Fig. 1.1
Methods for derivation of human embryonic stem cells (hESCs) from samples coming from anonymous donations of processes such as in vitro fertilization or the injection of sperm into the egg. The diagram shows the various moments in the embryonic development in vitro in which different groups have derived hESCs lines (Schematic adapted from Ref. [213])
After the discovery that hESCs could be easily isolated from human blastocysts, the scientific community pointed out that one of the main hurdles of blastomere- and ICM-derived hESC for clinical application is that transplantation of their differentiated derivatives might lead to allograft rejection. At that time several strategies were proposed to overcome such impediment, as the establishment of hESC bank containing cell lines covering the majority of human leukocyte antigen (HLA) genotypes. In this regard, tissues differentiated from homozygous hESCs express only one set of histocompatibility antigens, thus, being more readily matched to patients [15]. In addition, homozygous hESCs are routinely derived by parthenogenesis by the artificial activation of unfertilized metaphase II (MII) human oocytes into parthenogenetic ESCs (pESCs) [16–22]. Since all genetic material in parthenotes originates from the maternal genome, the resulting pESCs possess only maternal patterns of gene imprinting, becoming an instrumental platform for the study of the mechanisms regulating maternal epigenetic regulation, as well as to explore disease-related mechanisms. Lately, parthenogenesis has been used to create a pESC line for the common deletion associated with spinal muscular atrophy type 1 [23], paving the way for the generation of pESCs for disease modeling. Also very recently, Sagi and colleagues have generated a collection of hESCs with a normal haploid karyotype from pESC lines originating from haploid oocytes, opening the door to the development of genetic screenings [24].
Somatic cell nuclear transfer (SCNT) consists in the transplantation of the nucleus of a somatic cell into an enucleated oocyte. In this regard, SCNT-derived ESCs (NT-ESCs) are genetically autologous to the nuclear donor somatic cell, offering great potential in regenerative medicine, including disease modeling and cell replacement therapy. NT-ESCs were reported first in mice [25] and later in primates [26]. Recently, Mitalipov group has shown that it is possible to generate hESCs via SCNT. In their work, differentiated fetal and infant fibroblasts were used as nuclear donors [27]. More recently others have demonstrated that age-associated changes in the nucleus donor cell do not hamper NT-hESC derivation [28], and that is also possible to generate insulin-producing beta cells from NT-ESCs from a patient with long-standing diabetes [29]. These last findings pinpoint SCNT as a suitable platform for the generation of autologous cells for clinical purposes.
1.1.1.2 Induced Pluripotent Stem Cells (iPSCs)
The “reprogramming history” started in 1958, when Gurdon et al. [30] by using the technique of SCNT, originally described by Briggs and King [31], showed that the nuclei of intestinal epithelial cells from Xenopus laevis, after transplantation into enucleated eggs, could develop into normal and healthy tadpoles, thus demonstrating successful nuclear reprogramming. Taken together, these first advances pointed out that the process of cell differentiation could be reversible and did not require irreversible nuclear changes. One of the most important advances in this field of research was the publication by Wilmut et al. in 1998 of the birth of a cloned sheep (Dolly) by transplanting the nucleus of an adult somatic mammary gland cell into an enucleated oocyte [32]. In the last 15 years, progress has been made producing “clones” for reproductive purposes in several species—cattle, goats, mice, and pigs [33–39]––culminating this period with the creation of the first cloned human embryo in 2013 by Mitalipov and colleagues [27].
In 1987 Schneuwly group found that in Drosophila melanogaster, the overexpression of certain transcription factors in somatic cells could activate the expression of genes arising from another cell type [40]. This group together with others also found similar results in mammals [41]. Pursuing the idea of changing cell fate and inducing dedifferentiation, Takahashi and Yamanaka in 2006 discovered that the pluripotent state could be artificially induced in somatic cell types through the overexpression of just four transcription factors (OCT4, SOX2, c-Myc, and KLF4-OSKM) [42–45]. The produced cells were called induced pluripotent stem cells (iPSCs), and they exhibited all the molecular and functional features of hESCs.
While, at first, somatic reprogramming was described using mouse embryonic fibroblasts, the Japanese team could show that also a reduced formula of the original “Yamanaka cocktail” could be used to reprogram human somatic cells toward human iPSCs (hiPSCs) [46]. Since 2007 several aspects have been considered when identifying the best cell source to be reprogrammed for regenerative medicine approaches. This also has conditioned the number of Yamanaka transcription factors used in each specific case (i.e., progenitor cells expressing endogenously any of the Yamanaka factors can be reprogrammed in the absence of it, as neural stem cells in the absence of SOX2), the strategy for Yamanaka factor delivery (i.e., proliferating cells can be easily transduced with retroviral vectors for reprogramming, as fibroblasts, keratinocytes, among others), and the cell type used (i.e., cell amenability has sometimes limited reprogramming applications, as for neural stem cells or intestinal cells, among others). Besides all these factors, up to day a huge variety of somatic cell types, which included fibroblasts, blood, keratinocytes, liver and gastrointestinal cells, as well as cancer cells, can be used to derive iPSCs [42, 46–56].
Interestingly, during the last years, the generation of protocols avoiding the use of lentiviral or retroviral vectors for the expression of Yamanaka factors has involved the definition of novel strategies for hiPSC generation, including the use of recombinant proteins [57, 58], episomal vectors [59], or mRNAs [60, 61], among others [62]. Thus, the generation of hiPSCs, especially the generation of patient-derived iPSCs suitable for disease modeling in vitro, opens the door for the potential translation of patient-derived iPSCs into the clinic. Successful replacement or augmentation of the function of damaged cells by patient-derived differentiated stem cells would provide a novel cell-based therapy for skeletal muscle-related diseases.
Integrative Methods for Cell Reprogramming
The first generation of iPSCs was accomplished by retroviral-mediated ectopic expression of “OSKM or Yamanaka factors” into mouse fibroblasts [42]. This method has been successfully used for several cell types, such as mouse and human fibroblasts, neural stem cells, keratinocytes, adipose cells, liver cells, and blood cells, with efficiencies of reprogramming between 0.01 and 0.02% [46]. An alternative approach to transduce OSKM factors to derive iPSCs is the use of a lentiviral system which yields a higher efficiency (0.1–2%) than retroviral transduction [62]. Both platforms have been intensively used during the first years of the reprogramming decade; however, the disadvantage of viral integration into the host genome, together with the use of oncogenic factors as KLF4 or c-Myc, bound the application of iPSCs for clinical purposes [62–64].
Since viral integration can cause insertional mutagenesis, interference with gene transcription, and genome instability and induce malignant transformation [65–68], several non-integrating virus-mediated iPSC reprogramming methods have been currently established [62, 64, 69]. One example is the use of doxycycline (dox)-inducible lentiviral vector harboring OSKM factors flanked by LoxP sites that can be subsequently excised by the use of Cre recombinase [70]. Also, replication-defective adenoviral vectors expressing OSKM factors have proven useful for derivation of iPSCs because they do not integrate into chromosomal DNA [71, 72]. Adenoviral vectors have been mainly used to generate iPSCs from liver cells and fibroblasts without viral integration [69, 73]. While the non-integrating aspect of the adenoviral method is appealing, to be of significant use in translational medicine, optimization improving reprogramming efficiency is necessary [74].
Non-integrative Methods for Reprogramming
Lately, different laboratories have made use of episomal plasmids as another method for integration-free reprogramming of somatic cells into iPSCs [75, 76]. This procedure has also been used to derive iPSCs from cord blood and peripheral blood cells [77]. This technique yields a very low efficiency, but several modifications by different groups provide promising results for future use [75, 76, 78]. Interesting minicircle DNA vectors containing Lin28, Nanog, SOX2, and OCT4 factors have been described as a procedure to derive human iPSCs from human adipocytes with an efficiency of 0.005% [79].
Other approaches relay in the use of the single-stranded RNA Sendai virus (SV); this method allows for the generation of iPSCs with an efficiency around 0.1%, comparable to the lentiviral approach while avoiding transgene integration [80]. Similarly mRNA transfection has been proved as another appealing system for the generation of iPSCs avoiding transgene integration [81]. RNA-induced pluripotent stem cell procedures offer a safe and effective method to generate “safe iPSCs” providing a reduced immunogenic response. Using this method, Warren et al. [60] derived iPSCs from human keratinocytes, human neonatal fibroblasts, human fetal lung fibroblasts, and cystic fibrosis patient fibroblasts with conversion efficiencies and kinetics substantially superior to established viral protocols (around 2%).
Other strategies such as the use of bioactive OSKM proteins have also been tested for iPSC generation [57]. In this regard, Kim et al. demonstrated the successful generation of stable iPSCs from human fibroblasts by direct delivery of four reprogramming protein factors (OSKM) yielding an efficiency of 0.001% [58]. A major challenge of this procedure, however, stands in the efficient delivery of OSKM proteins [82]. In this regard others have shown the possibility to fuse OSKM proteins with a short basic segment with a high proportion of amino acids, namely, cell-penetrating peptide (CPP) [83, 84]. CPP-OSKM proteins, when delivered into somatic cells, can directly reprogram them successfully without genetic manipulation and/or chemical treatments [57, 58]. Nevertheless, bioactive reprogramming proteins are difficult to synthesize in large quantities, and reprogramming efficiencies by this method vary between 0.001 and 4%.
Cellular reprogramming using small molecules offers many advantages such as temporally and spatially manageable, reversible, cell permeability, and cost-effectiveness. Small molecules used to generate iPSCs are comprised of epigenetic modifiers, WNT signal modulators, cell senescence attenuators, metabolism modulators, and regulators of cell apoptosis/senescence pathways. Small molecules inducing iPSCs can be classified into three types: (1) small molecules that improve reprogramming efficiency [85], (2) compounds replacing one or more reprogramming factors [86–88], and (3) combinations of compounds that suffice for reprogramming [89, 90]. Small molecule methods have been successfully applied to reprogram mouse and human fibroblasts directly into iPSCs [62, 89–93].
1.2 General Approaches to Induce In Vitro Differentiation of Pluripotent Stem Cells (PSCs)
Both mouse and human PSCs are routinely cultivated in the presence of feeder layers (Fig. 1.2a). Initial studies made use of mouse embryonic fibroblasts mitotically inactivated as feeder cells in the presence of embryonic stem cell media for preserving hPSCs undifferentiated in culture. For mouse PSCs, LIF can substitute for feeder layers. However, since LIF is not needed for human PSC culture, in the last years, different chemically defined media have been produced in order to sustain human PSC culture and expansion in feeder-free substrates. PSCs grow on the feeder layers as colonies (Fig. 1.2b). Generally, human and mouse PSCs are enzymatically dissociated with different reagents as trypsin, acutase, or dispase; the obtained suspension of single cells is then transferred for subculture and expansion for differentiation purposes as guided differentiation, among others.
Fig. 1.2
Culture and propagation of human pluripotent stem cells (PSCs). (a) hPSCs can be cultured on top of irradiated human fibroblasts and grow as tight colonies that are manually expanded. (b) Lately, the culture of hPSCs is easily performed using defined matrices and medium sustaining pluripotency. hPSCs are routinely expanded by enzymatic methods allowing for the standardization of differentiation protocols worldwide. Scale bars 100 μm
As an option for culturing human PSCs without feeder cells, Matrigel™ has proven to be a useful alternative enabling the stable culture of human PSCs. Moreover, others and we have also shown that Matrigel™ allows the generation of hiPSCs for disease modeling purposes without animal-derived feeder cells [94]. Since Matrigel™ was derived from Engelbreth–Holm–Swarm mouse sarcoma cells [95], other types of matrices which do not contain animal-derived agents have been tested and used as feeder cell substitutes for the successful maintenance and generation of human PSCs, such as CellStart [96, 97], recombinant proteins [98–100], and synthetic polymers [101, 102].
The culture media used in the early generation of hESCs contained fetal bovine serum [4]. In order to remove unspecific agents that might cause spontaneous differentiation of hESCs, knockout serum replacement (KSR) has now been established as a defined material for maintaining hESCs [103] and is also traditionally used for hiPSC generation [46, 104–106]. In this regard, mTeSR1 medium was developed as a chemically defined medium for maintaining human PSCs [107]. Importantly, in the last years, several authors have reported the generation of commercially developed xeno-free media for maintaining hiPSCs, and such media have already been used successfully for iPSC generation. These media include TeSR2 [108], NutriStem [109], Essential E8 [99], and StemFit [110].
When factors that sustain PSC stemness are deprived from the media, PSCs spontaneously differentiate into derivatives of the three embryonic germ layers. This capacity has been profited for more than 30 years in order to direct PSCs to the desired cell product. In this regard, up to day, an infinite number of protocols have been established to promote the development of the cell type of interest.
The following are basic strategies to induce in vitro differentiation of PSCs:
- (a)
Embryoid bodies’ (EBs) formation: EBs are spherical structures that allow PSC culture in suspension when using nonadherent culture substrates (Fig. 1.3a). EBs can be induced from PSCs grown as monolayers by mechanical or enzymatic procedures. Interestingly, within the first 3 days of differentiation, PSCs propagated as EBs form three germ layers. The three-dimensional structure, including the establishment of complex cell adhesions and paracrine signaling within the EB microenvironment, enables differentiation and morphogenesis. The presence of ectoderm is manifested by the expression of fibroblast growth factor 5 (FGF5), endoderm by GATA-4, and mesoderm by Brachyury [111]. For all these reasons, the first protocols for muscle differentiation took advantage of EB induction, including those describing derivation of the first myogenic cells from mESCs and iPSCs [112, 113] and hESCs [114]. Although those first assays proved the feasibility of mouse and human PSCs to give rise to myogenic-like cells, lately, different works have proved the possibility to avoid the use of fetal bovine and/or horse serum in order to reduce potential contaminations of animal components by including serum-free-based media during the time course of differentiation.
Fig. 1.3
hPSC differentiation following embryoid body formation (EB). hPSCs are able to differentiate toward the three germ layers of the embryo. (a) The generation of EBs from hPSCs has been widely use in order to generate cells with myogenic potential (b) After several days grown in suspension, EBs are then transferred onto supporting cells (feeder cells) sustaining for myogenic differentiation. Scale bars 100 μm
Besides all the advances when using EB methodology for the derivation of muscle-like cells, their culture is laborious and time-consuming, limiting the control of growth factors/cytokines in a 3D setting. These limitations preclude the use of EB-based methods for the generation of PSC-derived muscle cells in a therapeutic setting, where large amounts of patient-derived muscle cells would need to be derived. Still EB-based methods may offer advantages when used as an intermediate step for the generation of myogenic cells from PSCs. In this regard, Hwang and coworkers have recently shown that cells differentiated as EBs and sorted for PDGF-αR expression could be successfully cultured in monolayer retaining the ability to undergo terminal myogenic differentiation despite culture pressure [115].
- (b)
Guiding muscle differentiation modifying medium composition: Traditionally monolayers of PSCs and/or EBs have been used as starting cellular populations to differentiate into specific lineages by mimicking developmental programs guiding tissue specification. Majorly, PSCs (grown as monolayers or EBs) have been subjected to changes in medium composition in order to induce their differentiation toward the desired cell type. With respect to myogenic differentiation, PSCs from mouse and human have been differentiated toward different stages of myogenic differentiation, i.e., paraxial mesodermal cells, muscle progenitor cells, satellite cells, myoblasts, and myotubes. In this regard myogenic cells at initial stages of differentiation (those expressing Pax3 and/or Pax7) were shown to be characterized by higher regenerative potential than cells that reached more advanced stages of differentiation and expressed myogenic transcription factors [115–119]. Although these works used serum and cell culture media with animal-derived components, they set the basis for the definition of serum-free protocols for myogenic differentiation. In general, the use of such specific cell media together with the control of the expression of myogenic transcription factors crucial for muscle determination and differentiation has demonstrated promising results when differentiating mouse or human PSCs toward myogenic cells (i.e., control of MyoD1 expression under the control of promoters responsive for tamoxifen/puromycin treatment).
- (c)
Genetic manipulation of PSCs: For a long time, PSCs have represented an unprecedented platform for controlling the expression of transcription factors aiming to direct the differentiation of PSCs toward the lineage of choice. PSCs can be kept in culture in the absence of feeders and expanded as single cells, favoring different manipulations such as electroporation and nucleofection, methods generally used when performing PSC genetic manipulation. Whereas the first studies for the generation of myogenic-like cells from mouse or human PSCs took advantage of integrative gene expression systems (i.e., lentivirus or retrovirus), nowadays the use of these tools is limited, since they incur uncertain risks for potential cell-based therapeutic applications [120]. In this regard, the use of excisable vectors (i.e., transposons [121, 122] or mRNAs [60]) offers an unprecedented opportunity for the derivation of differentiated PSCs suitable for regenerative medicine. Moreover, the recent discovery of DNA meganucleases, TAL effector nucleases, or clustered regularly interspaced short palindromic repeats (CRISPR) will offer the possibility to target specific loci determinant for muscle differentiation with fluorescent reporters leading to the definition of robust protocols of PSC differentiation.
- (d)
Coculture with supportive cells (feeder cells): Generally the coculture of mouse and human PSCs (either as monolayers or EBs) together with feeder cells has been used to induce PSC differentiation [123] (Fig. 1.3b). Different feeders have proven to commit PSC differentiation toward different lineages. In the context of muscle differentiation, Baghavati and coworkers showed that the coculture of EBs derived from mESCs together with primary muscle cells suffice for myogenic differentiation, since donor-derived myofibers could be occasionally found on the surface of the host muscle [124].
- (e)
Extracellular matrix (ECM) as an instructive scaffold for PSC differentiation: Extracellular matrix (ECM) is a dynamic and complex environment characterized by tissue-specific biophysical, mechanical, and biochemical properties. Different works have shown that muscle microenvironment (niche) enables freshly isolated muscle stem cells (MSCs) to contribute extensively to skeletal muscle regeneration when transplanted in dystrophic mice (i.e., mdx model, among others). On the contrary when MSCs are grown on standard conditions (i.e., plastic substrate) for several passages, they lose their “stemness” leading to progenitors with diminished regenerative potential [125, 126]. It has been also described that muscle regeneration in higher vertebrates depends on the capacity of the injured tissue for retaining ECM scaffolding, which serves as a template for the de novo formation of muscle fibers [127]. In this regard, the interaction between PSCs and ECM via integrins determines the expression of signaling molecules that affect PSC differentiation [123]. Of note, myogenesis (i.e., proliferation of myoblasts and further fusion into myotubes) has been positively induced when mouse iPSCs have been cultured in the presence of Matrigel™ [128]. Similar results have been observed when using collagen-based matrix for the differentiation of human iPSCs expressing a dox-inducible expression cassette of MyoD1 [129]. In order to control the organization and alignment of muscle fibers, both the composition of the ECM and its anisotropic architecture are essential. Self-organized myotubes have been generated by using topography-based approaches based on nanofibers [130], microabrated surfaces [131], and microcontact printing of ECM proteins [132, 133]. In a complementary approach, biochemical cues have also been introduced in order to promote cell alignment and differentiation. By using inkjet bioprinting, spatially defined patterns of myogenic and osteogenic cells were derived from primary MSCs as a response to growth factor patterning [134]. In order to mimic native tissue organization, topographical and biochemical signaling has also been explored [135]. The vast majority of these works present cells to static microenvironments. Latest trends point out the relevance of presenting cells to spatially and temporally dynamic microenvironments [136]. Surfaces with gradient concentrations of growth factors (BMP-2 and BMP-7) have shown to successfully drive cell differentiation [137, 138]. Overall, these strategies appear a promising way to direct the differentiation of PSCs [139].
Tissue engineering strategies are intended to provide synthetic and natural 3D scaffold materials to mimic the structural, biochemical, and mechanical properties of the stem cell niche [140, 141]. Natural scaffolds based on ECM proteins have been used to form hydrogels for musculoskeletal tissue engineering [142–144]. Commercially available ECM substitutes such as Matrigel™ hydrogels are also showing promising results in the differentiation of PSCs toward cardiomyocytes [145]. Lately, technologies such as electrospinning, which allows organizing the polymers into thin sheets of fibrous meshes, are promising in this field [146, 147]. The use of acellular tissue scaffolds is also being explored in muscle regeneration since they offer a native ECM with the optimal biochemical and mechanical properties for MSC culture preserving the architectural features of the tissue.
- (f)
The use of microfluidics for PSC differentiation: PSC differentiation is affected by chemical, topographic, and mechanical effects and conventional culture methods. Microfluidic culture platforms have shown to accurately mimic physiological conditions for stem cell growth [148].This emerging technology offers the possibility to (1) manipulate the environment controlling oxygen supply, pH, temperature, flow shear stress [149], material shear, topography, and stiffness [150, 151] and surface properties [149]; (2) identify, separate, and position desired cell types [152]; (3) stimulate cells through mechanical stretching [153] or electrically [154]; (4) develop screening of several parameters [155]; (5) apply gradients of chemical and soluble factors [156, 157]; (6) control fluid mixing through compartmentalized devices [158]; and (7) include sensors [159, 160]. Recently, Uzel and coworkers have shown that mechanical or electrical stimuli facilitate the differentiation of stem cells into myocytes [161]. Biochemical stimuli include the presence of several factors on the cell culture, as skeletal muscle differentiation factors promoting differentiation [162]. Mechanical stimuli are needed to induce desired interactions with cells or matrix, especially for muscle fibers. Passive mechanical stimuli, such as mechano-topographical cues [163], scaffold structure orientation [163, 164], and substrate stiffness or elasticity [165], have proved effect on myogenic differentiation. Active mechanical stimuli include stretching or forcing cells or the entire microfluidic chip [153, 166]. This stretch can be uniaxial or equiaxial, having different effects on stem cell differentiation, as reviewed by Watt and Huck [167].
Besides all these findings, differentiation toward myocytes is not enough to achieve physiologically relevant 3D models with fascicle-like sarcomere structure capable of contraction with uniform distribution of oxygen and nutrients or cell alignment. Several approaches have been developed on these regard, mostly trying to promote cell alignment, that include among others (1) the use of parallel linear microgroves [168, 169] or ECM molecule micropatterns on the surface [170, 171] in order to facilitate cell alignment; (2) the employment of microchannels for chemical delivery [172] or for 3D constructs of the skeletal muscle filled with hydrogels [173]; (3) the use of anchoring points for the ECM with Velcro anchors [174], tendon-like anchors [171], or steel mesh to induce cell alignment through a stretching freestanding construct; (4) and the use of capped pillar-based constructs to encourage freestanding muscle alignment and maturation through a controlled stress, enabling measurement of forces [175–178]. Despite all these improvements, self-aggregation of myoblasts happens frequently. Some studies developed by the group of Professor Asada [179, 180] have reported 3D fascicle-like muscle-on-a-chip devices without self-aggregation of cells, creating sarcomeric structures capable of contraction, with uniform distribution of oxygen and nutrients, spontaneous aligning stress, cell alignment along transmission axis encouraged by uniform tension, fibers with high length to diameter ratio, high cell density, and overall good mimic of motor units. Very recently, an integrated tissue–organ bioprinting procedure has been reported, which can fabricate stable, human-scale tissue constructs of any shape, such as the skeletal muscle [181]. In order to study the skeletal muscle in a biological and physiological context, it is necessary to include its interaction with motor neurons. Neuromuscular junction on a chip includes, mainly, the following three approaches: 3D coculture of neurospheres and muscle fibers [178], 3D coculture of motor neurons and muscle fibers [182], and the use of compartmentalized microfluidic chips with chambers and microchannels [183–185].
1.3 Generating Myogenic Cells from Mouse and Human PSCs
Skeletal muscles in higher organisms originate from different areas of the embryonic mesoderm [186]. Head muscles derive from the unsegmented cranial paraxial mesoderm. In turn, muscles of the trunk and limbs arise in two subsequent stages from the dorsal part of the segmented paraxial mesoderm, commonly referred to as dermomyotome. In a first stage, postmitotic myogenic precursors delaminate from the borders of the dermomyotome and migrate ventrally to form the primary myotome [187]. This primary myotome serves as a scaffold for the second stage of myogenesis but also secretes factors that trigger an epithelial-to-mesenchymal transition (EMT) among muscle progenitors in the central dermomyotome that eventually migrate into the myotome [187]. This secondary migration of EMT-derived precursors from the dermomyotome also generates the satellite cells (SCs), the adult stem cell pool in the skeletal muscle, which are responsible for postnatal muscle maintenance, repair, and growth.
Over the last decades, the understanding of the transcription factors and intrinsic and extrinsic signals that govern SCs or terminally differentiated myogenic cells has represented a good starting point for the definition of protocols for the generation of myogenic cells from PSCs (both from mouse and human ESCs/iPSCs). In the same manner, the generation of patient-derived cell platforms can help us to develop experimental strategies toward generating muscle stem cells, either by differentiating patient-specific iPSCs or by converting patient’s somatic cells toward myogenic cells (transdifferentiation). Overall, the possibility to generate disease-free patient iPSCs can help us to identify which are the mechanisms driving muscle disease and, more importantly, to develop new compounds for treating MDs.
1.3.1 Exogenous Expression of Muscle-Related Transcription Factors in PSCs: How to Generate Myogenic Precursors and/or Terminally Differentiated Multinucleated Myogenic Cells
The use of autologous derived muscle stem cells for restoring muscle function has been envisioned as a powerful therapeutic strategy for muscle degenerative diseases. Successful generation of myogenic precursors from mouse and human iPSCs has been achieved by exogenous expression of transcription factors crucial for myogenic differentiation. Since PSCs are an expandable source amenable for genome editing (i.e., they can undergo extensive tissue culture manipulations, such as drug selection and clonal expansion, while still maintaining, e.g., their pluripotency signature and genome stability), latest advances in this field will increase our knowledge in PSC differentiation toward skeletal muscle lineage. Early studies in the field have relayed in the use of viral vectors for the generation of stable PSC lines expressing the myogenic transcription factor of interest under the control of specific drugs (i.e., Pax7 or MyoD1, Magic F-1, among others). Transduced PSCs are then subsequently exposed to culture media conditions promoting muscle differentiation. Other methods involve the use of non-integrative vectors such as adenovirus, transposons, or excisable lentiviral vectors in order to avoid undesirable effects when working with integrative systems (i.e., retrovirus or lentivirus). Following these different approaches, several studies have shown that PSC monolayers or PSC-derived EBs could be converted with different efficiencies into myogenic-like cells (see below).
Early Studies of Myogenic Differentiation from mESCs
Dekel and colleagues described the first protocol describing the generation of skeletal muscle cells from mESCs early in 1992. In their hands when mESCs were electroporated with MyoD1 cDNA driven by the β-actin promoter, some cells could be converted to skeletal muscle cells [188]. Although myogenesis was associated with the activation of MRF4 and Myf5 genes, the transient expression of MyoD1 did not lead to the efficient conversion of mESCs toward skeletal muscle cells. However, authors showed that contracting skeletal muscle fibers could be generated when the transfected cells were allowed to differentiate in vitro after EB formation in the presence of low-mitogen-containing medium. After that first work, other authors provided fine-tuned systems aiming to control the expression of the myogenic factor of choice at a precise moment during the onset of myogenic differentiation. Alongside this line, Ozasa and colleagues [189] established a mESC line by introducing a MyoD transgene controlled by a Tet-Off system (ZHTc6-MyoD). Under those conditions and only after 7 days, primed cells started to fuse into myotubes, and occasionally light muscle contractions were recorded. Intramuscular injections of MyoD–mESC-derived cells into mdx resulted in the generation of clusters of dystrophin-positive myofibers in the injected area.
Myogenic Differentiation from Human PSCs
Within the last years, different research groups have demonstrated the possibility to generate myocytes and even multinuclear myotubes from both hESCs and patient-derived hiPSCs. Already in 2012 two different reports indicated that after MyoD overexpression, mesodermal [190] or mesenchymal cells [191] could be generated from iPSCs. Similarly, Rao and colleagues (2012) generated a transgenic Tet-inducible MyoD cassette in which all the transgenic elements were inserted in hESCs making use of lentiviral vectors. Later on, Yasuno and colleagues [122] generated terminal multinucleated cells from iPSCs derived from patients affected with carnitine palmitoyltransferase II (CPT II) by the transduction of a self-contained Tet-inducible MyoD1 expressing piggyBac vector (Tet–MyoD1 vector) and transposase into hiPSCs by lipofection. This system allowed the indirect monitoring of MyoD cells in response to doxycycline by co-expression of a red fluorescent protein (mCherry). Moreover, authors increased the purity of the generated myocytes by culturing the cells in low glucose conditions [192]. Also Abujarour and colleagues [129] found that it is possible to derive myotubes from control iPSC and iPSC lines from patients with either Duchenne or Becker muscular dystrophies using a lentiviral system expressing MyoD under the control of a Tet-inducible promoter.
Other factors apart from MyoD1 have been used to promote myogenic differentiation from hPSCs. In this regard, Iacovino and colleagues [193] integrated one single copy of Myf5 into mESCs and hESCs by means of a system that authors called inducible cassette exchange (ICE). Overall, Iacovino and colleagues showed that Myf5 expression is sufficient to promote the myogenic commitment of nascent mesoderm, thereby establishing a novel and rapid method of differentiating mESCs and hESCs into skeletal muscle tissue. Interestingly, Darabi and colleagues generated an improved version of ICE system in order to generate mESCs in which Pax7 expression was controlled under the control of doxycycline [194, 195]. Later on, the same group generated inducible Pax7 hPSCs by means of a doxycycline-inducible lentiviral vector encoding Pax7 incorporating an IRES–GFP reporter allowing for the monitoring of transplanted Pax7-derived myogenic progenitors into dystrophin-deficient mice (mdx). Interestingly, authors could show that after transplantation the differentiated cells led to long-term muscle regeneration [196].
1.3.2 Generation of Myogenic Precursors and/or Terminally Differentiated Multinucleated Myogenic Cells by Soluble Factors
The exogenous expression of muscle-specific transcription factors in PSCs by the methodologies described above has proved to be successful strategies to direct muscle differentiation. Although valuable, those strategies could not be applied in the context of clinics to treat compromised skeletal muscle tissues. For this reason, in the past years, many efforts have been also directed to the definition of specific culture media and conditions to produce myogenic precursor cells. Several groups have investigated the possibility to expose EBs or monolayers of mouse and human PSCs to stage-specific differentiation protocols based on the addition of soluble factors known to be crucial during embryonic myogenesis. Following such protocols authors have been able to derive different cell populations with myogenic potential (i.e., paraxial mesoderm) that could be further isolated using FACS-based selection strategies. In this manner, authors could evaluate the myogenic differentiation yield by quantifying the percentage of cells expressing specific myogenic markers. In the same manner, these works have characterized the myogenic differentiation process by analyzing the expression of myogenic-related markers by common techniques such as polymerase chain reaction or immunohistochemistry
Early Studies in Myogenic Differentiation from Mouse PSCs by Soluble Factors
mPSCs propagated as EBs are known to form the three germ layers within the first 3 days of in vitro differentiation in undefined culture media. However, transplantation of EBs without any induction to direct development along a specific pathway leads to a failure of integration into recipient tissues and often forms teratomas. Thus, successful derivation of myogenic cells from PSCs requires selective induction of the myogenic lineage in PSCs. In a pioneering study by Rohwedel and coworkers, the expression of myogenic-related factors (i.e., Myf5, MyoD, and myogenin) was identified in 7-day-old outgrowths obtained from EBs formed by differentiating mouse ESCs [112]. The EB system was also used in one of the first studies that addressed the myogenic differentiation potential of miPSC [197], in which Pax3 and Pax7 expression was followed by the expression of myogenic markers such as Myf5, MyoD, and myogenin, similarly as is observed during embryonic myogenesis. In an attempt to enhance the myogenic conversion of PSCs, Bhagavati and Xu [124] described the coculture of EBs with freshly isolated muscle cells as a novel method for myogenic differentiation. Although authors showed that differentiated cells generated by this method developed vascularized and muscle tissue when transplanted in dystrophic mice (mdx mice), still the number of engrafted cells was too low [124]. Others described that the temporarily supplementation of culture medium with retinoic acid [198] or ascorbic acid and activin A [199] could improve myogenic differentiation from mESC. Although these initial studies involving EBs and coculture methodologies yielded important information, they resulted to be rather inefficient and often used serum-containing medium hampering the experimental reproducibility and their further translation into the clinics, due to the presence of undefined factors in the medium. In this regard, many efforts have been directed to the development of defined culture conditions. Sakurai and colleagues [200] differentiated a mESC line toward paraxial mesodermal progenitors. Specifically, authors selected paraxial mesodermal progenitors based on the expression of platelet-derived growth factor receptor-α (PDGFR-α) and the absence of Flk-1—a lateral mesodermal marker. Later on, the same authors demonstrated that mESCs could be directed toward the paraxial mesodermal lineage by a combination of bone morphogenetic protein (BMP) and Wnt signaling under chemically defined conditions [201].
Generation of Myogenic Cells from Human PSCs by Soluble Factors
Myogenic differentiation from hPSCs forming EBs was also achieved by allowing the differentiation of cell outgrowths from human EBs exposed to medium supplemented with ITS (i.e., insulin, transferrin, selenium), dexamethasone, and epidermal growth factor (EGF) or to medium supplemented with horse serum [114]. In this manner, myogenic markers could be detected 2 and 4 weeks after EB plating. Interestingly, the treatment with the hemimethylating agent, 5-azacytidine for 24 h, led to significant increase in the number of cells expressing myogenic markers [114]. However, in vitro formation of myotubes could not be seen under none of these culture conditions. In contrast, when those hESC-derived myogenic precursors were transplanted in NOD-SCID mice, they could incorporate into the host muscle and became part of regenerating muscle fibers [114].
Given that the EB culture system is a laborious and time-consuming method that does not allow for generation of large quantities of differentiated cells for therapeutic purposes, researchers have developed alternative myogenic differentiation protocols by omitting the EB formation step. Myogenic differentiation of hPSCs in monolayer cultures has been also proved to be feasible [202–204]. Following feeder-free monolayer culture of hESCs, Barberi and colleagues derived multipotent mesenchymal precursors (MMPs) that could be further differentiated into myogenin-expressing cells [202, 203]. Their monolayer differentiation method involved a serial of cell culture steps in specific culture media and two purification steps based on FACS sorting of CD73-positive mesodermal precursors that after 2–4 days of subculturing were subsequently sorted for NCAM-bright expression, a marker of the embryonic skeletal muscle. Forty-six percent of NCAM-positive cells revealed expression of myogenin, and importantly they were able to fuse and form MyHC-expressing contracting myotubes [202, 203]. First, MMPs were maintained in inactivated fetal serum and in the presence of the mouse skeletal myoblast line C2C12 [202]. Later, Barberi and colleagues could avoid the use of C2C12 cells by using serum-free N2 medium, allowing for the expansion of hESC-derived myoblasts in a serum-free N2 medium in the presence of insulin [203].
Following a similar strategy, Sakurai and colleagues [200] [201] developed a defined protocol for the production of paraxial mesodermal progenitors from mESCs and miPSCs that they could apply to differentiate hiPSCs toward PDGFR-α+/KDR- cells. Those progenitors could be further differentiated into osteocytes, chondrocytes, and skeletal muscle cells, demonstrating the suitability of their procedures for the generation of myogenic cells for regenerative purposes.
Other authors have also shown the possibility to generate PDGFR-α+ from hESCs, although low engraftment was observed after transplantation of such hESC-derived myogenic cells into injured skeletal muscle [205]. Interestingly, the same authors have recently demonstrated that addition of Wnt3a in the culture medium promoted a rapid myogenic commitment of hESCs and, more significantly, that those hESC-derived myogenic cells could contribute to muscle regeneration in a NOD/SCID mice model of the cardiotoxin-injured skeletal muscle [206]. In the same line, other works have demonstrated that inhibition of GSK3β and treatment with FGF2 could specifically induce skeletal muscle differentiation. In particular, Xu and colleagues [207] have demonstrated that simultaneous inhibition of GSK3B, activation of adenyl cyclase, and stimulation with FGF2 during EB formation could promote the generation of myogenic precursors that terminally differentiated in vitro and showed some functional characteristics typical of satellite cells upon transplantation. Similarly, Borchin and colleagues [208] have described that hPSCs could be differentiated toward Pax3/Pax7 double-positive cells after GSK3β and FGF2 treatment.
Moreover, Xu and colleagues have developed a massive platform for the identification of soluble factors promoting muscle differentiation making use of zebra fish embryos [209]. Their system took advantage of zebra fish embryo culture system with reporters of early and late skeletal muscle differentiation, enabling for the examination of 2400 chemicals on myogenesis. Interestingly, authors identified six compounds expanding muscle progenitors, including three GSK3β inhibitors, two calpain inhibitors, and one adenylyl cyclase activator named forskolin. Of note, when bFGF, forskolin, and GSK3β inhibitor BIO were used in hiPSCs, they induced skeletal muscle differentiation and produced engraftable myogenic progenitors that contributed to muscle repair in vivo [209]. Taking advantage of these findings, the same group has recently demonstrated that the same protocol promoted the generation of myotubes from hiPSCs derived from patients affected from Donohue syndrome, offering the first model of human skeletal muscle insulin resistance [210].
1.4 How to Model Muscle Disease in the Petri Dish
Nowadays, the development of protocols to direct cell differentiation from human PSCs in a high range of cell types has set the basis to generate massive platforms for the study of differentiation procedures and disease progression. Furthermore, the correction of the genetic disorders in these cells with classical genetic engineering or emerged genome editing technologies not only allows molecular studies of MDs but also development of future strategies for gene and cellular therapies.
So far, different groups have demonstrated the suitability of patient iPSC approaches in order to model MDs. Abujarour and colleagues [129] have obtained myotubes by direct MyoD-mediated differentiation of hiPSCs from Duchenne muscular dystrophy (DMD) and Becker muscular dystrophy (BMD) patients. Authors validated the differentiated myotubes by a global expression profile that showed how they adopted the skeletal muscle program and the functional response to protein factors investigated as potential treatments for MD, in a similar manner to primary myotubes. These results prove that iPSC derived from DMD and BMD patients has no intrinsic barriers preventing from myogenesis. Although the delivery of MyoD by a lentiviral vector precludes the use of these iPSCs in a clinical setting, they still represented a scalable source of normal and dystrophic myoblast for immediate application in disease modeling and drug discovery.
Recently, Tedesco and colleagues [190] developed the first protocol for the differentiation of mesoangioblast-like cells from iPSCs generated from fibroblasts and myoblasts of limb–girdle muscular dystrophy 2D (LGMD2D) patients. After obtaining mesoangioblast-like cells, authors expanded and genetically corrected them by means of a lentiviral vector for the specific expression of human α-sarcoglycan in differentiated striated muscle cells. A tamoxifen-inducible lentiviral vector of MyoD–ER was also used to induce differentiation of the corrected cells into myotubes before its transplantation into α-sarcoglycan-null immunodeficient mice. Authors showed the engraftment of these cells in the dystrophic skeletal muscle and the related production of myofibers clusters expressing α-sarcoglycan. The amelioration of the dystrophic phenotype in terms of motor capacity was increased when the same experiments were conducted using mouse-derived mesoangioblasts. Overall, Tedesco and colleagues showed how to avoid the limited availability of adult tissue-specific muscle progenitor cells by deriving patient-specific iPSCs and expanding their differentiated progeny. Together with the in vitro genetically correction and later transplantation, this approach could be useful for gene and cell therapies.
In the same line, Tanaka and colleagues [121] developed a myogenic induction system to differentiate iPSCs from patients affected by Miyoshi myopathy (MM), a congenital distal myopathy caused by mutations in dysferlin (DYSF). Authors obtained myotubes that showed MM associated phenotype with impaired expression of DYSF and defective membrane repair. These features were rescued by the expression of full-length DYS by a piggyBac (PB)-based vector. A similar work was performed by Yasuno and colleagues [122], where authors generated iPSCs from patients affected by carnitine palmitoyltransferase II (CPT II) deficiency, an inherited disorder involving B oxidation of long-chain fatty acids (FAO). Differentiated myocytes recapitulated the increase accumulation of C16 (palmitoylcarnitine) that could be restored by bezafibrate, mimicking some clinical aspects of CPT II deficiency. All these data show how the patient-specific iPSCs and later differentiation result in the generation of validated in vitro models of both diseases.
Recently, Li and colleagues [211] have demonstrated the possibility to correct iPSCs derived from DMD patients by the use of genome editing technologies: TALEN and CRISPR/Cas9. Authors took advantage of the ability to expand iPSCs limitlessly to develop three different strategies: exon skipping, frameshifting, and exon knock-in, in order to correct the pathological mutation. The exon knock-in was the most effective approach to restore the full-length dystrophin protein in the iPSC-differentiated myocytes. In this context Turan and colleagues [212] have corrected limb–girdle muscular dystrophy 2B (LGMD2B) and 2D (LGMD2D) by DICE or TALEN-mediated integration of wild-type DYSF cDNA into the H11 safe harbor locus and single-stranded oligonucleotide-mediated gene editing by CRISPR/Cas9, respectively. These approaches resulted in the adequate protein expression for DYS and relocation of corrected α-sarcoglycan protein to the cell membrane in muscle progenitor cells differentiated from iPSC. These works demonstrate the capability of iPSC technology to provide in vitro muscle models and in combination with genome editing autologous corrected cells for ex vivo gene therapy approaches.
Very recently Salvatore Iovino and colleagues [210] derived iPSC from patients of Donohue syndrome related with insulin receptor mutations (IR-Mut). These cells were differentiated in myotubes that exhibited insulin resistance-like (IR) responses in vitro. IR-Mut myotubes fail to increase glucose uptake, glycogen synthase activity, or glycogen stores in response to insulin stimulation. Transcriptional regulation was also perturbed in IR-myotubes with reduced insulin-stimulated expression of insulin receptor protein and reduced insulin-stimulated phosphorylation of the receptor and downstream effectors. This work indicates an impairment of the insulin signaling to induce the expression of metabolic and early growth response genes. This data validated this model of skeletal muscle insulin resistance not only to dissect its genetic features related with Donohue syndrome but also to study epigenetic acquired features related with other insulin resistance states such type 2 diabetes. All these advances are summarized in Table 1.1.
Table 1.1
Summary of current approaches for modeling and correcting muscle diseases using patient-specific-induced pluripotent stem cells (iPSCs)
Differentiation | Disease correction | ||||||
|---|---|---|---|---|---|---|---|
Author | Disease | Gene affected | Exogenous Expresion | Cullture | Differenciated cells obtained | Molecular strategy | Rescued phenotype |
Abujarour et al. | Duchenne Muscular Distrophy (DMD) and Becker Muscular Distrophy (BMD) | Dystrophin (DMD) | Doxycycline induction of MYOD1 by lentiviral vector | · DAY -1: iPSCs culture media w/o bFGF+ROCKi+Collagen or Matrigel | Myotubes | Only Modeled | |
· DAY 0-4: Dox+DMEM+15 % FBS | |||||||
DAY 4-7: low-glucose DMEM and 5% HS | |||||||
Tedesco et al. | Limb-girdle muscular dystrophy 2D (LGMD2D) | α-sarcoglycan (SGCA) | — | · Week 1-2: Single cell, low O2+ROCKi+αMEM+matrigel. Split at week 1 | Mesoangioblast-like stem/progenitors (MABs) | · Lentiviral vector carrying wt SGCA under the muscle-specific myosin light chain 1F promoter and enhancer. | DMD expresion and muscle colonization of corrected cells in SGCA-null mice |
Week 3: Post trypsin, seed at 80% confuency, low O2+MegaCell or IMDM+matrigel | · Or human artificial chromosome containing the entire DMD locus. | ||||||
Week 3 to 4: Post trypsin, after 100% seed at 80% low O2+MegaCell or IMDM w/o matrigel | Transduced at MABs state and then myogenic differenciated to multinucleated myotubes by tamoxifen inducible lentiviral vector of MyoD-ER | ||||||
Tanaka A et al. | Miyoshi myopathy (MM) | Dysferlin (DYSF) | Doxycycline induction of MYOD1 by piggyBac (PB) based vector | · DAY 0-2: w/o bFGF+Dox (at D1) | Myocytes | EF1a promoter-driven constitutive expression of DYSF by a piggyBac (PB) based vector | Expression of DYSF in differenciated myocytes |
DAY 2-7: 5% KSR αMEM+Dox | |||||||
DAY 7-9: 5% HS +DMEM+IGF-1 | |||||||
Yasuno et al. | Carnitine palmitoyltransferase II (CPT II) deficiency | Carnitine palmitoyltransferase II (CPT2) | Doxycycline induction of MYOD1 by piggyBac (PB) based vector | · DAY 0-2: w/o bFGF+Dox (at D1) | Myocytes | Only Modeled | |
DAY 2-7: 5% KSR+αMEM+Dox | |||||||
DAY 7-9: 5% HS +DMEM+IGF-1· | |||||||
DAY 10-11: Low-glucose +MEM+0.4% BSA+L-carnitine+palmitic acid+P/S | |||||||
Li et al. | Duchenne Muscular Distrophy (DMD) | Dystrophin (DMD) | Doxycycline induction of MYOD1 by piggyBac (PB) based vector | · DAY 0-2: w/o bFGF+Dox (at 1)· | Myocytes | · TALENs and CRISPRs for Exon skipping, Frameshift or knock-In | Expresion of DMD in differenciated myocytes |
DAY 2-7: 5% KSR+αMEM+Dox· | |||||||
DAY 7-9: 5% HS+DMEM+IGF-1 | |||||||
Limb-girdle muscular dystrophy 2B (LGMD2B) | Dysferlin (DYSF) | — | · DAY -7-0: Embryoid bodies (EB)+APEL media+bFGF +BIO+forskolin· | Muscle Progenitors | · Integration of wt DYSF cDNA into the H11 safe harbor locus by DICE or TALEN. | Expresion of DMD or SGCA in differentiated muscle progenitor cells | |
Turan S. et al. | Limb-girdle muscular dystrophy 2D (LGMD2D) | α-sarcoglycan (SGCA) | DAY 0-21: matrigel+DMEM+2%HS | · Singlestranded oligonucleotide-mediated gene editing by CRIPR/Cas9 (HDR). | |||
Iovino S. et al | Donohue syndrome | Insulin Receptor (INSR) | · DAY 0-7: STEM Diff Apel medium+bFGF+Bio+Fordkolin | Myotubes | Only Modeled | ||
DAY 7-9: Matrigel+Media Serum Free
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