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Dissecting metabolic changes in muscle stem cells during activation
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Dissecting metabolic changes in muscle stem cells during activation
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Content
Copyright 2020 Sanjana Ahsan
DISSECTING METABOLIC CHANGES IN MUSCLE STEM CELLS DURING ACTIVATION
by
Sanjana Ahsan
A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
DEVELOPMENT, STEM CELLS, AND REGENERATIVE MEDICINE
December 2020
ii
DEDICATION
To my parents, for unconditionally supporting me to get to where I am today.
iii
ACKNOWLEDGEMENTS
When I began my research training in Fall 2015, I expected my doctoral experience to
come with challenges and surprises. However, like most graduate students, I would not have
imagined that I would be completing my work in the midst of a national public health crisis. This
pandemic has brought on unforeseen challenges for everyone who has been part of my training
for the past few years, and I thank them for their continued support during this difficult time.
I thank my mentor, Joe Rodgers, for the guidance and support he provided throughout the
course of my training. He instilled in his trainees the courage to be ambitious, creative, and
independent, while also practicing patience and perseverance. Not only did I receive support to
further my scientific expertise, but I also had the freedom to pursue my interests outside of the
laboratory. This enabled me to develop a wide range of skills crucial for my career development.
I was lucky enough to train in a truly collaborative lab. I thank my lab mates: Andrew
Chareunsouk, Max Ederer, Manmeet Raval, and Rajiv Tiwari ––you were all instrumental to my
success as PhD student. We supported each other and we had fun along the way.
I’d like to thank Gage Crump, Denis Evseenko, and Neil Segil for giving their time to provide
additional guidance on my thesis project. Without your encouragement, I would not have had
the confidence to explore ideas outside the boundaries of my project.
Thank you PIBBS and the Broad CIRM Center, for providing me the opportunity to train at USC,
and also to fulfill my goal of conducting stem cell research.
And to my family and friends – thank you for giving me the strength to keep going during
challenging times.
iv
TABLE OF CONTENTS
DEDICATION ii
ACKNOWLEDGEMENTS iii
TABLE OF CONTENTS iv
LIST OF FIGURES v
ABSTRACT vi
CHAPTER 1: INTRODUCTION 1
CHAPTER 1.1 SKELETAL MUSCLE 1
CHAPTER 1.2 SKELETAL MUSCLE STEM CELLS 3
CHAPTER 1.3 SKELETAL MUSCLE REGENERATION 7
CHAPTER 1.4 MUSCLE STEM CELL ACTIVATION 9
CHAPTER 1.5 MUSCLE STEM CELL METABOLISM 14
CHAPTER 2: METABOLISM OF GLUCOSE AND GLUTAMINE IS CRITICAL FOR 15
CHAPTER 2: ACTIVATION OF SKELETAL MSUCLE STEM CELLS
CHAPTER 2.1 INTRODUCTION 15
CHAPTER 2.2 RESULTS 16
CHAPTER 2.3 DISCUSSION 27
CHAPTER 2.4 METHODS 31
CHAPTER 3: CONCLUSIONS 35
CHAPTER 3.1 MITOCHONDRIAL AND GLYCOLYTIC METABOLISM 35
C CHAPTER 3.1 IN ADULT STEM CELLS
CHAPTER 3.2 SUBSTRATE METABOLISM IN ADULT STEM CELLS 39
CHAPTER 3.3 FURTHER INVESTIGATION 42
REFERENCES 44
v
LIST OF FIGURES
Figure 1 Cellular activity increases in MuSCs during activation 17
Figure 2 MuSCs require mitochondrial metabolism to activate 20
Figure 3 Glucose metabolism plays an important role in MuSC activation 22
Figure 4 Glutamine metabolism is important but not necessary for MuSC 24
activation
Figure 5 Fatty acid metabolism is not necessary for MuSC activation 26
Figure 6 Glucose and glutamine contribute to majority of ATP production in 38
activating MuSCs
vi
ABSTRACT
When muscle is injured, skeletal muscle stem cells (MuSCs) are induced to activate,
wherein they exit quiescence and enter the cell cycle. Although previous works have shown that
MuSCs undergo significant metabolic changes during activation, the substrates that MuSCs
consume to support activation remain poorly understood. Here, we show that MuSCs generate
the majority of energy through mitochondrial respiration, and that oxidative phosphorylation is
necessary for MuSC activation. Furthermore, we have found that while glucose, glutamine, and
fatty acid metabolism significantly, and roughly equally, contribute to ATP production in MuSCs
during activation, they do not have equal functional role in the dynamics of MuSC activation.
Pharmacologic suppression of glycolysis, using 2-deoxy-D-glucose, or glutaminolysis, using
BPTES, significantly impairs MuSC cell cycle entry. However, etomoxir-mediated inhibition of
mitochondrial fatty acid transport has no effect on MuSC cell cycle progression. Our findings
suggest that apart from their roles in fueling ATP production by the mitochondria, glucose and
glutamine may generate metabolic intermediates needed for MuSC activation.
1
CHAPTER 1: INTRODUCTION
1.1 SKELETAL MUSCLE
Skeletal muscle plays a significant role in maintaining overall health by contributing to
mobility and metabolic homeostasis
1
. The contractile function of skeletal muscle enables
movement and in turn influences posture and social activity, and is important for sustaining an
independent lifestyle. Systemic metabolism relies on the function of muscle as a storage unit for
essential nutrients like amino acids and glucose, especially under starvation. In addition,
skeletal muscle produces heat to support temperature balance. Understanding the underlying
mechanisms that maintain skeletal muscle integrity and function, therefore, is important for
maintaining quality of life.
Skeletal muscle is comprised of multinucleated muscle fibers (myofiber), each
surrounded by basal lamina, a layer of extracellular matrix
1
. The muscle fibers are bundled into
groups encased in connective tissue known as the perimysium, and the myofiber bundles,
which compose a whole muscle, are enclosed by the epimysium. Muscle is a highly protein-
dense tissue ––the average muscle fiber is ~80% protein in dry weight
2
, compared to 60% for
the average mammalian cell
3
. Each muscle fiber is constructed from repeating units of
sarcomere, which is the basic functional unit of muscle
1
. The two major myofilament proteins
that make up the sarcomeres are actin and myosin. During force generation, contraction of the
sarcomere depends on an intricate coordination in the interaction between the head of the
myosin filament and the actin filament. The force generated by skeletal muscle is not only
determined by the size of the muscle fibers, but also the fiber type
4
. There are two major types
of muscle fibers: slow-twitch Type I fibers and fast-twitch Type II fibers. Slow-twitch muscle
fibers contract more slowly and generate less force than fast-twitch fibers; however, slow-twitch
fibers are able to contract for longer and therefore are more fatigue-resistant. Expression of
distinct myofilament isoforms in each fiber type underlies the difference in contractile capability.
2
The basal lamina that encapsulates each muscle fiber is made of type IV collagen,
laminin, entactin, fibronectin, perlecan, glycoproteins, and proteoglycans
5
. Majority of the
components of the basal lamina are produced and secreted by interstitial fibroblasts
6
. The two
predominantly abundant components of the basal lamina are type IV collagen and laminin,
which self-assemble and create interconnected networks joined by entactin
5
. The interaction
between the basal lamina and the cytoskeleton of the myofiber cell membrane (known as the
sarcolemma) is stabilized by the transmembrane proteins dystroglycan and integrin
7,8
. Several
forms of muscular dystrophy are associated with loss of basal lamina components and the
receptors which bind to the basal lamina. Congenital muscular dystrophy, for example, is
characterized by deficiencies in laminin-α2, integrin-α7, and dystroglycan
9
. Skeletal muscles
develop normally in these diseases but atrophy, suggesting that the interaction between the
basal lamina and sarcolemma is essential for maintaining muscle integrity
5
.
In addition to muscle fibers and connective tissue, muscle tissue is supported by motor
neurons. Motor neurons transmit electric impulses to myofibers through the neuromuscular
junction (NMJ) synapse
10
. The muscle fiber membrane comprises the postsynaptic component
of the NMJ. The membrane folds in, amplifying the surface area available to accommodate
voltage-gated sodium channels and neural cell adhesion molecules, which localize to the
bottom of the fold, as well as nicotinic acetylcholine receptors (nAChRs), which concentrate at
the fold’s crest. Acetylcholine released from the motor neuron terminal activates nAChRs, which
propagates action potentials across the muscle fiber. Electric impulses from motor neurons not
only enable muscle contraction, but also maintain muscle strength
11
.
Like most tissues, the health of skeletal muscle relies on a rich vascular supply.
Myofibers are surrounded by capillary networks at the terminus of arteries that extend and
branch into muscle tissue
12
. As type I muscle fibers contain more mitochondria and therefore
are more metabolically oxidative, the microvascular density is 3 times greater than that of type II
fibers
12
. The flow of blood through different layers of muscle tissue significantly changes as the
3
diameter of the vessels change from arteries to the capillary networks. The ability of the vessels
to constrict and dilate in response to changes in blood flow is regulated by the sympathetic
nervous system innervating the muscle.
Skeletal muscles posses an undoubtedly complex composition that has been scrutinized
for centuries
13
. The function of skeletal muscle not only depends on the integration of its
numerous components during homeostasis, but also on the tissue’s ability to recover following
injury. Muscle tissue is endowed with a robust capability to heal. Although many cell types
coordinate to repair damaged muscle, one cell population is indispensable for muscle
regeneration: the skeletal muscle stem cells
14
. Since Alexander Mauro observed them nearly 60
years ago, muscle stem cells have deepened our understanding of the cellular and molecular
biology of skeletal muscle
15
. The discoveries made thus far demonstrate the significant
contribution of muscle stem cells to muscle tissue health.
1.2 SKELETAL MUSCLE STEM CELLS
The ability of muscle tissue to regenerate highly depends on the function of its tissue
resident muscle stem cells (MuSCs)
14
. MuSCs in uninjured muscle lie between the basal lamina
and sarcolemma of their host muscle fiber in a dormant quiescent state
15
. Quiescent MuSCs are
primarily identified by the expression of the paired box transcription factor, Pax7, as well as the
absence of the basic helix-loop-helix myogenic transcription factors, MyoD and myogenin
(MyoG)
16,17,18
. Maintenance of quiescence is crucial for preservation of the MuSC pool, and
consequently, long-term regeneration of muscle tissue
19
. The molecular regulation of
quiescence in MuSCs, therefore, has been extensively studied.
Studies using genetic mouse knockout models have shown that Notch signaling is
required to maintain MuSC quiescence. MuSC-specific deletion of RBP-J (a transcriptional
regulator of canonical Notch signaling), driven by CreER recombinase under the control of the
Pax7 promoter, results in spontaneous entry into the cell cycle in vivo, as indicated by
4
incorporation of 5-ethynyl-2′-deoxyuridine, or EdU; consequently, this contributes to apoptosis-
independent reduction of the quiescent MuSC pool
19, 20
. In addition to Notch signaling, miRNAs
play an important role in MuSC quiescence. MuSC-specific conditional deletion of Dicer, the
endonuclease that processes long double-stranded RNAs into miRNAs, contributes to
spontaneous exit from quiescence and entry into the cell cycle
21
. As biogenesis of miRNAs
depends on the expression of their host genes, it is expected that quiescence-regulating genes
in MuSCs control the synthesis of quiescence-specific miRNAs
22
. Indeed, the Ctr (calcitonin
receptor) gene, which has been shown to regulate MuSC quiescence, hosts miRNA-489, a
quiescence-regulating miRNA
23,21
. Inhibition of miRNA-489 induces MuSCs to spontaneously
activate and incorporate EdU
21
.
In a recent study, Notch signaling was shown to regulate quiescence-specific miRNAs in
MuSCs. RNA sequencing of freshly isolated MuSCs showed that the quiescence-regulating
gene, Odz4, hosts miRNA-708
24
. ChIP (chromatin immunoprecipitation) sequencing revealed
binding sites for the Notch intracellular domain (NICD) and RBP-J on proximal enhancers of
Odz4, suggesting that Notch controls Odz4 expression in MuSCs. As expected, activation of
Notch signaling using the Notch receptor ligand Dll1 increases Odz4 enhancer activity.
Additionally, MuSC-specific ablation of Rbpj results in significant decreases in miRNA-708, and
blocking miRNA-708 induces MuSCs to exit quiescence and activate
24
. These findings show
that Notch signaling regulates MuSC quiescence through downstream control of Ozd4/miRNA-
708 expression. Increased cell cycle entry in MuSCs has also been observed following MuSC-
specific deletion of the retinoblastoma (Rb) protein and the phosphatase and tensin homologue
(Pten), suggesting that Rb and Pten play a role in maintenance of MuSC quiescence
25, 26
.
In contrast to Notch, Rb, and Pten, which are activated in quiescent MuSCs, protein
translation is suppressed. Missense mutation of serine 51 in the translation initiation factor
eIF2α – which is phosphorylated to inhibit translation— to alanine promotes increased
translation, entry into the cell cycle, as well as expression of myogenic genes in MuSCs
27
.
5
Additionally, MuSC-specific deletion of PKR-like endoplasmic reticulum kinase, which
phosphorylates eIF2α, induces cell cycle entry
27
. These findings support that MuSCs must
maintain low levels of translation to remain quiescent. Among the genes targeted for
translational repression in quiescent MuSCs are myogenic factors. Interestingly, it has been
observed that quiescent MuSCs express Myod transcripts in vivo, but not the MyoD protein
28
.
Although the levels of MyoD protein increase in MuSCs during activation, Myod transcription
does not
28
. Analysis of potential translational repressors of MyoD has identified Staufen1, an
RNA-binding protein, as the candidate repressor. Immunoprecipitation experiments show
binding interaction between mature Myod transcripts and Staufen1
28
. In addition,
downregulation of Staufen1 in MuSCs results in increased MyoD protein levels as well as EdU
incorporation, suggesting that Staufen1 regulates MuSC quiescence through translational
repression of MyoD.
Regulation of heterochromatin structure also plays an important role in MuSC
quiescence. Heterochromatin forms as either facultative heterochromatin (fHC) or constitutive
heterochromatin (cHC)
29
. fHC primarily forms in regions that encode genes and can transition
between euchromatin and heterochromatin, whereas cHC comprises mostly non-coding,
repetitive regions. Characterization of heterochromatin in quiescent MuSCs has shown that both
cHC and fHC exist; however, compared to differentiated myotubes, MuSCs contain lower levels
of cHC
30
. Preservation of fHC is critical for MuSC quiescence, as deletion of Suv4-20h1, a
dimethyltransferase that promotes fHC formation, results in increased cell cycle entry. Thus,
chromatin condensation is important for MuSCs to remain quiescent.
MuSC quiescence is also extrinsically regulated by its microenvironment. Baghdadi and
colleagues showed that NICD and RBP-J occupy proximal enhancers of several collagen genes
in MuSCs, suggesting that Notch signaling controls the production of basal lamina collagens
31
.
Rbpj-deficient MuSCs exhibit reduced collagen gene expression. Additionally, the study found
that MuSCs bind to autonomously generated collagen V through the calcitonin receptor, and
6
that ablation of Col5a1 induces aberrant cell cycle entry
31
. Interaction with components of the
basal lamina, therefore, enables MuSCs to stay in a quiescent state. Muscle fibers also promote
MuSC quiescence. In recent work by Eliazer et al., it was shown that muscle fiber-derived Wnt4
regulates MuSC quiescence by binding to Frizzled receptors on MuSCs and inducing non-
canonical Wnt signaling through Rho-GTPase
32
. Downstream of Rho, cytoskeletal signaling
maintains MuSC cellular shape and tension, and prevents migration from the niche
32
. In vivo
downregulation of Rho in MuSCs results in expansion of the MuSC pool in uninjured muscle
and increased incorporation of BrdU. Similarly, in vitro inhibition of Rho in isolated MuSCs
accelerates entry into the cell cycle, as measured by EdU incorporation. Thus, the muscle stem
cell niche is essential for long-term preservation of the quiescent MuSC pool.
Cell-to-cell interaction also promotes MuSC quiescence. Endothelial cells (ECs), for
example, have been found to influence MuSC quiescence through the Notch signaling pathway.
Identification of surface proteins expressed by muscle-specific ECs shows significantly higher
levels of Dll4 (a Notch ligand) compared to ECs in non-muscle tissue
33
. Previous works have
shown that Dll4 expression can be induced by vascular endothelial growth factor A (VEGFA),
which is abundantly expressed by MuSCs in vivo
34, 33
. Consistent with this, overexpression of
Vegfa in MuSCs strongly correlates with increased expression of Dll4 in muscle ECs,
suggesting that MuSCs interact with capillaries in their niche
33
. In turn, EC-derived Dll4 supports
MuSC quiescence by activating Notch signaling, as MuSC-specific deletion of Vegfa results in
downregulation of Notch target genes. Expression of Notch target genes in MuSCs are
unchanged following in vitro inactivation of Vegfa; this implies that VEGFA-dependent Notch
signaling in MuSCs is mediated through a non-cell-autonomous mechanism. The interaction
between MuSCs and the vascular niche, therefore, regulates quiescence.
Both intrinsic and extrinsic mechanisms control MuSC quiescence and in turn contribute
to long-term preservation of the MuSC pool. Preservation of quiescent MuSCs is crucial for
proper maintenance of muscle health throughout life, as multiple works have shown that
7
impairments in the pathways that promote MuSC quiescence correlate with defects in muscle
repair. The contribution of MuSCs to muscle health is not only dependent on their ability to
remain dormant, but also on their ability to rapidly transform following an injury to the muscle
and generate new muscle fibers.
1.3 SKELETAL MUSCLE REGENERATION
When muscle is injured, the sarcolemma of damaged myofibers loses structural integrity
and becomes more permeable
15
. Consequently, this increases calcium influx into the affected
fibers and induces calcium-dependent proteolysis, which contributes to further fiber
degeneration
35
. Muscle injury also triggers inflammation and recruits macrophages to the
wounded area to clean up the debris generated from deteriorating fibers through
phagocytosis
36
. Interestingly, it has been shown that a scaffold of basal lamina remains
following this necrotic process and guides MuSCs in regenerating injured muscle fibers
37,38
.
MuSCs activate in response to signals originating from the damaged microenvironment
following muscle injury
15
. Upon activation, MuSCs exit quiescence, enter the cell cycle, and
proliferate. Proliferating MuSCs, or myogenic progenitor cells (MPCs), exhibit a tremendous
increase in cell movement, migrating between muscle fibers as well as muscle groups during
regeneration
39,40
. During the initial proliferation phase, MPCs continue to express Pax7, which
promotes rapid expansion by inducing genes that positively regulate cell proliferation
41
. MPCs
also begin expressing MyoD; however, Pax7 initially inhibits MyoD activity to sustain
proliferation
41-42
. The effect of Pax7-mediated MyoD inhibition on MPC expansion is consistent
with work showing that deletion of MyoD contributes to increased MPC proliferation
43,44
. MyoD
deletion, however, also results in impaired differentiation, indicating that it is required for MPCs
to progress through differentiation
43,44
. Based on the antagonistic dynamic between Pax7 and
MyoD, it has been proposed that MPC differentiation depends on the ratio of Pax7 to MyoD
8
expression
42
. In this model, MPCs gain the ability to differentiate as extracellular signals
downregulate the Pax7 to MyoD ratio.
Following several rounds of proliferation, MyoD-positive MPCs begin to terminally
differentiate. Commitment to terminal differentiation is facilitated by MyoD-mediated expression
of MyoG
45
. ChIP analyses of C2C12 myoblasts cultured in growth medium have shown that
prior to commitment, MyoD binds the Myog promoter in association with histone deacetylase 1
(HDAC1)
46
. The enrichment of methylated histone H3K9 as well as hypoacetylation in this
genomic region suggests that HDAC1 contributes to suppression of Myog transcription during
C2C12 proliferation. Induction of differentiation by switching to differentiation medium results in
reduced HDAC1 occupancy and increased acetyltransferase P/CAF at the MyoD-bound Myog
promoter; this corresponds with H3K9 hyperacetylation and increased myogenin expression
46
.
Late-stage myogenic differentiation continues to rely on MyoD activity. Expression of
MyoG target genes requires the recruitment of histone acetyltransferase by MyoD, as the
absence of functional MyoD results in reduced acetylation at promoters of MyoG targets, as well
as reduced expression
45
. Furthermore, genome-wide analysis of MyoD and MyoG-bound
promoters (in C2C12 myotubes) shows a significant overlap in target sites, indicating that MyoD
and MyoG cooperate to drive myogenesis
45
. In addition to MyoD and MyoG, the myocyte
enhancer factor 2 (MEF2) family of transcription factors is also needed for MPC differentiation
47
.
Simultaneous deletion of Mef2a, c, and d in MuSCs results in impaired differentiation and
muscle regeneration; deletion of the individual isoforms, however, has no effect, suggesting that
MEF2 transcription factors play redundant roles in muscle repair. Together, MyoD, MyoG, and
MEF2s control the expression of many genes essential for skeletal muscle structure and
function.
Regeneration of damaged muscle fibers not only requires MPCs to proliferate and
differentiate, but also fuse together. The rate of MPC migration slows down in order to increase
cell-to-cell adhesion and initiate fusion
48,49
. The formation of multinucleated muscle fibers begins
9
with differentiated MPCs fusing together to give rise to myotubes within the basal lamina
scaffold of the injured host muscle fiber
50
. Several cell surface proteins have been found to
regulate the fusion of individual MPCs ––these include β-1 integrin, VLA-4 integrin, and VLA-4’s
receptor, VCAM-1
51, 52
. The myotubes fuse to each other as well as with the damaged ends of
the parent fiber
53, 54
. This is followed by further fusion of MPCs with myotubes, known as nuclear
addition, to generate larger muscle fibers. Calcium-dependent pathways play a crucial role in
regulating the growth of myotubes during muscle regeneration. The transcription factor NFATC2
is a downstream target of the calcium-regulated phosphatase, calcineurin
55
. It has been found
that NFATC2 is necessary for nuclear addition, as muscle regeneration under loss of function of
NFATC2 results in reduced nuclear number in the repaired muscle fibers
56
. NFATC2 promotes
nuclear addition by inducing the expression of IL-4 in myotubes, which is then secreted to
recruit myoblasts via the IL-4 receptor α
57
.
Myogenesis is a complex process that requires the coordination of many pathways. The
cellular and molecular mechanism of muscle generation has been extensively studied to
improve muscle regeneration in contexts where it is impaired, such as advanced aging. Much of
our understanding of the mechanisms that regulate muscle regeneration centers on the stages
that follow the transition of MuSCs into proliferating MPCs. Recent works, however, have
expanded to dissect the mechanisms which control early stages of muscle repair.
1.4 MUSCLE STEM CELL ACTIVATION
In recent years, the ability to isolate primary MuSCs through fluorescence-activated cell
sorting (FACS) has enabled observation of MuSC cellular behavior during the initial activation
phase, wherein MuSCs complete the first cell division after exiting quiescence. FACS
purification has also enabled dissection of molecular pathways that regulate the early stages of
muscle regeneration before MuSCs begin to rapidly proliferate. In a study conducted by
Rodgers and colleagues, they demonstrated that MuSC activation is a critical step in muscle
10
repair, in that improvements in MuSC activation contribute to improvements in muscle
regeneration
58
. They found through time-lapse microscopy that MuSCs isolated from uninjured
muscles of animals given a priming injury (away from the muscle tissue of interest) complete the
first mitosis in a shorter period of time than MuSCs from animals not given a priming injury.
MuSCs, therefore, can be induced to activate faster. Muscle regeneration in animals given a
priming injury beforehand (away from the injury in the muscle tissue of interest) also progresses
at a faster rate than regeneration in non-primed animals; this is consistent with the observation
made 50 years ago that a distant injury preceding a second injury accelerates regeneration of
the subsequent injury
58,59
. These findings thus showed that there is a strong correlation between
the rate of MuSC activation and muscle regeneration
58
.
At the molecular level, the hepatocyte growth factor (HGF) signaling pathway, known to
systemically activate in response to tissue injury, has been shown to prime MuSCs to activate
58,
60
. MuSC-specific ablation of the HGF receptor, cMet, blocks the effects of priming injury on
MuSC activation speed
58
. In addition, MuSCs isolated from mice injected with the active form of
HGF activator activate faster than MuSCs from control mice
60
. Downstream of cMet, the
mammalian target of rapamycin complex 1 (mTORC1) regulates MuSC activation
58
. Compared
to MuSCs from control animals, a larger proportion of MuSCs from primed animals exhibit
phosphorylation of the ribosomal protein S6, an indicator of active mTORC1 signaling. In
contrast, phosphorylation of S6 is undetectable in cMet knockout MuSCs of primed animals,
suggesting that mTORC1 activation is dependent on HGF signaling. More importantly,
mTORC1 has been shown to play an indispensable role in the enhancement of MuSC
activation, as deletion of its positive regulator, Rptor, slows down MuSC activation speed.
Conversely, induction of mTORC1 activity in MuSCs through deletion of its negative regulator,
Tsc1, results in decreased time to completion of the first cell division; thus, mTORC1 is
sufficient to induce MuSC activation. These findings have significantly advanced our
understanding of the mechanisms that control the activation of quiescent tissue resident stem
11
cells. Further work is needed to dissect the pathways downstream of mTORC1 that regulate
activation.
In addition to HGF, high mobility group box 1 (HMGB1) is another secreted factor that
has been shown to prime MuSCs to activate
61
. Although commonly recognized as a nuclear
DNA-binding protein, HMGB1 can translocate out of the nucleus and be secreted by cytokine-
activated immune cells, as well as cells undergoing apoptosis
62
. Secreted HMGB1 forms a
heterocomplex with CXCL12, a chemokine known to bind the CXCR4 receptor, which is
expressed on MuSCs
63
. MuSCs isolated from mice pre-treated with HMGB1 exhibit faster entry
into the cell cycle
61
. Furthermore, muscle regeneration is accelerated in mice pre-treated with
HMGB1. These effects of HMGB1 are mTORC1-dependent, as they are blocked by the
mTORC1 inhibitor, rapamycin. Thus, multiple systemic molecules can prime MuSCs to activate.
The ability of MuSCs to activate and expand relies on Notch signaling. During activation,
the levels of NICD – an indicator of Notch signaling activity— increase in explants of myofiber-
attached MuSCs
64
. Insufficient induction of Notch signaling during MuSC activation results in
reduced proliferation of myogenic progenitors and impaired muscle regeneration
64,65
. In a recent
study, Liu and colleagues showed that p53 activity underlies the ability of Notch to promote
MuSC proliferation
66
. They showed that MuSCs are susceptible to mitotic catastrophe (cell
death during or following mitosis), a phenomenon that has been associated with expression of
the mutant form of p53
66,67
. Indeed, in vivo deletion of Trp53 in MuSCs correlates with increased
cell death during activation
66
. Conversely, pharmacologic inhibition of p53 degradation in
MuSCs promotes survival during early activation. These results suggest that p53 activity
mitigates mitotic catastrophe in MuSCs. In the same study, recombinant Dll1-treated Rbpj-null
MuSCs were transfected with a p53 luciferase reporter plasmid to test crosstalk between Notch
signaling and p53 activity
66
. Compared to control MuSCs, luciferase activity was blocked in
Rbpj-null MuSCs, suggesting that Notch target genes regulate p53 activity. Findings from these
studies demonstrate that Notch signaling plays an important role in MuSC activation.
12
Multiple signaling pathways regulate MuSC activation during the early phase of muscle
regeneration. These pathways coordinate and induce dramatic changes in the cellular function
of MuSCs, including transcription and metabolism. Recently, several works have focused on the
metabolic changes in MuSCs during activation.
1.5 MUSCLE STEM CELL METABOLISM
MuSC metabolism has been shown to be integrally linked to MuSC activation and
muscle regeneration. Measurement of ATP concentration in MuSCs cultured following FACS
shows that there is a significant increase in ATP levels during activation
58,68
. Additionally, fast-
activating MuSCs such as those isolated from animals given a priming injury, as well as TSC1
knockout MuSCs, display increased mitochondrial activity compared to control MuSCs
58
. The
importance of MuSC metabolism for muscle regeneration has been demonstrated through the
use of caloric restriction. Work by Cerletti et al. shows that caloric restriction enhances
mitochondrial oxygen consumption in MuSCs
69
. Mice subjected to caloric restriction also show
improvements in muscle repair following injury
69
. In addition to caloric restriction, autophagy
regulates the metabolic and activation function of MuSCs. siRNA-mediated inhibition of
autophagy in cultured MuSCs decreases ATP levels, which suggests that autophagic flux
contributes to bioenergetic homeostasis during activation
68
. As well, cell cycle entry is delayed
in FACS-purified MuSCs cultured under chemical and siRNA-mediated inhibition of autophagy.
Furthermore, MuSC-specific ablation of atg5 – a gene necessary for autophagic flux— results in
decreased EdU incorporation
68
. Therefore, autophagy is necessary for MuSC activation. The
NAD-dependent deacetylase, SIRT1, has been found to regulate autophagy in activating
MuSCs, as loss of SIRT1 activity leads to defects in autophagic flux
68
. Consistently, MuSC-
specific deletion of sirt1 results in delayed entry into the cell cycle.
In a study by Ryall and colleagues investigating the role of SIRT1 in MuSC activation, it
was shown that MuSC-specific loss of SIRT1 function leads to delays in muscle regeneration
70
.
13
However, contrary to work which shows that SIRT1 function is essential for MuSC activation
68
,
Ryall et al. report that SIRT1 activity declines after MuSCs exit quiescence and enter the cell
cycle
70
. The decline in SIRT1 activity during activation is proposed to result from the observed
decreases in NAD
+
levels
70
. In concordance with decreases in NAD
+
(which correlates with
decreased mitochondrial metabolism), RNA-sequencing, as well as measurement of oxygen
consumption and extracellular acidification (an indicator of glycolytic activity) in freshly-isolated
and cultured MuSCs suggest that MuSCs undergo a metabolic switch from mitochondrial
oxidative phosphorylation to glycolysis during activation
70
. Moreover, ChIP analyses show that
reduction in SIRT1 deacetylase activity promotes acetylation of H4K16, which in turn promotes
expression of myogenic genes
70
. This model of MuSC activation, therefore, contradicts previous
works correlating increased mitochondrial metabolism with activation.
Several other works also support the paradigm that MuSCs reprogram their metabolism
from oxidative phosphorylation to glycolysis during activation. Among these studies includes
work by Chen et al., which dissects the role of the transcription factor Ying Yang 1 (YY1), a
regulator of mitochondrial metabolism, in MuSC activation
71
. In their work, Chen and colleagues
find that deletion of YY1 in MuSCs results in upregulation of mitochondrial gene expression and
a reduced ability to enter the cell cycle
71
. This suggests that YY1-dependent repression of
mitochondrial genes is necessary for MuSC to be able to activate. However, functional
measurement of mitochondrial oxygen consumption shows that it is reduced in YY1-deficient
MuSCs
71
. Consistent with the oxygen consumption measurements, previous works have shown
that YY1 promotes mitochondrial metabolism in multiple cell types, including C2C12
myoblasts
72,73,74
. Although YY1 appears to be required for MuSC activation, further work may be
necessary to understand its effect on mitochondrial gene expression and function in activated
MuSCs.
Recent work by Yucel et al. also supports previous findings that activating MuSCs are
glycolytic
75
. Yucel et al. show that proliferating MuSCs express higher levels of glycolytic genes
14
and exhibit higher histone acetylation than quiescent MuSCs (including on H4K16)
75
. The
maintenance of acetylation depends on the availability of glucose-derived acetyl-CoA, as
measured by isotopic labeling using
13
C6-D-glucose
75
. Furthermore, increasing metabolism of
glucose-derived pyruvate (to generate acetyl-CoA) through chemical inhibition of pyruvate
dehydrogenase kinase (PDK), the negative regulator of pyruvate dehydrogenase (PDH) in
mitochondria, increases histone acetylation near transcription start sites
75
. Synthesis of glucose-
derived acetyl-CoA in the nucleus depends on nuclear transport of (glucose-derived) citrate
generated in the mitochondrial TCA cycle
76
. Therefore, based on their findings, Yucel et al.
propose that glucose metabolism is dispensable for mitochondrial respiration in proliferating
MuSCs and is regulated to generate acetyl CoA to maintain histone acetylation.
Currently, there are conflicting findings on the metabolic changes that MuSCs undergo
during activation. While the results from some studies show that mitochondrial metabolism
increases after MuSCs exit quiescence, others find that activating MuSCs are glycolytic. Further
investigation is therefore needed to better understand the metabolic profile of activating MuSCs.
15
CHAPTER 2: METABOLISM OF GLUCOSE AND GLUTAMINE IS CRITICAL FOR
ACTIVATION OF SKELETAL MUSCLE STEM CELLS
2.1 INTRODUCTION
Skeletal muscle has a robust capacity to regenerate after injury. The ability of muscle
tissue to regenerate highly depends on the function of its tissue resident muscle stem cells
(MuSCs), also known as satellite cells. Under homeostatic conditions, MuSCs exist in a
quiescent state characterized by small size, low transcription, and low metabolic
activity
58,68,70,75,77
. When muscle is injured, signals from the damaged tissue environment induce
MuSCs to activate, i.e. to exit the quiescent state, enter the cell cycle, and proliferate. MuSC
activation has been found to correlate with increases in cellular and metabolic activity
58,68
. In
addition, previous work has shown that activation is a critical step in muscle repair, in that
defects in activation contribute to impairments in regeneration
58,70,78
. Conversely, MuSCs with
improved activation function contribute to improvements in muscle regeneration
58
.
MuSC metabolism is integrally linked with MuSC activation and muscle regeneration.
Work by Cerletti et al. shows that caloric restriction enhances MuSC metabolic function, and
that mice subjected to caloric restriction exhibit improved muscle repair following injury
69
.
Additionally, deficiency in SIRT1 activity leads to defects in autophagic flux needed to meet the
bioenergetic demands of MuSC activation, and consequently delays MuSC activation and
muscle regeneration
68,70
. Furthermore, loss of function of pyruvate dehydrogenase kinase (PDK)
2 and PDK4 in MuSCs contributes to dysregulation of glucose metabolism during activation and
is associated with impaired muscle regeneration
75
. While these works have shown the important
role of metabolism in MuSC activation, there have been no systematic investigations of
metabolic substrate utilization by MuSCs. Here, we report that during activation, MuSCs display
a profound increase in ATP production that is essentially entirely produced by the mitochondria.
We find that glucose and glutamine are the primary substrates that account for the majority of
16
ATP production, with only a minor contribution by fatty acids. Moreover, we find that MuSC cell
cycle entry is specifically dependent upon glucose and glutamine oxidation. Collectively, these
data provide a new detailed analysis of substrate metabolism during MuSC activation.
2.2 RESULTS
Cellular activity significantly increases in MuSCs during activation
To gain a more detailed understanding of the metabolic function of MuSCs during
activation, we employed an ex vivo model of isolation-induced activation
58,68,70
. Immediately
after dissociation from the hind limb muscle of adult mice (3.5 - 7 months old) and FACS-
mediated purification, we found that essentially no freshly isolated (FI) MuSCs were positive for
phosphorylated Ser 807/811-RB (pRB), a marker of cells that have exited the G1 phase of the
cell cycle, or incorporated EdU nucleotide, suggesting that the vast majority of these cells were
in early G1 (Figures 1A and 1B). After culturing for 24 and 40 hours post-isolation (hpi), the
number of MuSCs positive for these markers progressively increased, demonstrating that
MuSCs enter the cell cycle following isolation. We used time-lapse microscopy to continuously
monitor cultures of MuSCs for the first 90 hours after isolation and found that MuSCs require a
median of 48 hours to complete cytokinesis after isolation (Figure 1C). Consistent with previous
work, these data show that MuSCs require about two days to enter and complete the first cell
cycle following isolation
58
. Readily apparent in our time-lapse microscopy experiments was that
MuSCs displayed tremendous changes in cell migration and size during activation. We found
that MuSCs migrated a cumulative distance of 419 µm in the first 38 hours after plating (Figure
1D). Interestingly, we noticed that the distance versus time graph (Figure 1D) of MuSC
migration was convex, suggesting that the speed of MuSC migration was also increasing. We
analyzed the average speed of MuSC migration by taking the average distance between each
2-hour time window, and found that the speed at which MuSCs move also increases during
17
18
Figure 1. Cellular activity increases in MuSCs during activation. (A & B) Proportion of MuSCs
positive for pRb (A) and EdU (B) when they are freshly isolated (FI), 24, and 40 hours post-
isolation (n = 3 - 4). (C) Cumulative proportion of dividing MuSCs that have completed the first
division following isolation (n = 5). (D) Cumulative distance traveled by MuSCs from 18 to 38 hpi
(n = 30 cells). (E) MuSC migration speed between 19 to 37 hpi (n = 30 cells). Significance was
calculated between 19 and 37 hpi. (F) Average MuSC area between 18 to 42 hpi (n = 40 cells).
Significance was calculated between 18 and 42 hpi. (G) Average cell volume of MuSCs at 0 (n
= 4), 24 and 40 (n = 3) hours after isolation (left) and representative images (right). (H) Protein
translation in MuSCs detected through incorporation of HPG into newly synthesized proteins at
0 (n = 87 cells), 20 (n = 101 cells), and 40 (n = 103 cells) hpi. (I) Immunofluorescence images of
MuSCs stained for DAPI and HPG.
activation, from 5.1 µm/hour at 19 hpi to 36.9 µm/hour at 37 hpi (Figure 1E). Similarly, MuSC
area increases during activation from 199.9 µm
2
at 18 hpi to 489 µm
2
at 50 hpi (Figure 1F).
To determine if the changes in cell area reflected changes in cell size, we approximated
cell volume from diameter measurements of trypsinized (spherical) MuSCs at various time
points during activation. We found that in the first 40 hpi, cell volume increases by nearly five-
fold, from 164 µm
3
to 774 µm
3
(Figure 1G). To determine if these changes in cell size reflected
cell growth or anabolism, we measured protein synthesis rates. To do this, we pulsed MuSCs
with HPG, an amino acid analog that incorporates into nascent protein synthesis and can be
detected using Click-It chemistry, to measure protein translation rates. We found that after a 2-
hour pulse with HPG, 40 hpi MuSCs incorporate about four-fold more HPG than freshly isolated
MuSCs, suggesting that protein translation rates increase during activation (Figures 1H and 1I).
Collectively, these data show that MuSCs display tremendous changes in cell size and activity
as they enter and progress through the cell cycle following isolation.
MuSC activation requires ATP production from the mitochondria
These dramatic increases in cellular activity during activation suggested that cellular
metabolism would also need to significantly change to fuel these processes
79
. To investigate
this, we performed a series of live cell metabolic flux analyses on MuSCs using a Seahorse
bioanalyzer. We found that MuSCs display a dramatic overall increase in oxygen consumption
19
rate (OCR) in the first 48 hours after isolation (Figure 2A). After subtracting the contribution of
non-mitochondrial OCR, we found that mitochondrial dependent OCR increases by roughly 16-
fold over 48 hours after isolation (Figure 2B). Additionally, we measured the extracellular
acidification rate (ECAR), or the rate at which metabolic activity in MuSCs acidifies the media.
Similar to OCR, we found that MuSC ECAR increases by ten-fold during the first 48 hours after
isolation (Figure 2C). These results show that there is a dramatic increase in MuSC metabolic
activity during activation.
Previous work has shown that OCR and ECAR measurements can be used to calculate
ATP production rate from glycolysis and the mitochondria
80
. Our analysis showed that ATP
production by the mitochondria displays very dramatic and significant increase in the 48 hours
after isolation (Figure 2D). ATP production rate from glycolysis also displayed a similar trend of
increase, but was not statistically significant (Figure 2E). Most interestingly, when we directly
compared ATP production rates from the mitochondria and glycolysis, our measurements
showed that the vast majority, ~80%, of cellular ATP production comes from the mitochondria
(Figure 2F). The relative contributions of glycolytic and mitochondrial ATP production did not
change over the course of our measures, despite the strong increase in magnitude. These
results show that metabolic contribution from mitochondria is significantly larger than that of
glycolysis in activating MuSCs.
The dramatic increase in mitochondrial ATP production rate suggested to us that
mitochondrial metabolism plays a significant role in MuSC activation. To determine the
requirement of mitochondrial ATP production during activation, we cultured cells in oligomycin,
20
21
Figure 2. MuSCs require mitochondrial metabolism to activate. (A) Changes in basal OCR in
MuSCs after inhibition of ATP synthase using oligomycin, followed by inhibition of Complex I
and III of the electron transport chain with rotenone and antimycin A (R + A) (n = 5). (B) Total
OCR derived from mitochondria at 0 (n = 4), 24 and 48 hpi (n = 5). (C) ECAR at 0, 24, and 48
hpi (n = 5). (D) Rate of mitochondrial ATP production in MuSCs at 0, 24, and 48 hpi (n = 5). (E)
Rate of glycolytic ATP production in MuSCs at 0, 24, and 48 hpi (n = 5). (F) Relative proportion
of MuSC total cellular ATP production derived from mitochondrial respiration and glycolysis at 0,
24, and 48 hpi (n = 5). (G) Experimental schematic of the EdU incorporation assay in MuSCs
treated with or without oligomycin. (H) Immunofluorescence images of MuSCs stained for DAPI,
EdU, and Live/Dead cell viability stain. (I) Proportion of control and oligomycin-treated MuSCs
that have incorporated EdU at 40 hpi (n = 3). (J) Cell death in MuSCs cultured with or without
oligomycin for 40 hpi (n = 3).
an inhibitor of the mitochondrial ATP synthase, for 40 hours post-isolation (Figure 2G).
Treatment of cells with oligomycin dramatically decreases OCR due to feedback caused by the
inability of cells to dissipate the mitochondrial proton gradient via ATP synthase (Figure 2A).
Interestingly, we found no obvious changes in cell death in MuSCs cultured in oligomycin for 40
hours (Figures 2H and 2J). However, we observed very strong changes in EdU incorporation:
essentially zero MuSCs treated with oligomycin incorporated EdU in the first 40 hpi (Figures 2H
and 2I). Our findings show that inhibition of mitochondrial ATP synthesis nearly completely
blocks the ability of MuSCs to enter the S-phase of the cell cycle, but does not significantly
affect the rates of cell death. These results suggest that mitochondrial ATP production is
necessary for activation.
Glucose oxidation is critical for MuSC activation
Given that mitochondrial ATP production is necessary for MuSC activation, we next
investigated which metabolic substrates were consumed in activating MuSCs. The three major
substrates that cells oxidize to generate ATP and biosynthetic intermediates are glucose,
glutamine, and fatty acids
81
. To investigate the contribution of glucose to MuSC activation, we
treated cells with 2-deoxy-D-glucose (2-DG), an inhibitor of glycolysis, and examined the
change in metabolic flux
82
. We found that MuSCs treated with 2-DG displayed significantly
reduced levels of total OCR, OCR-linked to ATP production (OCRATP), and ECAR
22
Figure 3. Glucose metabolism plays an important role in MuSC activation. (A & B) Changes in
total (A) and ATP-linked (B) oxygen consumption rate induced by injection of 2-DG at 24 and 48
hpi (n = 5). (C) Total ECAR before and after 2-DG injection at 24 and 48 hpi (n = 5). (D)
Mitochondrial ATP production derived from glucose and non-glucose substrates at 24 and 48
hpi (n = 5). (E) Proportion of MuSC mitochondrial ATP production derived from glucose and
non-glucose substrates at 24 and 48 hpi (n = 5). (F) Immunofluorescence images of control and
2-DG treated MuSCs stained for DAPI and EdU at 40 hpi. (G & H) Proportion of control and 2-
DG treated MuSCs that are positive for EdU (G; n = 4) and Caspase 3 (H; n = 4 for control, n =
3 for 2-DG) at 40 hpi.
(Figures 3A - C). Using the measurements from control and 2-DG treated MuSCs, we calculated
mitochondrial ATP synthesized from oxidation of glucose versus non-glucose substrates and
23
found that both increased roughly three-fold from 24 to 48 hpi (Figure 3D). When we normalized
to total mitochondrial ATP production (glucose + non-glucose), we found that glucose
contributes to 29.6% of ATP generated by mitochondria at 24 hpi and 30.2% at 48 hpi (Figure
3E).
Our finding that glucose accounts for nearly one-third of mitochondrial ATP production
suggested that glucose plays an important role in MuSC activation. We therefore tested the
ability of MuSCs to activate under suppression of glycolytic metabolism. To do this, we cultured
the cells in 2-DG for 40 hours following isolation and measured EdU incorporation. We found
that compared to MuSCs cultured under control conditions, a smaller proportion of MuSCs
activate under 2-DG treatment (Figures 3F and 3G). Detection of caspase 3-positive MuSCs
showed that inhibition of glycolysis does not significantly change the proportion of cells
undergoing apoptosis (Figure 3H). Thus, in contrast to the near complete block of activation by
oligomycin, the data on 2-DG indicate that glucose metabolism has an important role, but is not
required for MuSC activation.
Glutamine is a major metabolic substrate during MuSC activation
Glutamine is another major metabolic substrate that is oxidized by the mitochondria via
the TCA cycle. To examine the contribution of glutamine metabolism in MuSC activation, we
treated cells with Bis-2-(5-phenylacetamido-1, 3, 4-thiadiazol-2-yl)ethyl sulfide (BPTES), a small
molecule which inhibits glutaminase, which converts glutamine to glutamate
83
. Glutamate is a
precursor for the TCA cycle intermediate α-ketoglutarate; therefore, we expected BPTES
treatment to suppress mitochondrial oxygen consumption in activating MuSCs
84
. Interestingly,
we found that culturing cells with BPTES for 24 hours did not have a significant effect on total
OCR or OCRATP (Figures 4A - C). However, BPTES had a very pronounced effect at reducing
OCR, and consequently mitochondrial ATP production rate, at 48 hpi (Figure 4A - C, 4E).
BPTES did not have a significant effect on ECAR at either time point (Figure 4D). By comparing
24
25
Figure 4. Glutamine metabolism is important but not necessary for MuSC activation. (A)
Changes in OCR induced by oligomycin injection in MuSCs cultured with or without BPTES for
24 and 48 hpi (n = 3). (B - D) Total (B) and ATP-linked (C) OCR and ECAR (D) in MuSCs
cultured with or without BPTES for 24 and 48 hpi (n = 3). (E) Mitochondrial ATP production in
control and BPTES-treated MuSCs (n = 3). (F) Proportion of MuSC mitochondrial ATP
production derived from glutamine and non-glutamine substrates at 24 and 48 hpi (n = 3). (G)
Immunofluorescence images of control and BPTES-treated MuSCs stained for DAPI and EdU at
40 hpi. (H & I) Proportion of control and BPTES-treated MuSCs that are positive for EdU (G; n =
4) and Caspase 3 (H; n = 5 for control, n = 4 for BPTES) at 40 hpi. Data are presented as mean
± standard deviation.
the measurements from control and BPTES treated cells, we calculated the proportion of
mitochondrial ATP from glutamine and non-glutamine substrates and found that glutamine
accounts for 8% of mitochondrial ATP production at 24 hpi, and 38% at 48 hpi (Figure 4F).
Although exposure to BPTES for 24 hours did not have a significant effect on MuSC
mitochondrial ATP production, the magnitude of effect observed under 48-hour BPTES
treatment was similar to that observed under inhibition of glycolysis. Based on these results, we
predicted that inhibition of glutamine metabolism would impair MuSC activation similar to the
effect of 2-DG. To determine the role of glutaminolysis in activation, we cultured MuSCs for 40
hours with or without BPTES treatment and measured EdU incorporation. As expected, fewer
MuSCs had incorporated EdU at 40 hpi in cultures treated with BPTES than control (Figures 4G
and 4H). BPTES treatment induced a modest increase in apoptosis in MuSCs (Figure 4I).
These results suggest that suppression of glutaminolysis slows down MuSC activation, and that
similar to glycolysis, glutaminolysis is important but not necessary for activation.
Fatty acid metabolism is dispensable for MuSC activation
Finally, we investigated the role of fatty acid β-oxidation in MuSC activation. To do this,
we treated cells with etomoxir, an inhibitor of carnitine palmitoyltransferase I, to block the
transport of fatty acids into the mitochondria
85
. Similar to BPTES, treatment with etomoxir for 24
hours did not have a statistically significant effect on OCR or ECAR (Figures 5A - D).
26
27
Figure 5. Fatty acid metabolism is not necessary for MuSC activation. (A) Changes in OCR
induced by oligomycin injection in MuSCs treated with or without etomoxir for 24 (n = 3) and 48
(n = 5) hpi. (B - D) Total (B) and ATP-linked (C) OCR and ECAR (D) in MuSCs cultured with or
without etomoxir for 24 (n = 3) and 48 (n = 5) hpi. (E) Mitochondrial ATP production in control
and etomoxir-treated MuSCs at 24 (n = 3) and 48 (n = 5) hpi. (F) Proportion of MuSC
mitochondrial ATP production derived from fatty acid and non-fatty acid substrates at 24 (n = 3)
and 48 (n = 5) hpi. (G) Immunofluorescence images of control and etomoxir-treated MuSCs
stained for DAPI and EdU at 40 hpi. (H & I) Proportion of control and etomoxir-treated MuSCs
that are positive for EdU (H) and Caspase 3 (I) at 40 hpi (n = 3).
However, after 48 hours of etomoxir treatment, total OCR, OCRATP, and mitochondrial ATP
production rates were significantly reduced compared to control (Figures 5A-C, 5E). We
determined the relative contribution of fatty acids to mitochondrial ATP production and found
that there was a trend of increase from 26.2% at 24 hpi to 38.8% at 48 hpi (Figure 5F). These
results suggest that fatty acids may be an important metabolic substrate during MuSC
activation.
Based on the reduced ability of MuSCs to enter the S-phase under inhibition of glucose
and glutamine metabolism, we predicted that etomoxir, by blocking fatty acid entry into the
mitochondria, would also slow down MuSC activation. Surprisingly, however, MuSCs treated
with etomoxir for 40 hours had the same level of EdU incorporation as control (Figures 5G and
5H). We did not observe changes in apoptosis under etomoxir exposure (Figure 5I).
Collectively, these data suggest that while MuSCs utilize fatty acids as a fuel source during
activation, fatty acids are not critical for the entry of MuSCs into S-phase.
DISCUSSION
Metabolism has been shown to play a crucial role in the function of many tissue resident
stem cells
86
. Specifically in the context of MuSCs, recent work has shown that MuSCs display
an increase in glycolytic metabolism during activation and this is important for muscle
regeneration
70,75
. We similarly observed that rates of glycolysis increase during activation
(Figures 2C and E). Interestingly, we found that the ATP production from glycolysis actually
28
decreases, relative to ATP production from the mitochondria as (Figure 2F). Our data show that
glycolysis is clearly an important pathway
Figure 6. Glucose and glutamine contribute to majority of ATP production in activating MuSCs.
Changes in MuSC activity, mitochondrial ATP production, and substrate metabolism during
activation.
29
during MuSC activation, inhibition of glycolysis by 2-DG significantly blunts entry of MuSCs into
S-phase of the cell cycle (Figure 3G). However, our data suggest that a function of glycolysis
during activation is to provide substrates for further oxidation in the mitochondria. Our findings
are consistent with recent work showing that MuSC-specific deletion of the transcription factor
Ying Yang 1 (YY1), which promotes mitochondrial metabolism
72,73,74
contributes to reduction of
mitochondrial respiration and impairs the ability of MuSCs to enter the cell cycle
71
.
We also analyzed the metabolism of major circulating nutrients in MuSCs during
activation. Our analysis of MuSC substrate metabolism showed that glucose, glutamine, and
fatty acids can contribute to mitochondrial ATP synthesis at similar levels in activating MuSCs
(Figure 6). All three metabolic substrates, however, were not equally important for MuSCs to
enter the S-phase of the cell cycle. While suppression of glucose and glutamine metabolism
delayed entry into S-phase, inhibiting fatty acid metabolism had no effect. One possible
explanation for the lack of effect of etomoxir on S-phase entry is that MuSCs are able to
compensate by increasing other metabolic pathways. We analyzed ECAR and found that it did
not significantly change in response to etomoxir (Figure 5D), suggesting that glycolysis rates did
not increase to compensate for inhibition of fatty acid oxidation. As mitochondrial ATP
production rates only displayed a modest, not statistically significant decrease in response to
etomoxir at 24 hpi, this may suggest that MuSCs compensate by increasing glutamine
oxidation. Further work is needed to test the effect of etomoxir treatment on MuSC glutamine
metabolism. An alternative explanation is that fatty acid oxidation may not have an important
role in fueling the early phases of MuSC cell cycle entry. Indeed, the strongest effect of etomoxir
is after 48 hours of activation, a stage when the majority of MuSCs have already entered and
completed the first cell cycle after isolation
58
. This suggests that fatty acid oxidation may have
an important role in the daughter cells from MuSC division. These data show that while fatty
acid metabolism contributes to energy production in activating MuSCs, it is not necessary for
the cells to activate. Inhibition of individual metabolic pathways for each substrate only partially
30
reduced mitochondrial metabolism; it is likely that combined suppression of all three pathways
will block mitochondrial metabolism and cell cycle entry, similar to oligomycin.
Although glucose, glutamine, and fatty acids are able to support ATP synthesis in
MuSCs, only inhibition of glucose and glutamine metabolism delays MuSC activation. This
indicates that in addition to ATP, synthesis of other metabolites, specifically derived from
glucose and glutamine, are critical for MuSC activation. Indeed, isotopic labeling in MuSCs
using 13C6-D-glucose has shown that a significant level of histone acetylation marks derive
from glucose-derived acetyl-coA
75
. The mitochondrial pyruvate dehydrogenase (PDH) converts
glucose-derived pyruvate to acetyl-coA. Pharmacologic suppression of PDH kinase, the
negative regulator of PDH, correlates with increases in glucose-derived histone acetylation in
MuSCs. This suggests that glucose is an important source of epigenetic modification during
MuSC activation. Glutamine also supports cellular functions outside of oxidative
phosphorylation. Among the non-respiratory roles of glutamine include serving as a precursor
for synthesis of non-essential amino acids (NEAAs), as well as nucleotides
87
. Radiolabeling
experiments in C2C12 myoblasts show that glutamine predominantly contributes to protein
biomass through generation of NEAAs
88
. Glutamine metabolism has also been analyzed in
other tissue resident stem cells, such as hematopoietic progenitor cells (HPCs). Analysis of
glutamine-derived metabolites using high performance liquid chromatography mass
spectrometry shows significant contribution of glutamine to nucleotide biosynthesis in CD34+
HPCs, and that this contribution is abolished following suppression of glutaminolysis
89
. Inhibition
of glutaminolysis as well as downregulation of the ASCT2 glutamine transporter in HPCs
contributes to defects in erythroid differentiation and bias towards myeloid lineage commitment,
suggesting that glutamine metabolism is necessary for balanced hematopoiesis. Addition of
nucleosides to glutaminolysis-deficient HPCs restores erythropoiesis; it is possible that
nucleoside supplementation to BPTES-treated MuSCs will rescue activation. Given these
findings, MuSCs may metabolize glucose and glutamine not only to maintain bioenergetic
31
homeostasis, but also to support epigenetic regulation of MuSC activation and biosynthesis of
macromolecules. Further work is needed to dissect the diverse fates of glucose and glutamine
in MuSCs during activation.
METHODS
Mice
All experiments were performed on samples derived from 3.5 to 7 months old male C57Bl6j
mice obtained from Charles River. Mice were housed in the fully AAALAC-accredited animal
care facility in the basement of the Eli and Edythe Broad Center for Regenerative Medicine and
Stem Cell Research at USC. Animal use protocols were reviewed and approved by the USC
Institutional Animal Care and Use Committee (IACUC).
MuSC isolation and purification
MuSCs were isolated as previously described
21
. Immediately after CO 2 euthanasia, hind limb
skeletal muscles were extracted and minced. Minced skeletal muscles were enzymatically
digested into a single-cell solution by collagenase and dispase. The digested skeletal muscle
sample was stained with CD31-FITC (BioLegend) CD45-FITC (BioLegend), Sca-1-PerCP
(BioLegend), VCAM-PeCy7 (BD Biosciences), and Integrin α7-PE (Thermo Fischer Scientific).
MuSCs were purified on the BD FACSAria IIu as a population of CD31-, CD45-, Sca1-, VCAM+,
and Integrin-α7+ cells, and sorted into Ham’s F-10 plating medium (Cellgro) supplemented with
5 ng/mL basic fibroblast growth factor (Invitrogen), 10% FBS (Invitrogen), and 1X
penicillin/streptomycin (GIBCO).
Cell culture
Purified MuSCs in plating medium were seeded in 8-well chamber slides coated with ECM
(Sigma E1270) over poly-D-lysine (Millipore) at 10,000 cells per well and allowed to adhere for
32
approximately 1 hour post-isolation. After allowing MuSCs to adhere, plating medium was
switched to growth medium, or Ham’s F10 media supplemented with 10% FBS and 10% horse
serum. MuSCs were grown at 37 °C and 5% CO 2 and growth medium was replenished every 4
to 12 hours. Freshly isolated MuSCs were imaged at 1 hpi for cell diameter analysis. MuSCs
cultured for 24 and 40 hours were trypsinized then imaged for cell diameter measurement. To
quantify protein synthesis, cells were cultured in normal growth medium then switched to
cysteine- and methionine-free medium DMEM supplemented with 2 mM L-glutamine (VWR) and
100 µM HPG (Invitrogen), and cultured for 2 hours before fixation with 4% PFA. To measure
entry into S-phase of the cell cycle, MuSCs were cultured in 5 µM EdU (Invitrogen) for 40 hours.
HPG and EdU incorporation were detected using Alexa Fluor 488 azide. MuSCs were cultured
in growth medium treated with 200 nM oligomycin (Cayman Chemical), 1 mM 2-deoxy-D-
glucose (Sigma-Aldrich), 15 µM BPTES (Tocris), or 50 µM etomoxir (EMD Millipore) to block
mitochondrial ATP synthesis, glycolysis, glutaminolysis, and fatty acid oxidation, respectively.
The LIVE/DEAD Fixable Dead Cell Stain Kit (Invitrogen) was used according to the
manufacturer’s protocol to detect cell viability under oligomycin treatment.
Immunostaining
Immunocytochemistry staining was performed on MuSCs after fixation and permeabilization.
MuSCs were fixed with 4% PFA for 10 minutes at room temperature then permeabilized using
0.3% Triton-X PBS for 15 minutes. Cells were then blocked with 10% donkey serum in 0.3%
Triton-X PBS for 15 minutes. Cleaved caspase-3 was probed in MuSCs by incubating the cells
overnight in 1:100 Cleaved Caspase-3 (Asp175) (5A1E) rabbit mAb (Cell Signaling
Technology). MuSCs were stained for 3 hours with 1:500 donkey anti-rabbit Alexa 594
secondary antibody (Invitrogen) to detect cleaved caspase-3. pRB was stained for in MuSCs
following EdU staining. MuSCs were first permeabilized then blocked in donkey serum for 20
minutes. After blocking, the cells were stained using 1:100 phospho-Rb (Ser807/811) (D20B12)
33
XP rabbit mAb (Cell Signaling Technology). Secondary staining was performed using 1:500
donkey anti-rabbit Alexa 488 antibody (Invitrogen). All primary and secondary antibodies were
diluted in 10% donkey serum.
Time-lapse microscopy
MuSCs were plated on 8-well chamber slides coated with ECM and cultured for 90 hours at 37
°C and 5% CO2. MuSCs were then transferred to a temperature- and CO2-controlled chamber
mounted on the microscope stage of the Zeiss Axio Observer Z1. Bright-field images were
acquired every 10 minutes and cells were visualized using ZEN Black software. Time taken to
complete the first division following initiation of image acquisition was only analyzed for MuSCs
that remained in acquisition field.
Extracellular flux assay
One day prior to the assay, Seahorse XF sensor cartridges were hydrated in XF Calibrant at 37
°C and 0% CO2 overnight. On the day of the assay, MuSCs were purified into plating medium
and seeded immediately following isolation in 8-well Seahorse mini-plates coated with ECM.
Plating medium was changed to growth medium treated with or without 15 µM BPTES and 50
µM etomoxir for MuSCs cultured for 24 and 48 hours at 37 °C and 5% CO 2. For freshly isolated
(FI) MuSCs, plating medium was changed to Seahorse assay medium (Buffered XF DMEM
base medium containing 5 mM HEPES, 2 mM L-glutamine, and 10 mM glucose) after MuSCs
were seeded. MuSCs were incubated in Seahorse assay medium for 1 hour at 37 °C and 0%
CO2, then stained with propidium iodide and imaged to detect viable cells immediately before
loading in the XF analyzer. MuSCs were administered 1 µM oligomycin and 0.25 µM rotenone
and antimycin A during the assay. For measuring glycolytic metabolism, MuSCs were injected
with 50 mM 2-DG before oligomycin and rotenone and antimycin A. ATP production rates were
calculated from OCR based on the calculations described in Quantifying Cellular ATP
34
Production Rate Using Agilent Seahorse XF Technology. Glucose-linked mitochondrial ATP
production rate was calculated as the difference between baseline ATP production rate and
ATP production post-2-DG injection. Glutamine or fatty acid-linked ATP production rate was the
difference between baseline ATP production rate in control and BPTES- or etomoxir-treated
MuSCs.
Data analysis
All data are presented as mean ± standard deviation. For experiments that involved treatment of
MuSCs with pharmacological inhibitors, MuSC population isolated from one mouse was split
into control (untreated) and experimental (treated) groups post-isolation so that measurements
can be taken in parallel. Time to the first division data are presented as the cumulative
proportion of cells that have divided starting at 30 hpi and at every 10-hour interval afterwards.
Cell migration was analyzed starting at 18 hpi and the distance traveled was measured at 2-
hour intervals for the next 20 hours. Migration speed was calculated by dividing the distance
traveled during each 2-hour interval by 2 hours; the speed is plotted as the speed at the
midpoint of each time interval. Changes in MuSC area were analyzed starting at 18 hpi and
measurements were taken every 6 hours, up to 42 hpi.
Excluding Figures 1E - F and 3A - C, all statistical comparisons were performed using a two-
tailed, unpaired Student’s t-test; variance was tested using the F-test. A two-tailed, paired
Student’s t-test was used for Figures 1E-F and 3A-C.
35
CHAPTER 3: CONCLUSION
3.1 MITOCHONDRIAL AND GLYCOLYTIC METABOLISM IN ADULT STEM CELLS
Metabolism has been shown to play a crucial role in the function of various tissue
resident stem cells. In our study, we found that suppression of glycolysis impairs the ability of
skeletal muscle stem cells to activate (Figures 3F and 3G), while inhibition of mitochondrial
respiration completely blocks activation (Figure 2H and 2I). Our findings thus showed that both
glycolytic and mitochondrial metabolism are important for MuSC activation; however,
mitochondrial metabolism is absolutely essential. Metabolism is also important for MuSC self-
renewal. The AMP-activated protein kinase (AMPK) coordinates multiple metabolic pathways to
maintain energetic homeostasis
90
. Work by Theret et al. shows that MuSC-specific deletion of
AMPK results in an increased number of quiescent MuSCs following muscle regeneration, an
indicator of enhanced self-renewal; however, the fiber size of the newly regenerated muscles is
reduced, suggesting an impaired balance between commitment and self-renewal
91
. Metabolic
analysis of AMPK-null MuSCs shows increased lactate dehydrogenase activity – which
indicates increased glycolytic activity— as well as reduced mitochondrial respiration
91
. Inducing
MuSCs to utilize glycolytic or oxidative metabolism by culturing them in glucose or galactose
media, respectively, reveals that oxidative conditions reduce MuSC self-renewal, as measured
by expression of Pax7 and absence of Ki67 and MyoD. Further, AMPK-null MuSCs were
resistant to the effects of the galactose media and retained their self-renewal capability. These
findings suggest that AMPK regulates MuSC fate specification by modulating the balance of
glycolytic and mitochondrial metabolism during myogenesis.
As muscle tissue is highly metabolically active, it requires functional and morphological
development of mitochondria during myogenesis. The Dlk-Dio3 gene cluster encodes miRNAs
which suppress mitochondrial biogenesis
92,93
. RNA-sequencing analyses show that compared to
freshly isolated MuSCs, expression of the Dlk-Dio3 cluster is significantly reduced in myogenic
36
progenitors and myotubes
94
. miR-1/133a, the negative regulator of the Dlk-Dio3 cluster,
however, is highly expressed in myotubes. Upregulation of Dlk-Dio3 expression in MuSCs
through deletion of miR-1/133a contributes to defects in mitochondrial maturation and function
94
.
Whereas muscle fibers of control mice display evenly distributed mitochondria, fibers in miR-
1/133a knockout mice exhibit aberrant distribution, with some areas devoid of mitochondria and
other areas harboring mitochondrial aggregates. Similar mitochondrial network aberrations are
found in myotubes generated from miR-1/133a-null MuSCs. Additionally, the respiratory function
of mitochondria in muscles of miR-1/133a knockout mice is reduced compared to that of control.
Interestingly, MuSCs isolated from miR-1/133a knockout mice do not show defects in
proliferation or commitment to differentiate (as indicated by myogenin expression). These
findings show that although the Dlk-Dio3 cluster regulates mitochondrial homeostasis during
myogenesis, it is dispensable for MuSC proliferation and differentiation.
Studies on other stem cells also show that metabolism is crucial for their function.
Analysis of metabolism in hematopoietic stem cells (HSCs) reveals that quiescent HSCs
generate majority of ATP through anaerobic glycolysis, and that glycolysis is necessary for
maintenance of quiescence
95
. Indeed, maintenance of long-term HSCs relies on the
suppression of mitochondrial metabolism, mediated by the Dlk1-Gtl2 locus
93
. Although
quiescent HSCs are glycolytic, mitochondria remain essential for their survival. HSC-specific
deletion of LKB1, an upstream regulator of AMPK, contributes to defects in mitochondrial
biogenesis and consequently disruption of metabolic homeostasis in quiescent HSCs
96
. Further,
mice with LKB1-deficient HSCs display increased apoptosis and a reduced HSC pool compared
to control mice. These findings suggest that LKB1-mediated regulation of mitochondrial function
is necessary for maintenance of HSCs. Similar to MuSCs, metabolism also influences HSC
proliferation. Reduction of mitochondrial metabolism through suppression of the mitochondrial
pyruvate dehydrogenase has been shown to maintain HSCs in culture, but blocks proliferation
95
.
Surprisingly, and in contrast to our findings in respiration-deficient (i.e. oligomycin-treated)
37
MuSCs, defects in the mitochondrial respiratory chain induced through deletion of the Rieske
iron-sulfur protein, an essential component of the mitochondrial respiratory complex III, has no
effect on HSC proliferation
97
. The divergent effects from suppressing components of
mitochondrial activity suggest that only specific aspects of mitochondrial function are critical for
HSC proliferation.
Mitochondrial function also plays an important role in HSC differentiation. Defects in
polymerase gamma (POLG), the mitochondrial DNA polymerase, contribute to perturbations in
hematopoietic differentiation, indicated by an overabundance of erythroid committed
progenitors, as well as deficiency in B cell committed progenitors in POLG mutator mice
98
.
PTPMT1, a mitochondrial protein tyrosine phosphatase, also regulates HSC differentiation.
Measurement of real-time oxygen consumption shows that HSC-specific ablation of PTPMT1
reduces mitochondrial respiration, suggesting that PTPMT1 regulates oxidative
phosphorylation
99
. Furthermore, in vitro colony-forming unit assays show defects in myeloid and
lymphoid colony formation in PTPMT1-null HSCs
99
. In fact, cell cycle analysis reveals that a
higher proportion of PTPMT1-null HSCs are in the G1 phase compared to control HSCs
99
.
These findings support that mitochondrial function regulates HSC fate commitment.
Like MuSCs, hair follicle stem cells (HFSCs) also proliferate following activation.
Interestingly, HFSCs strongly rely on generation of lactate, a product of glycolysis. Flores and
colleagues have shown that deletion of lactate dehydrogenase in HFSCs results in a reduced
ability of HFSCs to proliferate – as measured via Ki67— as well as failure to initiate anagen
100
.
Conversely, induction of lactate production in HFSCs, through deletion of the mitochondrial
pyruvate carrier (MPC), improves the initiation of new hair cycles
100
. Although these findings
show that glycolysis is critical for HFSC activation and that mitochondrial pyruvate metabolism
may be dispensable, the precise role of mitochondria in HFSC function remains unclear.
Glycolysis also promotes the proliferation of intestinal stem cells (ISCs) and
mesenchymal stem cells (MSCs). Similar to HFSCs, induction of lactate biosynthesis through
38
ablation of MPC drives intestinal stem cells to proliferate
101
. Conversely, Rodriguez-Colman and
colleagues have shown that inhibition of glycolysis reduces ISC proliferation and enhances
differentiation into mature intestinal crypts
102
. In line with the effect of glycolytic inhibition on
ISCs, suppression of mitochondrial respiration impairs ISC differentiation
102
. Interestingly, work
by Rodriguez-Colman et al. also shows that metabolism during ISC differentiation relies on
lactate production by the supporting Paneth cells adjacent to ISCs
102
. They find that Paneth cell-
derived lactate is transported and converted to pyruvate in ISCs, which is then metabolized by
mitochondria. Inhibition of glycolysis or lactate transport in Paneth cells contributes to defects in
ISC differentiation. The reliance of HFSCs and ISCs on lactate is in agreement with the recent
finding that lactate serves as a major metabolic substrate in multiple tissues
103
.
Hypoxia, which increases glycolytic flux, promotes in vitro expansion of MSCs
104,105
.
Although MSCs proliferate under normoxia as well using oxidative metabolism, expansion under
these conditions results in increased senescence
104
. MSCs are multipotent and capable of
adipogenic, chondrogenic, and osteogenic differentiation, and the metabolic requirements of
MSC differentiation is lineage dependent. While chondrogenic differentiation relies on glycolytic
metabolism, adipogenic and osteogenic differentiation require oxidative metabolism
104,106
.
Adipogenic differentiation specifically requires mitochondria-derived reactive oxygen species
(ROS), as treatment of MSCs with mitochondria-targeted antioxidants impairs adipogenesis
106
.
ROS signaling also influences neurogenesis; however, the dynamics of mitochondrial
morphology, rather than metabolic activity, underlie the function of ROS in neural stem cell
(NSC) homeostasis. Mitochondria in uncommitted NSCs are elongated, while those in neural
progenitor cells (NPCs) are fragmented
107
. Although measurement of metabolic flux indicates
that NSCs are glycolytic, Western blot analysis shows that they express enzymes of the
mitochondrial respiratory chain, and can utilize mitochondrial metabolism under glucose-
deprivation, suggesting that mitochondrial metabolism is suppressed in NSCs. NPCs, on the
other hand, downregulate components of the glycolytic pathway and utilize oxidative
39
metabolism, and as such, exhibit higher levels of mitochondrial ROS (mtROS) compared to
NSCs
107
. Perturbation of mitochondrial fusion and elongation in NSCs through deletion of the
mitochondrial fusion proteins MFN1/2 and OPA1 induces oxidative metabolism and generation
of mtROS, impairs NSC self-renewal, and consequently, reduces the NSC pool. Conversely,
deletion of the mitochondrial fission protein DRP1, which induces mitochondrial fragmentation,
results in decreased mtROS and enhanced NSC self-renewal. These findings show that
although there is a metabolic switch from glycolytic to oxidative metabolism during
neurogenesis, this process depends on the structure of the mitochondria.
3.2 SUBSTRATE METABOLISM IN ADULT STEM CELLS
Regulation of stem cell metabolism is not only important for maintenance of energy
homeostasis, but also for production of intermediate metabolites critical for stem cell function.
The consumption of metabolic substrates in stem cells is thus temporally and differentially
regulated. We found in our work that suppression of glucose and glutamine metabolism impairs
MuSC activation (Figures 3F-G and 4G-H), while inhibition of fatty acid oxidation (FAO) has no
effect (Figures 5G-H). These findings suggest that metabolic intermediates generated from
glucose and glutamine consumption are important for the early stages of myogenesis, while
those generated from FAO are more likely needed at a later point. Consistent with our finding
that activating MuSCs utilize similar levels of glucose and glutamine, work by others show that
glucose and glutamine equally contribute to cell mass turnover in C2C12 myoblasts
88
. In
addition to glucose and glutamine, myogenic progenitor cells require serine and glycine to
proliferate, as absence of serine and glycine blocks the expansion of MPCs
108
. Addition of
serine and glycine restores MPC proliferation in a dose-dependent manner.
Like myogenesis, hematopoiesis also demands catabolism of glucose and glutamine.
Glutamine serves as a precursor for the TCA cycle intermediate, α-ketoglutarate, and also
supports nucleotide biosynthesis
109
. Analysis of glutamine-derived metabolites using high
40
performance liquid chromatography mass spectrometry shows significant contribution of
glutamine to the TCA cycle (anaplerosis) and nucleotide biosynthesis in CD34
+
hematopoietic
progenitors cells (HPCs), and that this contribution is abolished following suppression of
glutaminolysis. Additionally, inhibition of glutaminolysis as well as downregulation of the ASCT2
glutamine transporter in HPCs contributes to defects in erythroid differentiation and bias towards
myeloid lineage commitment, suggesting that glutamine metabolism is necessary for balanced
hematopoiesis
89
. Although addition of nucleosides to glutaminolysis-deficient HPCs restores
erythropoiesis, supplementation with dimethyl α-ketoglutarate is not sufficient to rescue defects
in hematopoietic differentiation; this suggests that the metabolic contribution of glutamine to
HPC function is linked to nucleotide synthesis and not anaplerosis. In contrast to suppression of
glutaminolysis, 2-DG-mediated inhibition of glycolysis in CD34
+
HPCs increases erythroid
specification. Furthermore, inhibition of the pentose phosphate pathway, which can generate
nucleotide precursors using glycolytic intermediates, blocks erythroid specification. These
findings suggest that glutamine- and glucose-derived nucleotide synthesis is indispensable for
homeostatic hematopoiesis. The essential amino acid, valine, is also required for HSC
homeostasis, as proliferation of HSCs is significantly reduced in valine-depleted media
110
.
HSC differentiation relies on fatty acid metabolism. Genetic deletion of PPAR-δ, a
regulator of mitochondrial FAO, contributes to defects in HSC repopulation post-
transplantation
111
. In addition, inhibition of mitochondrial fatty acid transport using etomoxir
reduces HSC colony formation in vitro
111
. Similar to our findings in MuSCs, etomoxir treatment
decreases ATP production in HSCs
111
. Pharmacological activation of PPAR-δ using GW-50156
results in enhanced repopulation capacity and increased HSC proliferation; however, these
effects of GW-50156 are attenuated by etomoxir. This suggests that PPAR-δ-driven regulation
of HSC function is dependent on downstream mitochondrial fatty acid metabolism. Non-
mitochondrial fatty acid metabolism is also critical for HSC function. Deletion of 12/15
lipoxygenase, which catalyzes production of bioactive lipid intermediates, contributes to
41
impairments in HSC repopulation capability, increased propensity to enter the cell cycle, and
reduced HSC self-renewal
112
. The findings on the metabolic regulation of HSC function show
that multiple metabolic pathways are tightly controlled to support HSC homeostasis. Much of the
work thus far focuses on HSC differentiation; further work is needed to understand the
metabolic requirements of HSC activation and proliferation.
Similar to HSCs, intestinal and neural stem cell function also require fatty acid
metabolism. Work by Mihaylova et al. shows that fasting enhances self-renewal of ISCs,
measured via organoid formation by crypts isolated from non-fasted and fasted mice
113
.
Inhibition of FAO using etomoxir, however, attenuates the effects of fasting on organoid
formation, suggesting that fasting-induced enhancement of ISC function relies on fatty acid
metabolism
113
. Similar observations have been made in MuSCs from calorie-restricted (CR)
mice, wherein CR-dependent improvements in MuSC proliferation are reversed by etomoxir
69
.
Furthermore, deletion of Cpt1a, the mitochondrial fatty acid transporter, contributes to a
reduction in the ISC pool as well as reduced organoid formation. In quiescent neural
stem/progenitor cells (NSPCs), radiolabeling experiments show that fatty acids are oxidized to
generate ATP, TCA cycle intermediates, and TCA cycle-derived amino acids at a higher level
than in proliferative NSPCs
114
. In line with this, etomoxir-mediated inhibition of FAO in quiescent
NSPCs induces significant cell death
114
. In proliferative NSPCs, FAO suppression impairs cell
cycle progression and proliferation. In vivo clonal analysis of Cpt1a-null NSPCs also reveals that
suppression of FAO contributes to defects in adult neurogenesis
114
. The effects of etomoxir on
NSPCs contrast our observations in MuSCs, where etomoxir had no effect on cell viability or cell
cycle progression. In addition to fatty acid consumption, NSPC function also requires de novo
lipid biogenesis. Short-term chemical inhibition of fatty acid synthase (Fasn), which catalyzes
fatty acid synthesis, decreases NSPC proliferation, while long-term Fasn suppression induces
cell death
115
. Furthermore, NSPC-specific deletion of Fasn results in impaired neurogenesis, as
42
indicated by a reduced frequency of differentiating NSPCs and fewer newly generated neurons.
Fatty acid metabolism is therefore necessary for intestinal and neural regeneration.
Substrate metabolism also regulates chondrogenic and osteogenic differentiation during
bone repair. As previously discussed, chondrogenic differentiation relies on glycolytic
metabolism
104
. Consistent with this, it was found in a recent study by van Gastel et al. that
limiting vascularization, and hence oxygenation, of the repair callus during bone regeneration
results in increased frequency of SOX9-positive early chondrogenic cells
116
. The group also
found that growth of periosteal cells – which include skeletal stem and progenitor cells— under
serum-deprived conditions or in lipid-reduced serum promotes increased chondrogenic
differentiation
116
. Re-addition of fatty acids to lipid-deprived serum rescues osteogenic
differentiation of periosteal progenitor cells. In addition, injection of fatty acids, as well as
GW9508, an agonist of free fatty acid receptors 1 and 4, into the repair callus reduces cartilage
formation without impairing bone regeneration. Furthermore, deletion of SOX9 in periosteal cells
contributes to enhanced fatty acid oxidation. These findings suggest that SOX9 suppresses fatty
acid metabolism to promote chondrogenic fate specification during fracture repair.
3.3 FURTHER INVESTIGATION
We found in our work that ATP production significantly decreases in response to 40-hour
inhibition of glucose, glutamine, or fatty acid metabolism; however, cell cycle progression is only
impaired under suppression of glycolysis and glutaminolysis. This suggests that glucose and
glutamine not only feed into oxidative phosphorylation, but also other pathways that support
MuSC activation. As Oburoglu et al. observed in hematopoietic progenitors, it is possible that
nucleotide synthesis in activating MuSCs also depends on glucose and glutamine catabolism
89
.
This can be tested by measuring activation of MuSCs cultured in BPTES or 2-DG-treated
growth medium supplemented with nucleosides. In addition, the role of fatty acids in MuSC
43
myogenic function remains unclear. Though treatment with etomoxir had no effect on MuSC
activation, it is likely that similar to hematopoiesis, neurogenesis, intestinal regeneration, and
osteogenic maturation, myogenic differentiation also requires fatty acid oxidation.
Muscle stem cell metabolism is a nascent but growing field within the larger field of
muscle biology. Although studies by several groups have improved our understanding of how
metabolism supports and regulates MuSC function, further work is needed to model the
metabolic landscape of muscle regeneration. Treatment of MuSCs with pharmacologic inhibitors
of metabolism ex vivo has indicated the substrates that MuSCs consume, however, steady-state
metabolic flux in quiescent MuSCs and myogenic progenitor cells remain elusive. In vivo
radiolabeling and differential metabolomics will help visualize the contribution of major nutrients
(e.g. glucose, glutamine, and fatty acids) and intermediary metabolites (e.g. lactate) to
metabolic pathways in MuSCs. Understanding the metabolic distribution of substrates can in
turn identify potential regulators of metabolism critical for long-term maintenance of quiescent
MuSCs and also for muscle repair. Dissecting the coordination among regulators of quiescence,
metabolism, and cell cycle progression will significantly advance our understanding of the
mechanism of muscle regeneration.
44
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Abstract (if available)
Abstract
When muscle is injured, skeletal muscle stem cells (MuSCs) are induced to activate, wherein they exit quiescence and enter the cell cycle. Although previous works have shown that MuSCs undergo significant metabolic changes during activation, the substrates that MuSCs consume to support activation remain poorly understood. Here, we show that MuSCs generate the majority of energy through mitochondrial respiration, and that oxidative phosphorylation is necessary for MuSC activation. Furthermore, we have found that while glucose, glutamine, and fatty acid metabolism significantly, and roughly equally, contribute to ATP production in MuSCs during activation, they do not have equal functional role in the dynamics of MuSC activation. Pharmacologic suppression of glycolysis, using 2-deoxy-D-glucose, or glutaminolysis, using BPTES, significantly impairs MuSC cell cycle entry. However, etomoxir-mediated inhibition of mitochondrial fatty acid transport has no effect on MuSC cell cycle progression. Our findings suggest that apart from their roles in fueling ATP production by the mitochondria, glucose and glutamine may generate metabolic intermediates needed for MuSC activation.
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Ahsan, Sanjana
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Dissecting metabolic changes in muscle stem cells during activation
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Keck School of Medicine
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Development, Stem Cells and Regenerative Medicine
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10/26/2020
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