the role of niche stiffness on muscle stem cell …...currently, there are three primary strategies...
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The Role of Niche Stiffness on Muscle Stem Cell Symmetric
Self-renewal
by
Richard Yi-Hsiu Cheng
A thesis submitted in conformity with the requirements
for the degree of Masters of Applied Science
Institute of Biomaterials & Biomedical Engineering
University of Toronto
© Copyright by Richard Yi-Hsiu Cheng 2015
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The Role of Niche Stiffness on Muscle Stem Cell
Symmetric Self-Renewal
Richard Yi-Hsiu Cheng
Masters of Applied Science
Institute of Biomaterials & Biomedical Engineering
University of Toronto
2015
Abstract
Reconstruction of the adult muscle tissue relies on stem cells situated in a niche between the
basal lamina and sarcolemma, but the molecular mechanisms of cell fate decisions, especially
self-renewal, during muscle regeneration remains unclear. We report that the local extracellular
matrix surrounding the muscle stem cell in healthy, uninjured tissue is permissive for planar and
apical-basal division orientations that lead to either symmetric or asymmetric cell fates, but that
the increased extracellular matrix deposition during injury physically alters the niche architecture
to significantly favour symmetric divisions in the planar orientation. We demonstrate that the
addition of soluble ligands such as Wnt7a leads to a significant increase of self-renewal divisions
instead of differentiation, but only when coupled with 3D culture in a stiff environment.
Together, these results suggest that biophysical cues such as niche stiffness work synergistically
with biochemical cues like Wnt7a signaling to regulate muscle stem cell self-renewal.
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Acknowledgments
I would especially like to thank my supervisor Dr. Penney Gilbert for her patience, motivation,
and mentorship during my Master’s training. I also want to thank the members of my committee
Dr. Boris Hinz, Dr. Jonathan Rocheleau, and Dr. Craig Simmons for their insightful comments
and challenging questions. My sincerest thanks go to the laboratory members of the Gilbert lab,
specifically Sadegh Davoudi, Mohsen Afshar, Aliyah Nissar, James Morrissey and Min Rui for
their stimulating talks; Erfan Farno and Kent Hsieh, who performed immunohistochemical
analyses; and Jing Yang and Gini Chin for myofiber isolation and cryosectioning. Thank you to
Dr. Libero Vitiello for training me experimental techniques such as isolating bulk skeletal
muscle from our mouse models, and Haijiao Liu for performing all the AFM measurements. Last
but not least, thank you to my friends and family for their continued support over these past two
years.
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Table of Contents
Introduction .....................................................................................................................................1
Hypothesis & Objectives ..............................................................................................................19
Results ...........................................................................................................................................20
Discussion .....................................................................................................................................26
Materials and Methods ..................................................................................................................36
Appendix .......................................................................................................................................41
References .....................................................................................................................................44
Figures ...........................................................................................................................................53
Appendix Figures ..........................................................................................................................63
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List of Figures
Figure 1. The skeletal muscle stem cell niche architecture ..........................................................53
Figure 2. The bulk stiffness of skeletal muscle is dynamic over the course of regeneration .......54
Figure 3. Systemic changes occur in the MuSC niche during regeneration .................................55
Figure 4. The stiffness of MuSC niche is greater after injury compared to healthy control ........56
Figure 5. The stiffness of plasmin-treated MuSC niche is significantly lowered compared to non-
treated control ...............................................................................................................................57
Figure 6. The stiffness of agarose gel corresponds with weight/volume percentage composition
........................................................................................................................................................58
Figure 7. Muscle fibers remain viable when 3D cultured in soft and stiff agarose ......................58
Figure 8. Muscle stem cell division orientation differs between 3D culture in soft versus stiff
environments .................................................................................................................................59
Figure 9. MuSC count and percentage of EdU+ MuSC population does not change in soft or stiff
environments .................................................................................................................................60
Figure 10. Biochemical cues synergize with niche architecture to support symmetric self-renewal
divisions ........................................................................................................................................61
Figure 11. Niche architecture impacts myoblast morphology ......................................................62
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List of Appendices
Appendix 1. The stiffness of bulk TA muscle is greater in aged mice compared
to young mice ................................................................................................................................63
Appendix 2. Mouse diaphragm as a platform to visualize in vivo changes to MuSC behavior after
exposure to injury or artificial stiffness ........................................................................................63
Appendix 3. Diaphragm viability in culture +/- cryoinjury ..........................................................64
1
INTRODUCTION
Form and Function of Skeletal Muscle
Skeletal muscle is a type of tissue that is necessary for movement, positioning, and
strength. In contrast to other major types of muscle such as smooth or cardiac muscle, the skeletal
muscle is striated and under control of the somatic nervous system1. Repeating functional units of
tubular muscle cells called the sarcomere provide the characteristic light and dark striated pattern
observed conventional microscopy2, and contractions in these striated muscles are voluntarily
controlled by the somatic nervous system to enable a wide range of physical motion. Motor
neurons, which are responsible for innervating multiple myofibers, generate synaptic input that
lead to skeletal muscle contractions3. Additionally, the skeletal muscle organ is organized into
individual bundles of muscle cells called the muscle fascicle. The muscle cells, also known as the
muscle fiber or myocyte, in turn contain myofibrils that contain actin and myosin filaments
essential for muscle contraction. The development of mature skeletal muscle relies on the
recruitment of multiple immature myoblasts that fuse into a long, cylindrical shape to give its
characteristic multi-nucleated phenotype4. While the primary component of muscle cells are
myofibrils, other organelles such as the mitochondria and sarcoplasmic reticulum provide
essential functions such as energy generation and storage. Individual cells are surrounded by basal
lamina, which provide biochemical signaling, mechanical interaction and a physical barrier to
other fibers5. Together, the anatomical components of the skeletal muscle organ work in tandem
to facilitate physiological functions such as motility, weight-bearing, and force generation.
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Skeletal Muscle Dysfunction and Disease
Diseases in the skeletal muscle often impact the form and function of the tissue which
may ultimately lead to a decrease in motility or death. Duchenne Muscular Dystrophy (DMD) is
a genetic disease characterized by the deletion of the dystrophin gene that affects 1:3,600 boys6.
The impaired stability of the dystroglycan complex on the skeletal muscle cell membrane
enables excessive calcium to penetrate the sarcolemma and damage the integrity of the myofiber.
The absence of myofibers coupled with aberrant microenvironment signaling leads to fat
infiltration, deposition of extracellular matrix, and an overall decrease in muscle quantity7. The
progressive deterioration of skeletal muscle tissue leads to loss of movement, paralysis, and in
some cases, death. Other types of skeletal muscle disease include neuromuscular junction
disorders like Amyotrophic lateral sclerosis (ALS) or Lou Gehrig’s disease, where the
communication between the central nervous system and peripheral muscles become disrupted8.
Not only can general motility be affected as a result, but the ability for the patient to breathe or
swallow may be also impaired. Skeletal muscle wasting and overall loss of strength can also
arise in the context of other diseases like cancer or AIDS (cachexia)9, or in the context of aging
(sarcopenia)10. Other chronic inflammatory myopathies associated with high levels of fibrosis
and severe muscle weakness often have no known cause11, thus highlighting the importance of
research in this field.
The Satellite Cell and Innate Skeletal Muscle Regenerative Capacity
Everyday motions for an active, healthy organism lead to minor wear and tear of
individual muscle fibers. Fortunately, the regenerative capacity of skeletal muscle enables
maintenance of the organ despite continual physical damage. The ability to repair and re-form
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intact muscle fibers is primarily due to the activity of muscle stem cells (MuSCs), which are a
rare population representing 2-10% of all myonuclei that is anatomically positioned in a niche
between the basal lamina and the sarcolemma of the skeletal muscle fiber12,13. Another name for
this cell is the “satellite cell”, a term coined by Alexander Mauro in 1961 due to its anatomical
location satellite to the muscle fiber14. Efforts over the last fifty years have established the role of
the MuSCs as the progenitor for myogenic cell differentiation in times of repair and growth.
Indeed, upon skeletal muscle damage such as exercise, injury, or degenerative disease, quiescent
MuSCs can activate, proliferate, and give rise to progenitor cells known as myoblasts. These
myoblasts terminally differentiate by exiting the cell cycle and fusing with damaged myofibers
or fusing with one another to form nascent fibers that integrate with the surrounding
environment15,16. However, activated MuSCs must self-renew in order to maintain the stem cell
pool and prevent stem cell depletion. Repeated injuries that prompt multiple rounds of stem cell
activation and division did not lead to an overall decreased amount of MuSCs17, suggesting that
these cells are able to balance quiescence, self-renewal, and activation to ensure long-term
maintenance. Muscle stem cells are necessary for the regeneration of skeletal muscle, as shown
by the complete lack of regeneration in Pax7-depleted tissues18,19.
The discovery of muscle-specific molecular markers has enabled the identification of cell
progression along the myogenic lineage. The paired-box transcription factor 7 (Pax7), which is a
protein important for MuSC function and survival, is uniformly used in the muscle field to
identify muscle stem cells17,19,20. In the initial stages of activation, Pax7 expression gradually
decreases to be replaced by Myogenic differentiation 1 (MyoD). Myogenic factor 5 (Myf5)
protein expression is found further along the myogenic lineage as activated MuSCs develop into
progenitor cells and myoblasts15. Due to inherent MuSC heterogeneity, there may be subtle
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differences in terms of cellular lineage progression and contribution to muscle regeneration
within a total population of MuSCs21–23. In contrast to MuSCs that proceed to muscle precursor
cells by hierarchical activation, the Rudnicki group has proposed that a subpopulation of MuSCs
maintain Pax7 levels and never express markers indicative of myogenic progression. Using a
Myf5-Cre/ROSA26-YFP mouse where cells that express Myf5 at any point during their
development will be permanently labeled with yellow fluorescent protein (YFP), they concluded
that ~10% of total satellite cells never express Myf5 during development24. In addition to
possessing higher self-renewal capacity and an increased tendency to commit to myogenic
progenitors, this subpopulation leads to long-term engraftment in transplantation experiments
instead of maturation or fusion into adult fibers. Together, these results suggest that satellite cells
are a heterogeneous population where some are predisposed to progress through the myogenic
lineage compared to others that self-renew and maintain the stem cell pool.
Current Therapies and Future Regenerative Medicine Strategies
Current treatment for muscle diseases often involve the use of corticosteroids which may
improve muscle strength and delay disease progression, but prolonged use have been shown to
lead to weakened bones and increased fracture risk25. Non-invasive assistive therapy and devices
such as range-of-motion and stretching exercises or braces have been demonstrated to increase
the quality of life for patients afflicted with muscular dystrophies26. Nevertheless, many of
current treatment options are supportive in nature and do not address the molecular mechanisms
underlying the disease. Future therapies will be based on knowledge generated from researching
the basic biology of muscle stem cell regulation and interaction with its surrounding
microenvironment.
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Currently, there are three primary strategies in muscle regenerative medicine under
investigation in the laboratory or clinical setting: transplanting skeletal muscle stem cells,
engineering replacement tissue, and manipulating the endogenous stem cell population.
Transplantation of muscle stem cells have been somewhat successful in animal models, with
studies indicating that some transplanted cells successfully integrate with the host tissue to
differentiate into mature myofibers23. However, the process is highly inefficient due to
significant levels of cell death upon transplantation, and the inability for the cells to migrate
beyond the point of injection. Existing projects explore the possibility of delivering the cells
using different types of hydrogel carriers to limit harmful shear forces to increase viability and
enable better dispersal of stem cells27. The loss of a large area of muscle may require
transplantation of engineered muscle tissue instead of individual stem cells, and several
laboratories are investigating the possibility of creating skeletal muscle maintained in culture
with the ultimate goal of surgical engraftment28. Before that can be a reality, more research needs
to be done to address issues with myofiber innervation and integration with the existing
vasculature. Manipulating the endogenous stem cell population within an organism, for example
through delivering soluble ligands to activate specific signaling pathways to encourage
regeneration of damaged tissue may also prove to be an effective strategy, but more experiments
need to be performed to reduce off-target effects and eliminate the possibility of aberrant cell
signaling. Regardless of the different approaches, further understanding the basic biology behind
muscle stem cell regulation is required before effective treatments to treat muscular dystrophy
are developed.
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Regulation of stem cell fate
The mechanisms directing MuSC self-renewal or differentiation in homeostasis or as a
response to injury has been an important subject for scrutiny. Changes in cell state may differ
due to intrinsic differences arising from the organism’s age, health, or genetics. For example,
during aging, the accumulation of genomic damage may lead to deregulation of critical pathways
essential for maintaining MuSC quiescence29. To complement intrinsic cell changes, it was
traditionally thought that MuSC cell fate is also regulated by interactions with surrounding
immune cells, motor neurons, and vascular neighbors that signal via paracrine biochemical
mechanisms30. While extracellular matrix (ECM) ligand availability plays a crucial role31,32,
emerging studies investigating biophysical cues have found that compression or stretch induced
by muscle contraction are important regulators of MuSC fate33,34.
Biochemical regulation of the stem cell fate
Muscle stem cell activation following injury or over the course of aging leads to intrinsic
changes that ultimately lead to proliferation, differentiation, and regeneration of tissue.
Investigation of isolated fibers have demonstrated that p38α/β MAPK (Mapk14/11) translocates
to the nucleus after activation and inhibition of this leads to the inability of the MuSC to enter
cell cycle35. Intrinsic MAPK pathways essential for myogenic progression include FGF2-
dependent ERK1/2, which was shown to be critical for G1 to S-phase transitions36, and JNK,
whose interaction with cyclin D1 enables cell cycle progression37. During aging, elevated levels
of p38α/β MAPK is associated with the increase in senescence markers and contribute to a
reduced capacity to repair myofibers and repopulate the stem cell reservoir in vivo following
transplantation29. Genome wide analysis of freshly isolated MuSCs from young and old muscles
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have also revealed that the JAK/STAT signaling pathway intrinsically upregulated in the aged
population38. Altogether, these results indicate that the decline in the regenerative capacity of
MuSCs is due in part to alterations in the intrinsic properties of the stem cell.
MicroRNAs (miRNAs) have also been shown to play an important role in the regulation
of muscle stem cell fate. Deletion of miRNA-processing endoribonuclease DICER1 resulted in
the precocious activation of MuSCs leading to an overall decline in muscle regeneration39.
Additional miRNAs that regulate the cell cycle and prevent premature differentiation such as
miR-489 are present in quiescent MuSCs where they post-transcriptionally block oncogene Dek
to prohibit entry into cell cycle. Quiescence of satellite cells can also be achieved by sequestering
Myf5 into messenger ribonucleoprotein particle (mRNP) granules through the activity of miR-
3140. Together, these results highlight the importance of microRNA activity in the intrinsic
regulation of muscle stem cell fate.
During resting conditions, the muscle stem cell is at a quiescent, or “g0 state”, where its
metabolism is significantly reduced and the possibility of DNA damage minimized. Long-term
labeling experiments using transgenic doxycyclin-inducible H2B-GFP animals showed that a
population of Pax7+ cells can maintain quiescence throughout the lifetime of the organism
despite alterations in the stem cell niche41. Since this state is essential for long-term maintenance
of skeletal muscle, any disruption of quiescence leads to an increased tendency of differentiation
and a general reduction in satellite cell numbers over time. In addition, the capacity for the stem
cell to rapidly enter cell cycle after activation suggests that quiescence is a highly regulated state
and that stem cells in this state are primed for immediate activity. Using microarray analysis, 500
genes were found to be highly upregulated in quiescent muscle stem cells compared to myoblasts
actively in cycle42, which included numerous negative regulators of cell cycle such as cyclin-
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dependent kinase inhibitor 1B (Cdkn1b), retinoblastoma tumour suppressor protein (Rb), and
peripheral myelin protein 22, among others. Conditional knockouts of Cdkn1b and Rb proteins
resulted in the disruption of satellite cell expansion and maintenance of the primitive state,
suggesting that the disruption of MuSC quiescence leads to impairment of self-renewal and
overall regeneration.
Soluble factors such as the wingless family member Wnt7a released by regenerating
fibers in the muscle stem cell niche also play a role in dictating stem cell fate. The interaction
between Wnt7a with the frizzled 7 (Fzd7) receptor located on the satellite cell membrane
activates small Rho-GTPase Rac1, which is a member of the planar cell polarity (PCP) pathway.
This leads to the symmetric polarization of downstream effectors such as Vangl2 at opposite
ends of the dividing stem cell and ultimately supports symmetric expansion of the satellite cell
population. Ex vivo treatment of satellite cells with Wnt7a showed increased engraftment in
transplantation experiments32. Interactions between soluble ligands like Wnt7a with extracellular
matrix components such as fibronectin have been shown to influence symmetric expansion43.
Additionally, the control of MuSC quiescence and activity is in part regulated by the Notch
signaling pathway where Delta or Jagged ligands on neighboring cells or ECM interact with
Notch receptor and Syndecan-1 co-receptor on the satellite cell. Expression of Notch1 and
Notch3 receptors are activated by forkhead transcription factor FOXO3 enriched in satellite cells
in quiescence44, and downstream effectors of Notch signaling include transcription of Hes and
Hey family proteins. Knockout of downstream effectors such as Hey1 and Heyl led to increased
differentiation of MuSC, suggesting that Notch signaling is important for maintenance of stem
cell state. Importantly, myogenic cell progression during regeneration requires regulated
decrease in Notch expression as shown by Brack et al.45
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While signaling factors in the Notch and Wnt families are the most extensively studied,
recent studies have shown that a variety of biochemical factors influence MuSC fate. Other
factors released in the context of injury include insulin-like growth factor (Igf1), which
inactivates FOXO1 to downregulate cell cycle inhibitor Cdkn1b to force cell cycle entry46.
Single fiber experiments have shown that fibroblast growth factor 2 (FGF2), whose secretion is
increased during regeneration, is responsible for activating specific MAPK pathways in cells.
Interestingly, proteins located in the extracellular matrix within the satellite cell niche have also
been shown to influence cell fate choice. Collagen VI in the ECM has been shown to be a ligand
essential for reducing myogenic commitment and promotion of self-renewal31. Fibronectin, a
significant component of the ECM in the MuSC niche, also promotes stem cell expansion by
interacting with MuSC surface proteins such as syndecan 4 (Sdc4) and frizzled 7 (Fzd7) to form
a complex for downstream signaling. Knockdown of fibronectin leads to a significant decline in
engraftment potential and self-renewal ability. Together, these results suggest that biochemical
cues from neighbouring cells, ECM, and circulation work in tandem to influence satellite cell
quiescence, activation, and proliferation.
Extracellular matrix protein composition
Not only is the extracellular matrix in the skeletal muscle critical for maintaining the
structure of the basal lamina47,48 and for providing mechanical support31, but they also serve as a
biochemical ligand that can signal directly to cells32. The ECM has been shown to change
dynamically following injury and may serve as scaffold for migratory satellite cells fated for
tissue regeneration49–51. In vitro experiments exploring the effects of principle ECM components
used in the growth substrate have shown the importance of individual building blocks of ECM
such as collagen, fibronectin, and laminin to be essential for cell viability and differentiation
10
behaviour52–54. However, how these ECM building blocks impact cell fate at the molecular level
after still remains speculative.
Collagen is the most abundant protein in mammals, and also the main structural protein
in the extracellular matrix. Its triple helical shape enables covalent cross-linking to produce
structures with physical properties ranging from loosely packed collagen fibrils to dense, highly
insoluble aggregates. In the skeletal muscle, where the bulk stiffness is often correlated with
increased collagen content, the basement membrane is enriched for collagen IV in addition to
collagen I and III55. Deficiencies in collagen VI production has been linked to a lower elastic
modulus of bulk muscle tissue compared to wild-type animal controls, suggesting that local
alterations in the ECM may impact physical characteristics at the tissue level56. Excessive
collagen production, on the other hand, has been shown to impact muscle cell fate by inhibiting
myoblast differentiation and fusion, perhaps due to the mechanical restraint that restricts
morphological changes57. Taken together, these findings indicate a dynamic environment where
the production of collagen is carefully balanced by stem and other support cells prevent aberrant
cell behaviour.
Fibronectin is another ECM component integral to the development58,59 and regenerative
process of skeletal muscle60–62. Although it is detected at low levels in the homeostatic adult
tissue, the expression level of this protein is increased in myogenic regions and necrotic sites32.
MuSCs and muscle progenitor cells both interact directly with this ECM component, for example
through the Wnt7a-fibronectin-syndecan4 complex where downstream effects include
cytoskeletal actin assembly and lamellapodia formation63. Importantly, these downstream effects
reflect an alteration in cellular geometry64, which has been observed in other studies where the
stem cell adopted a round shape65, with a rigid cytoskeleton66. Additionally, increased
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fibronectin may lead to reduced cellular locomotion and adhesion67, possibly by physically
restricting the mobility of the stem cell to the basal lamina. As a result, the cell may be more
inclined to maintain the stem cell pool instead of exiting the basal lamina to differentiate.
Together, these results show how an ECM component like fibronectin can support specific cell
fates by modulating intracellular cytoskeleton activity.
Laminin, another component of the extracellular matrix surrounding the muscle stem cell,
have been found to be essential for the organization of the basal lamina. Disruption of laminin
leads to impaired stem cell proliferation by affecting cell locomotion and morphology68–70, but
only when presented on substrate instead of in solution66. This indicates that the physical
organization of ECM is important, perhaps hinting at the requirement of a specific ligand
configuration for effective interaction. In support of this notion, muscle stem cell expression of
laminin-binding a7b1-integrin only on the basal surface but not on the myofiber-facing apical
side is reflective of the importance of ligand polarity in determining cell fate24.
Taken together, these findings suggest that the MuSC is sensitive to the ECM
composition in the niche which may lead to downstream changes in cell fate. The production and
dissociation of ECM is carefully maintained by both stem and neighbouring support cells, and
disruption may lead to impairments in viability and proliferation. Furthermore, stem cells can
create complexes with the ECM ligand and integrin receptor to alter intracellular cytoskeleton
activity and modulate cell motility and adhesion. The directional availability of the ECM also
highlights the importance of ligand presentation in promoting downstream asymmetric cell fates.
Biophysical regulation of the muscle stem cell
The contribution of biophysical cues from the surrounding extracellular matrix is
emerging as a critical regulator of MuSC fate. In addition to serving as a ligand for MuSC fate-
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determing signaling cascades, the ECM can also be a biophysical signal depending on its protein
composition, stiffness, and mechanical loading capacity. Recently, the focus has been on
uncovering the niche cues that drive the activation and silencing of signal transduction pathways
responsible for satellite cell fate decisions. The dynamic changes in the physical properties of the
skeletal muscle may provide insight to understand the regulation of satellite cells from the niche.
Substrate stiffness
The stiffness of mammalian tissues varies widely and often plays a role on its functional
behaviour, such as the importance of a rigid skeletal system to accommodate load bearing.
However, these mechanical properties in the bulk tissue may not necessarily match the
mechanical properties at the cellular level. Substrate stiffness, or the stiffness directly felt by
individual cells, is considered a biophysical cue that influences cell shape, adhesion, motility,
and even differentiation71,72. Generally, cells can respond to rigid substrates by spreading their
plasma membrane to cover a larger surface area. Integrins have been shown to be critical for
mechanotransduction, defined as the mechanism by which cells can convert mechanical stimulus
into electro-chemical activity to influence cell behaviour. Signaling cascades known to be
sensitive to substrate stiffness include the focal adhesion kinase (FAK) – protein kinase B (Akt)
complexes, extracellular signal-regulated kinases (ERK)73,74, and Yes-associated protein (YAP)
– transcriptional coactivator with PDZ domain (TAZ) complexes75,76. During
mechanotransduction, membrane-bound integrin receptors on the cell interact with the
surrounding stiffness to provide a mechanical stimulus that translates into an intracellular cue to
regulate gene expression for cell fate determination.
In the skeletal muscle, waves of regeneration and degeneration that occur during injury
and aging are associated with fluctuating mechanical properties such as bulk tissue stiffness.
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Changes in the local environment surrounding the MuSC, such as the deposition of extracellular
matrix proteins and infiltration of adipose may lead to further alterations of niche properties.
Previous studies investigating the effects of fibrosis on Mdx mice that models Duchenne
muscular dystrophy have revealed that there is an increase in fibrin deposition, collagen
deposition, and muscle fiber rigidity77–79, all of which contribute to the stiffening of the stem cell
niche. The sensitivity of the stem cell to local stiffness reflects the necessity of using culture
conditions that mimic mechanical properties of the muscle tissue. In contrast to extremely stiff
tissue culture plastic where reincorporation of MuSCs into the murine muscle is significantly
impaired80,81, cell cultures on hydrogels tuned to the stiffness of healthy muscle leads to
increased MuSC viability and self-renewal82,83. Furthermore, lowering the collagen content has
been reported to increase satellite cell differentiation to later stages of the myogenic lineage,
including Pax7+/MyoD+ myoblasts and mature myofibers83. Taken together, these findings
provide evidence that the MuSC is highly sensitive to mechanical properties of the niche, and
that substrate stiffness is a fundamental biomechanical cue that regulates cell fate.
Mechanical loads as regulators of stem cell fate
Contraction and relaxation of the skeletal muscle during routine activities impose
mechanical loading on the skeletal muscle, such as shear and stretch force. Using in vitro, in
vivo84–86, and computational modelling87, the movements between the muscle fiber and the
enveloping connective tissue have been demonstrated to enable force transduction leading to
myofiber adaptation33,34. However, there are currently no studies to determine whether the
MuSCs situated on top of myofibers can directly sense and respond to endomysium shear
loading. In addition to the possibility of shear stress imposed on the satellite cell in the apical
direction, it may be possible that the myofiber itself provides a stretching force as it shortens and
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elongates during muscle movement. Cyclic strain has been demonstrated previously to be
involved in both satellite cell activation88,89 and signal transduction through the p38, JNK, and
ERK1/2 pathways90–95. While much of the focus in elucidating biomechanical regulators has
been limited to identifying the contribution from ECM composition and substrate stiffness, these
results give weight to the idea that mechanical loading may be equally relevant.
The association of muscle stem cell fate with division orientation
In general, stem cells are defined as having two main properties: self-renewal, or the
ability to go through numerous cycles of cell division while maintaining an undifferentiated
state, and potency, which is the capacity of the cell to differentiate into a specialized cell type.
Many stem cell types including those of hematopoietic lineage or skeletal muscle can divide
asymmetrically, where one daughter cell remains identical to the mother cell and the other cell
proceeds to differentiate. Alternatively, the stem cell can divide in a symmetric fashion where
both daughter cells are the same as the mother cell, but can then activate and differentiate if the
need arises. Severe injuries which require extensive regeneration from stem cell activity typically
require a balance between asymmetric and symmetric divisions to effectively respond to the
demands of tissue repair, yet preserve the stem cell pool for future use. Previously, we have
shown that biophysical cues such as extracellular matrix composition, substrate stiffness, and
mechanical loading regulate MuSC fate by activating cell-fate specific signaling pathways linked
to stem cell viability, motility, and differentiation. Here, we discuss the idea that these physical
characteristics of the niche modulate cell morphology to control symmetric or asymmetric
division.
The ability for stem cells to self-renew or differentiate has been observed in a variety of
cell types. In skeletal muscle, Rudnicki et al. has demonstrated on isolated fibers that MuSC can
15
divide symmetrically to rapidly expand the myogenic progenitor population by producing two
Pax7+/Myf5- daughter cells24. Alternatively, the MuSC can divide asymmetrically to produce
one Pax7+/Myf5+ cell committed to differentiation in addition to one Pax7+/Myf5- cell to
maintain the available MuSC pool96. Importantly, they highlights the importance of cell polarity
to cell divisions by attributing symmetrical divisions with planar division orientation, where cells
divide in plane with the muscle fiber and both daughter cells maintain contact with the
sarcolemma. In contrast, asymmetrical divisions are more often associated with cell division
orientation in the apical-basal plane relative to the muscle fiber, where the daughter cell that
loses contact with the basement membrane proceeds along the myogenic progression and the
daughter cell that maintains a connection with the basement membrane remains an
undifferentiated stem cell.
Additionally, their lab has established various biochemical which interact synergistically
with intrinsic cues to direct MuSC to either self-renew or differentiate43,45,97. The planar cell
polarity (PCP) pathway promotes symmetric division in MuSCs by distributing polarity effectors
such as Vangl2 in the daughter cells. Removal of effectors as demonstrated in Vangl2 knock-
down mice led to a decrease in total satellite cell numbers but increased levels of differentiated
progenitors, suggesting that symmetric division is essential for expanding the satellite cell pool
and self-renewal. Asymmetric division, on the other hand, relies on the distribution of Notch3
receptor to the daughter cell maintaining the stem cell fate, and the allocation of Notch ligand
Delta1 and Notch antagonist Numb to the committed daughter cell98. Downstream effectors such
as MyoD expression was also found to be asymmetrical in nature, with the committed daughter
cell expressing high levels of MyoD compared to the daughter cell adopting the stem cell fate
with low levels of MyoD but maintained levels of Pax799. Similar patterns were observed for
16
protein expression of differentiated fates, such as Myog and Myf5100. Other experiments
investigating the distribution of DNA using BrdU incorporation assays revealed that the daughter
cells receiving the parent’s template DNA expressed SCA-1 compared to the daughter cell with
the new template where desmin levels was increased instead101. Taken together, these results
suggest that symmetric and asymmetric division of MuSCs rely on the unequal distribution of
polarity proteins that dictate cell fate. The response of the MuSC to divide either symmetrically
or asymmetrically depends on the needs of the regenerating muscle. For example, routine
activities that do not demand a high level of regeneration would likely result in asymmetric cell
divisions where both the stem cell pool is maintained and myogenic progenitors are generated.
Alternatively, more serious injuries that demand high stem cell activity would likely lead to
symmetric expansion, where there is a rapid expansion of available satellite cells. The stem cell
must be poised to receive an integrated signal, whether it be biochemical or biophysical, and
generate an appropriate response by regulating self-renewal or differentiation.
The role of spindle pole orientation in directing cell polarity
The establishment of different cell fates can arise through inheritance from one cell
generation to the next due to the unequal distribution of polarity proteins and cell fate
determinants102. The orientation of spindle poles not only dictates the direction of stem cell
division, but it can also influence the fate of the resulting daughter cells. Since misaligned
mitotic spindles have been shown to be associated with disease103, we can reason that regulation
of mitotic direction is critical for tissue morphogenesis. The molecular principles of spindle
orientation during asymmetric cell division have primarily been established in invertebrate
model systems like Drosophila and C. elegans, but recent work with mammalian models has
demonstrated that these mechanisms are highly conserved104. Briefly, the process of spindle
17
orientation first requires the identification of the cortical regions responsible for capturing the
mitotic spindle. Next, the interaction between the mitotic spindle and the cortical region of the
cell must be established, typically through the nucleation of astral microtubes by centrosomes at
the spindle poles. Microtubule binding proteins such as G alphai, LGN, and other cellular
machinery associated with spindle positioning then operate to exert torque on the spindle104. The
Numa/Mud complex interacts with cytoplasmic Dynein to control the orientation of the mitotic
spindle, as demonstrated in the chicken neural tube, mouse neocortex, and the developing
epithelia105–107. Regardless, how the orientation of the mitotic spindle leads to downstream
changes in cell fate is not well understood108. The potential for cell morphology that integrates
with extrinsic signaling to control division axis of stem cells may prove to be an interesting
mechanism to explain the relationship between spindle orientation and cell fate. Studies in c.
elegans epithelial stem cells have revealed that disordered orientation of cell division only
occurred when both extrinsic Wnt signaling and cell shape were modified109. Future studies in
skeletal muscle field will aim to adopt approaches previously identified in other model
organisms and understand how extrinsic biochemical and biophysical cues integrate with
intrinsic mechanisms to manipulate mitotic spindle movement during cell division.
Niche architecture as a biophysical regulator of stem cell fate
Biochemical cues that synergize with intrinsic signaling pathways in the context of
regeneration have been thoroughly studied in the muscle field, but the contributions from
biophysical cues are emerging as critical regulators. As introduced previously, ECM
composition, substrate stiffness, and mechanical loading are all considered biophysical regulators
that influence MuSC fate. In this thesis, we explore the notion that the niche architecture is a
novel biophysical cue that plays a crucial role in determining division orientation to drive muscle
18
stem cell self-renewal. Here, the architecture of the niche is defined as the organization and the
composition of the extracellular space surrounding the MuSC that can either be pliant or rigid
(Figure 1).
19
HYPOTHESIS & OBJECTIVES
We hypothesize that the increased deposition of ECM during tissue regeneration modifies
the MuSC niche architecture to support symmetric self-renewal divisions in the planar
orientation. Specifically, the objectives of this study are 1) to characterize the physical properties
of the regenerating MuSC niche, 2) to quantify MuSC division orientations in the regenerating
niche with an agarose gel model, and 3) to determine synergistic effects between niche
architecture and biochemical cues in regulating MuSC fate.
To investigate the effects of ECM deposition on niche mechanics during regeneration, we
used atomic force microscopy on local regions surrounding the MuSC. Furthermore, 3D
immunohistochemistry on fibers embedded in soft and stiff agarose gels enabled us to determine
MuSC fate in healthy and injured niche environments. We found that the local ECM surrounding
the MuSC in healthy, uninjured tissue is permissive for planar and apical-basal division
orientations that lead to symmetric or asymmetric cell fates, and that the increased ECM
deposition during injury physically alters the niche architecture to significantly favour symmetric
divisions in the planar orientation. Lastly, we observed that stem cell morphology is altered
depending on division orientations, suggesting a role of spindle pole arrangement in niche-
regulated fate determination. We report that the increased deposition of extracellular matrix
during regeneration modifies the niche architecture to support planar division orientations
leading to symmetric self-renewal of MuSCs, and postulate that the resulting distribution of
mitotic spindles plays a crucial role in determining cell fate.
20
RESULTS
Stiffness of bulk muscle increases during regeneration
Mechanical changes in the bulk muscle stiffness during regeneration were analyzed by
performing compression machine on regenerating muscle following BaCl2 injury. After injection
of BaCl2 directly in the middle of the tibialis anterior muscle, the animals were sacrificed at time
points ranging from immediate to seven days following injection to assess different stages of
muscle regeneration (Figure 2A). Compared to the uninjured control with a bulk stiffness of
13kPa, the bulk muscle stiffness 1 day after injury increased to 24kPa. The increased stiffness
persisted for another 2 days with a value of 26kPa at day 2 after injury and 24kPa at day 3 after
injury. After the fourth day, however, bulk muscle stiffness returned to pre-injury levels ranging
from 11kPa to 13kPa from days four to seven. The bulk stiffness values were statistically
significant between uninjured control and the first three days of regeneration (p<0.05), but was
not statistically significant when compared to days four to seven (p>0.05). No statistical
significance was reported between the first three days of regeneration or between the last four
days of regeneration. Standard deviation for all samples ranged from 0.5kPa to 4kPa, with
experiments each performed with three biological replicates (Figure 2B). Taken together, this
data suggests a significant increase in bulk muscle stiffness during the first three days of injury,
then a reduction of bulk muscle stiffness to pre-injury levels from days four to seven.
The muscle stem cell niche stiffens during injury repair
To determine if the niche surrounding the muscle stem cell has an altered physical
property during injury repair, we evaluated cryosectioned regenerating bulk muscles in addition
21
to local stiffness of muscle stem cells on isolated regenerating fibers. Immunostaining
cryosectioned bulk muscles during the process of regeneration enabled us to visualize dynamic
changes in the ECM composition following injury. Indeed, we observed using Pax7 and
collagen-I staining that there was an increase in the amount of cell infiltration and ECM
deposition at three to four days following BaCl2, which dwindled after two weeks (Figure 3).
This qualitative assessment of the local ECM environment surrounding MuSCs led us to utilize
more accurate methods to probe local depositions immediately enveloping the stem cell.
The use of atomic force microscopy (AFM) enables the analysis of physical properties
such as stiffness at a nanoscale level by utilizing a sensitive rounded tip that repeatedly indents
the surface of the sample. Muscle fibers were selected as the experimental sample due to the fact
that fiber isolation harvests the muscle fiber and associated Pax7+ MuSCs in addition to
maintaining the presence of the enveloping basement membrane. Since the objective was to
investigate local stiffness of regenerating samples, we injected barium chloride (BaCl2) directly
into the tibialis anterior muscle of the mouse, and isolated the neighboring extensor digitorum
longus muscle, seven days post-injury (Figure 4A). Using AFM, we probed regions local to the
MuSC using a 5nN indentation force and found that regenerating fibers had an average of 0.5
kPa stiffness in contrast to the non-injured muscle fiber at 0.2 kPa. These values were calculated
to be statistically significant (p<0.05) with three biological replicates and standard deviations of
0.07 kPa for both injured and non-injured fibers. In other words, the local stiffness surrounding
MuSCs as measured using AFM was found to be 2.7-fold greater in comparison to an uninjured
control mouse (Figure 4B). Immunostaining of laminin and Pax7 on regenerating myofibers
revealed a subtle increase in laminin staining surrounding MuSCs in the regenerating fiber
compared to the uninjured control (Figure 4C).
22
To validate that the measurements were reflective of the extracellular matrix (ECM)
surrounding the MuSC, we treated the fibers in a separate experiment with plasmin to degrade
laminin, fibrin, fibronectin, and other major components of the ECM (Figure 5A). After 1 hour
treatment of muscle fibers with plasmin, qualitative assessments of immunohistochemistry with
antibodies specific to laminin revealed a decrease in ECM protein expression relative to fibers
that were not treated with plasmin (Figure 5B). Furthermore, using AFM, we found that the
stiffness of plasmin-treated decreased a statistically significant amount of 4.2-fold (p<0.05)
compared to the control. Experiments were performed with three biological replicates with a
standard deviation of 0.05kPa (Figure 5C). The plasmin-treated experiments provide evidence
that the ECM significantly contributes to the stiffness of the fiber. Together, these results
utilizing the AFM to detect local stiffness surrounding the MuSC reveals that the niche stiffens
during regeneration from the contribution of ECM components in the basement membrane.
A stiff niche constrains spindle pole orientation to support planar divisions
To mimic the soft, uninjured niche and the stiff niche experienced by MuSC in the setting
of regeneration, we embedded muscle fibers in varying weight/volume (w/v) compositions of
agarose gel (Figure 6A). This type of gel was selected primarily due to its ability to be
mechanically tuned depending on percentage agarose composition, and its relative biological
inertness which is important since we were investigating the contribution from niche stiffness
instead of cell-ECM interactions. To demonstrate that the stiffness of agarose gel can be robustly
controlled, compression testing analysis in collaboration with the Simmons lab was performed.
The Young’s Modulus values ranged from 5.9, 9.8, and 21.7 kPa which corresponded to the
0.5%, 1%, and 3% weight/volume agarose gels (Figure 6B). These distinct values indicated that
23
the stiffness of the agarose gel can be accurately achieved by preparing the appropriate
weight/volume mix.
After the muscle fiber was isolated, it was allowed to equilibrate overnight in growth
culture. After 24 hours, the fiber was embedded in a soft 0.5% w/v agarose gel with a Young’s
Modulus of 5.9kPa or a stiff 3% w/v agarose gel with a Young’s Modulus of 21.7kPa, as
measured by compression with a tensile machine. Using ethidium homodimer staining to analyze
cell viability (Figure 7A), we demonstrate that MuSCs found on fibers that were 3D-embedded
24 to 48 hours after isolation maintained 84.8% viability in soft gels and 84.5% viability in stiff
gels with a standard deviation of 0.54% and 1.85%, respectively, when quantifying three
biological replicates with at least 100 MuSCs analyzed each (Figure 7B).
Using timelapse microscopy over 48 hours of 3D culture, we also observed that the
MuSC divided in both planar and apical-basal orientations with respect to the fiber (Figure 8A).
Apical-basal division orientation occurred 72% of the time in the soft agarose gel and 16% of the
time in stiff agarose gel (Figure 8B). In contrast, planar division orientation occurred 28% of the
time in soft agarose gel and 84% of the time in the stiff agarose gel (Figure 8C). These first-
division events were recorded 111 times from at least three biological samples. Statistical
significance was observed between the percentage of apical-basal divisions between culture in
0.5% agarose compared to 3% agarose (p<0.05), in addition to the percentage of planar divisions
between culture in 0.5% and 3% agarose. These results suggest that MuSC have the flexibility to
divide in both apical-basal and planar division orientations when embedded in a soft 3D
environment, in contrast to a stiffened environment where they are restricted to divisions in the
planar orientation.
24
Biochemical cues synergize with niche architecture to support symmetric self-renewal
divisions
To determine cell fate after MuSC division in the apical-basal or planar orientation, we
performed 3D immunohistochemistry on fibers 48 hours post-isolation and 3D culture. Prior to
this, we had to demonstrate that potential changes in Pax7 and MyoD expression was not due to
differences in stem cell division or cells entering cell cycle. 5-ethynyl-2′-deoxyuridine (EdU)
was added to the culture media between 36 and 48 hours post-isolation to differentiate MuSCs
that have already divided from MuSCs exiting quiescence and activating, but not actively
dividing. We recorded the average number of MuSCs per myofiber and the percentage of EdU+
MuSCs in the soft and stiff condition, and we report that the average number of MuSCs per
myofiber in the soft and stiff environments were 8.5 and 8.6, respectively, and the average % of
EdU+ cells in the two environments were 64% and 66%, respectively (Figure 9A, B). No
statistical significance was found (p>0.05).
Once myofibers were fixed after allowed to divide (Figure 10A), antibodies specific to
Pax7 and MyoD were used to determine cell fate where MuSCs expressed Pax7 only
(Pax7+/MyoD-), co-expressed Pax7 and MyoD (Pax7+/MyoD+), or expressed MyoD only
(Pax7-/MyoD+), depending on its progression along the myogenic lineage (Figure 10B). For
fibers that were embedded in soft agarose, we report that out of all the EdU+ MuSCs that have
undergone cell division, 14% were Pax7+/MyoD- and 86% were Pax7+/MyoD+. In contrast,
EdU+ MuSCs on fibers that were embedded in stiff agarose resulted in 10% were Pax7+/MyoD-
and 90% were Pax7+/MyoD+. The percentage of MuSCs that were EdU+ were 64% and 66% in
the soft and stiff environment, respectively (Figure 10C). Together, this data shows that there is
no significant difference in fate determination between soft and stiff agarose gel environments.
25
The addition of Wnt3a or Wnt5a ligands to the culture media between 36 and 48 hours led to no
significant increase in symmetric divisions. Notably, despite no significant differences in fate
determination and cell cycling in soft or stiff environments, the addition of Wnt7a ligand led to a
specific increase in Pax7+/MyoD- in the stiff niche as shown by the increase of Pax7+/MyoD-
cells to 61% and the decrease of Pax7+/MyoD+ cells to 39%. To demonstrate that this effect is
synergistic and requires both the stiff environment and the presence of Wnt7a ligand, we
embedded MuSCs on myofibers in a soft environment and supplied the Wnt7a ligand. The
percentage of Pax7+/MyoD- cells decreased significantly to only 35% and the percentage of
Pax7+/MyoD+ cells increased to 65%. Together, this data supports that the synergy between
biochemical cues and niche architecture is necessary to significantly increase MuSC symmetric
self-renewal divisions.
Niche architecture impacts myoblast morphology
To determine if niche architecture impacts cell morphology, green fluorescent protein
(GFP) transgenic myoblasts were cultured in soft and stiff agarose for 24 hours in standard
culturing conditions. After fixing and staining with nuclear marker Draq5, a 3-D image was
produced by taking 1um z-stack slices using the FITC channel with the confocal microscope.
Qualitative assessments of GFP-myoblasts in soft agarose revealed an increase in cell size and
cytoplasmic area (Figure 11A) compared to cells cultured in stiff agarose, where cells were small
and spherical with low cytoplasmic content (Figure 11B). Taken together, these preliminary
results suggest that the niche architecture alone is sufficient to impact the morphology of muscle
progenitor cells.
26
DISCUSSION
Our first set of experiments, where a standard compression machine was used to apply
mechanical force on normal and injured bulk tissues, revealed that the stiffness of bulk muscle
changes over the course of regeneration. Acute injury in the form of barium chloride injection
was performed at day zero and animals were sacrificed at timepoints one to seven days after,
enabling us to analyze the progression of muscle regeneration over a period of a week. The
significant increase in muscle stiffness sustained over three days following initial injury suggests
physiological changes beyond a biochemical response. Indeed, immunohistochemical analysis of
cryosectioned regenerating muscle revealed a massive infiltration of immune cells in the first
days of regeneration, perhaps contributing to the increase in bulk muscle stiffness. The more
likely explanation for the immediate increase in stiffness is edema, where excess interstitial fluid
trapped in the injured regions causes high levels of swelling, combined with the presence of
fibrin clots to act as a hemostatic plug over the wound site. The return to pre-injury levels of
stiffness at days four and onwards reflects the clearance of immune cells, fibrin clots, and excess
interstitial fluid, but how this impact the local niche surrounding the MuSC remain to be seen.
The contribution of extracellular matrix deposition to the increased stiffness in the bulk tissue in
the first couple of days following injury is likely minimal, since it has previously been shown
that ECM-secreting fibroblasts require days to become activated and remodel the matrix110.
According to immunohistochemical analysis where the expression of laminin, collagen, and
fibronectin was observed to be increased at days five onwards but normalized at day twenty-one,
we postulate that the increase in ECM begins at a later time point approximately five to seven
days post injury and is cleared over three weeks to resemble a non-injured tissue. The use of bulk
stiffness measurements coupled with immunohistochemistry provides an indirect way to
27
understand the highly complex process skeletal muscle during regeneration as exemplified by the
time-dependent process of fibrin clot formation, interstitial fluid buildup, immune cell
infiltration, and deposition of extracellular matrix. While the use of compression testing is useful
for revealing bulk changes in muscle stiffness, perhaps more sensitive approaches are necessary
to capture the subtle changes in ECM deposition at the niche level.
To address this issue, we used an atomic force microscope performed by Haijiao Liu
from Dr. Craig Simmons’ lab at the University of Toronto to determine local changes in ECM
deposition at the niche level during regeneration. AFM was used to indent the regions
surrounding normal and injured MuSCs on fibers, and led to the observation that the stem cell
niche stiffens during regeneration. The use of muscle fibers enabled us to investigate the impact
of ECM on MuSC properties since the basement membrane containing ECM proteins such as
laminin remains associated following extraction. By comparing injured fibers that were allowed
to regenerate for seven days with normal fibers that were isolated without injury, we were able to
make direct comparisons of substrate stiffness. Since the AFM only indented the less than 1um
of the surface, or less than 10% of the total height of the myofiber, we were confident that only
the extracellular matrix was being measured. Deeper indentations would possibly reveal the
stiffness of the underlying sarcolemma region, but for the purposes of this experiment we were
only interested in the contribution of the ECM to mechanical properties of the niche.
Since it has previously been established that the extracellular matrix has quantifiable
mechanical properties such as rigidity, we postulated that alterations in the niche during
regeneration must correspond to changes in stiffness measurements. The use of AFM enabled us
to measure the stiffness of the thin layer of ECM local to the muscle stem cell with high
sensitivity. Indeed, our observations that the Young’s Modulus value was more than two-fold
28
higher in the regenerating tissue suggested that the mechanical properties of the ECM were
modified. However, increases in the stiffness could be indicative of several changes, including
increased protein-protein crosslinking, altered protein configuration, or heightened levels of
protein production. Here, we focused on investigating whether ECM production increased in the
fiber model based on previous work on immunostained transverse cryosections of muscle tissue,
where we found and increased level of ECM protein expression in BaCl2 injured muscles. Using
a qualitative approach, we performed immunohistochemistry on isolated fibers from healthy or
injured animals and probed for ECM protein expression, and observed that injured fibers had
subtle increases in ECM protein immunostaining compared to the healthy control. Both
qualitative assessments using immunohistochemistry and quantitative analysis using AFM
suggested that the increase in stiffness is in part due to heightened levels of ECM protein
production.
If the increase in ECM protein levels corresponds to the increase in niche stiffness, then
removal of the ECM must lead to a significant decrease in niche stiffness. To support this notion,
we treated healthy fibers with plasmin, which degrades laminin, fibronectin, and fibrin
associated with the ECM. Since the results indicated that the plasmin-treated fiber was
significantly softer than a non-treated control, we concluded that the ECM plays a significant
role in determining the stiffness of the muscle fiber. Thus, these results indirectly suggest that the
increase in ECM deposition as observed previously contributes to the increase in fiber stiffness
during muscle regeneration. Future investigations would likely involve plasmin treatment on
injured fibers compared to non-plasmin treated injured fibers to determine if the Young’s
Modulus levels will be reduced to softer levels exhibited by non-injured fibers. Additionally, it
would be interesting for additional experiments to identify the contribution of ECM protein
29
configuration and crosslinking to substrate stiffness, possibly through the use of second
harmonics generation microscopy to directly visualize collagen composition in the niche without
immunohistochemistry. It is likely that an ECM characterized by a soft, strand-like composition
of collagen fibrils, for example in a healthy niche, will have drastically different mechanical
properties compared to bundles of cross-linked collagen aggregates postulated to be found in a
regenerating niche. Since muscle fibers are isolated from muscle bundles through treatment with
collagenase which digests surrounding collagen, further optimization experiments will need to be
performed to find a balance between sufficiently weakening the ECM to enable fiber isolation
with preserving as much of the ECM for SHG analysis. Regardless, our first result here provides
evidence that an increase in ECM protein quantity directly leads to a stiffened MuSC niche in the
context of muscle regeneration.
Our first set of experiments demonstrated that during the process of regeneration, the
niche transforms from a soft environment with low levels of ECM protein to a stiffened
environment with increased levels of ECM protein. To model these two modalities, our next sets
of experiments utilized an in vitro culturing platform using agarose gel, prepared at 0.5%
weight/volume to mimic the soft environment, and prepared at 3% weight/volume to mimic the
stiffened environment. We recognize that MuSCs are likely exposed to niche stiffness that range
between the values that we prepared, but we were interested in observing cell behaviour at
extreme measurements first before teasing out subtle differences closer to physiological ranges.
According to our analysis using a standard compression machine on thin layers of agarose gels,
we found that the value of 0.5% w/v agarose gel was 12 kPa, and the value of the 3% w/v
agarose gel was 24 kPa, which corresponded to the stiffness of healthy and injured bulk muscle,
respectively. While our measurements focused on a more narrow range of agarose gel stiffness,
30
our measurements corroborated with values reported by other groups111. The decision to use
agarose gels as opposed to commercially available poly-ethylene glycol (PEG) or collagen/fibrin
gels was due to our commitment to investigate only the contribution of niche architecture. While
previous literature has established that cells can sense an increase in ECM deposition or altered
crosslinking of ECM components through integrin signaling, we were interested in whether the
increase in stiffness could indicate a decrease in the pliability of the niche architecture. The
biologically inert property of agarose gels prevents cell attachment to the surrounding ECM
aside from its own basal lamina, so any reasons for an altered cell fate would be limited purely to
the contributions of stiffness and substrate pliability, and independent from ECM ligand-
mediated mechanotransduction.
Viability assays where MuSCs were embedded in soft and stiff agarose gels indicated
that more than 85% of cells remained alive over 36 hours of 3D culture, highlighting that the
agarose gel is sufficiently porous for nutrient exchange and waste management. No statistically
different results for viability were reported between soft and stiff environments, suggesting that
3D niche architecture is not a contributing factor to cell viability. However, it is important to
acknowledge that a portion of cells (~15%) were consistently dying compared to fibers cultured
in growth media without embedding in 3D agarose where viability is close to 100%, indicating
that either being in a 3D environment or being surrounded with agarose negatively impacts
survival of the muscle stem cell in culture. Regardless, the high viability of MuSCs in 3D
agarose culture opened up the possibility of tracking cell behaviour over an extended period of
time using confocal timelapse microscopy.
We observed both types of division when tracking individual MuSCs in 3D agarose:
apical-basal orientation division, where the daughter cell emerge in a stacked formation and
31
planar orientation, where both daughter cells emerge next to another relative to the fiber. After
quantifying our observations, we found that the first division of MuSC in soft substrates was
typically apical-basal division in orientation, while cells in stiff environments predominantly
underwent division in the planar orientation. We observed that the first division occurred usually
at the 40-hour mark after isolation from the bulk tissue, which is consistent with other reports in
the literature65. Additionally, we noticed that the cell adopted a round phenotype and increased in
size prior to division, which is also consistent with reports involving other cell types112. Since the
only variable was cell culture in different % w/v preparation of agarose gel, these results suggest
that niche architecture alone can impact division orientation of muscle stem cells. If we
extrapolate this information to address physiological changes during regeneration, we can
conclude that the systematic deposition of ECM in the stem cell niche is supportive of specific
division orientations. Specifically, a soft niche with low levels of ECM protein as modelled by a
0.5% agarose gel platform is sufficiently pliable to enable both apical-basal and planar division
orientations, whereas a stiff niche with high levels of ECM protein as modelled by a 3% agarose
gel platform has a restrictive niche that prevents cells to stack on top of each other during apical-
basal orientation division, and is instead only supportive of planar type divisions.
Lastly, we show that biochemical cues synergize with biophysical characteristics such as
the niche architecture to support symmetric self-renewal divisions. MuSCs in fibers embedded in
soft and stiff agarose gel without the addition of any Wnt ligands displayed no significant
difference in cell fate despite variation in cell division orientation. Specifically, out of all MuSCs
quantified, equal proportions of Pax7+/MyoD-, Pax7+/MyoD+, and Pax7-/MyoD+ was observed
in both environments, suggesting that niche architecture alone was insufficient to impact MuSC
fate determination.
32
However, the addition of soluble ligand Wnt7a in the culture media in the stiff
environment during the cell division timeframe shifted the balance to favor symmetric self-
renewal divisions, as indicated by the increased proportion of Pax7+/MyoD- cells. Our
observations are in agreement with previous reports suggesting that Wnt7a acts as the primary
ligand for the non-canonical Wnt pathway to distribute Vangl2 in the poles of the cells to
increase symmetrical expansion. The slight variation in the proportions of Wnt7a-treated
Pax7+/MyoD- cells compared to previous papers adding the soluble factor without the context of
3D suggest that the rigid niche architecture and increased availability of soluble ligands during
regeneration may have a cumulative impact on increased self-renewal divisions. In contrast, the
addition of Wnt3a to the MuSCs in the stiff gel had no impact on the proportion of
Pax7+/MyoD+ cells or quantity of Pax7+/MyoD- cells, indicating that this particular
biochemical cue does not have an effect on stem cell self-renewal or differentiation in this
context. Previous reports investigating the role of the Wnt3a ligand in determining cell fate have
reported an increase in differentiated cells, but it is important to note that those experiments were
performed in a 2D environment. Lastly, the addition of Wnt5a to the MuSC in 3D culture
resulted in no change in the ratio between Pax7+/MyoD+ and Pax7+/MyoD- cells reported in the
control with no added ligands, suggesting that the ligand also does not synergize with niche
architecture to promote stem cell self-renewal or differentiation. The Wnt5a has been reported in
the literature to be the ligand for an ‘integrated’ Wnt pathway that utilizes aspects from the
canonical and non-canonical Wnt pathways to drive gene expression, but more experiments need
to be performed to understand its role in MuSC self-renewal.
The results from this section add on to existing literature investigating the effects of 3D
culture and soluble ligands activating signaling pathways important for MuSC self-renewal and
33
differentiation. Previous reports investigating uninjured MuSC activity in 3D collagen culture
has found that 65% of cells undergoing their first division were in the planar orientation113.
Studies analyzing injured MuSC on fibers in floating media culture have found that the amount
of planar divisions increases to 91%24. In this study, culturing healthy fibers in a stiff
environment as opposed to soft changed the percentage of planar divisions from 40% to 80%,
which suggests that our stiff agarose system is effective at mimicking the ECM-rich, injured
environment found in regenerating muscle. MuSCs on healthy myofibers cultured in collagen
gels have reported a similar ratio of planar and apical-basal division orientation, but here we
clarified the contribution from biophysical cues such as stiffness and niche architecture through
the use of biological inert agarose gels. Furthermore, in support of existing literature, the
addition of soluble ligands Wnt7a, Wnt3a, or Wnt5a in a 3D environment shifted MuSC identity
to various fates reflecting both symmetrical and asymmetrical cell divisions. Here, we
demonstrate that biophysical cues such as niche architecture synergize with biochemical cues
like Wnt ligands to regulate MuSC self-renewal and differentiation.
While we have demonstrated a synergistic effect between niche architecture and soluble
ligands, the mechanism regulating this phenomenon remains elusive. Our preliminary
observations of myoblast increase in roundness and decrease in size when cultured in stiff
agarose compared to soft gel led us to postulate that niche architecture may restrict cell
morphology to potentially regulating mitotic spindle orientation. The pliability of a soft agarose
gel may enable increased cell stretching and permit movement of mitotic spindles to apical-basal
poles of the stem cell. In contrast, we speculate that the stiff environment present in high
weight/volume agarose gel not only reduces the cytoplasmic space but also limits movement of
spindle pole to only allow divisions in the planar orientation. Specifically, we would expect that
34
the majority of MuSCs in the soft environment to have mitotic spindles oriented parallel to the
underlying sarcolemma, with microtubules pulling towards the top and bottom of the cell. In
contrast, we would expect the majority of MuSCs in the stiff environment to have mitotic
spindles perpendicular to the underlying sarcolemma, with microtubules pulling towards the
sides of the cell. Our previous observations showing an increased amount of apical-basal
divisions in the soft 3D environment in contrast to the stiff environment where planar division
dominate would then support this idea that spindle pole orientation and microtubule organization
is associated with division orientation. We assume that parallel spindle pole orientation coupled
with microtubule arrangement towards the top and bottom of the cell enables the physical split of
the cell to support division in the apical-basal orientation. Conversely, perpendicular spindle pole
orientation associated with microtubule arrangement towards the sides of the cell enables the
physical split of the cell to support division in the planar orientation. Future studies will likely
involve identifying the mechanism between cell shape and spindle pole orientation in the context
of MuSC divisions. Additionally, it would be interesting to see if the physical arrangement of
spindle poles due to the niche architecture is associated with specific sites of soluble ligand
interaction. If so, this may point towards a mechanism that explains the synergistic effect that we
observe in our experiments.
The fate of muscle stem cells has traditionally been believed to be regulated by
interactions with surrounding support cells that signal via paracrine biochemical mechanisms.
However, more support for the biomechanical contribution of the ECM has emerged as
regulators of MuSC fate. Here, we highlight the importance of niche architecture in providing
structural support in tandem with soluble cues to direct cell division orientation, spindle
alignment, and myogenic progression. Using both immunohistochemistry and atomic force
35
microscopy on muscle stem cells on myofibers, we show that the stem cell niche stiffens during
the process of tissue repair. Our use of agarose gel to mimic the stiff injured environment and the
soft healthy environment led us to determine that a stiff niche constrains muscle stem cell
division and spindle pole orientation. Lastly, using confocal timelapse microscopy and 3D
immunostaining, we show that biochemical cues in the form of Wnt ligands synergize with niche
architecture to regulate symmetric or asymmetric cell divisions. Together, our results contribute
to unveiling the intricate biophysical and biochemical regulatory network that regulates stem cell
division orientation and fate determination in the context of tissue regeneration.
36
MATERIALS AND METHODS
Animals
All animal protocols and experimental procedures were approved by the Division of
Comparative Medicine at the University of Toronto. C57BL/6 wild type mice were obtained
from Jackson laboratories, and Pax7-zsGreen reporter mice were gifted from the Kyba lab in
Minnesota. We validated all generated animal genotypes by PCR-based strategies. All mice used
in these studies were female and between 6-10 weeks of age with a median age of 8 weeks. For
all muscle depolarization injuries, a single injection of 30ul barium chloride (1.2% w/v; Bio
Basic) was administered directly in the center of the tibialis anterior muscle of the recipient
animal. For injury experiments, muscles were isolated up to 7 days after the initial barium
chloride injection.
Bulk muscle compression testing
Compression testing was performed on freshly isolated bulk TA muscle from young (8-week
old), aged (20 month-old), uninjured (8-week old, no BaCl2 injection), and injured (BaCl2
injected) mice. Bulk muscle was gently placed onto a flat platform and compressed to 10% of
maximum thickness. Dimensions of the isolated bulk TA muscle were visualized using the
Navitar camera. Five cycles of addition 10% compression was then immediately performed
using TestResources tensile machine. Mechanical stress and strain was calculated using
TestResources software and Microsoft Excel to give the Apparent Modulus from the respective
tissues according to the following formula:
37
𝐸 =𝑇𝑒𝑛𝑠𝑖𝑙𝑒 𝑆𝑡𝑟𝑒𝑠𝑠
𝐸𝑥𝑡𝑒𝑛𝑠𝑖𝑜𝑛𝑎𝑙 𝑆𝑡𝑟𝑎𝑖𝑛=
𝐹/𝐴O
∆𝐿/𝐿o
where
E is the Young’s Modulus;
F is the force exerted on an object;
Ao is the cross-sectional area of sample;
ΔL is the amount of length change;
Lo is the original length of sample.
For regeneration experiments, injected inorganic compound BaCl2 dissolved in Phosphate
Buffered Saline (PBS) caused damage to the bundled fibers within the TA muscle via fiber
depolarization, and the animal was allowed to recover for one to seven days after injury and then
sacrificed for tissue compression experiments.
Muscle fiber isolation and plasmin treatment
Both left and right extensor digitorum longus muscles were dissected from mice and subjected to
45 minutes of collagenase (630 units/ml; Sigma) in Dulbecco’s modified eagle medium (DMEM)
digestion (Life Technologies). The bulk muscle was then repeatedly flushed with warmed media
to separate individual strands of muscle fibers. Every 5-10 minutes of this process, the muscle
was returned to the incubator to maintain the culture temperature close to 37 degrees Celsius.
Isolated fibers were then transferred to a plate containing pre-warmed growth media (80%
Ham’s F12 media; Life Technologies, 20% Fetal Bovine Serum from Gibco; Life Technologies)
for 24 hours. For plasmin-treatment experiments, 1:100 dilution of plasmin (2 units/mg; Sigma-
Aldrich) was added to the culture media containing the isolated fibers for 1 hour prior to
38
measurements. Muscle fibers from regenerating EDL muscles were isolated from animals that
received BaCl2 injection in the TA muscle seven days prior to tissue isolation.
Atomic force microscopy
A layer of collagen solution was placed on top of charged glass microscope slides and incubated
overnight in 4 °C to create a collagen-coated surface to facilitate fiber attachment. Isolated fibers
with 500 µl of warmed media were then gently placed on top of the slide surface and allowed to
attach for 30 minutes. Muscle fibers were tested using a commercial AFM (BioScope Catalyst,
Bruker) mounted on an inverted optical microscope (Nikon Eclipse-Ti). The force-indentation
measurements were done with a spherical tip at more than three distinct locations of sarcolemma
regions local to the MuSC per fiber. The measurements were repeated on three biological
samples. The indentation rate was set to be 1 Hz. The spherical tips were made by carefully
assembling a borosilicate glass microsphere (15 μm radius) onto an AFM cantilever with epoxy
glue. The cantilever (MLCT-D, Bruker) had a nominal spring constant of 0.03 N/m. The trigger
force applied to the fiber was varied from 2 nN to 5 nN, 10 nN for comparison of effects of
indentation depth. No explicit correction for finite sample thickness effects was made here. The
Hertz model was applied to the force curves to estimate the Young's modulus and contact point.
We repeated indentation at the same location of the fiber five times and observed no significant
change in the Young’s moduli. Since the Young's modulus calculated from the Hertz model is
sensitive to the spring constant, cantilever spring constants were calibrated before running the
experiment by measuring the power spectral density of the thermal noise fluctuation of the
unloaded cantilever. Detailed method in using spherical tip and data analysis were described
elsewhere114,115. All AFM measurements were done in the fluid environment at room
temperature.
39
3D muscle fiber culture preparation
0.5% and 3% agarose gels were prepared from agarose powder (Bioshop) dissolved in F-12
media. A 200ul base layer was created on a 12-well dish and covered with growth media for 24
hours, then aspirated. Isolated fibers were then gently placed on the gel and allowed to
equilibrate for 30 minutes. A second layer of agarose gel cooled to 40 degrees Celsius was then
added on top of the fibers. Once the second layer solidified, growth media was added and the
plate was placed in a 37 degrees Celsius incubator. To track MuSC proliferation, EdU (1:1000
from Click-iT EdU Imaging Kit, Invitrogen) was added to the media between 36 and 48 hours
after fiber isolation. Soluble Wnt ligands 3, 5, or 7 was added to the media during this period.
3D immunostaining
Embedded fibers were first washed with PBS three times to remove residual culture media, then
fixed with 4% paraformaldehyde for 30 minutes. After further washing to remove fixative,
samples were then blocked with blocking solution (1% BSA, 0.5% Triton-X, in PBS) rocking
overnight in 4 degrees Celsius to minimize non-specific staining and background. Individual
islands of gels containing fibers were excised and placed in a 1.5ml Eppendorf tube containing
primary antibodies and placed overnight in 4 degrees Celsius. Primary antibodies used included
mouse anti-Pax7 (1:5, gifted from Dr. Libero Vitiello, University of Padova), rabbit anti-MyoD
(1:200, Abcam), and washed overnight at 4 degrees Celsius. Subsequently, the cells were
incubated with goat anti-mouse AlexaFluor488-conjugated (1:500, Life Technologies) and goat
anti-rabbit546-conjugated (1:500, Life Technologies) secondary antibodies, for 30 minutes at
room temperature. After further washes, fibers were analyzed under confocal microscopy, where
40
images from a single z-plane were obtained. Image analysis was performed using ImageJ
software and data was compiled using Microsoft Excel and Graphpad.
Statistical Analysis
All experiments, including AFM analyses, were performed with at least three biological
replicates with at least three technical replicates per animal. For single-cell viability,
proliferation, and immunofluorescence assays, we report the number of individual cells
quantified in the legends to reach statistical significance. We used unpaired, two-tailed student t-
tests to compare conditions for injured versus uninjured, plasmin-treated versus control, and fiber
culture in soft versus stiff environments. For all tests, we used a significance level of α= 0.05.
41
APPENDIX
Increase of bulk muscle stiffness in aged muscle
The systematic deposition of extracellular matrix between muscle fibers as individuals
age has been established by previous literature. However, the link between changes in local
environment with bulk muscle stiffness has yet to be determined. To establish if there are
mechanical changes in the bulk tissue between aged and young tissue, we performed
compression testing using a standard tensile machine on freshly isolated bulk TA muscle from
young and aged mice (Appendix 1A). Compression testing analysis with three biological
replicates revealed that young bulk TA muscle had an average Young’s Modulus value of 14kPa
compared to the aged bulk muscle average value of 28kPa. Standard deviation was 4.8kPa and
2.6kPa for young and aged tissues, respectively, and statistical significance was reported with
p<0.05 at n=3 (Appendix 1B). These results suggest that the cells, depending on age, may be
responding to different physical cues such as stiffness. Nevertheless, additional experiments need
to be performed to determine if these age-related differences exist at the local stem cell level. We
postulate that the deposition of ECM for an aged organism may be so severe that the niche
architecture becomes significantly stiffer even in comparison to a regenerating niche, leading to
the stem cell’s impaired ability to divide.
Diaphragm as a long-term ex vivo model for MuSC behaviour
While MuSCs on fibers are advantageous in modelling the contributions of niche
architecture to stem cell fate, it fails to capture many key interactions in vivo. As previously
mentioned, a myriad of signal sources ranging from neighbouring cells, surrounding vasculature,
42
and other paracrine signals are missing in the 3D agarose in vitro assay outlined in this thesis. To
address this issue, we propose the use of the isolated diaphragm as a model for ex vivo imaging,
which can enable long-term monitoring of MuSC behaviour following controllable stimuli such
as physical injury. Furthermore, the diaphragm is a well-established tissue in the muscle field,
with numerous links to diseases such as Duchenne Muscular Dystrophy. The presence of Pax7+
stem cells has been extensively outlined in the literature, and we have confirmed this by
successfully immunostaining Pax7+ MuSCs in the cryosectioned diaphragm (Appendix 2).
However, in order to proceed with cell tracking over time, it is critical to establish that the tissue
remains alive in culture.
Preliminary studies utilizing ethidium homodimer assays have revealed that the excised
diaphragm, which is still attached to the surrounding costal cartilage to maintain fiber tension,
remains viable for at least four days in culture media consisting of DMEM + 10% FBS with
daily media exchange. Specifically, the quantity of cells that stained negative for ethidium
homodimer remained at above 90% for four days compared to the 60% viability for injured
tissue (Appendix 3). For these viability experiments, three randomly chosen regions of interest
were collected via confocal microscopy and assessed for cell viability.
To study regeneration, a local cryoinjury was inflicted onto a region of the isolated
diaphragm tissue. For these experiments, a metal pin with rounded end was frozen in liquid
nitrogen and applied to the surface of the left costal muscle region of the diaphragm for ten
seconds without puncture. Appropriate media was immediately added and the tissue placed in 37
degree incubation. This method of injury was determined to be a better model compared to a
mechanical puncture wound since it more closely resembles atrophy in muscle tissue as it is
highly unlikely that an organism can recover from a puncture wound in the costal area.
43
Cryoinjury experiments revealed that the injured region had 51% average viability, and distal
regions showed 61% viability compared to the 87% viability of the non-injured control,
suggesting long-range effects of cryoinjury (Appendix 3). Together, these results point to a
reliable method to locally induce injury without causing total cell death.
By demonstrating that tissue viability of isolated diaphragm tissue can be maintained ex
vivo in tissue culture conditions and that reproducible local injury is possible, future experiments
will focus on tracking individual cell fates over time. Through the use of transgenic animals such
as the Pax7zsGreen mouse gifted by the Kyba lab, it may be possible to track cell behaviour as it
responds to injuries. Since the diaphragm tissue is relatively thin compared to other bulk muscles
responsible for strength and weight lifting, this allows for effective confocal microscopy. We
envision future experiments to track Pax7zsGreen-positive cells in real time without the need for
fixation and immunostaining after inflicting cryoinjury to stimulate regeneration. However,
many hurdles need to be overcome, such as reducing the high levels of background from auto-
fluorescing dead cells, and finding the right immunological stimuli since there is a lack of an
active vasculature system in this model. Nevertheless, we believe that the use of ex vivo systems
as exemplified by the isolated diaphragm will aid in revealing the intricate timings and
contributions of multiple cell types to the complex, but fascinating muscle stem cell-based tissue
regeneration.
44
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FIGURES
Figure 1. The skeletal muscle stem cell niche architecture. The niche architecture is defined
as the organization and the composition of the extracellular space surrounding the MuSC that can
either be pliant or rigid. In a healthy niche, the architecture is permissive for cell division in both
apical-basal orientations. In a regenerating niche, the stiffened environment is supportive of
primarily planar division orientations.
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Figure 2. The bulk stiffness of skeletal muscle is dynamic over the course of regeneration.
(A) Schematic of ex vivo compression testing experiments. Dissected tibialis anterior muscles at
daily timepoints following day 0 barium chloride injection were subject to compression testing
and analyzed using TestResources software. Mean (SD) of n=3 per timepoint was calculated
with both left and right muscles isolated amounting to 48 muscles total. (B) Side and top-down
view of isolated TA muscle, ruler in millimeters. (C) Representative stress vs. strain test of a
complete cycle (top) and linear region only (bottom) to obtain apparent modulus by calculating
slope (D) Black bars represent average Apparent Modulus values of regenerating muscles for
each date, with significant difference only between control and each of the first three days
following injury. p<0.05 (*) by One-way ANOVA.
55
Figure 3. Systemic changes occur in the MuSC niche during regeneration. Dissected tibialis
anterior muscles were fixed, cryosectioned, and immunostained for laminin at daily timepoints
up to fourteen days following day 0 barium chloride injection with experimental control.
Representative images were stained green for collagen-I and blue for cell nucleus. Images were
taken at 20X magnification using fluorescence microscopy. Scale bar: 100um.
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Figure 4. The stiffness of MuSC niche is greater after injury compared to healthy control.
(A) Schematic of AFM experiments on quiescent and regenerating myofibers. Isolated myofibers
from extensor digitorum longus muscles from healthy or injured muscle allowed seven days to
regenerate following day 0 barium chloride were analyzed using atomic force microscopy with
an indentation force of 5nN injury. Mean (SD) of n=3 myofibers per experimental group were
determined from at least three regions local to distinct muscle stem cells probed per myofiber.
(B) Left panel: average Young’s Modulus values of quiescent or regenerating fibers. Right panel:
fold-change of Young’s Modulus value of myofibers after injury. p<0.05 (*) by Student’s T-test.
(C) Immunofluorescent images of quiescent and regenerating myofibers stained for laminin in
red and Pax7 in green. Magnification: 20X, scale bar: 50um.
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Figure 5. The stiffness of plasmin-treated MuSC niche is significantly lowered compared to
non-treated control. (A) Schematic of AFM experiments on myofibers treated with plasmin.
Isolated myofibers from extensor digitorum longus muscles from healthy non-injured animals
were treated with or without 100nM plasmin for one hour and analyzed using atomic force
microscopy with an indentation force of 5nN. (B) Immunofluroescent images of control and
plasmin-treated myofibers stained for Pax7 in green and laminin in red. Magnification: 20X,
scale bar: 50um. (C) Left panel: average Young’s Modulus values of control or plasmin-treated
fibers. Right panel: fold-change of Young’s Modulus value of myofibers after injury. Mean (SD)
of n=3 myofibers per experimental group were determined from at least three regions local to
distinct muscle stem cells probed per myofiber. Not significant (n.s.), p<0.05 (*) by Paired T-
test.
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Figure 6. The stiffness of agarose gel corresponds with weight/volume percentage
composition. (A) Side and top view of agarose gels. Agarose gels were prepared with 0.5, 1, and
3 % weight per volume and subjected to compression testing and analysis using TestResources
software. Mean (SD) of n=5 gels were calculated per experimental condition. (B) Average
Young’s Modulus values of 0.5%, 1%, and 3% agarose gels, with significant differences
between each % w/v preparations. p<0.05 by Student’s T-test.
Figure 7. Muscle fibers remain viable when 3D cultured in soft and stiff agarose. Isolated
myofibers from extensor digitorum longus muscles from healthy Pax7-zsGreen mice where
Pax7+ MuSCs fluoresced green were treated with ethidium homodimer after 48 hours of 3D
culture in soft (0.5%) or stiff (3%) agarose gel. (A) Merged images Pax7zsG (green) and
ethidium homodimer (red) staining representing live and dead MuSCs in the left and right
panels, respectively. (B) Graph indicating average percentage of ethidium homodimer negative
staining in each experimental condition. Mean (SD) of n=3 animals with at least 10 fibers per
animal included with all muscle stem cells analyzed. Not significant (n.s.) by Student’s T-test.
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Figure 8. Muscle stem cell division orientation differs between 3D culture in soft versus stiff
environments. (A) Schematic of timelapse microscopy analysis of MuSC division orientation.
Isolated myofibers from extensor digitorum longus muscles from healthy mice were embedded
in soft (0.5%) or stiff (3%) agarose gel 36 to 48 hours following isolation. Confocal timelapse
microscopy was used during this time period to capture first division events of individual MuSCs
on myofibers (n=3 animals, at least 50 division events experimental conditions quantified). (B)
Representative stills of MuSC undergoing apical-basal division orientation. Right panel: analysis
of the percentage of apical-basal division orientations in soft or stiff gel. (C) Representative stills
of MuSC undergoing planar division orientation. Right panel: analysis of the percentage of
planar division orientations in soft or stiff gel. Mean (SD) of n = at least 6 MuSC on 10 fibers,
repeated with 3 animals, with a total of 61 and 53 division instances in soft and stiff gels,
respectively. P<0.05 (*) by Student’s t-test.
60
Figure 9. MuSC count and percentage of EdU+ MuSC population does not change in soft
or stiff environments. Isolated myofibers from extensor digitorum longus muscles from healthy
mice were embedded in soft (0.5%) or stiff (3%) agarose gel 36 to 48 hours following isolation.
At t=48 hours after isolation, fibers were immunostained for Pax7, MyoD, and EdU expression.
(A) Average quantity of MuSCs per myofiber in soft and stiff conditions. (B) Percentage of
EdU+ MuSCs in soft and stiff conditions. Mean (SD) of n=3 animals, with at least 50 MuSCs per
animal for each experimental condition analyzed. No significance (n.s.) by Student’s t-test.
61
Figure 10. Biochemical cues synergize with niche architecture to support symmetric self-
renewal divisions. (A) Schematic of timelapse microscopy analysis of MuSC division
orientation with the addition of Wnt soluble ligands. (B) Representative immunostaining of Pax7
(green), MyoD (red), and EdU (grey) on MuSCs from isolated myofibers embedded in agarose
gel. Top panels: Pax7+/MyoD-/EdU+, bottom panels: Pax7+/MyoD+/EdU+. Scale bar, 50 µm.
(C) Quantification of EdU+, Pax7+/MyoD+ and Pax7+/MyoD- MuSCs in soft versus stiff
environments. Mean (SD) of n = at least 41 cells per condition, repeated with 3 animals each. (D)
Quantification of EdU+, Pax7+/MyoD+ and Pax7+/MyoD- MuSCs in stiff environments with
the addition of Wnt3a, Wnt5a, and Wnt7a ligands, with Wnt7a in soft gel control. Mean (SD) of
n = at least 41 cells per condition, repeated with 3 animals each. (c, d) Not significant (NS), P <
0.05 (*) by Student’s t test.
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Figure 11. Niche architecture impacts myoblast morphology. Green fluorescent protein
(GFP) transgenic myoblasts fixed and stained with nuclear stain Draq5 after 24 hours in culture
in soft (left panel) or stiff (right panel) agarose gels. Yellow: nuclear staining, Red: cytoplasm of
GFP myoblast. Magnification: 40X. Scale bar: 30um.
63
APPENDIX FIGURES
Appendix 1. The stiffness of bulk TA muscle is greater in aged mice compared to young
mice. (A) Dissected tibialis anterior muscles were subject to standard compression testing and
analyzed using TestResources software. Mean (SD) was determined from n=3 animals per age
group, with both left and right muscles isolated amounting to 12 muscles total. (C) Young’s
Modulus of young (2 month old) and aged (20+ month old) mice, respectively. The Young’s
Modulus value of aged mice was significantly greater than young. p<0.05 (*) by Student’s T-
test.
Appendix 2. Mouse diaphragm as a platform to visualize in vivo changes to MuSC
behavior after exposure to injury or artificial stiffness. Representative image of isolated
murine diaphragm in 35mm plastic dish, aerial vs. side (left panel) view. Middle panel: TD1
laser absorption visualized at 4X magnification demonstrates capacity to visualize fiber integrity
and surrounding vasculature. Right panel: Pax-7+ cells are present in the murine diaphragm.
10um Transverse section of immunostained mouse diaphragm. Green cells represent MuSC,
counterstained with Hoechst for nuclear staining and anti-laminin for visualization of the
extracellular matrix.
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Appendix 3. Diaphragm viability in culture +/- cryoinjury. Top left panel: representative
image of uninjured diaphragm at 20X magnification stained for cell nuclei (blue, Hoechst) or
dead cells (red, ethidium homodimer). Bottom left: ethidium homodimer assay shows that cells
at three random regions in the uninjured diaphragm have greater than 90% viability, compared to
an injured region where 65% viability is observed (p<0.05). Top right: non-injury control is a
ROI from an uninjured diaphragm, cryoinjured is ROI of injury, and cryoinjure control is a
region not directly cryoinjured. Bottom left: cryoinjured ROI has significantly less viability
compared to non-injury control. Mean (SD) with 3 biological replicates. Not significant (n.s.),
p<0.05 (*) by Student’s T-test.