Skeletal Muscle Cell Induction from Pluripotent Stem Cells

Kodaka, Yusaku; Rabu, Gemachu; Asakura, Atsushi

doi:https://doi.org/10.1155/2017/1376151

Stem Cells International

On this page

Abstract Introduction Conclusions Conflicts of Interest Acknowledgments References Copyright Related Articles

Special Issue

Skeletal Muscle Cells Generated from Pluripotent Stem Cells

View this Special Issue

Review Article | Open Access

Volume 2017 | Article ID 1376151 | https://doi.org/10.1155/2017/1376151

Skeletal Muscle Cell Induction from Pluripotent Stem Cells

Yusaku Kodaka,^1,2,3Gemachu Rabu,^1,2,3and Atsushi Asakura^1,2,3

Academic Editor: Silvia Brunelli

Received15 Jan 2017

Accepted28 Mar 2017

Published26 Apr 2017

Abstract

Embryonic stem cells (ESCs) and induced pluripotent stem cells (iPSCs) have the potential to differentiate into various types of cells including skeletal muscle cells. The approach of converting ESCs/iPSCs into skeletal muscle cells offers hope for patients afflicted with the skeletal muscle diseases such as the Duchenne muscular dystrophy (DMD). Patient-derived iPSCs are an especially ideal cell source to obtain an unlimited number of myogenic cells that escape immune rejection after engraftment. Currently, there are several approaches to induce differentiation of ESCs and iPSCs to skeletal muscle. A key to the generation of skeletal muscle cells from ESCs/iPSCs is the mimicking of embryonic mesodermal induction followed by myogenic induction. Thus, current approaches of skeletal muscle cell induction of ESCs/iPSCs utilize techniques including overexpression of myogenic transcription factors such as MyoD or Pax3, using small molecules to induce mesodermal cells followed by myogenic progenitor cells, and utilizing epigenetic myogenic memory existing in muscle cell-derived iPSCs. This review summarizes the current methods used in myogenic differentiation and highlights areas of recent improvement.

1. Introduction

Duchenne muscular dystrophy (DMD) is a genetic disease affecting approximately 1 in 3500 male live births [1]. It results in progressive degeneration of skeletal muscle causing complete paralysis, respiratory and cardiac complications, and ultimately death. Normal symptoms include the delay of motor milestones including the ability to sit and stand independently. DMD is caused by an absence of functional dystrophin protein and skeletal muscle stem cells, as well as the exhaustion of satellite cells following many rounds of muscle degeneration and regeneration [2]. The dystrophin gene is primarily responsible for connecting and maintaining the stability of the cytoskeleton of muscle fibers during contraction and relaxation. Despite the low frequency of occurrence, this disease is incurable and will cause debilitation of the muscle and eventual death in 20 to 30 year olds with recessive X-linked form of muscular dystrophy. Although there are no current treatments developed for DMD, there are several experimental therapies such as stem cell therapies.

Skeletal muscle is known to be a regenerative tissue in the body. This muscle regeneration is mediated by muscle satellite cells, a stem cell population for skeletal muscle [3, 4]. Although satellite cells exhibit some multipotential differentiation capabilities [5], their primary differentiation fate is skeletal muscle cells in normal muscle regeneration. Ex vivo expanded satellite cell-derived myoblasts can be integrated into muscle fibers following injection into damaged muscle, acting as a proof-of-concept of myoblast-mediated cell therapy for muscular dystrophies [6–9]. However, severe limitations exist in relation to human therapy. The number of available satellite cells or myoblasts from human biopsies is limited. In addition, the poor cell survival and low contribution of transplanted cells have hindered practical application in patients [6, 8, 9]. Human-induced pluripotent stem cells (hiPSCs) are adult cells that have been genetically reprogrammed to an embryonic stem cell- (ESC-) like state by being forced to express genes and factors important for maintaining the defining properties of ESCs. hiPSCs can be generated from a wide variety of somatic cells [10, 11]. They have the ability to self-renew and successfully turn into any type of cells. With their ability to capture genetic diversity of DMD in an accessible culture system, hiPSCs represent an attractive source for generating myogenic cells for drug screening.

The ESC/iPSC differentiation follows the steps of embryonic development. The origin of skeletal muscle precursor cells comes from the mesodermal lineage, which give rise to skeletal muscle, cardiac muscle, bone, and blood cells. Mesoderm subsequently undergoes unsegmented presomitic mesoderm followed by segmented compartments termed somites from anterior to caudal direction. Dermomyotome is an epithelial cell layer making up the dorsal part of the somite underneath the ectoderm. Dermomyotome expresses Pax3 and Pax7 and gives rise to dermis, skeletal muscle cells, endothelial cells, and vascular smooth muscle [12]. Dermomyotome also serves as a tissue for secreted signaling molecules to the neural tube, notochord, and sclerotome [13, 14]. Upon signals from the neural tube and notochord, the dorsomedial lip of dermomyotome initiates and expresses skeletal muscle-specific transcription factors such as MyoD and Myf5 to differentiate into myogenic cells termed myoblasts. Myoblasts then migrate beneath the dermomyotome to form myotome. Eventually, these myoblasts fuse with each other to form embryonic muscle fibers. ESCs/iPSCs mimic these steps toward differentiation of skeletal muscle cells. Many studies utilize methods of overexpression of muscle-related transcription factors such as MyoD or Pax3 [15], or the addition of small molecules which activate or inhibit myogenic signaling during development. Several studies show that iPSCs retain a bias to form their cell type of origin due to an epigenetic memory [16–19], although other papers indicate that such epigenetic memory is erased during the reprogramming processes [20–22]. Therefore, this phenomenon is not completely understood at the moment. In light of these developments, we have recently established mouse myoblast-derived iPSCs capable of unlimited expansion [23]. Our data demonstrates that these iPSCs show higher myogenic differentiation potential compared to fibroblast-derived iPSCs. Thus, myogenic precursor cells generated from human myoblast-derived iPSCs expanded ex vivo should provide an attractive cell source for DMD therapy. However, since DMD is a systemic muscle disease, systemic delivery of myoblasts needs to be established for efficient cell-based therapy.

2. Myogenic Master Transcription Factors for Skeletal Muscle Development (Figure 1)

Figure 1

Hierarchal master transcription factor cascade for myogenesis. For myogenic differentiation during early embryogenesis, Mesogenin1 works as a master regulator for unsegmented presomitic mesoderm formation. Then, segmented somites are formed. Pax3 and Pax7 are activated in presomitic mesoderm, which generates somite-derived dermomyotome. Pax3 and Pax7 then work as master regulators for myogenic progenitor cell induction. Finally, MyoD and Myf5 are upregulated in the dorsomedial lip of dermomyotome and function as master regulators for myogenic specification to generate myoblasts. Eventually, myoblasts stop cell proliferation and express myogenin, which induces terminal differentiation of myoblasts to form multinucleated myotubes.

During developmental myogenesis, presomitic mesoderm is first formed by Mesogenin1 upregulation, which is a master regulator of presomitic mesoderm [24]. Then, the paired box transcription factor Pax3 gene begins to be expressed from presomitic mesoderm to dermomyotome [25]. Following Pax3 expression, Pax7 is also expressed in the dermomyotome [26], and then Myf5 and MyoD, skeletal muscle-specific transcription factor genes, begin to be expressed in the dorsomedial lip of the dermomyotome in order to give rise to myoblasts which migrate beneath the dermomyotome to form the myotome. Subsequently, Mrf4 and Myogenin, other skeletal muscle-specific transcription factor genes, followed by skeletal muscle structural genes such as myosin heavy chain (MyHC), are expressed in the myotome for myogenic terminal differentiation (Figure 1) [27, 28]. Pax3 directly and indirectly regulates Myf5 expression in order to induce myotomal cells. Dorsal neural tube-derived Wnt proteins and floor plate cells in neural tube and notochord-derived sonic hedgehog (Shh) positively regulate myotome formation [13, 29]. Neural crest cells migrating from dorsal neural tubes are also involved in myotome formation: Migrating neural crest cells come across the dorsomedial lip of the dermomyotome, and neural crest cell-expressing Delta1 is transiently able to activate Notch1 in the dermomyotome, resulting in conversion of Pax3/7(+) myogenic progenitor cells into MyoD/Myf5(+) myotomal myoblasts [30, 31]. By contrast, bone morphogenetic proteins (BMPs) secreted from lateral plate mesoderm are a negative regulator for the myotome formation by maintaining Pax3/Pax7(+) myogenic progenitor cells [29, 32]. Pax3 also regulates cell migration of myogenic progenitor cells from ventrolateral lip of dermomyotome to the limb bud [33]. Pax3 mutant mice lack limb muscle but trunk muscle development is relatively normal [34]. Pax3/Pax7 double knockout mice display failed generation of myogenic cells, suggesting that Pax3 and Pax7 are critical for proper embryonic myogenesis [35]. Therefore, both Pax3 and Pax7 are also considered master transcription factors for the specification of myogenic progenitor cells. Importantly, MyoD was identified as the first master transcription factor for myogenic specification since MyoD is directly able to reprogram nonmuscle cell type to myogenic lineage when overexpressed [36–38]. In addition, genetic ablation of MyoD family gene(s) via a homologous gene recombination technique causes severe myogenic developmental or regeneration defects [39–45]. Finally, genetic ablation of combinatory MyoD family genes demonstrates that MyoD^−/−:Myf5^−/−:MRF4^−/− mice do not form any skeletal muscle during embryogenesis, indicating the essential roles in skeletal muscle development of MyoD family genes [28, 46]. It was proven that Pax3 also possesses myogenic specification capability since ectopic expression of Pax3 is sufficient to induce myogenic programs in both paraxial and lateral plate mesoderm as well as in the neural tube during chicken embryogenesis [47]. In addition, genetic ablation of Pax3 and Myf5 display complete defects of body skeletal muscle formation during mouse embryogenesis [48]. Finally, overexpression of Pax7 can convert CD45(+)Sca-1(+) hematopoietic cells into skeletal muscle cells [49]. From these notions, overexpression of myogenic master transcription factors such as MyoD or Pax3 has become the major strategy for myogenic induction in nonmuscle cells, including ES/iPSCs.

3. Overexpression Approaches of Myogenic Master Transcription Factors in ESCs/iPSCs

The overexpression of MyoD approach to induce myogenic cells from mESCs was first described by Dekel et al. in 1992. This has been a standard approach for the myogenic induction from pluripotent stem cells (Table 1). Ozasa et al. first utilized Tet-Off systems for MyoD overexpression in mESCs and showed desmin(+) and MyHC(+) myotubes in vitro [50]. Warren et al. transfected synthetic MyoD mRNA in to hiPSCs for 3 days, which resulted in myogenic differentiation (around 40%) with expression of myogenin and MyHC [51]. Tanaka et al. utilized a PiggyBac transposon system to overexpress MyoD in hiPSCs. The PiggyBac transposon system allows cDNAs to stably integrate into the genome for efficient gene expression. After integration, around 70 to 90% of myogenic cells were induced in hiPSC cultures within 5 days [52]. This study also utilized Miyoshi myopathy patient-derived hiPSCs for the MyoD-mediated myogenic differentiation. Miyoshi myopathy is a congenital distal myopathy caused by defective muscle membrane repair due to mutations in dysferlin gene. The patient-derived hiPSC-myogenic cells will be able to provide the opportunity for therapeutic drug screening. Abujarour et al. also established a model of patient-derived skeletal muscle cells which express NCAM, myogenin, and MyHC by doxycycline-inducible overexpression of MyoD in DMD patient-derived hiPSCs [53]. Interestingly, MyoD-induced iPSCs also showed suppression of pluripotent genes such as Nanog and a transient increase in the gene expression levels of T (Brachyury T), Pax3, and Pax7, which belong to paraxial mesodermal/myogenic progenitor genes, upstream genes of myogenesis. It is possible that low levels of MyoD activity in hiPSCs may initially suppress their pluripotent state while failing to induce myogenic programs, which may result in transient paraxial mesodermal induction. Supporting this idea, BAF60C, a SWI/SNF component that is involved in chromatin remodeling and binds to MyoD, is required to induce full myogenic program in MyoD-overexpressing hESCs [54]. Overexpression of MyoD alone in hESC can only induce some paraxial mesodermal genes such as Brachyury T, mesogenin, and Mesp1 but not myogenic genes. Co-overexpression of MyoD and BAF60C was now able to induce myogenic program but not paraxial mesodermal gene expression, indicating that there are different epigenetic landscapes between pluripotent ESCs/iPSCs and differentiating ESC/iPSCs in which MyoD is more accessible to DNA targets than those in pluripotent cells. The authors then argued that without specific chromatin modifiers, only committed cells give rise to myogenic cells by MyoD. These results strongly indicate that nuclear landscapes are important for cell homogeneity for the specific cell differentiation in ESC/iPSC cultures. Similar observations were seen in overexpression of MyoD in P19 embryonal carcinoma stem cells, which can induce paraxial mesodermal genes including Meox1, Pax3, Pax7, Six1, and Eya2 followed by muscle-specific genes. However, these MyoD-induced paraxial mesodermal genes were mediated by direct MyoD binding to their regulatory regions, which was proven by chromatin immunoprecipitation (ChIP) assays, indicating the novel role for MyoD in paraxial mesodermal cell induction [55].

hESCs/iPSCs have been differentiated into myofibers by overexpression of MyoD, and this method is considered an excellent in vitro model for human skeletal muscle diseases for muscle functional tests, therapeutic drug screening, and genetic corrections such as exon skipping and DNA editing. Shoji et al. have shown that DMD patient-derived iPSCs were used for myogenic differentiation via PiggyBac-mediated MyoD overexpression. These myogenic cells were treated with morpholinos for exon-skipping strategies for dystrophin gene correction and showed muscle functional improvement [56]. Li et al. have shown that patient-derived hiPSC gene correction by TALEN and CRISPR-Cas9 systems, and these genetically corrected hiPSCs were used for myogenic differentiation via overexpression of MyoD [57]. This work also revealed that the TALEN and CRISPR-Cas9-mediated exon 44 knock-in approach in the dystrophin gene has high efficiency in gene-editing methods for DMD patient-derived cells in which the exon 44 is missing in the genome.

Along this line of the strategy, Darabi et al. first performed overexpression of Pax3 gene, which can be activated by treatment with doxycycline in mESCs, and showed efficient induction of MyoD/Myf5(+) skeletal myoblasts in EB cultures [15]. Upon removing doxycycline, these myogenic cells underwent MyHC(+) myotubes. However, teratoma formation was observed after EB cell transplantation into cardiotoxin-injured regenerating skeletal muscle in Rag2^−/−:γC^−/− immunodeficient mice [15]. This indicates that myogenic cell cultures induced by Pax3 in mESCs still contain some undifferentiated cells which gave rise to teratomas. To overcome this problem, the same authors separated paraxial mesodermal cells from Pax3-induced EB cells by FACS using antibodies against cell surface markers as PDGFRα(+)Flk-1(−) cell populations. After cell sorting, isolated Pax3-induced paraxial mesodermal cells were successfully engrafted and contributed to regenerating muscle in mdx:Rag2^−/−:γC^−/− DMD model immunodeficient mice without any teratoma formations. Darabi et al. also showed successful myogenic induction in mESCs and hES/iPSCs by overexpression of Pax7 [58, 59]. Pax3 and Pax7 are not only expressed in myogenic progenitor cells. They are also expressed in neural tube and neural crest cell-derived cells including a part of cardiac cell types in developmental stage, suggesting that further purification to skeletal muscle cell lineage is crucial for therapeutic applications for muscle diseases including DMD.

Taken together, overexpression of myogenic master transcription factors such as MyoD or Pax3/Pax7 is an excellent strategy for myogenic induction in hESCs and hiPSCs, which can be utilized for in vitro muscle disease models for their functional test and drug screening. However, for the safe stem cell therapy, it is essential to maintain the good cellular and genetic qualities of hESC/hiPSC-derived myogenic cells before transplantation. Therefore, random integration sites of overexpression vectors for myogenic master transcription factors and inappropriate expression control of these transgenes may diminish the safety of using these induced myogenic cells for therapeutic stem cell transplantation.

4. Supplement with Defined Factors for Myogenic Induction in ESCs/iPSCs

Stepwise induction protocols utilizing small molecules and growth factors have been established as alternative myogenic induction approaches and a more applicable method for therapeutic situations. As described above, during embryonic myogenesis, somites and dermomyotomes receive secreted signals such as Wnts, Notch ligands, Shh, FGF, BMP, and retinoic acid (RA) with morphogen gradients from surrounding tissues in order to induce the formation of myogenic cells (Figure 2). The canonical Wnt signaling pathway has been shown to play essential roles in the development of myogenesis. In mouse embryogenesis, Wnt1 and Wnt3a secreted from the dorsal neural tube can promote myogenic differentiation of dorsomedial dermomyotome via activation of Myf5 [31, 32, 60]. Wnt3a is able to stabilize β-catenin which associates with TCF/LEF transcription factors that bind to the enhancer region of Myf5 during myogenesis [61]. Other Wnt proteins, Wnt6 and Wnt7a, which emerge from the surface ectoderm, induce MyoD [62]. BMP functions as an inhibitor of myogenesis by suppression of some myogenic gene expressions. In the lateral mesoderm, BMP4 is able to increase Pax3 expression which delays Myf5 expression in order to maintain an undifferentiated myogenic progenitor state [63]. Therefore, Wnts and BMPs regulate myogenic development by antagonizing each other for myogenic transcription factor gene expression [64, 65]. Wnt also induces Noggin expression to antagonize BMP signals in the dorsomedial lip of the dermomyotome [66]. In this region, MyoD expression level is increased, which causes myotome formation. Notch signaling plays essential roles for cell-cell communication to specify the different cells in developmental stages. During myotome formation, Notch is expressed in dermomyotome, and Notch1 and Notch2 are expressed in dorsomedial lip of dermomyotome. Delta1, a Notch ligand, is expressed in neural crest cells which transiently interact with myogenic progenitor cells in dorsomedial lip of dermomyotome via Notch1 and 2. This contact induces expression of the Myf5 or MyoD gene in the myogenic progenitor cells followed by myotome formation. The loss of function of Delta1 in the neural crest displays delaying skeletal muscle formation [67]. Knockdown of Notch genes or use of a dominant-negative form of mastermind, a Notch transcriptional coactivator, clearly shows dramatically decrease of Myf5 and MyHC(+) myogenic cells. Interestingly, induction of Notch intracellular domain (NICD), a constitutive active form of Notch, can promote myogenesis, while continuous expression of NICD prevents terminal differentiation. Taken together, transient and timely activation of Notch is crucial for myotome formation from dermomyotome [30].

Current studies for myogenic differentiation of ESCs/iPSCs have utilized supplementation with some growth factors and small molecules, which would mimic the myogenic development described above in combination with embryoid body (EB) aggregation and FACS separation of mesodermal cells (Table 2). To induce paraxial mesoderm cells from mESCs, Sakurai et al. utilized BMP4 in serum-free cultures [68]. Three days after treatment with BMP4, mESCs could be differentiated into primitive streak mesodermal-like cells, but the continuous treatment with BMP4 turned the ESCs into osteogenic cells. Therefore, they used LiCl after treatment with BMP4 to enhance Wnt signaling, which is able to induce myogenic differentiation. After treatment with LiCl, PDGFRα(+) E-cadherin(−) paraxial mesodermal cells were sorted by FACS. These sorted cells were cultured with IGF, HGF, and FGF for two weeks in order to induce myogenic differentiation. Hwang et al. have shown that treatment with Wnt3a efficiently promotes skeletal muscle differentiation of hESCs [69]. hESCs were cultured to form EB for 9 days followed by differentiation of EBs for additional 7 days, and then PDGFRα(+) cells were sorted by FACS. These PDGFRα(+) cells were cultured with Wnt3a for additional 14 days. Consequently, these Wnt3a-treated cells display significantly increased myogenic transcription factors and structural proteins at both mRNA and protein levels. An interesting approach to identify key molecules that induce myogenic cells was reported by Xu et al. [70]. They utilized reporter systems in zebrafish embryos to display myogenic progenitor cell induction and myogenic differentiation in order to identify small compounds for myogenic induction. Myf5-GFP marks myogenic progenitor cells, while myosin light polypeptide 2 (mylz2)-mCherry marks terminally differentiated muscle cells. They found that a mixed cocktail containing GSK3β inhibitor, bFGF, and forskolin has the potential to induce robust myogenic induction in hiPSCs. GSK3β inhibitors act as a canonical Wnt signaling activator via stabilizing β-catenin protein, which is crucial for inducing mesodermal cells. Forskolin activates adenylyl cyclase, which then stimulates cAMP signaling. cAMP response element-binding protein (CREB) is able to stimulate cell proliferation of primary myoblasts in vitro, suggesting that the forskolin-cAMP-CREB pathway may help myogenic cell expansion [71], However the precise mechanisms for CREB-mediated myogenic cell expansion remain unclear. The adenylyl cyclase signaling cascade leads to CREB activation [71]. During embryogenesis, phosphorylated CREB has been found at dorsal somite and dermomyotome. CREB gene knockout mice display significantly decreased Myf5 and MyoD expressions in myotomes. While activation of Wnt1 or Wnt7a promotes Pax3, Myf5, and MyoD expressions, inhibition of CREB eliminates these Wnt-mediated myogenic gene expressions without altering the Wnt canonical pathway, suggesting that CREB-induced myogenic activation may be mediated through noncanonical Wnt pathways. Several groups also utilized GSK3β inhibitors for inducing mesodermal cells from ESCs and iPSCs [72, 73]. These mesodermal cell-like cells were expanded by treatment with bFGF, and then ITS (insulin/transferrin/selenite) or N2 medium were used to induce myogenic differentiation. Finally, bFGF is a stimulator for myogenic cell proliferation. Caron et al. demonstrated that hESCs treated with GSK3β inhibitor, ascorbic acid, Alk5 inhibitor, dexamethasone, EGF, and insulin generated around 80% of Pax3(+) myogenic precursor cells in 10 days [74]. Treatment with SB431542, an inhibitor of Alk4, 5, and 7, PDGF, bFGF, oncostatin, and IGF was able to induce these Pax3(+) myogenic precursor cells into around 50–60% of MyoD(+) myoblasts in an additional 8 days. For the final step, treatment with insulin, necrosulfonamide, an inhibitor of necrosis, oncostatin, and ascorbic acid was able to induce these myoblasts into myotubes in an additional 8 days. Importantly, the same authors utilized ESCs from human facioscapulohumeral muscular dystrophy (FSHD) to demonstrate the myogenic characterization after myogenic induction by using the protocol described above. Hosoyama et al. have shown that hESCs/iPSCs with high concentrations of bFGF and EGF in combination with cell aggregation, termed EZ spheres, efficiently give rise to myogenic cells [75]. After 6-week culture, around 40–50% of cells expressed Pax7, MyoD, or myogenin. However, the authors also showed that EZ spheres included around 30% of Tuj1(+) neural cells. Therefore, the authors discussed the utilization of molecules for activation of mesodermal and myogenic signaling pathways such as BMPs and Wnts.

Taken together, it is likely that the induced cell populations from ESCs/iPSCs may contain other cell types such as neural cells or cardiac cells because neural cells share similar transcription factor gene expression with myogenic cells such as Pax3, and cardiac cells also develop from mesodermal cells. To overcome this limitation, Chal et al. treated ESCs/iPSCs with BMP4 inhibitor, which prevents ESCs/iPSCs from differentiating into lateral mesodermal cells [76, 77]. To identify what genes are involved in myogenic differentiation in vivo, they performed a microarray analysis which compared samples of dissected fragments in mouse embryos, which are able to separate tail bud, presomitic mesoderm, and somite regions. From microarray data, the authors focused on Mesogenin1 (Msgn1) and Pax3 genes. Importantly, they utilized three lineage tracing reporters, Msgn1-repV (Mesogenin1-Venus) marking posterior somitic mesoderm, Pax3-GFP marking anterior somitic mesoderm and myogenic cells, and Myog-repV (Myogenin-Venus) marking differentiated myocytes, allowing the authors to readily detect different differentiation stages during ESC/iPSC cultures. Treatment with GSK3β inhibitors and then BMP inhibitors in ESC cultures induced Msgn1(+) somitic mesoderm with 45 to 65% efficiencies, Pax3(+) anterior somitic mesoderm with 30 to 50% efficiencies, and myogenin(+) myogenic cells with 25 to 30% efficiencies. Furthermore, the authors examined differentiation of mdx ESCs into skeletal muscle cells and revealed abnormal branching myofibers. Current protocols were also published and described more details for hiPSC differentiation [77].

5. Induction of Skeletal Muscle Cells from iPSC-Derived Mesoangioblast-Like Cells

Some nonmuscle cell populations such as mesoangioblasts have the potential to differentiate into skeletal muscle [6]. Mesoangioblasts were originally isolated from embryonic mouse dorsal aorta as vessel-associated pericyte-like cells, which have the ability to differentiate into a myogenic lineage in vitro and in vivo [6, 78]. Mesoangioblasts possess an advantage for the clinical cell-based treatment because they can be injected through an intra-arterial route to systemically deliver cells, which is crucial for therapeutic cell transplantation for muscular dystrophies [79]. Tedesco et al. successfully generated human iPSC-derived mesoangioblast-like stem/progenitor cells called HIDEMs by stepwise protocols without FACS sorting [80, 81]. They displayed similar gene expression profiles as embryonic mesoangioblasts. However, HIDEMs do not spontaneously differentiate into skeletal muscle cells, and thus, the authors utilized overexpression of MyoD to differentiate into skeletal muscle cells. Similar to mesoangioblasts, HIDEM-derived myogenic cells could be delivered to injured muscle via intramuscular and intra-arterial routes. Furthermore, HIDEMs have been generated from hiPSCs derived from limb-girdle muscular dystrophy (LGMD) type 2D patients and used for gene correction and cell transplantation experiments for the potential therapeutic application.

6. Enrichment of ESC/iPSC-Derived Myogenic Precursor Cells

Myogenic precursor cells derived from ESCs/iPSCs by various methods may contain nonmuscle cells. Therefore, further purification is mandatory for therapeutic applications. Barberi et al. isolated CD73(+) multipotent mesenchymal precursor cells from hESCs by FACS, and these cells underwent differentiation into fat, cartilage, bone, and skeletal muscle cells [82]. Barberi et al. also demonstrated that hESCs cultured on OP9 stroma cells generated around 5% of CD73(+) adult mesenchymal stem cell-like cells [83]. After FACS, these CD73(+) mesenchymal stem cell-like cells were cultured with ITS medium for 4 weeks and then gave rise to NCAM(+) myogenic cells. After FACS sorting, these NCAM(+) myogenic cells were purified by FACS and transplanted into immunodeficient mice to show their myogenic contribution to regenerating muscle.

It has been shown that many genes are associated with myogenesis. In addition, exhaustive analysis, such as microarray, RNA-seq, and single cell RNA-seq supplies much gene information in many different stages. Chal et al. showed key signaling factors by microarray from presomitic somite, somite, and tail bud cells [76]. They found that initial Wnt signaling has important roles for somite differentiation. Furthermore, mapping differentiated hESCs by single cell RNA-seq analysis is useful to characterize each differentiated stage [84].

As shown above, cell sorting of mesodermal progenitor cells, mesenchymal precursor cells, or myogenic cells is a powerful tool to obtain pure myogenic populations from differentiated pluripotent cells. Sakurai et al. have been able to induce PDGFRα(+)Flk-1(−) mesodermal progenitor cells by FACS followed by myogenic differentiation [85]. Chang et al. and Mizuno et al. have been able to sort SMC-2.6(+) myogenic cells from mouse ESCs/iPSCs [86, 87]. These SMC-2.6(+) myogenic cells were successfully engrafted into mouse regenerating skeletal muscle. However, this SMC-2.6 antibody only recognizes mouse myogenic cells but not human myogenic cells [86, 88]. Therefore, Borchin et al. have shown that hiPSC-derived myogenic cells differentiated into c-met(+)CXCR4(+)ACHR(+) cells, displaying that over 95% of sorted cells are Pax7(+) myogenic cells [72]. Taken together, current myogenic induction protocols utilizing small molecules and growth factors, with or without myogenic transcription factors, have been largely improved in the last 5 years. It is crucial to standardize the induction protocols in the near future to obtain sufficient myogenic cell conversion from pluripotent stem cells.

7. Epigenetic Myogenic Memory in Myoblast-Derived iPSCs

Recent work demonstrated that cells inherit a stable genetic program partly through various epigenetic marks, such as DNA methylation and histone modifications. This cellular memory needs to be erased during genetic reprogramming, and the cellular program reverted to that of an earlier developmental stage [16, 22, 89]. However, iPSCs retaining an epigenetic memory of their origin can readily differentiate into their original tissues [16–19, 90–100]. This phenomenon becomes a double-edged sword for the reprogramming process since the retention of epigenetic memory may reduce the quality of pluripotency while increasing the differentiation efficiency into their original tissues. DNA methylation levels are relatively low in the pluripotent stem cells compared to the high levels of DNA methylation seen in somatic cells [101]. Global DNA demethylation is required for the reprogramming process [102]. In the context of these observations, recent work demonstrates that activation-induced cytidine deaminase AID/AICDA contributing to the DNA demethylation can stabilize stem-cell phenotypes by removing epigenetic memory of pluripotent genes. This directly deaminates 5-methylcytosine in concert with base-excision repair to exchange cytosine in genomic DNA [103]. MicroRNA-155 has been identified as a key player for the retention of epigenetic memory during in vitro differentiation of hematopoietic progenitor cell-derived iPSCs toward hematopoietic progenitors [104]. iPSCs that maintained high levels of miR-155 expression tend to differentiate into the original somatic population more efficiently.

Recently, we generated murine skeletal muscle cell-derived iPSCs (myoblast-derived iPSCs) [23] and compared the efficiency of differentiation of myogenic progenitor cells between myoblast-derived iPSCs and fibroblast-derived iPSCs. After EB cultures, more satellite cell/myogenic progenitor cell differentiation occurred in myoblast-derived iPSCs than that in fibroblast-derived-iPSCs (unpublished observation and Figure 3), suggesting that myoblast-derived iPSCs are potential myogenic and satellite cell sources for DMD and other muscular dystrophy therapies (Figure 4). We also noticed that MyoD gene suppression by Oct4 is required for reprogramming in myoblasts to produce iPSCs (Figure 3) [23]. During overexpression of Oct4, Oct4 first binds to the Oct4 consensus sequence located in two MyoD enhancers (a core enhancer and distal regulatory region) [105–107] preceding occupancy at the promoter in myoblasts in order to suppress MyoD gene expression. Interestingly, Oct4 binding to the MyoD core enhancer allows for establishment of a bivalent state in MyoD promoter as a poised state, marked by active (H3K4me3) and repressive (H3K27me3) modifications in fibroblasts, one of the characteristics of stem cells (Figure 3) [23, 108]. It should be investigated whether the similar bivalent state is also established in Oct4-expressing myoblasts during reprogramming process from myoblasts to pluripotent stem cells. It remains to be elucidated whether Oct4-mediated myogenic repression only relies on repression of MyoD expression or is just a general phenomenon of functional antagonism between Oct4 and MyoD on activation of muscle genes. Nevertheless, myoblast-derived iPSCs will enable us to produce an unlimited number of myogenic cells, including satellite cells that could form the basis of novel treatments for DMD and other muscular dystrophies (Figure 4).

Figure 3

Schematic model for chromatin status of myoblast versus fibroblast-derived iPSCs for myogenic induction. In myoblasts, MyoD binds to the two MyoD enhancers (core and DRR) and promoter (PRR), and histone marks show the open chromatin state characteristic. During iPSC reprogramming via expression of Oct4, Sox3, Klf4, and cMyc, exogenous Oct4 binds to both MyoD enhancers which may lead to the bivalent state characteristic of pluripotent stem cells. In fibroblast, both MyoD enhancers and promoter show the closed chromatin state characteristic. During iPSC reprogramming, exogenous Oct4 binds to both MyoD enhancers which may lead to the bivalent state characteristic of pluripotent stem cells. However, myoblast-derived iPSCs may maintain the more open bivalent state characteristic, and thus, myogenic conversion efficiency is increased upon induction.

Figure 4

Myogenic cells induced from myoblast-derived iPSCs for DMD therapy. DMD patient-derived fibroblasts or myoblasts will be reprogramed into iPSCs by reprogramming factors (Oct4, Sox3, Klf4, and cMyc). These fibroblast- and myoblast-derived iPSCs will be induced to myogenic cells via combinatory small molecules and factors such as GSK3βi, forskolin, ascorbic acid, BMPi, ALK5i, Dex, EGF, oncostatin, insulin, bFGF, IGF, HGF, and Wnt3a. These iPSC-derived myogenic cells will be used for autologous cell therapy. Myoblast-derived iPSCs maintain epigenetic myogenic memory.

8. Conclusions

There are pros and cons of transgene-free small molecule-mediated myogenic induction protocols. In the transgene-mediated induction protocols, integration of the transgene in the host genome may lead to risk for insertional mutagenesis. To circumvent this issue, there is an obvious advantage for transgene-free induction protocols. Some key molecules such as Wnt, FGF, and BMP have used signaling pathways to induce myogenic differentiation of ES/iPSCs. However, these molecules are also involved in induction of other types of cell lineages, which makes it difficult for ES/iPSCs to induce pure myogenic cell populations in vitro. By contrast, transgene-mediated myogenic induction is able to dictate desired specific cell lineages. In any case, it is necessary to intensively investigate these myogenic induction protocols for the efficient and safe stem cell therapy for patients.

For skeletal muscle diseases, patient-derived hiPSCs, which possess the ability to differentiate into myogenic progenitor cells followed by myotubes, can be a useful tool for drug screening and personalized medicine in clinical practice. However, there are still limitations for utilizing hiPSC-derived myogenic cells for regenerative medicine. For cell-based transplantation therapies such as a clinical situation, animal-free defined medium is essential for stem cell culture and skeletal muscle cell differentiation. Therefore, such animal-free defined medium needs to be established for optimal myogenic differentiation from hiPSCs. Gene correction in DMD patient iPSCs by TALENs and CRISPR-Cas9 systems are promising therapeutic approaches for stem cell transplantation. However, there are still problems for DNA-editing-mediated stem cell therapy such as safety and efficacy. Since iPSC-derived differentiated myotubes do not proliferate, they are not suited for cell transplantation. Therefore, a proper culture method needs to be established for hiPSCs in order to maintain cells in proliferating the myogenic precursor cell stage in vitro in order to expand cells to large quantities of transplantable cells for DMD and other muscular dystrophies. For other issues, it is essential to establish methods to separate ES/iPSC-derived pure skeletal muscle precursor cells from other cell types for safe stem cell therapy that excludes tumorigenic risks of contamination with undifferentiated cells. In the near future, these obstacles will be taken away for more efficient and safe stem cell therapy for DMD and other muscular dystrophies.

Conflicts of Interest

The authors declare that they have no conflicts of interest.

Acknowledgments

This work was supported by the NIH R01 (1R01AR062142) and NIH R21 (1R21AR070319). The authors thank Conor Burke-Smith and Neeladri Chowdhury for critical reading.

References

K. J. Nowak and K. E. Davies, “Duchenne muscular dystrophy and dystrophin: pathogenesis and opportunities for treatment,” EMBO Reports, vol. 5, no. 9, pp. 872–876, 2004.
View at: Publisher Site | Google Scholar
L. Heslop, J. E. Morgan, and T. A. Partridge, “Evidence for a myogenic stem cell that is exhausted in dystrophic muscle,” Journal of Cell Science, vol. 113, Part 12, pp. 2299–2308, 2000.
View at: Google Scholar
P. Seale, A. Asakura, and M. A. Rudnicki, “The potential of muscle stem cells,” Developmental Cell, vol. 1, no. 3, pp. 333–342, 2001.
View at: Publisher Site | Google Scholar
A. Asakura, P. Seale, A. Girgis-Gabardo, and M. A. Rudnicki, “Myogenic specification of side population cells in skeletal muscle,” The Journal of Cell Biology, vol. 159, no. 1, pp. 123–134, 2002.
View at: Publisher Site | Google Scholar
A. Asakura, M. Komaki, and M. Rudnicki, “Muscle satellite cells are multipotential stem cells that exhibit myogenic, osteogenic, and adipogenic differentiation,” Differentiation, vol. 68, no. 4-5, pp. 245–253, 2001.
View at: Publisher Site | Google Scholar
B. Peault, M. Rudnicki, Y. Torrente et al., “Stem and progenitor cells in skeletal muscle development, maintenance, and therapy,” Molecular Therapy, vol. 15, no. 5, pp. 867–877, 2007.
View at: Publisher Site | Google Scholar
S. Perie, C. Trollet, V. Mouly et al., “Autologous myoblast transplantation for oculopharyngeal muscular dystrophy: a phase I/IIa clinical study,” Molecular Therapy, vol. 22, no. 1, pp. 219–225, 2014.
View at: Publisher Site | Google Scholar
D. Skuk and J. P. Tremblay, “Cell therapy in muscular dystrophies: many promises in mice and dogs, few facts in patients,” Expert Opinion on Biological Therapy, vol. 15, no. 9, pp. 1307–1319, 2015.
View at: Publisher Site | Google Scholar
E. Negroni, A. Bigot, G. S. Butler-Browne, C. Trollet, and V. Mouly, “Cellular therapies for muscular dystrophies: frustrations and clinical successes,” Human Gene Therapy, vol. 27, no. 2, pp. 117–126, 2016.
View at: Publisher Site | Google Scholar
K. Takahashi and S. Yamanaka, “Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors,” Cell, vol. 126, no. 4, pp. 663–676, 2006.
View at: Publisher Site | Google Scholar
K. Takahashi, K. Tanabe, M. Ohnuki et al., “Induction of pluripotent stem cells from adult human fibroblasts by defined factors,” Cell, vol. 131, no. 5, pp. 861–872, 2007.
View at: Publisher Site | Google Scholar
M. Buckingham, L. Bajard, P. Daubas et al., “Myogenic progenitor cells in the mouse embryo are marked by the expression of Pax3/7 genes that regulate their survival and myogenic potential,” Anatomy and Embryology (Berlin), vol. 211, Supplement 1, pp. 51–56, 2006.
View at: Google Scholar
A. E. Munsterberg, J. Kitajewski, D. A. Bumcrot, A. P. McMahon, and A. B. Lassar, “Combinatorial signaling by Sonic hedgehog and Wnt family members induces myogenic bHLH gene expression in the somite,” Genes & Development, vol. 9, no. 23, pp. 2911–2922, 1995.
View at: Publisher Site | Google Scholar
A. Asakura and S. J. Tapscott, “Apoptosis of epaxial myotome in Danforth’s short-tail (Sd) mice in somites that form following notochord degeneration,” Developmental Biology, vol. 203, no. 2, pp. 276–289, 1998.
View at: Publisher Site | Google Scholar
R. Darabi, K. Gehlbach, R. M. Bachoo et al., “Functional skeletal muscle regeneration from differentiating embryonic stem cells,” Nature Medicine, vol. 14, no. 2, pp. 134–143, 2008.
View at: Publisher Site | Google Scholar
K. Kim, A. Doi, B. Wen et al., “Epigenetic memory in induced pluripotent stem cells,” Nature, vol. 467, no. 7313, pp. 285–290, 2010.
View at: Publisher Site | Google Scholar
J. M. Polo, S. Liu, M. E. Figueroa et al., “Cell type of origin influences the molecular and functional properties of mouse induced pluripotent stem cells,” Nature Biotechnology, vol. 28, no. 8, pp. 848–855, 2010.
View at: Publisher Site | Google Scholar
K. Hynes, D. Menicanin, K. Mrozik, S. Gronthos, and P. M. Bartold, “Generation of functional mesenchymal stem cells from different induced pluripotent stem cell lines,” Stem Cells and Development, vol. 23, no. 10, pp. 1084–1096, 2014.
View at: Publisher Site | Google Scholar
M. Quattrocelli, M. Swinnen, G. Giacomazzi et al., “Mesodermal iPSC-derived progenitor cells functionally regenerate cardiac and skeletal muscle,” The Journal of Clinical Investigation, vol. 125, no. 12, pp. 4463–4482, 2015.
View at: Publisher Site | Google Scholar
K. Y. Tan, S. Eminli, S. Hettmer, K. Hochedlinger, and A. J. Wagers, “Efficient generation of iPS cells from skeletal muscle stem cells,” PloS One, vol. 6, no. 10, article e26406, 2011.
View at: Publisher Site | Google Scholar
K. J. Hewitt and J. A. Garlick, “Cellular reprogramming to reset epigenetic signatures,” Molecular Aspects of Medicine, vol. 34, no. 4, pp. 841–848, 2013.
View at: Publisher Site | Google Scholar
T. A. Hore, F. von Meyenn, M. Ravichandran et al., “Retinol and ascorbate drive erasure of epigenetic memory and enhance reprogramming to naive pluripotency by complementary mechanisms,” Proceedings of the National Academy of Sciences of the United States of America, vol. 113, no. 43, pp. 12202–12207, 2016.
View at: Publisher Site | Google Scholar
S. Watanabe, H. Hirai, Y. Asakura et al., “MyoD gene suppression by Oct4 is required for reprogramming in myoblasts to produce induced pluripotent stem cells,” Stem Cells, vol. 29, no. 3, pp. 505–516, 2011.
View at: Publisher Site | Google Scholar
R. B. Chalamalasetty, R. J. Garriock, W. C. Dunty Jr. et al., “Mesogenin 1 is a master regulator of paraxial presomitic mesoderm differentiation,” Development, vol. 141, no. 22, pp. 4285–4297, 2014.
View at: Publisher Site | Google Scholar
C. M. Fan and M. Tessier-Lavigne, “Patterning of mammalian somites by surface ectoderm and notochord: evidence for sclerotome induction by a hedgehog homolog,” Cell, vol. 79, no. 7, pp. 1175–1186, 1994.
View at: Publisher Site | Google Scholar
A. Otto, C. Schmidt, and K. Patel, “Pax3 and Pax7 expression and regulation in the avian embryo,” Anatomy and Embryology (Berlin), vol. 211, no. 4, pp. 293–310, 2006.
View at: Publisher Site | Google Scholar
T. Braun and H. H. Arnold, “Inactivation of Myf-6 and Myf-5 genes in mice leads to alterations in skeletal muscle development,” The EMBO Journal, vol. 14, no. 6, pp. 1176–1186, 1995.
View at: Google Scholar
L. Kassar-Duchossoy, B. Gayraud-Morel, D. Gomes et al., “Mrf4 determines skeletal muscle identity in Myf5:Myod double-mutant mice,” Nature, vol. 431, no. 7007, pp. 466–471, 2004.
View at: Publisher Site | Google Scholar
C. Marcelle, M. R. Stark, and M. Bronner-Fraser, “Coordinate actions of BMPs, Wnts, Shh and Noggin mediate patterning of the dorsal somite,” Development, vol. 124, no. 20, pp. 3955–3963, 1997.
View at: Google Scholar
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.
View at: Publisher Site | Google Scholar
E. Nitzan and C. Kalcheim, “Neural crest and somitic mesoderm as paradigms to investigate cell fate decisions during development,” Development, Growth & Differentiation, vol. 55, no. 1, pp. 60–78, 2013.
View at: Publisher Site | Google Scholar
S. E. Patterson, N. C. Bird, and S. H. Devoto, “BMP regulation of myogenesis in zebrafish,” Developmental Dynamics, vol. 239, no. 3, pp. 806–817, 2010.
View at: Publisher Site | Google Scholar
L. Bajard, F. Relaix, M. Lagha, D. Rocancourt, P. Daubas, and M. E. Buckingham, “A novel genetic hierarchy functions during hypaxial myogenesis: Pax3 directly activates Myf5 in muscle progenitor cells in the limb,” Genes & Development, vol. 20, no. 17, pp. 2450–2464, 2006.
View at: Publisher Site | Google Scholar
E. Dahl, H. Koseki, and R. Balling, “Pax genes and organogenesis,” BioEssays, vol. 19, no. 9, pp. 755–765, 1997.
View at: Publisher Site | Google Scholar
F. Relaix, D. Rocancourt, A. Mansouri, and M. Buckingham, “A Pax3/Pax7-dependent population of skeletal muscle progenitor cells,” Nature, vol. 435, no. 7044, pp. 948–953, 2005.
View at: Publisher Site | Google Scholar
R. L. Davis, H. Weintraub, and A. B. Lassar, “Expression of a single transfected cDNA converts fibroblasts to myoblasts,” Cell, vol. 51, no. 6, pp. 987–1000, 1987.
View at: Publisher Site | Google Scholar
H. Weintraub, S. J. Tapscott, R. L. Davis et al., “Activation of muscle-specific genes in pigment, nerve, fat, liver, and fibroblast cell lines by forced expression of MyoD,” Proceedings of the National Academy of Sciences of the United States of America, vol. 86, no. 14, pp. 5434–5438, 1989.
View at: Google Scholar
H. Weintraub, R. Davis, S. Tapscott et al., “The myoD gene family: nodal point during specification of the muscle cell lineage,” Science, vol. 251, no. 4995, pp. 761–766, 1991.
View at: Publisher Site | Google Scholar
M. A. Rudnicki, T. Braun, S. Hinuma, and R. Jaenisch, “Inactivation of MyoD in mice leads to up-regulation of the myogenic HLH gene Myf-5 and results in apparently normal muscle development,” Cell, vol. 71, no. 3, pp. 383–390, 1992.
View at: Publisher Site | Google Scholar
T. Braun, M. A. Rudnicki, H. H. Arnold, and R. Jaenisch, “Targeted inactivation of the muscle regulatory gene Myf-5 results in abnormal rib development and perinatal death,” Cell, vol. 71, no. 3, pp. 369–382, 1992.
View at: Publisher Site | Google Scholar
Y. Nabeshima, K. Hanaoka, M. Hayasaka et al., “Myogenin gene disruption results in perinatal lethality because of severe muscle defect,” Nature, vol. 364, no. 6437, pp. 532–535, 1993.
View at: Publisher Site | Google Scholar
P. Hasty, A. Bradley, J. H. Morris et al., “Muscle deficiency and neonatal death in mice with a targeted mutation in the myogenin gene,” Nature, vol. 364, no. 6437, pp. 501–506, 1993.
View at: Publisher Site | Google Scholar
L. A. Megeney, B. Kablar, K. Garrett, J. E. Anderson, and M. A. Rudnicki, “MyoD is required for myogenic stem cell function in adult skeletal muscle,” Genes & Development, vol. 10, no. 10, pp. 1173–1183, 1996.
View at: Publisher Site | Google Scholar
L. A. Sabourin, A. Girgis-Gabardo, P. Seale, A. Asakura, and M. A. Rudnicki, “Reduced differentiation potential of primary MyoD−/− myogenic cells derived from adult skeletal muscle,” The Journal of Cell Biology, vol. 144, no. 4, pp. 631–643, 1999.
View at: Publisher Site | Google Scholar
A. Asakura, H. Hirai, B. Kablar et al., “Increased survival of muscle stem cells lacking the MyoD gene after transplantation into regenerating skeletal muscle,” Proceedings of the National Academy of Sciences of the United States of America, vol. 104, no. 42, pp. 16552–16557, 2007.
View at: Google Scholar
M. A. Rudnicki, P. N. Schnegelsberg, R. H. Stead, T. Braun, H. H. Arnold, and R. Jaenisch, “MyoD or Myf-5 is required for the formation of skeletal muscle,” Cell, vol. 75, no. 7, pp. 1351–1359, 1993.
View at: Publisher Site | Google Scholar
M. Maroto, R. Reshef, A. E. Munsterberg, S. Koester, M. Goulding, and A. B. Lassar, “Ectopic Pax-3 activates MyoD and Myf-5 expression in embryonic mesoderm and neural tissue,” Cell, vol. 89, no. 1, pp. 139–148, 1997.
View at: Publisher Site | Google Scholar
S. Tajbakhsh, D. Rocancourt, G. Cossu, and M. Buckingham, “Redefining the genetic hierarchies controlling skeletal myogenesis: Pax-3 and Myf-5 act upstream of MyoD,” Cell, vol. 89, no. 1, pp. 127–138, 1997.
View at: Publisher Site | Google Scholar
P. Seale, J. Ishibashi, A. Scime, and M. A. Rudnicki, “Pax7 is necessary and sufficient for the myogenic specification of CD45+:Sca1+ stem cells from injured muscle,” PLoS Biology, vol. 2, no. 5, article E130, 2004.
View at: Publisher Site | Google Scholar
S. Ozasa, S. Kimura, K. Ito et al., “Efficient conversion of ES cells into myogenic lineage using the gene-inducible system,” Biochemical and Biophysical Research Communications, vol. 357, no. 4, pp. 957–963, 2007.
View at: Publisher Site | Google Scholar
L. Warren, P. D. Manos, T. Ahfeldt et al., “Highly efficient reprogramming to pluripotency and directed differentiation of human cells with synthetic modified mRNA,” Cell Stem Cell, vol. 7, no. 5, pp. 618–630, 2010.
View at: Publisher Site | Google Scholar
A. Tanaka, K. Woltjen, K. Miyake et al., “Efficient and reproducible myogenic differentiation from human iPS cells: prospects for modeling Miyoshi myopathy in vitro,” PloS One, vol. 8, no. 4, article e61540, 2013.
View at: Publisher Site | Google Scholar
R. Abujarour, M. Bennett, B. Valamehr et al., “Myogenic differentiation of muscular dystrophy-specific induced pluripotent stem cells for use in drug discovery,” Stem Cells Translational Medicine, vol. 3, no. 2, pp. 149–160, 2014.
View at: Publisher Site | Google Scholar
S. Albini, P. Coutinho, B. Malecova et al., “Epigenetic reprogramming of human embryonic stem cells into skeletal muscle cells and generation of contractile myospheres,” Cell Reports, vol. 3, no. 3, pp. 661–670, 2013.
View at: Publisher Site | Google Scholar
P. J. Gianakopoulos, V. Mehta, A. Voronova et al., “MyoD directly up-regulates premyogenic mesoderm factors during induction of skeletal myogenesis in stem cells,” The Journal of Biological Chemistry, vol. 286, no. 4, pp. 2517–2525, 2011.
View at: Publisher Site | Google Scholar
E. Shoji, H. Sakurai, T. Nishino et al., “Early pathogenesis of Duchenne muscular dystrophy modelled in patient-derived human induced pluripotent stem cells,” Scientific Reports, vol. 5, p. 12831, 2015.
View at: Google Scholar
H. L. Li, N. Fujimoto, N. Sasakawa et al., “Precise correction of the dystrophin gene in Duchenne muscular dystrophy patient induced pluripotent stem cells by TALEN and CRISPR-Cas9,” Stem Cell Reports, vol. 4, no. 1, pp. 143–154, 2015.
View at: Publisher Site | Google Scholar
R. Darabi, F. N. Santos, A. Filareto et al., “Assessment of the myogenic stem cell compartment following transplantation of Pax3/Pax7-induced embryonic stem cell-derived progenitors,” Stem Cells, vol. 29, no. 5, pp. 777–790, 2011.
View at: Publisher Site | Google Scholar
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.
View at: Publisher Site | Google Scholar
M. Ikeya and S. Takada, “Wnt signaling from the dorsal neural tube is required for the formation of the medial dermomyotome,” Development, vol. 125, no. 24, pp. 4969–4976, 1998.
View at: Google Scholar
U. Borello, B. Berarducci, P. Murphy et al., “The Wnt/beta-catenin pathway regulates Gli-mediated Myf5 expression during somitogenesis,” Development, vol. 133, no. 18, pp. 3723–3732, 2006.
View at: Publisher Site | Google Scholar
S. Tajbakhsh, U. Borello, E. Vivarelli et al., “Differential activation of Myf5 and MyoD by different Wnts in explants of mouse paraxial mesoderm and the later activation of myogenesis in the absence of Myf5,” Development, vol. 125, no. 21, pp. 4155–4162, 1998.
View at: Google Scholar
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.
View at: Google Scholar
N. Itasaki and S. Hoppler, “Crosstalk between Wnt and bone morphogenic protein signaling: a turbulent relationship,” Developmental Dynamics, vol. 239, no. 1, pp. 16–33, 2010.
View at: Publisher Site | Google Scholar
K. Kuroda, S. Kuang, M. M. Taketo, and M. A. Rudnicki, “Canonical Wnt signaling induces BMP-4 to specify slow myofibrogenesis of fetal myoblasts,” Skeletal Muscle, vol. 3, no. 1, p. 5, 2013.
View at: Publisher Site | Google Scholar
R. Reshef, M. Maroto, and A. B. Lassar, “Regulation of dorsal somitic cell fates: BMPs and Noggin control the timing and pattern of myogenic regulator expression,” Genes & Development, vol. 12, no. 3, pp. 290–303, 1998.
View at: Publisher Site | Google Scholar
K. Schuster-Gossler, R. Cordes, and A. Gossler, “Premature myogenic differentiation and depletion of progenitor cells cause severe muscle hypotrophy in Delta1 mutants,” Proceedings of the National Academy of Sciences of the United States of America, vol. 104, no. 2, pp. 537–542, 2007.
View at: Google Scholar
H. Sakurai, Y. Inami, Y. Tamamura, T. Yoshikai, A. Sehara-Fujisawa, and K. Isobe, “Bidirectional induction toward paraxial mesodermal derivatives from mouse ES cells in chemically defined medium,” Stem Cell Research, vol. 3, no. 2-3, pp. 157–169, 2009.
View at: Publisher Site | Google Scholar
Y. Hwang, S. Suk, Y. R. Shih et al., “WNT3A promotes myogenesis of human embryonic stem cells and enhances in vivo engraftment,” Scientific Reports, vol. 4, p. 5916, 2014.
View at: Publisher Site | Google Scholar
C. Xu, M. Tabebordbar, S. Iovino et al., “A zebrafish embryo culture system defines factors that promote vertebrate myogenesis across species,” Cell, vol. 155, no. 4, pp. 909–921, 2013.
View at: Publisher Site | Google Scholar
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, 2005.
View at: Publisher Site | Google Scholar
B. Borchin, J. Chen, and T. Barberi, “Derivation and FACS-mediated purification of PAX3+/PAX7+ skeletal muscle precursors from human pluripotent stem cells,” Stem Cell Reports, vol. 1, no. 6, pp. 620–631, 2013.
View at: Publisher Site | Google Scholar
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.
View at: Publisher Site | Google Scholar
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.
View at: Publisher Site | Google Scholar
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 culture,” Stem Cells Translational Medicine, vol. 3, no. 5, pp. 564–574, 2014.
View at: Publisher Site | Google Scholar
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.
View at: Publisher Site | Google Scholar
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.
View at: Publisher Site | Google Scholar
M. G. Minasi, M. Riminucci, L. De Angelis et al., “The meso-angioblast: a multipotent, self-renewing cell that originates from the dorsal aorta and differentiates into most mesodermal tissues,” Development, vol. 129, no. 11, pp. 2773–2783, 2002.
View at: Google Scholar
M. Sampaolesi, Y. Torrente, A. Innocenzi et al., “Cell therapy of alpha-sarcoglycan null dystrophic mice through intra-arterial delivery of mesoangioblasts,” Science, vol. 301, no. 5632, pp. 487–492, 2003.
View at: Publisher Site | Google Scholar
F. S. Tedesco, M. F. Gerli, L. Perani et al., “Transplantation of genetically corrected human iPSC-derived progenitors in mice with limb-girdle muscular dystrophy,” Science Translational Medicine, vol. 4, no. 140, p. 140ra189, 2012.
View at: Google Scholar
M. F. Gerli, S. M. Maffioletti, Q. Millet, and F. S. Tedesco, “Transplantation of induced pluripotent stem cell-derived mesoangioblast-like myogenic progenitors in mouse models of muscle regeneration,” Journal of Visualized Experiments, no. 83, article e50532, 2014.
View at: Publisher Site | Google Scholar
T. Barberi, L. M. Willis, N. D. Socci, and L. Studer, “Derivation of multipotent mesenchymal precursors from human embryonic stem cells,” PLoS Medicine, vol. 2, no. 6, article e161, 2005.
View at: Publisher Site | Google Scholar
T. Barberi, M. Bradbury, Z. Dincer, G. Panagiotakos, N. D. Socci, and L. Studer, “Derivation of engraftable skeletal myoblasts from human embryonic stem cells,” Nature Medicine, vol. 13, no. 5, pp. 642–648, 2007.
View at: Publisher Site | Google Scholar
K. M. Loh, A. Chen, P. W. Koh et al., “Mapping the pairwise choices leading from pluripotency to human bone, heart, and other mesoderm cell types,” Cell, vol. 166, no. 2, pp. 451–467, 2016.
View at: Publisher Site | Google Scholar
H. Sakurai, Y. Okawa, Y. Inami, N. Nishio, and K. Isobe, “Paraxial mesodermal progenitors derived from mouse embryonic stem cells contribute to muscle regeneration via differentiation into muscle satellite cells,” Stem Cells, vol. 26, no. 7, pp. 1865–1873, 2008.
View at: Publisher Site | Google Scholar
H. Chang, M. Yoshimoto, K. Umeda et al., “Generation of transplantable, functional satellite-like cells from mouse embryonic stem cells,” The FASEB Journal, vol. 23, no. 6, pp. 1907–1919, 2009.
View at: Publisher Site | Google Scholar
Y. Mizuno, H. Chang, K. Umeda et al., “Generation of skeletal muscle stem/progenitor cells from murine induced pluripotent stem cells,” The FASEB Journal, vol. 24, no. 7, pp. 2245–2253, 2010.
View at: Publisher Site | Google Scholar
S. Fukada, S. Higuchi, M. Segawa et al., “Purification and cell-surface marker characterization of quiescent satellite cells from murine skeletal muscle by a novel monoclonal antibody,” Experimental Cell Research, vol. 296, no. 2, pp. 245–255, 2004.
View at: Publisher Site | Google Scholar
R. Jaenisch and A. Bird, “Epigenetic regulation of gene expression: how the genome integrates intrinsic and environmental signals,” Nature Genetics, vol. 33, pp. 245–254, 2003.
View at: Publisher Site | Google Scholar
Q. Hu, A. M. Friedrich, L. V. Johnson, and D. O. Clegg, “Memory in induced pluripotent stem cells: reprogrammed human retinal-pigmented epithelial cells show tendency for spontaneous redifferentiation,” Stem Cells, vol. 28, no. 11, pp. 1981–1991, 2010.
View at: Publisher Site | Google Scholar
M. Quattrocelli, G. Palazzolo, G. Floris et al., “Intrinsic cell memory reinforces myogenic commitment of pericyte-derived iPSCs,” The Journal of Pathology, vol. 223, no. 5, pp. 593–603, 2011.
View at: Publisher Site | Google Scholar
Y. Ohi, H. Qin, C. Hong et al., “Incomplete DNA methylation underlies a transcriptional memory of somatic cells in human iPS cells,” Nature Cell Biology, vol. 13, no. 5, pp. 541–549, 2011.
View at: Publisher Site | Google Scholar
K. Kim, R. Zhao, A. Doi et al., “Donor cell type can influence the epigenome and differentiation potential of human induced pluripotent stem cells,” Nature Biotechnology, vol. 29, no. 12, pp. 1117–1119, 2011.
View at: Publisher Site | Google Scholar
O. Bar-Nur, H. A. Russ, S. Efrat, and N. Benvenisty, “Epigenetic memory and preferential lineage-specific differentiation in induced pluripotent stem cells derived from human pancreatic islet beta cells,” Cell Stem Cell, vol. 9, no. 1, pp. 17–23, 2011.
View at: Publisher Site | Google Scholar
N. Pfaff, N. Lachmann, S. Kohlscheen et al., “Efficient hematopoietic redifferentiation of induced pluripotent stem cells derived from primitive murine bone marrow cells,” Stem Cells and Development, vol. 21, no. 5, pp. 689–701, 2012.
View at: Publisher Site | Google Scholar
R. Rizzi, E. Di Pasquale, P. Portararo et al., “Post-natal cardiomyocytes can generate iPS cells with an enhanced capacity toward cardiomyogenic re-differentation,” Cell Death and Differentiation, vol. 19, no. 7, pp. 1162–1174, 2012.
View at: Publisher Site | Google Scholar
K. Shao, C. Koch, M. K. Gupta et al., “Induced pluripotent mesenchymal stromal cell clones retain donor-derived differences in DNA methylation profiles,” Molecular Therapy, vol. 21, no. 1, pp. 240–250, 2013.
View at: Publisher Site | Google Scholar
D. Sareen, M. Saghizadeh, L. Ornelas et al., “Differentiation of human limbal-derived induced pluripotent stem cells into limbal-like epithelium,” Stem Cells Translational Medicine, vol. 3, no. 9, pp. 1002–1012, 2014.
View at: Publisher Site | Google Scholar
D. Nukaya, K. Minami, R. Hoshikawa, N. Yokoi, and S. Seino, “Preferential gene expression and epigenetic memory of induced pluripotent stem cells derived from mouse pancreas,” Genes to Cells, vol. 20, no. 5, pp. 367–381, 2015.
View at: Publisher Site | Google Scholar
J. Phetfong, A. Supokawej, M. Wattanapanitch, P. Kheolamai, Y. U-Pratya, and S. Issaragrisil, “Cell type of origin influences iPSC generation and differentiation to cells of the hematoendothelial lineage,” Cell and Tissue Research, vol. 365, no. 1, pp. 101–112, 2016.
View at: Publisher Site | Google Scholar
B. Nashun, P. W. Hill, and P. Hajkova, “Reprogramming of cell fate: epigenetic memory and the erasure of memories past,” The EMBO Journal, vol. 34, no. 10, pp. 1296–1308, 2015.
View at: Publisher Site | Google Scholar
P. W. Hill, R. Amouroux, and P. Hajkova, “DNA demethylation, Tet proteins and 5-hydroxymethylcytosine in epigenetic reprogramming: an emerging complex story,” Genomics, vol. 104, no. 5, pp. 324–333, 2014.
View at: Publisher Site | Google Scholar
R. Kumar, L. DiMenna, N. Schrode et al., “AID stabilizes stem-cell phenotype by removing epigenetic memory of pluripotency genes,” Nature, vol. 500, no. 7460, pp. 89–92, 2013.
View at: Publisher Site | Google Scholar
M. Vitaloni, J. Pulecio, J. Bilic, B. Kuebler, L. Laricchia-Robbio, and J. C. Izpisua Belmonte, “MicroRNAs contribute to induced pluripotent stem cell somatic donor memory,” The Journal of Biological Chemistry, vol. 289, no. 4, pp. 2084–2098, 2014.
View at: Publisher Site | Google Scholar
A. Asakura, G. E. Lyons, and S. J. Tapscott, “The regulation of MyoD gene expression: conserved elements mediate expression in embryonic axial muscle,” Developmental Biology, vol. 171, no. 2, pp. 386–398, 1995.
View at: Publisher Site | Google Scholar
D. J. Goldhamer, B. P. Brunk, A. Faerman, A. King, M. Shani, and C. P. Emerson Jr., “Embryonic activation of the myoD gene is regulated by a highly conserved distal control element,” Development, vol. 121, no. 3, pp. 637–649, 1995.
View at: Google Scholar
H. Hirai, M. Verma, S. Watanabe, C. Tastad, Y. Asakura, and A. Asakura, “MyoD regulates apoptosis of myoblasts through microRNA-mediated down-regulation of Pax3,” The Journal of Cell Biology, vol. 191, no. 2, pp. 347–365, 2010.
View at: Publisher Site | Google Scholar
P. C. Taberlay, T. K. Kelly, C. C. Liu et al., “Polycomb-repressed genes have permissive enhancers that initiate reprogramming,” Cell, vol. 147, no. 6, pp. 1283–1294, 2011.
View at: Publisher Site | Google Scholar
I. Dekel, Y. Magal, S. Pearson-White, C. P. Emerson, and M. Shani, “Conditional conversion of ES cells to skeletal muscle by an exogenous MyoD1 gene,” The new Biologist, vol. 4, no. 3, pp. 217–224, 1992.
View at: Google Scholar
J. Rohwedel, V. Horak, M. Hebrok, E. M. Fuchtbauer, and A. M. Wobus, “M-twist expression inhibits mouse embryonic stem cell-derived myogenic differentiation in vitro,” Experimental Cell Research, vol. 220, no. 1, pp. 92–100, 1995.
View at: Publisher Site | Google Scholar
K. Prelle, A. M. Wobus, O. Krebs, W. F. Blum, and E. Wolf, “Overexpression of insulin-like growth factor-II in mouse embryonic stem cells promotes myogenic differentiation,” Biochemical and Biophysical Research Communications, vol. 277, no. 3, pp. 631–638, 2000.
View at: Publisher Site | Google Scholar
A. Myer, E. N. Olson, and W. H. Klein, “MyoD cannot compensate for the absence of myogenin during skeletal muscle differentiation in murine embryonic stem cells,” Developmental Biology, vol. 229, no. 2, pp. 340–350, 2001.
View at: Publisher Site | Google Scholar
V. M. Sumariwalla and W. H. Klein, “Similar myogenic functions for myogenin and MRF4 but not MyoD in differentiated murine embryonic stem cells,” Genesis, vol. 30, no. 4, pp. 239–249, 2001.
View at: Publisher Site | Google Scholar
L. Caron, F. Bost, M. Prot, P. Hofman, and B. Binetruy, “A new role for the oncogenic high-mobility group A2 transcription factor in myogenesis of embryonic stem cells,” Oncogene, vol. 24, no. 41, pp. 6281–6291, 2005.
View at: Publisher Site | Google Scholar
H. Kamochi, M. S. Kurokawa, H. Yoshikawa et al., “Transplantation of myocyte precursors derived from embryonic stem cells transfected with IGFII gene in a mouse model of muscle injury,” Transplantation, vol. 82, no. 4, pp. 516–526, 2006.
View at: Publisher Site | Google Scholar
A. M. Craft, D. M. Krisky, J. B. Wechuck et al., “Herpes simplex virus-mediated expression of Pax3 and MyoD in embryoid bodies results in lineage-related alterations in gene expression profiles,” Stem Cells, vol. 26, no. 12, pp. 3119–3129, 2008.
View at: Publisher Site | Google Scholar
F. Meier-Stiegen, R. Schwanbeck, K. Bernoth et al., “Activated Notch1 target genes during embryonic cell differentiation depend on the cellular context and include lineage determinants and inhibitors,” PloS One, vol. 5, no. 7, article e11481, 2010.
View at: Publisher Site | Google Scholar
M. Iacovino, D. Bosnakovski, H. Fey et al., “Inducible cassette exchange: a rapid and efficient system enabling conditional gene expression in embryonic stem and primary cells,” Stem Cells, vol. 29, no. 10, pp. 1580–1588, 2011.
View at: Publisher Site | Google Scholar
E. C. Thoma, K. Maurus, T. U. Wagner, and M. Schartl, “Parallel differentiation of embryonic stem cells into different cell types by a single gene-based differentiation system,” Cellular Reprogramming, vol. 14, no. 2, pp. 106–111, 2012.
View at: Publisher Site | Google Scholar
S. Goudenege, C. Lebel, N. B. Huot et al., “Myoblasts derived from normal hESCs and dystrophic hiPSCs efficiently fuse with existing muscle fibers following transplantation,” Molecular Therapy, vol. 20, no. 11, pp. 2153–2167, 2012.
View at: Publisher Site | Google Scholar
L. Rao, W. Tang, Y. Wei et al., “Highly efficient derivation of skeletal myotubes from human embryonic stem cells,” Stem Cell Reviews, vol. 8, no. 4, pp. 1109–1119, 2012.
View at: Publisher Site | Google Scholar
T. Yasuno, K. Osafune, H. Sakurai et al., “Functional analysis of iPSC-derived myocytes from a patient with carnitine palmitoyltransferase II deficiency,” Biochemical and Biophysical Research Communications, vol. 448, no. 2, pp. 175–181, 2014.
View at: Publisher Site | Google Scholar
S. Albini and P. L. Puri, “Generation of myospheres from hESCs by epigenetic reprogramming,” Journal of Visualized Experiments, no. 88, article e51243, 2014.
View at: Publisher Site | Google Scholar
S. M. Maffioletti, M. F. Gerli, M. Ragazzi et al., “Efficient derivation 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.
View at: Publisher Site | Google Scholar
J. E. Dixon, G. Osman, G. E. Morris et al., “Highly efficient delivery of functional cargoes by the synergistic effect of GAG binding motifs and cell-penetrating peptides,” Proceedings of the National Academy of Sciences of the United States of America, vol. 113, no. 3, pp. E291–E299, 2016.
View at: Publisher Site | Google Scholar
E. Shoji, K. Woltjen, and H. Sakurai, “Directed myogenic differentiation of human induced pluripotent stem cells,” Methods in Molecular Biology, vol. 1353, pp. 89–99, 2016.
View at: Publisher Site | Google Scholar
T. Akiyama, S. Wakabayashi, A. Soma et al., “Transient ectopic expression of the histone demethylase JMJD3 accelerates the differentiation of human pluripotent stem cells,” Development, vol. 143, no. 20, pp. 3674–3685, 2016.
View at: Publisher Site | Google Scholar
A. Magli, T. Incitti, and R. C. Perlingeiro, “Myogenic progenitors from mouse pluripotent stem cells for muscle regeneration,” Methods in Molecular Biology, vol. 1460, pp. 191–208, 2016.
View at: Google Scholar
Y. ZhuangC. G. Kim, S. Bartelmez, P. Cheng, M. Groudine, and H. Weintraub, “Helix-loop-helix transcription factors E12 and E47 are not essential for skeletal or cardiac myogenesis, erythropoiesis, chondrogenesis, or neurogenesis,” Proceedings of the National Academy of Sciences of the United States of America, vol. 89, no. 24, pp. 12132–12136, 1992.
View at: Google Scholar
J. Dinsmore, J. Ratliff, T. Deacon et al., “Embryonic stem cells differentiated in vitro as a novel source of cells for transplantation,” Cell Transplantation, vol. 5, no. 2, pp. 131–143, 1996.
View at: Publisher Site | Google Scholar
J. Rohwedel, T. Kleppisch, U. Pich et al., “Formation of postsynaptic-like membranes during differentiation of embryonic stem cells in vitro,” Experimental Cell Research, vol. 239, no. 2, pp. 214–225, 1998.
View at: Publisher Site | Google Scholar
T. Sasaki, H. Matsuoka, and M. Saito, “Generation of a multi-layer muscle fiber sheet from mouse ES cells by the spermine action at specific timing and concentration,” Differentiation, vol. 76, no. 10, pp. 1023–1030, 2008.
View at: Publisher Site | Google Scholar
H. F. Teng, Y. L. Kuo, M. R. Loo et al., “Valproic acid enhances Oct4 promoter activity in myogenic cells,” Journal of Cellular Biochemistry, vol. 110, no. 4, pp. 995–1004, 2010.
View at: Publisher Site | Google Scholar
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.
View at: Publisher Site | Google Scholar
H. Sakurai, Y. Sakaguchi, E. Shoji et al., “In vitro modeling of paraxial mesodermal progenitors derived from induced pluripotent stem cells,” PloS One, vol. 7, no. 10, article e47078, 2012.
View at: Publisher Site | Google Scholar
D. Kuraitis, D. Ebadi, P. Zhang et al., “Injected matrix stimulates myogenesis and regeneration of mouse skeletal muscle after ischaemic injury,” European Cells & Materials, vol. 24, pp. 175–195, 2012, discussion 195-176.
View at: Publisher Site | Google Scholar
M. Leung, A. Cooper, S. Jana, C. T. Tsao, T. A. Petrie, and M. Zhang, “Nanofiber-based in vitro system for high myogenic differentiation of human embryonic stem cells,” Biomacromolecules, vol. 14, no. 12, pp. 4207–4216, 2013.
View at: Publisher Site | Google Scholar

Copyright

Copyright © 2017 Yusaku Kodaka 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.

PDF Download Citation

Download other formats

Order printed copies

Views

5332

Downloads

1926

Citations