Current Progress and Challenges for Skeletal Muscle Differentiation from Human Pluripotent Stem Cells Using Transgene-Free Approaches

Abstract

Neuromuscular diseases are caused by functional defects of skeletal muscles, directly via muscle pathology or indirectly via disruption of the nervous system. Extensive studies have been performed to improve the outcomes of therapies; however, effective treatment strategies have not been fully established for any major neuromuscular disease. Human pluripotent stem cells have a great capacity to differentiate into myogenic progenitors and skeletal myocytes for use in treating and modeling neuromuscular diseases. Recent advances have allowed the creation of patient-derived stem cells, which can be used as a unique platform for comprehensive study of disease mechanisms, in vitro drug screening, and potential new cell-based therapies. In the last decade, a number of methods have been developed to derive skeletal muscle cells from human pluripotent stem cells. By controlling the process of myogenesis using transcription factors and signaling molecules, human pluripotent stem cells can be directed to differentiate into cell types observed during muscle development. In this review, we highlight signaling pathways relevant to the formation of muscle tissue during embryonic development. We then summarize current methods to differentiate human pluripotent stem cells toward the myogenic lineage, specifically focusing on transgene-free approaches. Lastly, we discuss existing challenges for deriving skeletal myocytes and myogenic progenitors from human pluripotent stem cells.

Full text

Review Article
Current Progress and Challenges for Skeletal Muscle
Differentiation from Human Pluripotent Stem Cells Using
Transgene-Free Approaches
Nunnapas Jiwlawat,
1
Eileen Lynch,
1
Jeremy Jeffrey ,
1
Jonathan M. Van Dyke,
1
and Masatoshi Suzuki
1,2
1
Department of Comparative Biosciences, University of Wisconsin, Madison, WI, USA
2
The Stem Cell and Regenerative Medicine Center, University of Wisconsin, Madison, WI, USA
Correspondence should be addressed to Masatoshi Suzuki; masatoshi.suzuki@wisc.edu
Received 21 December 2017; Revised 11 February 2018; Accepted 18 February 2018; Published 11 April 2018
Academic Editor: Zhaohui Ye
Copyright © 2018 Nunnapas Jiwlawat et al. This is an open access article distributed under the Creative Commons Attribution
License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is
properly cited.
Neuromuscular diseases are caused by functional defects of skeletal muscles, directly via muscle pathology or indirectly via
disruption of the nervous system. Extensive studies have been performed to improve the outcomes of therapies; however,
effective treatment strategies have not been fully established for any major neuromuscular disease. Human pluripotent stem
cells have a great capacity to differentiate into myogenic progenitors and skeletal myocytes for use in treating and modeling
neuromuscular diseases. Recent advances have allowed the creation of patient-derived stem cells, which can be used as a
unique platform for comprehensive study of disease mechanisms, in vitro drug screening, and potential new cell-based
therapies. In the last decade, a number of methods have been developed to derive skeletal muscle cells from human pluripotent
stem cells. By controlling the process of myogenesis using transcription factors and signaling molecules, human pluripotent
stem cells can be directed to differentiate into cell types observed during muscle development. In this review, we highlight
signaling pathways relevant to the formation of muscle tissue during embryonic development. We then summarize current
methods to differentiate human pluripotent stem cells toward the myogenic lineage, specifically focusing on transgene-free
approaches. Lastly, we discuss existing challenges for deriving skeletal myocytes and myogenic progenitors from human
pluripotent stem cells.
1. Introduction
Recent advances in stem cell biology hold great promise for
use in treating and modeling neuromuscular diseases [1].
Neuromuscular diseases affecting the function or develop-
ment of skeletal muscle can arise directly via muscle pathology
or indirectly via disruption of the nervous system. Despite
devastating consequences, no effective treatment strategies
exist in many cases, including muscular dystrophy. Attractive
therapeutic strategies include the replacement of affected
muscle cells with healthy myocytes or progenitor cells,
thereby restoring skeletal muscle function.
Human pluripotent stem cells (PSCs), which include
embryonic stem cells (ESCs) and induced pluripotent stem
cells (iPSCs), represent a robust cell source for developing
cell-based therapies targeting degenerating muscles as well
as modeling neuromuscular disease conditions and for
drug screening in culture. Particularly, iPSC technology
allows creation of patient-derived stem cells, which can
simulate pathophysiological conditions in vitro [2]. These
in vitro models are expected to work as a unique platform
for drug screening and allow comprehensive study of dis-
ease mechanisms.
In the last decade, a number of culture methods for myo-
genic differentiation from human PSCs have been published
[3]. These include (1) transgene methods employing the direct
manipulation of gene expression and (2) transgene-free
methods employing pharmacologic inhibitors and agonists
Hindawi
Stem Cells International
Volume 2018, Article ID 6241681, 18 pages
https://doi.org/10.1155/2018/6241681

as well as isolated cytokines or other protein-based signals
[3]. In this review, we discuss relevant pathways and events
during skeletal muscle development which have been stud-
ied and manipulated in an effort to derive myogenic cell
types from human PSCs. We then overview recent prog-
ress of the methods for myogenic derivation from human
PSCs, specifically focusing on transgene-free approaches.
Finally, we discuss the limitations and potential of these
approaches for future treatment and modeling of neuro-
muscular diseases.
2. Skeletal Muscle Development and
Molecular Networks
2.1. Embryonic Myogenesis and Terminal Differentiation into
Myofibers. During early embryogenesis, the formation of
skeletal muscle begins when the paraxial mesoderm segments
form somites in response to signals from the notochord, neu-
ral tube, and surface ectoderm [4]. The developing somite
then forms the dermomyotome, myotome, and sclerotome.
The cells in the dermomyotome express the paired box
transcription factors Pax3 and Pax7 [4–7]. The dorsomedial
and ventrolateral portions of the dermomyotome give rise
to the epaxial (primaxial) and hypaxial (abaxial) myotomes,
respectively. Myf5-positive cells in the epaxial myotomes
differentiate and form the trunk and back muscles. In con-
trast, MyoD-positive progenitors delaminate and migrate
from the hypaxial myotome into the developing limb as
the source of limb muscles. Myf5 and MyoD are expressed
in committed muscle cells and are located in the myotome,
which is formed from the maturation of dermomyotome
lips [8–10].
The terminal differentiation of progenitors and myo-
blasts initiates when myogenic progenitors in the dermo-
myotome stop dividing and exit the undifferentiated stage
(Figure 1). Pax3- and/or Pax7-positive proliferating progeni-
tors withdraw from the cell cycle once the differentiation step
is initiated. These progenitors then become committed myo-
blasts expressing Myf5 and/or MyoD and form the nascent
myotubes expressing myogenin and myosin heavy chain
(MHC) (Figure 2(a)). Two waves of myotube formation
occur during skeletal muscle development, sequentially giv-
ing rise to primary and secondary myotubes [4, 11]. Primary
myotubes are generated from the fusion of early myoblasts
and are aligned between muscle tendons to form the basis
for embryonic muscle development. Late-stage myoblasts
proliferate alongside primary myotubes and fuse to form
secondary myotubes. As the secondary myotubes form,
motor axons begin to innervate the embryonic muscle [11].
Single-nucleated myoblasts then fuse with the nearby myo-
tubes to form multinucleated myotubes. Thick-myosin and
thin-actin filaments within the myotube begin organizing
and form sarcomeres, the functional units of muscle contrac-
tion. Sequential chains of sarcomeres, called myofibrils, align
in maturing myotubes. Mature myotubes contain well-
organized and aligned myofibrils which give rise to the char-
acteristic striated pattern of skeletal myocytes (Figures 2(b)
and 2(c)).
2.2. Signaling Molecules for Myogenesis. Myogenesis is deli-
cately regulated by signaling events that influence prolifera-
tion and differentiation of stem cells and progenitor cells
[4]. These events are driven by paracrine and/or autocrine
signaling molecules that pattern and generate specificcel-
lular lineages. A number of signaling molecules have been
characterized to play critical roles for specification and
differentiation from the somite to the myotomes [12, 13]. Sig-
naling molecules can also contribute to terminal differentia-
tion of myoblasts and myotube formation. These molecules
regulate the expression of myogenic genes and proteins and
influence the growth and fusion of MHC-positive myotubes.
This section will introduce several signaling molecules criti-
cal for myogenesis; however, this is not an exhaustive list.
Wnt signaling plays a significant role in the development
of myogenic progenitors in the somite and the formation
of committed myoblasts in later stages of myogenesis. A
diverse family of Wnt proteins is secreted from the neural
tube and ectoderm. Wnt1 [12] and Wnt3a [14, 15] are pro-
duced in the dorsal neural tube, while Wnt7a is expressed
in the dorsal ectoderm [12], and Wnt5a is localized in the
dorsal ectoderm and limb mesenchyme [14]. Wnt ligands
bind to Frizzled (Fzd) receptors and take action through a
canonical (β-catenin) pathway or noncanonical pathways
[16]. In mouse explant cultures, Wnt1 can enhance Myf5
expression and affects epaxial muscle formation. In contrast,
Wnt7a promotes MyoD expression and influences hypaxial
myogenesis [12, 17]. The initial expression of Pax3 and
Myf5 was decreased in mice lacking both Wnt1 and Wnt3a
[15]. A Wnt antagonist Frzb1 inhibits myogenesis in preso-
mitic mesoderm, but not in mature somites. When Frzb1
was injected in a pregnant mouse, the process of myogenesis
was disturbed by the reduction of Myf5 expression [18].
An inhibitor of Wnt/β-catenin signaling (IWR1-endo)
inhibits myotube formation in murine myotube culture
[19]. Additionally, an inhibition of glycogen synthase kinase
3β(GSK3β) can promote mesoderm differentiation via acti-
vating Wnt pathways [20–22].
Sonic hedgehog (Shh) is secreted from the notochord and
floor plate of the neural tube [23] and regulates myogenic
progenitor proliferation and differentiation [24]. In zebrafish,
the number of Pax3- and Pax7-positive cells was significantly
increased by a knockdown of the Shh gene [24]. Shh shows
positive effects on muscle development by directing progen-
itor cells to Myf5-/MyoD-positive committed myocytes in
the myotome by downregulating Pax3/Pax7 expression
[25]. A reduced level of Myf5 expression was observed in
Shh-null mice, resulting in a loss of distal limb structures
[26]. Shh also enhances myogenic differentiation by increas-
ing MyoD expression. An implantation experiment using
Affi-Gel agarose beads soaked with 100 μg/ml N-Shh in the
lumen of the neural tube showed that Shh activates both
MyoD and a sclerotomal marker, Pax1, in quail embryos
[27]. Shh also promotes sclerotome formation while inhibit-
ing dermatome formation [23].
Fibroblast growth factors (FGFs), including FGF2 (or
basic FGF, bFGF), are critical factors for controlling prolif-
eration and differentiation of myogenic progenitors and
myoblasts during myogenesis. FGF2 is known to inhibit
2 Stem Cells International

the differentiation of myogenic progenitors into myotubes
[28, 29], implying that FGF2 could be used to maintain the
progenitors at an immature stage. Interestingly, in murine
myoblast C2C12 cells, inhibition of the mitogen-activated
protein kinase (MAPK) pathway, which is downstream of
FGF, increased the expression of MyoD, myogenin, and
MHC and led to more myoblast fusion [29]. Both paracrine
and autocrine effects of FGFs are proposed, as myocytes have
been found to express both FGF ligands and FGF receptors.
FGF ligands can bind to four FGF receptors (FGFR1–4) with
varying levels of affinity. FGFR1–4 are transmembrane tyro-
sine kinase receptors capable of activating various down-
stream signaling cascades. FGFR1, 2, and 4 are expressed in
immortalized myoblast cell lines such as mouse Sol 8 cells.
Inhibitory effects of myocyte differentiation by FGF mole-
cules were only observed when FGFR1 and 2 were presented
in Sol 8 cells. Myogenic differentiation was stimulated when
FGFR1 signals were inhibited by overexpressing truncated
FGFR1 molecules [28]. Another study using chromatin
immunoprecipitation-on-chip analyses demonstrated that
FGFR4 is a direct downstream target of Pax3 in mouse
embryo [30]. Further studies are necessary to elucidate which
FGFRs are involved in modulating myogenesis. In addition,
application of FGF2 or forskolin to C2C12 mouse myoblasts
resulted in phosphorylation and activation of cyclic AMP
response binding (CREB) protein. A gain-of-function muta-
tion in CREB increased myoblast proliferation [31], indicat-
ing involvement of CREB signaling in myogenesis. Loss of
CREB activity significantly decreased Pax3, Myf5, and MyoD
expression in mouse embryos [17].
Both bone morphogenetic protein 4 (BMP4) and Notch
enhance progenitor proliferation but inhibit muscle differ-
entiation [25]. BMP4, secreted from the lateral plate meso-
derm, sustains Pax3 expression and delays Myf5 and MyoD
expression in chicken embryos [32]. An increased level of a
BMP4 inhibitor Noggin in the dorsomedial lip of the
dermomyotome of chick embryos inhibits BMP signaling
and increases medial, rather than lateral, somite patterning
[33]. Noggin-soaked bead implants promote muscle differ-
entiation in chick embryos [34]. While BMP4 works as a
secreted factor, an activation of Notch signaling requires
direct cell-cell contacts. The Notch receptor is a single-
pass transmembrane protein. Notch ligands bind to the
extracellular domain of the receptor and then lead to pro-
teolytic cleavage at the intracellular domain. After the intra-
cellular domain is released, it migrates toward nucleases
and modulates the expression of downstream genes [35].
A subset of migrating neural crest cells expresses a Notch
ligand, Delta1. When chick embryo dermomyotomal cells
transiently contact Delta1-expressing cells, expression of
Myf5 and MyoD is activated. However, a prolonged contact
with Delta1-expressing cells reverses the myogenic process
resulting in Pax7-positive progenitor cells [36]. Notch sig-
naling increases proliferation of myogenic progenitors but
inhibits muscle differentiation by blocking MyoD transcrip-
tional activity [37].
Transforming growth factor beta (TGF-β) and a TGF-β
superfamily protein, myostatin, are known to modulate myo-
genic differentiation. TGF-βinhibits myogenic differentia-
tion by suppressing the activity of myogenin [38]. However,
a potent and selective inhibitor for TGF-βtype I receptor
(SB431542) and retinoic acid have been shown to rescue
the negative effect of TGF-βon MHC
+
myotube formation
in C2C12 mouse myoblasts [39]. In mouse embryonic stem
cells, a combination of TGF-βinhibitor (SB431542), a Wnt
activator (BIO), and a Shh inhibitor (erismodegib) increased
the expression of Pax7, Myf5, MyoD, and myogenin and the
number of MHC
+
myotubes [40]. Myostatin (also known as
growth and differentiation factor-8, GDF-8) affects muscle
cell differentiation in a manner similar to that of TGF-β.
Dorsomorphin and LDN193189, which inhibit myostatin
activity, significantly enhance myotube formation when
Myf5 +
MyoD
MyoD+
Myogenin
+
(MyoG)
MHC+
IGF-I
Myostatin
TGF-
BMP
+
Myoblast Committed
myocyte
Nascent
myotube Myobril
Multinucleated
myotube
Myogenic
progenitor
Fusion
hypertrophy
Myober
Pax3+
Pax7+
Sarcomere
organized
myolaments
−
Figure 1: Skeletal muscle differentiation in vitro. The terminal differentiation starts when Pax3
+
and/or Pax7
+
progenitors begin to express
Myf5 or MyoD as committed myoblasts. These myoblasts gradually express myogenin (MyoG) and form single-nucleated nascent
myotubes with myosin heavy chain (MHC
+
). Insulin-like growth factor-I (IGF-I), TGF-β1 inhibitor, and myostatin inhibitors induce
myotube fusion to form multinucleated myotubes. Actin, myosin, and elastic myofilaments are arranged to form organized sarcomeres
within the myotubes. Organized sarcomere structures give rise to a striated pattern in the myotubes and represent the functional
contraction unit of muscles.
3Stem Cells International

applied to primary human myotubes and murine myotubes
[41]. Follistatin, another myostatin inhibitor, increased
fusion index and myogenic protein expression (including
MyoD, Myf5, and myogenin) in C2C12 cells [42]. Another
myostatin inhibitor, growth and differentiation factor-
associated serum factor protein 1 (GASP-1), also enhances
myogenin expression and fusion index in myotubes differen-
tiated from C2C12 cells [43].
Insulin-like growth factor-I (IGF-I) is produced and
secreted from myogenic cells and regulates muscle differen-
tiation and growth. Both IGF-I receptors and IGF binding
proteins are dramatically increased in mouse C2 myoblast
cells during muscle differentiation [44]. IGF-I triggers termi-
nal differentiation of myoblasts through the MAPK signaling
pathway and increases protein expression of myogenin in
murine C2C12 myotubes [29]. IGF-I, but not IGF-II,
Pax7 MyoG MHC
(A) (B) (C)
MyoD MyoG MHCPax3 MyoG MHC
25 m
(a)
MHC
Hoech st 10 m
Titin
Hoech st 20 m
(b)
10 m
2 m
1 m
Z
I
ZM
A
(c)
Figure 2: Derivation of skeletal myocytes and matured myotubes from human iPSCs using a transgene-free protocol.Human iPSCs can be
sufficiently differentiated into myogenic progenitors and myotubes in a defined culture without genetic modification using free-floating
spheres (EZ spheres) [59, 67]. (a) Human iPSC-derived myotubes were labeled with multiple myogenic proteins Pax3, Pax7, MyoD,
myogenin (MyoG), and myosin heavy chain (MHC), demonstrating colocalization of those proteins in the same field. Some MyoD
+
nuclei
were overlapped with MyoG
+
nuclei and fused on MHC
+
myotubes (double arrow: MyoD
+
/MyoG
+
). The other nuclei were not
overlapping with MHC but expressed either MyoD or MyoG (arrow: MyoD
+
/MyoG
−
; or arrow head: MyoD
−
/MyoG
+
) (C). Neither Pax3
+
nuclei (A) nor Pax7
+
nuclei (B) showed any localization with MyoG
+
nuclei, which mostly fused on MHC
+
myotubes. (b) Sarcomere
formation in iPSC-derived myotubes. Titin staining revealed that striated patterns were clearly visible in the myotubes at 12 weeks MHC
staining in the same cell preparations used for titin labeling. (c) Ultrastructures of iPSC-derived myotubes. After 12 weeks of terminal
differentiation, mature sarcomeres were observed to be assembled into myofibrils. Morphological hallmarks, including I-band of actin
filaments and A-band with distinct M line across myosin filaments, were clearly visible. Sarcomere Z lines appeared to be reasonably
aligned and gave rise to a striated pattern. This figure is reproduced from Jiwlawat et al. [67] (under the Creative Commons Attribution
license/public domain).
4 Stem Cells International

promotes myofiber fusion and hypertrophy in avian myo-
tubes. This hypertrophy was promoted by increased synthesis
and lower degradation of MHC proteins [45]. Interestingly,
the steroid testosterone can stimulate fusion and hypertro-
phy of primary human myotubes via the IGF-I signaling
pathway [46].
3. Derivation of Skeletal Muscle Cells from
Human PSCs
Cell signaling plays a critical role in all stages of myogenesis.
The timing of expression and the levels of signaling mole-
cules are tightly controlled in order for the different stages
of myogenesis to occur smoothly [12, 13]. Accumulated
knowledge of the signaling pathways guiding myogenesis
has aided the creation of a number of methods for deriving
myogenic progenitors and myocytes from human PSCs.
Current methods can be broadly categorized into two
approaches: (1) induction of myogenic differentiation by
overexpression of myogenic genes (transgene methods) and
(2) derivation of myogenic progenitors under defined culture
using growth factors and/or signaling molecules without
transgenes (transgene-free methods).
3.1. Transgene-Based Approaches to Enhance Myogenic
Differentiation. Selective induction of myogenic genes, such
as the overexpression of PAX3, PAX7, and MYOD1, has been
used in order to increase the efficiency of myogenic differen-
tiation [3]. As discussed above, these transcription factors
play critical roles in promoting proliferation and differentia-
tion of myogenic progenitors and myoblasts during embry-
onic myogenesis. Different systems of gene expression, such
as lentiviral and piggyback-based approaches, have been
applied to transduce PAX7 [47, 48] and MYOD1 [49–52]
genes into human PSCs. The transcription of myogenic genes
can also be controlled by inducible gene expression systems
such as tetracycline or tamoxifen [47–52]. These progenitors
can be sufficiently enriched by fluorescence-activated cell
sorting (FACS) if the transgene construct contains a fluoro-
phore reporter gene like green fluorescent protein (GFP)
and mCherry [47, 49].
One notable advantage of the transgene method is that
transgene-based approaches can secure high efficiency of
progenitor preparation (more than 90% in several methods).
Typically, transgene methods yield progenitors more rapidly
than transgene-free methods. However, as these approaches
require an introduction of exogenous genes to the cells, the
resulting cells may not fully reflect the normal processes of
progenitor proliferation, differentiation, and maturation.
Additionally, genetic modification remains a regulatory con-
cern if the progenitors are to be used for cell-based therapy
in patients. As such, myogenic progenitors prepared by
transgene-free methods may be more suitable for transplan-
tation in patients.
3.2. Transgene-Free Approaches: Myogenic Derivation under
Defined Culture Conditions. Recent attempts have been made
to derive myogenic progenitors from human iPSCs and ESCs
under defined culture conditions using specific molecules
secreted as paracrine factors that play important roles in
muscle development (Table 1). These molecules control pro-
liferation, migration, and differentiation from mesodermal
cells into somite and dermomyotome [25]. FGF2 has been
used at varying concentrations (5–100 ng/ml) to direct and
enhance myogenic differentiation [20, 53–61]. Although 10–
20 ng/ml FGF2 is commonly used to maintain proliferation
in cell lines or primary cells, during our recent study, we found
that a high concentration of FGF2 (100 ng/ml) significantly
increased the number of Pax7-positive myogenic progenitors
from human PSCs [59]. Other growth factors such as
insulin-like growth factor-I (IGF-I), epidermal growth factor
(EGF), hepatocyte growth factor (HGF), and platelet-derived
growth factor (PDGF) have also been known to promote myo-
genic progenitor expansion and differentiation in human
PSCs [57]. IGF-I can enhance myotube hyperplasia and fusion
[62, 63]. IGF-I has been used at a concentration of 2–50 ng/ml
to enhance terminal differentiation [55–57, 61, 64].
Small molecule inhibitors have also been used to direct
and enhance myogenic differentiation. GSK3βinhibitors,
such as CHIR99021 [55, 61] and BIO (6-bromoindirubin-
3′-oxime) [20], can promote mesoderm induction during
differentiation by activating Wnt pathways. CHIR99021 sig-
nificantly enhances the expression of mesoderm genes such
as T,TBX6, and MSGN1 in human PSCs [54, 55, 65], indicat-
ing that this selective GSK3βinhibitor can promote meso-
derm differentiation. While CHIR99021 has proven useful
for in vitro mesoderm differentiation, it should be noted that
it should only be used in culture for short periods and at a low
concentration due to its toxicity [54, 55]. In fact, a longer
exposure (more than 3 mM for 4 days) or a higher concentra-
tion (10 μM for 2 days) of CHIR99021 results in toxicity in
human PSC cultures [54, 55]. By contrast, a potent and
reversible GSK3βinhibitor (BIO) demonstrates the lowest
toxicity among other GSK3βinhibitors [20]. Further, an ade-
nylyl cyclase activator, forskolin, has been used in a triple
cocktail with FGF2 and a GSK3βinhibitor (BIO) to promote
muscle differentiation [20].
Inhibitors of BMP type I receptors or TGF-βtype I
receptors, such as LDN193189 [56, 61, 64] and SB431542
[57], have been used to enhance derivation of a myogenic
population from human PSCs. In some protocols, basal
medium supplement of insulin-transferrin-selenium (com-
monly known as ITS) has been used to induce the initial
step of mesodermal specification [53, 55, 66]. Oncostatin,
necrosulfonamide, ascorbic acid, insulin, and dexametha-
sone were recently used in combination with growth factors
and TGF-β1 inhibitors to increase skeletal myocyte deriva-
tion efficiency. These small molecules promoted a high per-
centage of skeletal muscle differentiation (up to 70% MHC
+
myotubes) and shortened the differentiation period to less
than a month [57]. A Notch antagonist DAPT (γ-secretase
inhibitor) increased MyoD and myogenin gene expression
[65]. A combination of CHIR99021 and DAPT synergisti-
cally enhanced myogenic differentiation [65]. Additionally,
the rescue effect of LDN193189 and SB431542 mixture was
demonstrated by the reduction of BMP4 levels and an
increase of fusion index when applied to myotubes prepared
from patient iPSCs with Duchenne muscular dystrophy [65].
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Table 1: Transgene-free methods of skeletal muscle differentiation using human pluripotent stem cells.
Reference PSC
type PSC culture
Myogenic progenitor
derivation and
proliferation
Progenitor
purification
Terminal
differentiation Efficiency of myogenic
differentiation In vivo engraftment Use of disease-specific
iPSCs
Culture condition Culture condition
Barberi
et al. 2007
[66]
ESCs
Feeder-
dependent
protocol
using MEF
Monolayer cells
were plated at 1 ×103
cells/cm
2
on MEF, or
3×10
3
cells/cm
2
on
feeder-free
gelatin-coated plates,
for 3-4 days under
standard ESC culture
conditions. The cells
were then switched
in DMEM/F12
supplemented with
ITS for 20 days, and
in αMEM, 10% FBS for
an additional 14 days.
FACS based on
CD73
+
/NCAM
+
.
Sorted NCAM
+
cells
were grown in
αMEM, 10% FBS
until confluence.
The cells were
differentiated in serum-
free N2 medium.
60–80% of sorted
NCAM
+
cells were
MyoD
+
. At 24 hours
after exposure
to N2 medium,
approximately 7%
and 46% of the total
cells expressed Pax7
+
and MyoG
+
,
respectively.
Upon terminal
differentiation, MyoG,
desmin, skeletal muscle
actin, and myosin
(MHC) were
identified.
Spontaneous twitching
of myotubes was
confirmed.
ESC-derived cells
(5 ×10
5
cells, CD73
+
/
NCAM
+
cells) were
transplanted into a
muscle injury model
in SCID/Beige mice.
The expression of
reporter proteins
(luciferase and GFP),
human cell-specific
nuclei, and laminin-
positive myofibers
were identified in the
grafted muscles.
Awaya
et al. 2012
[53]
ESCs
and
iPSCs
Feeder-
dependent
protocol,
using human
ES cell
maintenance
medium
(hESM)
EBs were formed
by suspension in
hESM for 7 days
and then plated
onto gelatin-coated
tissue culture plates
in ITS medium for
an additional
14 days.
EBs were differentiated
in skeletal muscle
induction medium
containing 10% FCS
and 5% HS until day
112 of differentiation.
In some experiments,
dissociated EB cells
(3 ×10
3
cells/cm
2
) were
seeded on collagen type
I-coated plates. On day
49, the medium was
changed to ITS
medium.
xIn the cells migrating
out of the EBs, the
clusters Pax3
+
and
Pax7
+
cells were
randomly distributed at
day 21. Skeletal
myosin-positive
multinucleated
myofibers had appeared
within most of the
attached EBs at day 63.
Progenitors (1–5×10
5
cells) were transplanted
into the muscle of
immunodeficient NOG
(NOD/Shiscid/IL-
2Rγnull) mice
following cardiotoxin
injury. Four weeks after
transplant, human
cell-specific laminin
A/C-positive nuclei
were detected in the TA
muscles. Further, the
detection of human-
specific laminin a2
proved that the
transplanted cells
produced human
protein around the
muscle fibers into
which they had
integrated.
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Table 1: Continued.
Reference PSC
type PSC culture
Myogenic progenitor
derivation and
proliferation
Progenitor
purification
Terminal
differentiation Efficiency of myogenic
differentiation In vivo engraftment Use of disease-specific
iPSCs
Culture condition Culture condition
Xu et al.
2013 [20] iPSCs
Feeder-free
protocol
using mTeSR
on Matrigel-
coated plates
EB culture in STEMdiff
APEL Medium
supplemented with
10 ng/ml FGF2, 0.5 μM
GSK3βinhibitor BIO,
20 μM forskolin (“triple
cocktail”) for 7 days.
EB cells were
then cultured on
Matrigel-coated
plates in DMEM,
2% HS for an
additional 29 days.
Under terminal
differentiation
procedures (day 36),
most of the cells
expressed desmin
(72%) and MyoG
(92%), forming
multinucleated
myofibers. Sarcomere
structures were also
confirmed by electron
microscopy.
iPSC-derived myogenic
progenitors (1 ×10
5
cells at day 14 of
differentiation) were
transplanted into
cardiotoxin-injured
muscles in NSG (NOD/
SCID/IL-2Rγnull)
mice. Human δ-
sarcoglycan expression
in myofibers and
colocalization of
human-specific histone
H2A and Pax7 were
characterized in the
grafted muscles.
Borchin
et al. 2013
[55]
ESCs
and
iPSCs
Feeder-free
protocol
using
mTeSR1 on
Matrigel-
coated plates
PSC colonies were
cultured in ITS medium
(DMEM/F12
supplemented with
ITS) in the presence of
3μM GSK3βinhibitor
CHIR99021 for 4 days,
and then in ITS
medium containing
20 ng/ml FGF2 for an
additional 14 days. The
cells were then
maintained in ITS
medium alone for a
further 17 days of
culture in ITS medium
alone.
FACS based on the
expression of HNK,
AChR, CXCR4,
C-MET, following
the differentiation
for 35 days.
FACS-sorted AChR
+
myocytes and CXCR4
−
/
C-MET
+
and CXCR4
+
/
C-MET
+
precursors
were plated onto
fibronectin/laminin-
coated tissue culture
wells in ITS medium
supplemented with
10 μM ROCK inhibitor
Y-27632. The myocytes
were maintained in ITS
medium with 50 ng/ml
IGF-I. The progenitor
cells were differentiated
in ITS medium.
In presorting cultures
of CXCR4
−
/C-MET
+
and CXCR4
+
/C-MET
+
cells isolated at day 35
of differentiation, >18%
Pax3
+
/Pax7
+
and >8%
MF20
+
muscle cells
were identified. In
postsorting cultures at
day 35, 97% in
CXCR4
−
/C-MET
+
and
98% in CXCR4
+
/C-
MET
+
were PAX3
+
;
84% in CXCR4
−
/C-
MET
+
and 96% in
CXCR4
+
/C-MET
+
were
PAX7
+
. After 3 days of
culture, few cells
retained PAX7
expression, whereas all
cells expressed MYH5.
In postsorting cultures
of AChR
+
myocytes, all
AChR
+
cells were
MyoG
+
and MHC
+
at
24 hours after plating.
7Stem Cells International

Table 1: Continued.
Reference PSC
type PSC culture
Myogenic progenitor
derivation and
proliferation
Progenitor
purification
Terminal
differentiation Efficiency of myogenic
differentiation In vivo engraftment Use of disease-specific
iPSCs
Culture condition Culture condition
Hwang
et al. 2013
[58]
ESCs
(OCT4-
GFP
reporter
line)
Feeder-
dependent
protocol
using MEF
Single cells were
cultured in suspension
on ultra-low
attachment plates to
form EBs for 9 days, in
high glucose DMEM
containing 5% FBS,
2mML-glutamine,
100 nM
dexamethasone,
100 μM
hydrocortisone, 1%
penicillin/
streptomycin, 1 mM
transferrin, 86.1 μM
recombinant insulin,
2 mM progesterone,
10.01 mM putrescine,
and 3.01 mM selenite.
The EBs were then spilt,
transferred, and
cultured on a Matrigel-
coated dish for an
additional 8 days.
The cells growing
out of the EBs were
concentrated by
FACS based on
PDGFRA and
OCT4-GFP.
PDGFRA
+
and
PDGFRA
−
cells
were expanded in
growth medium,
containing high
glucose DMEM,
10% FBS, 2 mM
L-glutamine, and
1% penicillin/
streptomycin.
PDGFRA
+
and
PDGFRA
−
cells (1 ×10
4
cells/cm
2
) were plated
on gelatin-coated
culture plates and
differentiated in high
glucose DMEM
containing 2 mM
L-glutamine, 100 nM
dexamethasone,
100 mM
hydrocortisone, 1%
penicillin/
streptomycin, 1 mM
transferrin, 86.1 mM
insulin, 2 mM
progesterone,
10.01 mM putrescine,
and 3.01 mM selenite
with 10% FBS or
without FBS.
The morphology of
PDGFRA
+
cells
progressively became
more spindle-like and
fused and formed
multinucleated
myotubes
(approximately 30%
MHC
+
) after 14 days of
terminal
differentiation. In
contrast, little or no
myogenic
differentiation was
observed in PDGFRA
−
cells population.
ESC-derived PDGFRA
+
cells were transplanted
into the muscle of
NOD/SCID mice
following cardiotoxin
injury. Human
laminin
+
myofibers
were identified after 14
days
posttransplantation.
Hosoyama
et al. 2014
[59]
ESC and
iPSCs
Feeder-
dependent
with MEF
(ESCs) or
feeder-
independent
protocols
(iPSCs)
Sphere-based culture
(EZ sphere) was
maintained in Stemline
medium containing
100 ng/ml FGF2,
100 ng/ml EGF, 5 ng/ml
heparin sulfate for 6–12
weeks (42–84 days).
The spheres were
passaged by mechanical
chopping every week.
Monolayer culture in
high glucose DMEM,
2% B27 serum-free
supplement on poly-L-
lysine/laminin-coated
coverslips.
Before terminal
differentiation,
progenitors were
approximately 40% and
56% Pax7
+
,
respectively. After 14
days of differentiation,
the prevalence of Pax7
+
,
MyoD
+
,36–61%
MyoG
+
, 24-25% MHC
+
after 14 days of
differentiation.
Spontaneous
contraction and AChR
+
in myotubes were
confirmed after 25 days
of terminal
differentiation.
The protocol
was applied to
patient-derived
iPSCs (ALS with
SOD1 or VAPB
mutation, BMD,
and SMA iPSC
lines) for
myogenic
differentiation.
8 Stem Cells International

Table 1: Continued.
Reference PSC
type PSC culture
Myogenic progenitor
derivation and
proliferation
Progenitor
purification
Terminal
differentiation Efficiency of myogenic
differentiation In vivo engraftment Use of disease-specific
iPSCs
Culture condition Culture condition
Shelton
et al. 2014
[54] and
2016 [22]
ESCs
Feeder-free
protocol in
E8 medium
Monolayer cells
(1.5 ×10
5
cells per well)
were plated on
Matrigel-coated dishes
in E8 medium
supplemented with
10 μM Y-27632
overnight. Cells were
grown in E6 medium
supplemented with
0.1% CHIR99021,
BMP4, or activin-A for
2 days and then
switched in
unsupplemented E6
medium until day 12.
From day 12 to 20, the
medium was replaced
with StemPro-34
media. Cells were then
returned to E6 medium
from day 20 to 35.
The cells were
differentiated in N2
medium until the
endpoint of the
experiment.
Skeletal myocytes
were prominent
approximately 37%
Pax7
+
and 14% MHC
+
by day 40 following 5
days of growth in N2
medium. Skeletal
muscle contractions
could be observed at
this time point. When
the cells were left in N2
media until day 50, 43%
Pax7
+
and 47% MHC
+
were identified.
Chal et al.
2015 [56]
and 2016
[64]
ESCs
and
iPSCs
Feeder-free
protocol in
mTeSR1
media
Single cells from PSC
colonies were seeded on
Matrigel-coated plates
(15,000–18,000 cells/
cm
2
) in mTeSR
medium supplemented
with Y-27632 for 1 day.
The medium was
changed to a DMEM-
based medium
supplement with ITS,
3μM CHIR99021, and
0.5 μM LDN193189 for
2 days. At day 3, 20 ng/
ml FGF2 was added for
an additional 3 days. At
day 6, the cells were
changed to a DMEM-
based medium
At day 8 in culture, the
medium was changed
to DMEM, 15% KSR,
supplemented with
2 ng/ml IGF-I for 4
days and then
supplemented with
both 10 ng/ml HGF and
2 ng/ml IGF-I after day
12.
After 20 days, the
cultures contained large
fields comprising
MHC
+
and MyoG
+
fibers and PAX7
+
cells.
By 4 weeks, ~22%
nuclei were MyoG
+
and
23% of nuclei were
Pax7
+
. The muscle
fibers showed
sarcomeres, as
demonstrated by titin
and fast MHC staining.
These striated fibers
exhibited spontaneous
twitching. The diameter
of the muscle fibers was
~3.5 μm.
9Stem Cells International

Table 1: Continued.
Reference PSC
type PSC culture
Myogenic progenitor
derivation and
proliferation
Progenitor
purification
Terminal
differentiation Efficiency of myogenic
differentiation In vivo engraftment Use of disease-specific
iPSCs
Culture condition Culture condition
supplemented with
10 ng/ml HGF, 2 ng/ml
IGF-I, 20 ng/ml FGF-2,
and 0.5 mM
LDN193189 for 2 days.
Caron et al.
2016 [57]
ESCs
and
iPSCs
Feeder-free
protocol in
serum-free
M2 medium
Monolayer culture at
2500 cells/cm
2
on
collagen-coated plates
and maintained in
skeletal muscle
induction medium
containing 5% HS,
3μM CHIR99021,
2μM Alk5 inhibitor,
10 ng/ml EGF, 10 μg/ml
insulin, 0.4 μg/ml
dexamethasone, and
200 μM ascorbic acid
for 10 days.
At day 10, cells were
dissociated with trypsin
and replated at 2500
cells/cm
2
onto
collagen-coated plates
and maintained for 8
days in skeletal
myoblast medium
containing 5% HS,
10 mg/ml insulin,
10 ng/ml EGF, 20 ng/ml
HGF, 10 ng/ml PDGF,
20 ng/ml FGF2, 20 mg/
ml oncostatin, 10 ng/ml
IGF-I, 2 μM SB431542,
and 200 μM ascorbic
acid. After 18 days of
differentiation, cells
were maintained in
myotube medium,
containing 10 μg/ml
insulin, 20 μg/ml
oncostatin, 50 mM
necrosulfonamide, and
200 μM ascorbic acid,
for 7 days.
After 10 days in
skeletal muscle
induction medium,
80% Pax3
+
, 20%
Pax7
+
, and
30–40% CD56
+
cells
were identified. At
day 18 (after the second
step of differentiation),
50–60% of the cells
were MyoD1
+
and 20%
desmin
+
. At day 26
(after the third and final
stage of the process),
50–80% of the cells
formed elongated and
multinucleated
myotubes that stained
positive for MyoG,
MHC, dystrophin,
and α-actinin.
FSHD1- or BMD-
affected ESCs were
differentiated into
MHC
+
myotubes
expressing MyoG.
Patient-derived iPSCs
with FSHD1 were also
tested.
Choi et al.
2016 [65]
ESCs
and
iPSCs
Feeder-
dependent
protocol
using MEF
At day 0, nonadherent
cells were plated on a
gelatin-coated dish (at
1.5 ×10
5
cells per well
of a 24-well plate), in
MEF-conditioned N2
media containing
10 ng/ml FGF2 and
10 μM Y-27632. At day
1, N2 media with 3 μM
To determine the
presence of fusion
component myoblasts,
the dissociated cells
from the CHIR99021-
DAPT culture (days
25–30) were also
replated.
At day 30 in the
CHIR99021-DAPT
culture, approximately
63% of cells were
MHC
+
and 61% were
MyoG
+
. The culture
resulted in
differentiation of
myoblasts into
multinucleated and
The dissociated
CHIR99021-DAPT
culture cells (1–3×10
6
cells) were transplanted
into the injured TA
muscle of NRG mice.
At 6 weeks after
transplantation, human
nuclei (human-specific
lamin A/C
+
) and
Disease-specific
cellular characteristics
were characterized in
the myotubes from
patient-derived iPSC
lines (FSHD, ALS with
C9orf72 repeats, and
DMD).
10 Stem Cells International

Table 1: Continued.
Reference PSC
type PSC culture
Myogenic progenitor
derivation and
proliferation
Progenitor
purification
Terminal
differentiation Efficiency of myogenic
differentiation In vivo engraftment Use of disease-specific
iPSCs
Culture condition Culture condition
GSK3βinhibitor
CHIR99021 was added.
At day 4, N2 media
with 10 mM γ-secretase
inhibitor DAPT were
added until day 12. The
resulting cells
(“CHIR99021-DAPT
culture”) were
maintained in defined
N2 media until day 30.
spontaneously
contractile myotubes
with sarcomere
structures. When the
cells from the
CHIR99021-DAPT
culture were replated,
the attached and
surviving cells were
mono-nucleated at day
2 after replating and
then formed
multinucleated
myotubes at day 10
after replating with
typical striations and
expression of 35%
dystrophin
+
, 37% titin
+
,
and 40% α-actinin
+
.
human-specific
laminin
+
myofibers
were detected in the
grafted muscles.
Swartz
et al. 2016
[60]
iPSCs
Feeder-free
culture on
vitronectin-
coated plates
in TeSR-E8.
When iPSC colonies
were ~250–400 μmin
diameter (day −1), 1.5%
DMSO in TeSR-E8
medium was added. On
day 0, cells were
cultured in chemically
defined medium
(CDM) supplemented
with 20 ng/ml FGF2,
10 μM LY294002,
10 ng/ml BMP4, 10 μM
CHIR99021 (FLyBC)
for 36 hours. Cells were
then cultured in CDM
supplemented with
20 ng/ml FGF-2 and
10 μM LY294002 (FLy)
for an additional
5.5 days. On day 7, cells
were cultured in MB-1
and 15% FBS for 6 days.
After 10 days of fusion
medium (22 days total
from the start of
differentiation), the
cells were changed to
N2 medium (DMEM/
F12 supplemented with
1% N2 supplement and
1% ITS).
At day 5, <5% of the
total cells were Pax3
+
mesodermal
progenitors. At day 36,
up to 64% (median
44.8%) of nuclei were
MyoG
+
. A mix of
intermediate- and late-
stage muscle cells as
demonstrated by
desmin
+
and MHC
+
.
After 63 total days in
fusion medium, brief
and spontaneous
contractions in a small
set of myotubes were
observed. Seven to 10
days after the addition
of N2 medium, robust
spontaneous
contractions
throughout the cell
11Stem Cells International

Table 1: Continued.
Reference PSC
type PSC culture
Myogenic progenitor
derivation and
proliferation
Progenitor
purification
Terminal
differentiation Efficiency of myogenic
differentiation In vivo engraftment Use of disease-specific
iPSCs
Culture condition Culture condition
On day 12, the cells
were cultured in fusion
medium (2% HS in
DMEM).
cultures were observed.
Titin
+
striation was
displayed.
Xi et al.
2017 [61]
ESCs
and
iPSCs
Feeder-free
culture on
Matrigel-
coated plates
in mTeSR1
medium.
On day −1, single cells
from PSC colonies
(25,000 cells/cm
2
) and
seeded on Matrigel-
coated plates in
mTeSR1 medium
containing 10 μM
Y-27632. On day 0, cells
were switched to basal
differentiation medium
(BDM; DMEM/F12, 1%
ITS and 0.5%
penicillin–
streptomycin)
supplemented with
3μM CHIR99021 for 2
days. On day 2, cells
were switched to BDM
supplemented with
200 nM LDN193189
and 10 μM SB431542
for another 2 days. On
day 4, culture medium
was changed to BDM
supplemented with
3μM CHIR99021 and
20 ng/ml FGF2 for 2
days. On day 6,
medium was switched
to a KSR/HGF/IGF-I-
based differentiation
medium (DMEM,
0.5% penicillin–
streptomycin and 15%
KSR, 10 ng/ml HGF,
2 ng/ml IGF-I) for
14–21 days.
At day 29, cell
suspension was filtered
through cell strainers to
exclude cell aggregates.
Filtered cells were
resuspended in SkGM2
medium supplemented
with 20 ng/ml FGF-2
and replated at 15,000–
20,000 cells onto
Matrigel-coated plates.
Cells were cultured for
7–10 days until
reaching >70%
confluency, and then
medium was switched
to N2 medium (BDM
containing 1% N2
supplement) for 5 days.
At day 2, ~80% cells
were Pax3
+
. Expression
of myogenic markers
was gradually increased
toward day 20. At day
27, large areas of MHC
+
cells emerged
throughout the culture,
and the majority also
expressed titin. A high
proportion of Pax7
+
,
MyoD
+
, and MyoG
+
was also identified. At
day 44, approximately
58% MHC
+
myocytes
and myotubes were
identified, as well as
cells outside MHC
+
area (6.5% Pax7
+
/
MyoD
−
, 9.1% Pax7
−
/
MyoD
+
, and 4.9%
Pax7
+
/MyoD
+
).
12 Stem Cells International

Table 1: Continued.
Reference PSC
type PSC culture
Myogenic progenitor
derivation and
proliferation
Progenitor
purification
Terminal
differentiation Efficiency of myogenic
differentiation In vivo engraftment Use of disease-specific
iPSCs
Culture condition Culture condition
Hicks et al.
2017 [79]
ESCs
and
iPSCs
Feeder-free
culture on
Matrigel-
coated plates
in mTeSR1
medium.
For direct
differentiation from
PSCs, two published
protocols (Shelton et al.
2014 [54] and Chal
et al. 2015 [56], listed in
Table 1) were used.
Myogenic progenitors
from day 50 culture
were dissociated and
filtered through
100 mm meshes.
FACS based on
HNK
−
/NCAM
+
or
ERBB3
+
/NGFR
+
were performed.
Sorted cells could be
grown in SkBM-2
media.
Myogenic progenitors
were induced to
differentiate in N2
media for 7 days. In
some experiments,
TGF-βinhibitors
(SB431542 or A83-01)
were added in
differentiation media.
HNK
−
/NCAM
+
enrichment increased
PAX7 and MYF5
expression by ~1.7-fold
in comparison to
unsorted SMPCs.
When differentiated in
culture, the number of
MHC
+
cells was
increased in HNK
−
/
NCAM
+
cells
compared to replated/
unsorted cells. ERBB3
+
/
NGFR
+
progenitors
were enriched for PAX7
and MYF5 by 20-fold in
comparison to
ERBB3
−
/NGFR
−
cells.
ERBB3
+
/NGFR
+
progenitors could form
homogenous myotubes
following terminal
differentiation. TGF-β
inhibitors (SB431542 or
A83-01) significantly
facilitated myotube
fusion and maturation,
as demonstrated by
MHC protein levels
(MYH1 and MYH8)
and sarcomere
formation.
The enriched cells
(1 ×106 cells per 5 μl,
injected 5–10 μlof
cells) were transplanted
into the injured or
irradiated TA muscle of
mdx-NRG mice. At 30
days after
transplantation,
human-specific lamin
A/C
+
, spectrin
+
, and
dystrophin
+
myofibers
were detected in the
grafted muscle. HNK
−
/
NCAM
+
sorted cells
survived in the grafted
muscle but did not
improve engraftment in
comparison to unsorted
cells. ERBB3
+
/NGFR
+
progenitors
significantly increased
the number of
engrafted myofibers in
comparison to NCAM
+
sorted cells.
Disease-specific iPSCs
from DMD patients
were used in this study.
The mutation in
DMD-iPSC lines was
corrected by CRISPR-
Cas9 gene editing,
which could restore
dystrophin expression.
AChR: acetylcholine receptor; αMEM: alpha minimum essential medium; ALS: amyotrophic lateral sclerosis; BMD: Becker muscular dystrophy; CXCR4: C-X-C chemokine receptor 4; DMD: Duchenne muscular
dystrophy; DMEM: Dulbecco’s modified Eagle’s medium; DMSO: dimethyl sulfoxide; EB: embryoid body; EGF: epidermal growth factor; ERBB3: receptor tyrosine-protein kinase erbB-3; SC: embryonic stem cell;
FACS: fluorescence-activated cell sorting; FBS: fetal bovine serum; FCS: fetal calf serum; FGF-2: fibroblast growth factor 2; FSHD: facioscapulohumeral muscular dystrophy; FTD: frontotemporal dementia; GFP:
green fluorescent protein; GSK3β: glycogen synthase kinase 3b; HNK: human natural killer; HS: horse serum; HGF: hepatocytegrowth factor; ITS: insulin-transferrin-selenium; iPSC: induced pluripotent stem cell;
KSR: knockout serum replacement; MEF: mouse embryonic fibroblasts; MHC: myosin heavy chain; MYH1, MYH5, or MYH8: myosin heavy chain type 1, 5, or 8; MyoG: myogenin; NCAM: neural cell adhesion
molecule (or CD56); NGFR: nerve growth factor receptor; NOD: nonobese diabetic; PDGF: platelet-derived growth factor receptor; PDGFRA: platelet-derived growth factor receptor-α; PSC: pluripotent stem cell;
SCID: severe combined immunodeficiency; SMA: spinal muscular atrophy; SOD1: superoxide dismutase 1; TA: tibialis anterior; TGF-β: transforming growth factor beta; VAPB: vesicle-associated membrane
protein/synaptobrevin-associated membrane protein B.
13Stem Cells International

4. Challenges for the Derivation of Skeletal
Myocytes from Human
PSCs Using Transgene-Free Methods
The evaluation of differentiation efficiency and myocyte
maturity has been inconsistent between studies that focus
on differentiating skeletal myocytes from stem cells. It would
be of great benefit to the field to establish standards for these
evaluations in order to more directly compare differentiation
methods. Another challenge facing the field is that in vitro
stem cell-derived skeletal myocytes often have an embryonic
or perinatal phenotype. Additional bioengineering methods
may be necessary in order to achieve skeletal muscle that is
fully mature and therefore more physiologically relevant to
in vivo skeletal muscle. In this section, we will discuss existing
concerns of the current methods for preparing skeletal myo-
cytes and myogenic progenitors from human PSCs, specifi-
cally related to transgene-free methods. However, several
concerns are also applicable to transgene methods.
4.1. Differentiation Efficiency. Compared to when using
transgene protocols, differentiation efficiency of skeletal
myocytes overall still remains low when using transgene-
free approaches. In order for the field to move forward
toward goals of disease modeling, drug testing, and thera-
peutic development, differentiation efficiency should be
improved. Currently, there is a wide range of reported effi-
ciencies due to differences in reporting methods and the def-
initions used to describe the maturity of myogenic cell types.
It is common to use stains for myogenic markers such as
Pax7, MyoD, myogenin, and MHC. However, there is varia-
tion in how these stains are used to determine efficiency.
Some protocols claim a very high efficiency rate of myogenic
differentiation but often use a pooled percentage of Pax7,
MyoD, myogenin, and/or MHC-positive cells. Others with
lower efficiency may only be using one of the markers, which
could be different from the marker chosen in another study.
Along with the usage of immunocytochemistry for MHC,
the counting of MHC
+
cells in a field of view, the number
of nuclei per myocyte, and the percentage of nuclei within
myocytes (fusion index) have all been used to evaluate dif-
ferentiation efficiency. Often, myocyte density and/or differ-
entiation efficiency varies across a culture. Therefore, it is
important to report the number of fields counted and how
they were selected—specifically noting how bias was con-
trolled. Overall, there is a need to standardize methods of
calculating differentiation efficiency in order to facilitate
comparisons between differentiation protocols.
4.2. Defining and Measuring the Extent of Myotube
Maturation. In recent years, there have been a number of
culture methods developed that yield MHC-positive skeletal
myocytes from human pluripotent cells. Many of them
require an extended culture period in comparison to methods
for deriving other cell types. A method yielding myogenic
progenitors or mature myocytes after a relatively short time
would be of high significance to the field. However, it is also
important to evaluate the maturity of cells yielded from rapid
preparations. To date, it is difficult to compare the maturation
state of myocytes generated by different methods due to dif-
ferences in how each study defines maturity. Some focus on
anatomical features, while others examine physiological func-
tionality. Ideally, both aspects should be considered when
evaluating myotube maturity. Studies taking an anatomical
approach tend to use immunocytochemistry or electron
microscopy to evaluate sarcomere formation and myofibril
alignment as indicators of myotube maturity. Immunocyto-
chemistry using antibodies against MHC or titin is a relatively
accessible method to detect striations (Figure 2(b)); however,
electron microscopy makes it possible to visualize sarcomeres
at an ultrastructural level and examine sarcomeric organiza-
tion and alignment (Figure 2(c)). It should be noted that
in some preparations of maturing human PSC-derived
myocytes, the results of immunocytochemical labeling of
sarcomeric proteins (such as titin) may not correlate with
ultrastructural results obtained through electron micros-
copy [67].
Some, but not all, studies take a physiological approach to
determine myocyte maturation by examining the functional-
ity of the cells. One method is to measure the frequency and
coordination of spontaneous contractions observed in the
differentiating myocytes. Contractions can be stimulated by
a calcium flux or with addition of acetylcholine to the culture.
Spontaneous contractions can also be observed shortly after
culture medium changes [59, 67]. To properly examine spon-
taneous contractions, it should be taken into consideration
when the cells were last given fresh media or were supple-
mented with the compounds that can promote contractions.
Electrophysiology has also been used to monitor contractions
and record contraction frequency and strength in cultured
myotubes [48, 68]. Further, calcium imaging using dyes such
as Fluo-3AM can be applied as an alternative method to elec-
trophysiology. With a wide variety of methods currently
being used, it is necessary to establish a preferred method
of assessing physiological maturity of in vitro myocytes to
better compare derivation methods. Another aspect of myo-
cyte maturity is the fiber type expressed. During myogenesis,
embryonic and slow type I MHC are expressed first. Then at
later stages of maturation, myofibers develop glycolytic fast
twitch MHC types IIa, IIb, and IIx [24, 69]. Commonly,
MHC expression is examined using an antibody that reacts
to all isoforms of MHC (such as MF20 clone), but a more
detailed evaluation of MHC type would be useful for describ-
ing derived myocyte maturation.
In order to use human PSC-derived myocytes for in vitro
modeling for adult-onset neuromuscular diseases, it is neces-
sary to generate fully matured myotubes. However, iPSC-
derived skeletal myocytes prepared using current methods
typically are of an embryonic or perinatal phenotype. In
addition to better understanding signaling molecules and
the timing required for generating mature myocytes, bioengi-
neering techniques will be needed to create surfaces recog-
nized by human PSC-derived myocytes as appropriate for
growth and maturation. Differentiation efficiency can likely
be improved by controlling features such as surface coatings,
adhesion ligands, and/or growth surfaces that encourage
directionality and elongation. For instance, micropatterned
surfaces can give myocytes much needed directionality [70].
14 Stem Cells International

It is likely that most two-dimensional culture environments
are not similar enough to in vivo and that three-dimensional
constructs will become necessary to encourage further stages
of maturation [67]. Cocultures with motor neurons may
support myotube maturation, as stimulation is required for
proper contractility in vivo [71].In the absence of motor
neurons, myotubes can be chemically stimulated to contract
by adding acetylcholine to the culture [72]. Electrical stimu-
lation can also induce contractions and enhance maturation
of myotubes [49, 73]. Further, mechanical stimulation may
accelerate muscle differentiation and maturation [74].
4.3. Cell Enrichment and Large-Scale Expansion. While the
way we report the efficiency of myogenic differentiation is
valuable, it is also important to improve upon current
methods to gain a pure population of myogenic progenitors
and skeletal myocytes in culture. When prepared by a
transgene-free method, the cultures commonly contain a
heterogeneous cell population with myocytes and other cell
types. Such heterogeneity influences the efficiency of in vivo
engraftment following transplantation [75]. In order to
improve differentiation efficiency, there is a need for more pre-
cise definition of which signal molecules to use and the timing
of their use, but improved cell sorting techniques will also be
necessary to further enrich derived myocytes. Fluorophore-
labeled progenitors can easily be purified by FACS, if genetic
modification is used [47, 49]. Also, several combinations of
specific cell surface markers can be used to enrich myogenic
progenitors and skeletal myocytes [55, 76–78]. Examples
include combinations of CD54
+
/integrin α9β1
+
/SDC2
+
[76],
CD45
−
/CD11b
−
/GlyA
−
/CD31
−
/CD34
−
/CD56
int
/ITGA7
hi
[77],
CD56
+
/CD15
−
[78], CXCR4
+
/C-MET
+
[55], and HNK
−
/
NCAM (CD56)
+
[65, 79]. The most recent study indicated
that a combination of two surface markers (ERBB3 and
NGFR) can be applied to sufficiently purify a specific cellular
population of human PSC-derived myogenic progenitors by
FACS [79].
Another important consideration when developing deri-
vation methods is whether they are adaptable to a large-scale
expansion of myogenic progenitors and skeletal myocytes.
Limited scalability seems to be a continued challenge among
methods [75], which limits practical application and transla-
tion to patients as cell-based therapies. Often, cells are main-
tained in small quantities as a monolayer culture that is not
always suitable for passaging. A recent study indicated that
animal serum could promote cell expansion in PSC-derived
myogenic progenitors, but the culture condition remained
less defined [56]. However, a sphere-based culture may work
to overcome this concern [59, 67]. As demonstrated in our
recent study, human PSC-derived spherical cultures can be
expanded for several weeks with specific signaling molecule
supplementation in the medium [59, 67].
5. Conclusions
Valuable knowledge regarding the differentiation of myo-
genic progenitors and myotubes from human PSCs has been
gradually accumulating [1, 3, 80–82]. Signaling molecules
significantly contribute to generating a sufficient number of
myogenic progenitors and myocytes from human ESCs and
iPSCs without genetic modification. In addition to directing
and enhancing differentiation of myogenic cells using signal-
ing molecules, recent bioengineering approaches such as
two-dimensional or three-dimensional culture, micropat-
terning, controlled stiffness, and mechanical, chemical, or
electrical stimulation have enabled us to more accurately
mimic the physiological environment of cultured cells while
improving throughput, accuracy, and efficiency of in vitro
analyses. A combination of signaling molecules and bioengi-
neering approaches may further enhance the differentiation
and maturation of human PSCs-derived myotubes for use in
disease modeling, drug testing, and therapeutic development.
Finally, in vitro cell models should represent similar morpho-
logical and physiological characteristics compared to tissues
in vivo. In the skeletal muscle, fully mature myotubes have
well-organized sarcomeres and the ability to contract in
response to stimulation. In order to assess the maturity of
human PSC-derived myotubes, it will be necessary to evaluate
them using both anatomical and functional approaches.
Conflicts of Interest
The authors declare that there is no conflict of interest
regarding the publication of this paper.
Acknowledgments
The first author (Nunnapas Jiwlawat) would like to thank
the Royal Thai Government Scholarship for the financial
support. This work was supported by grants from the ALS
Association (15-IIP-201, Masatoshi Suzuki), NIH/NINDS
(R01NS091540, Masatoshi Suzuki), and the University of
Wisconsin Foundation (Masatoshi Suzuki).
References
[1] T. Hosoyama, J. Van Dyke, and M. Suzuki, “Applications of
skeletal muscle progenitor cells for neuromuscular diseases,”
American Journal of Stem Cells, vol. 1, no. 3, pp. 253–263,
2012.
[2] I. H. Park, N. Arora, H. Huo et al., “Disease-specific induced
pluripotent stem cells,”Cell, vol. 134, no. 5, pp. 877–886, 2008.
[3] Y. Kodaka, G. Rabu, and A. Asakura, “Skeletal muscle cell
induction from pluripotent stem cells,”Stem Cells Interna-
tional, vol. 2017, Article ID 1376151, 16 pages, 2017.
[4] J. Chal and O. Pourquie, “Making muscle: skeletal myogenesis
in vivo and in vitro,”Development, vol. 144, no. 12, pp. 2104–
2122, 2017.
[5] B. Jostes, C. Walther, and P. Gruss, “The murine paired box
gene, Pax7, is expressed specifically during the development
of the nervous and muscular system,”Mechanisms of Develop-
ment, vol. 33, no. 1, pp. 27–37, 1990.
[6] M. Goulding, A. Lumsden, and A. J. Paquette, “Regulation of
Pax-3 expression in the dermomyotome and its role in muscle
development,”Development, vol. 120, no. 4, pp. 957–971,
1994.
[7] M. Buckingham and F. Relaix, “PAX3 and PAX7 as upstream
regulators of myogenesis,”Seminars in Cell & Developmental
Biology, vol. 44, pp. 115–125, 2015.
15Stem Cells International

[8] D. Sassoon, G. Lyons, W. E. Wright et al., “Expression of two
myogenic regulatory factors myogenin and MyoD1 during
mouse embryogenesis,”Nature, vol. 341, no. 6240, pp. 303–
307, 1989.
[9] Y. Cinnamon, N. Kahane, I. Bachelet, and C. Kalcheim, “The
sub-lip domain—a distinct pathway for myotome precursors
that demonstrate rostral-caudal migration,”Development,
vol. 128, no. 3, pp. 341–351, 2001.
[10] J. C. Kiefer and S. D. Hauschka, “Myf-5 is transiently expressed
in nonmuscle mesoderm and exhibits dynamic regional
changes within the presegmented mesoderm and somites
I-IV,”Developmental Biology, vol. 232, no. 1, pp. 77–90, 2001.
[11] M. Zhang and I. S. McLennan, “During secondary myotube
formation, primary myotubes preferentially absorb new nuclei
at their ends,”Developmental Dynamics, vol. 204, no. 2,
pp. 168–177, 1995.
[12] S. Tajbakhsh, U. Borello, E. Vivarelli et al., “Differential activa-
tion of Myf5 and MyoD by different Wnts in explants of
mouse paraxial mesoderm and the later activation of myogen-
esis in the absence of Myf5,”Development, vol. 125, no. 21,
pp. 4155–4162, 1998.
[13] H. Amthor, B. Christ, and K. Patel, “A molecular mechanism
enabling continuous embryonic muscle growth—a balance
between proliferation and differentiation,”Development,
vol. 126, no. 5, pp. 1041–1053, 1999.
[14] B. A. Parr, M. J. Shea, G. Vassileva, and A. McMahon, “Mouse
Wnt genes exhibit discrete domains of expression in the early
embryonic CNS and limb buds,”Development, vol. 119, no. 1,
pp. 247–261, 1993.
[15] M. Ikeya and S. Takada, “Wnt signaling from the dorsal neural
tube is required for the formation of the medial dermomyo-
tome,”Development, vol. 125, no. 24, pp. 4969–4976, 1998.
[16] R. van Amerongen and R. Nusse, “Towards an integrated view
of Wnt signaling in development,”Development, vol. 136,
no. 19, pp. 3205–3214, 2009.
[17] A. E. Chen, D. D. Ginty, and C. M. Fan, “Protein kinase A
signalling via CREB controls myogenesis induced by Wnt
proteins,”Nature, vol. 433, no. 7023, pp. 317–322, 2004.
[18] U. Borello, M. Coletta, S. Tajbakhsh et al., “Transplacental
delivery of the Wnt antagonist Frzb1 inhibits development
of caudal paraxial mesoderm and skeletal myogenesis in
mouse embryos,”Development, vol. 126, no. 19, pp. 4247–
4255, 1999.
[19] A. Suzuki, R. C. Pelikan, and J. Iwata, “WNT/β-catenin sig-
naling regulates multiple steps of myogenesis by regulating
step-specific targets,”Molecular and Cellular Biology, vol. 35,
no. 10, pp. 1763–1776, 2015.
[20] C. Xu, M. Tabebordbar, S. Iovino et al., “A zebrafish embryo
culture system defines factors that promote vertebrate myo-
genesis across species,”Cell, vol. 155, no. 4, pp. 909–921, 2013.
[21] C. Patsch, L. Challet-Meylan, E. C. Thoma et al., “Generation
of vascular endothelial and smooth muscle cells from human
pluripotent stem cells,”Nature Cell Biology, vol. 17, no. 8,
pp. 994–1003, 2015.
[22] M. Shelton, A. Kocharyan, J. Liu, I. S. Skerjanc, and W. L.
Stanford, “Robust generation and expansion of skeletal mus-
cle progenitors and myocytes from human pluripotent stem
cells,”Methods, vol. 101, pp. 73–84, 2016.
[23] R. L. Johnson, E. Laufer, R. D. Riddle, and C. Tabin, “Ectopic
expression of sonic hedgehog alters dorsal-ventral patterning
of somites,”Cell, vol. 79, no. 7, pp. 1165–1173, 1994.
[24] X. Feng, E. G. Adiarte, and S. H. Devoto, “Hedgehog acts
directly on the zebrafish dermomyotome to promote myo-
genic differentiation,”Developmental Biology, vol. 300, no. 2,
pp. 736–746, 2006.
[25] C. F. Bentzinger, Y. X. Wang, and M. A. Rudnicki, “Building
muscle: molecular regulation of myogenesis,”Cold Spring Har-
bor Perspectives in Biology, vol. 4, no. 2, 2012.
[26] C. Chiang, Y. Litingtung, E. Lee et al., “Cyclopia and defective
axial patterning in mice lacking sonic hedgehog gene func-
tion,”Nature, vol. 383, no. 6599, pp. 407–413, 1996.
[27] A. G. Borycki, L. Mendham, and C. P. Emerson Jr., “Control of
somite patterning by sonic hedgehog and its downstream sig-
nal response genes,”Development, vol. 125, no. 4, pp. 777–
790, 1998.
[28] K. A. Scata, D. W. Bernard, J. Fox, and J. L. Swain, “FGF recep-
tor availability regulates skeletal myogenesis,”Experimental
Cell Research, vol. 250, no. 1, pp. 10–21, 1999.
[29] L. L. Tortorella, D. J. Milasincic, and P. F. Pilch, “Critical
proliferation-independent window for basic fibroblast growth
factor repression of myogenesis via the p42/p44 MAPK signal-
ing pathway,”The Journal of Biological Chemistry, vol. 276,
no. 17, pp. 13709–13717, 2001.
[30] M. Lagha, J. D. Kormish, D. Rocancourt et al., “Pax3 regulation
of FGF signaling affects the progression of embryonic progen-
itor cells into the myogenic program,”Genes & Development,
vol. 22, no. 13, pp. 1828–1837, 2008.
[31] R. Stewart, L. Flechner, M. Montminy, and R. Berdeaux,
“CREB is activated by muscle injury and promotes muscle
regeneration,”PLoS One, vol. 6, no. 9, article e24714, 2011.
[32] O. Pourquie, M. Coltey, C. Breant, and N. M. le Douarin,
“Control of somite patterning by signals from the lateral
plate,”Proceedings of the National Academy of Sciences of the
United States of America, vol. 92, no. 8, pp. 3219–3223, 1995.
[33] E. Hirsinger, D. Duprez, C. Jouve, P. Malapert, J. Cooke, and
O. Pourquié, “Noggin acts downstream of Wnt and sonic
hedgehog to antagonize BMP4 in avian somite patterning,”
Development, vol. 124, no. 22, pp. 4605–4614, 1997.
[34] J. Gerhart, J. Elder, C. Neely et al., “MyoD-positive epiblast
cells regulate skeletal muscle differentiation in the embryo,”
The Journal of Cell Biology, vol. 175, no. 2, pp. 283–292, 2006.
[35] H. Wang, C. Zang, X. S. Liu, and J. C. Aster, “The role of Notch
receptors in transcriptional regulation,”Journal of Cellular
Physiology,vol. 230, no. 5, pp. 982–988, 2015.
[36] A. C. Rios, O. Serralbo, D. Salgado, and C. Marcelle, “Neural
crest regulates myogenesis through the transient activation of
NOTCH,”Nature, vol. 473, no. 7348, pp. 532–535, 2011.
[37] S. Jarriault, C. Brou, F. Logeat, E. H. Schroeter, R. Kopan, and
A. Israel, “Signalling downstream of activated mammalian
Notch,”Nature, vol. 377, no. 6547, pp. 355–358, 1995.
[38] T. J. Brennan, D. G. Edmondson, L. Li, and E. N. Olson,
“Transforming growth factor beta represses the actions of
myogenin through a mechanism independent of DNA bind-
ing,”Proceedings of the National Academy of Sciences of the
United States of America, vol. 88, no. 9, pp. 3822–3826, 1991.
[39] C. Krueger and F. M. Hoffmann, “Identification of retinoic
acid in a high content screen for agents that overcome the
anti-myogenic effect of TGF-beta-1,”PLoS One, vol. 5,
no. 11, article e15511, 2010.
[40] H. Lee, C. Haller, C. Manneville et al., “Identification of small
molecules which induce skeletal muscle differentiation in
embryonic stem cells via activation of the Wnt and inhibition
16 Stem Cells International

of Smad2/3 and sonic hedgehog pathways,”Stem Cells, vol. 34,
no. 2, pp. 299–310, 2016.
[41] D. Horbelt, J. H. Boergermann, A. Chaikuad et al., “Small mol-
ecules dorsomorphin and LDN-193189 inhibit myostatin/
GDF8 signaling and promote functional myoblast differentia-
tion,”The Journal of Biological Chemistry, vol. 290, no. 6,
pp. 3390–3404, 2015.
[42] J. Zhu, Y. Li, A. Lu et al., “Follistatin improves skeletal mus-
cle healing after injury and disease through an interaction
with muscle regeneration, angiogenesis, and fibrosis,”The
American Journal of Pathology, vol. 179, no. 2, pp. 915–
930, 2011.
[43] C. Brun, O. Monestier, S. Legardinier, A. Maftah, and
V. Blanquet, “Murine GASP-1 N-glycosylation is not essential
for its activity on C2C12 myogenic cells but alters Its secre-
tion,”Cellular Physiology and Biochemistry, vol. 30, no. 3,
pp. 791–804, 2012.
[44] S. E. Tollefsen, R. Lajara, R. H. McCusker, D. R. Clemmons,
and P. Rotwein, “Insulin-like growth factors (IGF) in muscle
development. Expression of IGF-I, the IGF-I receptor, and
an IGF binding protein during myoblast differentiation,”The
Journal of Biological Chemistry, vol. 264, no. 23, pp. 13810–
13817, 1989.
[45] H. H. Vandenburgh, P. Karlisch, J. Shansky, and R. Feldstein,
“Insulin and IGF-I induce pronounced hypertrophy of skeletal
myofibers in tissue culture,”American Journal of Physiology-
Cell Physiology, vol. 260, no. 3, pp. C475–C484, 1991.
[46] C. Serra, S. Bhasin, F. Tangherlini et al., “The role of GH and
IGF-I in mediating anabolic effects of testosterone on
androgen-responsive muscle,”Endocrinology, vol. 152, no. 1,
pp. 193–206, 2011.
[47] R. Darabi, R. W. Arpke, S. Irion et al., “Human ES- and iPS-
derived myogenic progenitors restore DYSTROPHIN and
improve contractility upon transplantation in dystrophic
mice,”Cell Stem Cell, vol. 10, no. 5, pp. 610–619, 2012.
[48] G. Skoglund, J. Laine, R. Darabi, E. Fournier, R. Perlingeiro,
and N. Tabti, “Physiological and ultrastructural features of
human induced pluripotent and embryonic stem cell-derived
skeletal myocytes in vitro,”Proceedings of the National Acad-
emy of Sciences of the United States of America, vol. 111,
no. 22, pp. 8275–8280, 2014.
[49] A. Tanaka, K. Woltjen, K. Miyake et al., “Efficient and repro-
ducible myogenic differentiation from human iPS cells: pros-
pects for modeling Miyoshi myopathy in vitro,”PLoS One,
vol. 8, no. 4, article e61540, 2013.
[50] T. Yasuno, K. Osafune, H. Sakurai et al., “Functional analysis
of iPSC-derived myocytes from a patient with carnitine palmi-
toyltransferase II deficiency,”Biochemical and Biophysical
Research Communications, vol. 448, no. 2, pp. 175–181, 2014.
[51] R. Abujarour, M. Bennett, B. Valamehr et al., “Myogenic dif-
ferentiation of muscular dystrophy-specific induced pluripo-
tent stem cells for use in drug discovery,”Stem Cells
Translational Medicine, vol. 3, no. 2, pp. 149–160, 2014.
[52] S. M. Maffioletti, M. F. Gerli, M. Ragazzi et al., “Efficient deri-
vation and inducible differentiation of expandable skeletal
myogenic cells from human ES and patient-specific iPS cells,”
Nature Protocols, vol. 10, no. 7, pp. 941–958, 2015.
[53] T. Awaya, T. Kato, Y. Mizuno et al., “Selective development
of myogenic mesenchymal cells from human embryonic and
induced pluripotent stem cells,”PLoS One, vol. 7, no. 12,
article e51638, 2012.
[54] M. Shelton, J. Metz, J. Liu et al., “Derivation and expansion of
PAX7-positive muscle progenitors from human and mouse
embryonic stem cells,”Stem Cell Reports, vol. 3, no. 3,
pp. 516–529, 2014.
[55] B. Borchin, J. Chen, and T. Barberi, “Derivation and FACS-
mediated purification of PAX3+/PAX7+ skeletal muscle pre-
cursors from human pluripotent stem cells,”Stem Cell Reports,
vol. 1, no. 6, pp. 620–631, 2013.
[56] J. Chal, M. Oginuma, Z. Al Tanoury et al., “Differentiation of
pluripotent stem cells to muscle fiber to model Duchenne
muscular dystrophy,”Nature Biotechnology, vol. 33, no. 9,
pp. 962–969, 2015.
[57] L. Caron, D. Kher, K. L. Lee et al., “A human pluripotent stem
cell model of facioscapulohumeral muscular dystrophy-
affected skeletal muscles,”Stem Cells Translational Medicine,
vol. 5, no. 9, pp. 1145–1161, 2016.
[58] Y. Hwang, S. Suk, S. Lin et al., “Directed in vitro myogenesis of
human embryonic stem cells and their in vivo engraftment,”
PLoS One, vol. 8, no. 8, article e72023, 2013.
[59] T. Hosoyama, J. V. McGivern, J. M. Van Dyke, A. D. Ebert,
and M. Suzuki, “Derivation of myogenic progenitors directly
from human pluripotent stem cells using a sphere-based cul-
ture,”Stem Cells Translational Medicine, vol. 3, no. 5,
pp. 564–574, 2014.
[60] E. W. Swartz, J. Baek, M. Pribadi et al., “A novel protocol for
directed differentiation of C9orf72-associated human induced
pluripotent stem cells into contractile skeletal Myotubes,”
Stem Cells Translational Medicine, vol. 5, no. 11, pp. 1461–
1472, 2016.
[61] H. Xi, W. Fujiwara, K. Gonzalez et al., “In vivo human somito-
genesis guides somite development from hPSCs,”Cell Reports,
vol. 18, no. 6, pp. 1573–1585, 2017.
[62] E. Latres, A. R. Amini, A. A. Amini et al., “Insulin-like growth
factor-1 (IGF-1) inversely regulates atrophy-induced genes via
the phosphatidylinositol 3-kinase/Akt/mammalian target of
rapamycin (PI3K/Akt/mTOR) pathway,”The Journal of Bio-
logical Chemistry, vol. 280, no. 4, pp. 2737–2744, 2005.
[63] J. Tureckova, E. M. Wilson, J. L. Cappalonga, and P. Rotwein,
“Insulin-like growth factor-mediated muscle differentiation:
collaboration between phosphatidylinositol 3-kinase-Akt-sig-
naling pathways and myogenin,”The Journal of Biological
Chemistry, vol. 276, no. 42, pp. 39264–39270, 2001.
[64] J. Chal, Z. Al Tanoury, M. Hestin et al., “Generation of human
muscle fibers and satellite-like cells from human pluripotent
stem cells in vitro,”Nature Protocols, vol. 11, no. 10,
pp. 1833–1850, 2016.
[65] I. Y. Choi, H. Lim, K. Estrellas et al., “Concordant but varied
phenotypes among Duchenne muscular dystrophy patient-
specific myoblasts derived using a human iPSC-based model,”
Cell Reports, vol. 15, no. 10, pp. 2301–2312, 2016.
[66] T. Barberi, M. Bradbury, Z. Dincer, G. Panagiotakos, N. D.
Socci, and L. Studer, “Derivation of engraftable skeletal myo-
blasts from human embryonic stem cells,”Nature Medicine,
vol. 13, no. 5, pp. 642–648, 2007.
[67] S. Jiwlawat, E. Lynch, J. Glaser et al., “Differentiation and sar-
comere formation in skeletal myocytes directly prepared from
human induced pluripotent stem cells using a sphere-based
culture,”Differentiation, vol. 96, pp. 70–81, 2017.
[68] X. Guo, K. Greene, N. Akanda et al., “In vitro differentiation of
functional human skeletal myotubes in a defined system,”Bio-
materials Science, vol. 2, no. 1, pp. 131–138, 2014.
17Stem Cells International

[69] J. Talbot and L. Maves, “Skeletal muscle fiber type: using
insights from muscle developmental biology to dissect targets
for susceptibility and resistance to muscle disease,”Wiley
Interdisciplinary Reviews: Developmental Biology, vol. 5,
no. 4, pp. 518–534, 2016.
[70] M. R. Salick, B. N. Napiwocki, J. Sha et al., “Micropattern
width dependent sarcomere development in human ESC-
derived cardiomyocytes,”Biomaterials, vol. 35, no. 15,
pp. 4454–4464, 2014.
[71] S. Ostrovidov, V. Hosseini, S. Ahadian et al., “Skeletal muscle
tissue engineering: methods to form skeletal myotubes and
their applications,”Tissue Engineering. Part B, Reviews,
vol. 20, no. 5, pp. 403–436, 2014.
[72] L. Rao, Y. Qian, A. Khodabukus, T. Ribar, and N. Bursac,
“Engineering human pluripotent stem cells into a functional
skeletal muscle tissue,”Nature Communications, vol. 9, no. 1,
p. 126, 2018.
[73] S. Ahadian, S. Ostrovidov, V. Hosseini et al., “Electrical stimu-
lation as a biomimicry tool for regulating muscle cell behav-
ior,”Organogenesis, vol. 9, no. 2, pp. 87–92, 2013.
[74] S. Rangarajan, L. Madden, and N. Bursac, “Use of flow, electri-
cal, and mechanical stimulation to promote engineering of
striated muscles,”Annals of Biomedical Engineering, vol. 42,
no. 7, pp. 1391–1405, 2014.
[75] J. Kim, A. Magli, S. S. K. Chan et al., “Expansion and purifica-
tion are critical for the therapeutic application of pluripotent
stem cell-derived myogenic progenitors,”Stem Cell Reports,
vol. 9, no. 1, pp. 12–22, 2017.
[76] A. Magli, T. Incitti, J. Kiley et al., “PAX7 targets, CD54, integ-
rin α9β1, and SDC2, allow isolation of human ESC/iPSC-
derived myogenic progenitors,”Cell Reports, vol. 19, no. 13,
pp. 2867–2877, 2017.
[77] A. Castiglioni, S. Hettmer, M. D. Lynes et al., “Isolation of pro-
genitors that exhibit myogenic/osteogenic bipotency in vitro
by fluorescence-activated cell sorting from human fetal mus-
cle,”Stem Cell Reports, vol. 2, no. 1, pp. 92–106, 2014.
[78] D. F. Pisani, N. Clement, A. Loubat et al., “Hierarchization of
myogenic and adipogenic progenitors within human skeletal
muscle,”Stem Cells, vol. 28, no. 12, pp. 2182–2194, 2010.
[79] M. R. Hicks, J. Hiserodt, K. Paras et al., “ERBB3 and NGFR
mark a distinct skeletal muscle progenitor cell in human devel-
opment and hPSCs,”Nature Cell Biology, vol. 20, no. 1, pp. 46–
57, 2018.
[80] H. M. Blau, “Cell therapies for muscular dystrophy,”The New
England Journal of Medicine, vol. 359, no. 13, pp. 1403–1405,
2008.
[81] F. Rinaldi and R. C. Perlingeiro, “Stem cells for skeletal muscle
regeneration: therapeutic potential and roadblocks,”Transla-
tional Research, vol. 163, no. 4, pp. 409–417, 2014.
[82] I. Roca, J. Requena, M. J. Edel, and A. B. Alvarez-Palomo,
“Myogenic precursors from iPS cells for skeletal muscle cell
replacement therapy,”Journal of Clinical Medicine, vol. 4,
no. 12, pp. 243–259, 2015.
18 Stem Cells International