Int, 1992. 41 (3): p. 560 5. 64. Chu, L. and D.K. Robinson, Industrial choices for protein production by large scale cell culture. Curr Opin Biotechnol, 2001. 12 (2): p. 180 7. 65. Marolt, D., et al., Engineering bone tissue from human embryonic stem cells. Proc Natl Acad Sci U S A, 2012. 109 (22): p. 8705 9. 66. de Peppo, G.M., et al., Human embryonic mesodermal progenitors highly resemble human mesenchymal stem cells and display high potential for tissue engineering applications. Tissue Eng Part A, 2 010. 16 (7): p. 2161 82. 67. Trappmann, B., et al., Extracellular matrix tethering regulates stem cell fate. Nat Mater, 2012. 11 (7): p. 642 9. 68. Bustin, S.A., et al., The MIQE guidelines: minimum information for publication of quantitative real time PCR e xperiments. Clin Chem, 2009. 55 (4): p. 611 22.

PAGE 62

57 69. Hoemann, C.D., H. El Gabalawy, and M.D. McKee, In vitro osteogenesis assays: influence of the primary cell source on alkaline phosphatase activity and mineralization. Pathol Biol (Paris), 2009. 57 (4): p. 3 18 23. 70. Bruderer, M., et al., Role and regulation of RUNX2 in osteogenesis. Eur Cell Mater, 2014. 28 : p. 269 86. 71. Born, A.K., S. Lischer, and K. Maniura Weber, Watching osteogenesis: life monitoring of osteogenic differentiation using an osteocalcin reporter. J Cell Biochem, 2012. 113 (1): p. 313 21. 72. Yousfi, M., et al., Increased bone formation and decreased osteocalcin expression induced by reduced Twist dosage in Saethre Chotzen syndrome. J Clin Invest, 2001. 107 (9): p. 1153 61. 73. Bialek, P., e t al., A twist code determines the onset of osteoblast differentiation. Dev Cell, 2004. 6 (3): p. 423 35. 74. Kronenberg, H.M., Twist genes regulate Runx2 and bone formation. Dev Cell, 2004. 6 (3): p. 317 8. 75. Pfaffl, M.W., A new mathematical model for relative quantification in real time RT PCR. Nucleic Acids Res, 2001. 29 (9): p. e45. 76. Gregory, C.A., et al., An Alizarin red based assay of mineralization by adherent cells in culture: comparison with cetylpyridinium chlorid e extraction. Anal Biochem, 2004. 329 (1): p. 77 84. 77. Song, W., Kawazoe, Naoki, Chen, Guoping, Dependence of Spreading and Differentiation of Mesenchymal Stem Cells on Micropatterned Surface Area. Journal of Nanomaterials, 2011. 2011 78. McBeath, R., et al., Cell shape, cytoskeletal tension, and RhoA regulate stem cell lineage commitment. Dev Cell, 2004. 6 (4): p. 483 95. 79. Owen, T.A., et al., Progressive development of the rat osteoblast phenotype in vitro: reciprocal relationships in expression of gen es associated with osteoblast proliferation and differentiation during formation of the bone extracellular matrix. J Cell Physiol, 1990. 143 (3): p. 420 30. 80. Phillips, J.E., et al., Glucocorticoid induced osteogenesis is negatively regulated by Runx2/Cbf a1 serine phosphorylation. J Cell Sci, 2006. 119 (Pt 3): p. 581 91. 81. Weng, J.J. and Y. Su, Nuclear matrix targeting of the osteogenic factor Runx2 is essential for its recognition and activation of the alkaline phosphatase gene. Biochim Biophys Acta, 201 3. 1830 (3): p. 2839 52.

PAGE 63

58 82. Golub, E.E., Boesze Battaglia, Kathleen, The role of alkaline phosphatase in mineralization. Current Opinion in Orthopaedics, 2007. 18 (5): p. 5. 83. Leboy, P.S., et al., Ascorbic acid induces alkaline phosphatase, type X collage n, and calcium deposition in cultured chick chondrocytes. J Biol Chem, 1989. 264 (29): p. 17281 6. 84. Dorst, K., Derek Rammelkamp, Michael Hadjiargyrou, and Yizhi Meng, The Effect of Exogenous Zinc Concentration on the Responsiveness of MC3T3 E1 Pre Osteob lasts to Surface Microtopography: Part II (Differentiation). Materials, 2014. 7 (2): p. 1097 1112.