Skeletal muscle formation and regeneration

Skeletal muscle formation, a process called myogenesis, occurs during embryonic development, and during postnatal muscle growth, regeneration and repair. Myoblasts, the early muscle cells with a single nucleus, proliferate and express muscle-specific genes leading to their fusion and the formation of multinucleated myotubes, which will ultimately form the mature myofibers [5]. Muscle maintenance is a continuous process depending on physical demand, injury and/or disease. Normal growth and replacement are mediated by a stem cell population termed satellite cells (SCs) (Fig. ​(Fig.1).1). These cells can be found beneath the basal lamina of muscle fibers and are stimulated after muscle injury to proliferate and differentiate to myoblasts, which then fuse to form new myotubes or fuse to existing myofibers, thereby contributing to muscle regeneration, which is further explained below [5, 6].

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Fig. 1

Skeletal muscle-resident cells. Schematic cross section of a healthy skeletal muscle bundle, containing more than a dozen individual muscle fibers (light red; nuclei at the periphery). Satellite cells (grey) are muscle-lineage committed progenitors that are located beneath the basal lamina of the muscle fibers, near the vasculature. In between the fibers are a variety of interstitial cells. Pericytes (purple) are one type and can be found wrapped around blood capillaries (insert). All these muscle-resident cell populations contribute to muscle repair and regeneration

The disease state in several myopathies affects the functional capacity of SCs and consequently impedes muscle regeneration. Stimulating regeneration by the addition of myogenic cell types has therefore been the main goal of cell therapy for muscle diseases. This development started with the use of myoblasts and SCs, which led to disappointing clinical results due to limited survival and scarce migration of injected cells. Next, the field moved to other myogenic cell types, each with their own advantages and disadvantages. Most recently, mesoangioblasts (MABs) and pericytes have been pursued for muscle cell therapy. These myogenic progenitor cells possess various beneficial characteristics needed for an effective cell therapy approach, i.e. simple isolation, the ability to proliferate in vitro, systemic application, and the capacity to differentiate efficiently into skeletal muscle fibers in vitro and in vivo. Before we will discuss MABs and pericytes in more detail, we will review other myogenic progenitor cell types and cell populations that have been studied for their therapeutic potential over the last few years.

Myoblasts and satellite cells: how the mighty have fallen?

In 1989, the groups of Louis Kunkel and Terence Partridge pioneered muscle cell transplantation in MD models [27, 28]. Allogenic neonatal mouse myoblasts were the first cells to be intramuscularly transplanted into a dystrophin-deficient DMD model with the goal to form new dystrophin-positive fibers. Although encouraging, the results in mice could not be recapitulated in patients, which were injected with muscle stem cells harvested from healthy human immunocompatible donors [18, 29–32]. Only low expression percentages of normal dystrophin were detected posttransplantation in muscle biopsies [29–32] and, with the exception of one study [33], no functional muscle improvement in the transplanted limb was measured. Functional effects in these studies were limited by cell death after transplantation and immune rejection, together resulting in a limited number of cells that survived in vivo, and a scarce migration of injected cells. Recently, in an attempt to increase engraftment, limit rejection and restore dystrophin expression, fused myoblasts from healthy donors and DMD-affected donors were i.m. injected into mdx/scid mice [34].

SCs are able to maintain their stemness due to their self-renewal capacity, implying that SCs are preferred over myoblasts for a successful therapy. They are considered to be the bona fide stem cell of skeletal muscle. SCs are positioned between the basal lamina and the sarcolemma of muscle fibers (Fig. ​(Fig.1)1) [6]. In this niche, quiescent SCs can respond to damage or disease due to the local release of cytokines, growth factors or cell differentiation factors like NOTCH and WNT. The factors are secreted by muscle tissue itself and nearby macrophages or fibroblasts. High expression of NUMB, an antagonist of NOTCH signaling, leads cells to go back in quiescence [5, 35]. Quiescent SCs express PAX7, MYF5, CD34 and generally also PAX3 [36, 37]. SCs in which p38α/β MAPK is asymmetrically activated, undergo myogenic commitment [5, 38]. A group of highly conserved myogenic regulatory factors (MRFs) is responsible for the entrance in the cell cycle and further myogenic differentiation. Activated SCs loose CD34 expression and instead start expressing MYOD, a well-known member of the MRF family. Co-expression with myogenin (MYOG) activates terminal differentiation of the muscle cells [37]. The cascade responsible for muscle differentiation is additionally influenced by a multitude of cytokines, circulating hormones and exosome-secreted signals [5, 37]. This process of skeletal muscle formation, growth and maintenance in healthy subjects or patients with MD is elaborately discussed elsewhere [5, 26, 39].

The therapeutic value of SCs has been examined in several transplantation studies [40, 41]. However, many of the same practical limitations and safety concerns seen for myoblasts appear to restrict the use of human SCs in the clinic. The low proliferative capacity in vivo, limited migration through muscle tissue [42], in addition to the immune reactions against donor cells [43, 44], were highlighted as the main concerns (reviewed in [15, 45]). Trying to overcome these challenges, researchers delivered a large number of allogeneic myogenic cells under immunosuppressive conditions using multiple injections [15, 46]. However, such high-density injection protocols seem to be feasible only for dystrophies where small muscles are locally affected, e.g. in OPMD [19, 47]. Accordingly, the search for a cell type more suitable for a systemic muscle cell therapy continued.

The microvasculature as niche and angiogenesis as distribution route

MABs are mesenchymal-like cells associated with the wall of the aorta. These myogenic cells are thus from non-somite, vascular origin [63, 74, 75, 79–84]. MABs use the microvasculature and angiogenesis events as progenitor cell niche and distribution route, respectively, during which progenitors are generated and spread throughout various postnatal tissues [20, 85]. MABs leave the blood vessels and enter the surrounding mesenchyme where they are exposed to local signals, allowing them to integrate and fully differentiate into mesodermal tissues such as blood, cartilage, bone and smooth, skeletal and cardiac muscle [73]. The idea that MABs, isolated from explant cultures of embryonic dorsal aorta, are blood borne is supported by the presence of known vascular endothelial markers such as VE-cadherin, β3-integrin and P-selectin, in addition to myogenic markers (MYOD, MYF5, DESMIN, MNF, c-MET and M-cadherin) [74].

Myogenic fate regulation of MABs is Pax3-dependent

The muscular fate determination of myogenic progenitors is influenced by the paired box/homeodomain transcription factors of the Pax family. Mouse SCs express Pax3 and Pax7 and once they become myoblasts, Pax7 expression goes down, while MyoD and Myf5 expression remain. Surprisingly, Pax7 expression is not found in MABs. The skeletal muscle fate of these cells is only Pax3-dependent [86, 87]. During embryonic development, skeletal muscle is formed from the dorsal somite, the dermomyotome, Pax3 is expressed in the multipotent cells of this structure [88]. Pax3 is not only essential for the survival of cells in the dermomyotome, but also plays an important role in the migration of these cells outside the somite [86]. When outside, in the limb buds for example, the myogenic determination gene Myf5 is activated [89]. Genetic labeling showed that vessel-derived progenitor cells and cells of the myotome both originate from Pax3⁺ multipotent cells of the paraxial mesoderm [90].

Pericyte, the postnatal equivalent of MAB

Interestingly, tracing Pax3⁺ GFP-labeled MABs also revealed Pax3⁺ cells in postnatal blood vessels. Although their frequency appears to be lower than the embryonic correlate, these Pax3⁺ progenitors, called pericytes, are considered the descendants of the multi-potent Pax3⁺ cells present during embryogenesis [86, 88, 90–92]. The cells delivered through fetal angiogenesis remain dormant within steady-state tissues and can be activated during postnatal events.

Pericytes are contractile cells in close proximity with the endothelial cells of small blood vessels [93]. They can be found wrapped around blood capillaries, precapillary arterioles, postcapillary venules, and collecting venules (Fig. ​(Fig.1)1) [94], where they regulate capillary barriers, endothelial proliferation and capillary diameter by integrating and coordinating neighboring endothelial cell response [92]. In contrast to MABs, pericytes have no unlimited self-renewal capacity in vitro. They adopt a large, flat morphology after approximately 20-30 population doublings and undergo senescence [20, 91]. Unlike MABs, pericytes spontaneously differentiate into skeletal muscle myotubes in low-serum culture conditions [95]. Skeletal muscle tissue appears to be the most effective source for pericytes with myogenic potential [22].

Establishing a comprehensive pericyte identity

It is difficult to discriminate pericytes from other interstitial progenitor cells. Pericyte abundance, morphology and marker identity differ throughout the body [96–98]. Moreover, there is lack of a specific pericyte maker, which makes it challenging to exactly characterize these progenitor cells. In contrast to MABs, pericytes lack endothelial markers CD31, CD34, and KDR. Markers such as M-cadherin, N-CAM, cytokeratins or neurofilaments (with the exception of nestin) are also not present. Neither are myogenic markers, MYOD, MYF5 and MYOG, apart from PAX3 [20]. The difference in marker expression between muscle progenitors may be correlated with the different localization of the cell types. SCs are located next to the basal lamina of muscle fibers, while MABs occupy an endothelial position. Pericytes, on the other hand, are located underneath the basal lamina of the small vessel, where they are completely embedded within the endothelial cell basement membrane.

Markers that have been used to identify pericytes include smooth muscle α-actin (αSMA) [99], desmin (DES) [100], high molecular weight melanoma antigen (HMW-MMA) (called NG2 in the mouse) [101, 102], platelet-derived growth factor receptor (PDGFR)-β [103], aminopeptidase A and N [104, 105], the regulator of G-protein signaling-5 (RGS5) [106, 107], and the promoter trap transgene XLACZ4 [108] (Table ​(Table4).4). The subfraction of pericytes with skeletal myogenic potential can be recognized by the absence of PDGFRα and the presence of alkaline phosphatase (ALP) in murine and human tissues [20, 91, 95, 109, 110]. Moreover, expression of PW1/PEG3 is related to progenitor cell competence, since silencing PW1/Peg3 leads to MYOD degradation and consequently inhibits myogenic potential in vitro. Additionally, without PW1/PEG3 cells are unable to modulate the junctional adhesion molecule-A (JAM-A) and are therefore incapable of migrating across the vessel wall and engrafting into damaged myofibers [111]. It is good to keep in mind that all markers nowadays used for selection are dynamic in their expression. They may be up- or down-regulated between organs, in conjunction with developmental phases, pericyte maturity [112, 113], diverse pathological states and in vitro culturing [114, 115].

Myogenic fate regulation of pericytes begins at the onset of differentiation

When investigating the differentiation process, it was found that mouse pericytes never expressed Pax7, Myf5 nor MyoD during proliferation, but activated these genes at the final stages of differentiation. This activation was concomitant with that of myogenin and just before the expression of myosin heavy chain proteins in myotubes. The temporal expression pattern is different in SCs, which do express Pax7, Myf5 or MyoD during proliferation and activate myogenin before myosin heavy chain [20].

ALP⁺ pericyte-mediated myogenesis seems modest in adult muscle as the ALP⁺ cells mainly contribute to skeletal muscle during the first few weeks of postnatal growth. The process of ALP⁺ cell contribution becomes infrequent in adult mice [109]. However, pericytes can respond to postnatal repair events. Vascular ALP⁺ pericyte progenitors mainly contributed to the growth of muscle fibers by entering the SC niche, albeit direct fusion with developing muscle fibers occurred as well. The ALP⁺ pericyte-derived SCs could self-renew and maintained a steady contribution to the SC pool [109].

Acute skeletal muscle regeneration and chronic skeletal muscle regeneration showed increased ALP⁺ cell counts at these sites, respectively five and three times more ALP⁺ cells compared to controls. Most studies indeed found an increase of pericytes in in myopathic muscle biopsies, especially when regeneration is present [116]. However, some studies discovered a reduced number of pericytes in specific forms of MD [117].

The role of trophic factors in myogenic fate regulation by pericytes

The functional benefit of stem cell therapies relies on the in situ differentiation of the grafted cells. Besides intrinsic cellular properties, differentiation potential is dependent on trophic factors. Paracrine factors secreted by pericytes may act on adjacent muscle stem cells and exert regulation of their postnatal fate. An ELISA for 121 cytokines and growth factors on pericyte culture supernatant showed that it contained two factors for receptors highly abundant in muscle progenitor cells: insulin-like growth factor 1 (IGF1), an important factor in muscle development and growth, and angiopoietin 1 (ANGPT1), a factor that regulates stem cell quiescence [118–120]. It seems that postnatal myofiber growth and regeneration is influenced by SC quiescence through ANGPT1 and differentiation and growth-promoting effects of IGF1, both produced by pericytes [120].

Preclinical research with MABs

To test whether MABs were suitable for skeletal muscle regeneration and restoration of protein expression, the cells were transplanted in a variety of MD animal models. Morphological and functional recovery was seen in adult immunocompetent α-SG null mice, an animal model for LGMD, after i.a. delivery of MABs transduced with a lentiviral vector expressing α-sarcoglycan (SG). After three consecutive injections, embryonic vessel-derived MABs restored α-SG expression in >20% of the muscle fibers in four different muscles. Furthermore, a decrease in fibrosis and injured muscle fibers allowed treated mice to run three times longer as untreated mice on a rotarod [121].

Delivery was optimized by exposure of MABs to combined pretreatment with SDF-1 or TNFα and expression of α4-integrin [122]. This led to α-SG expression in >90% of muscle fibers in the tibialis anterior muscle of α-SG-null mice four months after only one i.a. injection. It restored protein expression to roughly 60% of wild-type levels.

After this promising result, the possibility to use an allogenic donor therapeutic cell pool was assessed. H2-mismatched MABs from BalbC mice were transplanted in the same dystrophic mice. To assess the immune response against the donor cells, only half of the α-SG-null mice were treated with different immunosuppression. Under immune suppressive regime, donor cells formed more α-SG expressing fibers then did syngeneic MABs [123]. This observation indicates that with the correct immune modulation transplantation of MABs into an immunologically unrelated host can lead to long-term survival of donor cells that are able to form α-SG expressing fibers.

Finally, the formation of de novo muscle fibers was established when MABs derived from juvenile C57Bl/6 mice were i.m. injected into dystrophin- and utrophin-deficient double knockout (mdx|utrn-/-) mice, a phenotypic model for DMD. Dystrophin expression was restored to approximately 50%, as compared to wild-type control mice, in the injected gastrocnemius [124].

Ex vivo gene editing or gene augmentation

Many forms of MDs are monogenic disorders, which makes gene replacement a promising concept. Clustered regularly-interspaced short palindromic repeats (CRISPR)/CRISPR-associated system (Cas) is a powerful tool to accurately alter a target in the genome of eukaryotes [132]. However, unwanted off-target cleavage events by CRISPR/Cas remain a bottleneck and especially unintended germline modifications are worrisome [133, 134].

In an in vivo approach, it is impossible to select for the correctly edited cells and thus the accuracy, efficiency and safety of the gene editing process needs to be exceptionally high. Ex vivo gene editing comes with the considerable benefit of selection of correctly edited cells. A drawback of this approach, however, can be the size of the correctly edited cell pool that is needed. Freshly isolated myogenic precursor cells have the highest regenerative potential and culture conditions influence transplantation efficiency in a negative manner [135, 136]. The prolonged culture to amplify the few correctly edited cells will likely also lead to a less effective cell pool.

In vitro expansion of muscle precursor cells while maintaining therapeutic potential

The clinical trials included in this review (Table ​(Table3)3) were mostly focused on safety and feasibility [59, 78]. When functional effects are envisioned, larger cell doses are needed. Obtaining more cells is challenging since in vitro expansion under traditional culture conditions is often hampered by changes in phenotypic expression of cells, which in turn affects transplantation efficiency.

Cells in an artificial in vitro environment are exposed to non-physiological conditions such as the non-elastic plastic culture substrate, high oxygen supply, additional supplements and loss of contact with other cells. These circumstances affect cell behavior at that moment, but also have long-lasting effects on cells after transplantation. It is well established that expansion of myogenic cells before engraftment reduces their regenerative capacity [136, 137]. This is probably caused by a more activated and differentiated state of the grafted cells, which may diminish their regenerative potential, since freshly isolated progenitor cells are not activated at the time of grafting [136, 137].

One way to reduce culture-induced modifications is via the use of soft hydrogel substrates to mimic tissue rigidity, a biophysical property of the skeletal muscle microenvironment [138, 139]. The altered substrate rigidity is thought to preserve stemness by altering cell shape. Mechanical sensing signals are transmitted via focal adhesions towards nonmuscle myosin II isoforms (NMM IIA, B, and C). These non-muscle myosins signal cells to deform their matrix which results in cytoskeletal rearrangements and altered signaling [140, 141] [142, 143]. MSCs, for example, express no baseline levels of lineage-specific markers however the substrate rigidness influences lineage specification of these naive stem cells. Softer matrices of polyacrylamide coated with collagen I that mimic brain tissue are neurogenic for MSCs, while similar but stiffer matrices induce myogenic differentiation and rigid matrices give rise to osteogenic cells [140]. When cells are in identical serum conditions, the matrix stiffness is a stronger driving force for reprogramming MSCs than are soluble factors. However, the combination of correct elasticity and soluble induction factors synergistically cause an even more complete myogenesis [140].

Typically used polystyrene plastic has an elastic modus of ~3 GPa which is five orders of magnitude more rigid than skeletal muscle [138, 144]. Using tunable polyethylene glycol (PEG) hydrogel and laminin as an adhesion ligand, MuSC division rate remained unchanged and MuSC stemness was retained, as seen in the gene expression [138]. Hydrogels are effective in keeping the proliferative capacity stable in expansion cultures up to 20 passages in vitro [138, 139]. However the extent of engraftment from cultures MuSC is still not as high as that of freshly isolated cells, suggesting that additional chemical and physical cues may be required to establish an optimal in vitro muscle microenvironment to keep the maximal therapeutic potency [138].

Besides tissue rigidity, specific extracellular matrix (ECM) protein coatings can be used to mimic the SC niche. Matrigel, the most widely used commercialized ECM mixture, albeit without FDA approval, contains critical grow factors and cytokines [145, 146]. The most prevalent proteins in the mixture are laminins, collagens I and IV, and fibronectin. These proteins are also present in the ECM of the myofiber niche [147]. Laminin mainly affects proliferation and migration of myogenic progenitor cells, while fibronectin plays a role in remodeling after muscle damage [148]. Under culture conditions fibronectin can modulate cellular expansion by potentiating the Wnt7a-dependent signaling pathway [149]. Collagen is more of a structural component, it improved the self-renewal of SC [147, 150]. Several peptides released from ECM proteins called matricryptins have been suggested to be useful in the proliferation, migration, and survival rates of myoblasts [151, 152] although specific research is missing.

Other than mimicking the in vivo environment to aid cell expansion, proliferation of cells can be influenced by the use of nitric oxide, calorie mimicking drugs, or genetic modification of specific targets. Nitric oxide (NO) contributes to myogenesis by the formation of S-nitrosothiols (RSNO). An increase in RSNO inhibits the S-nitrosoglutathione reductase (GSNOR) pathway and leads to an increase in the number of myoblasts, followed by a decrease in the myoblast fusion index [153].

Myoblasts cultured in the presence of calorie restriction mimetics behave differently. These drugs, for example metformin and ursolic acid, regulate myogenic differentiation and cell proliferation depending on the dose. By varying the doses of the calorie restriction mimicking drugs one can induce a shift between proliferating and differentiating status [151, 154, 155]. Genetic modification of cells is also advocated to promote expansion, engraftment and sustained regeneration. Transcriptome studies revealed that activation of p38 signaling correlated with myogenic cell differentiation, while inhibition of p38 reversibly prevented differentiation and could be used ex vivo to promote expansion. Pharmacologic manipulation of p38 signaling can be leveraged for enhancement of both ex vivo expansion and subsequent in vivo engraftment of cells [135].

The value of using iPSCs in muscle cell therapy

A different approach to circumvent the limited availability of freshly isolated primary cells, low proliferation capacity and early senescence, is the use of induced pluripotent stem cells (iPSCs). Generation of iPSCs is achieved by reprogramming somatic cells with a defined set of transcription factors (OCT3/4, SOX2, c-MYC, KLF4) [156]. These so-called Yamanaka factors can be supplied by integrating and non-integrating viral vectors or non-viral episomal vectors to mitigate the risk for insertional mutagenesis. Next to their unlimited replicative capacity, another advantage of the use of iPSCs is the possibility to select correctly edited cells after CRISPR/Cas treatment. These can then be clonally expanded and differentiated into suitable muscle progenitor cells [157].

Genetically corrected human iPSC-derived MAB-like cells (HIDEMs) from LGMD patient fibroblasts only restored α-SG in 2% of skeletal myofibers in α-SG-KO mice after i.m. injection [117]. The relatively low engraftment of donor cells suggests that autologous therapy using genetically corrected HIDEMs might be difficult. Efficiency of transplantation of these cells seems to be low, indicating a delicate balance between proliferative capacity and differentiation potential for the ultimate transplantable cell [117].

As mentioned before, progeny-specific imprinting plays a role in differentiation [24, 25]. Stronger myogenic commitment can be achieved via the generation of iPSCs from prospectively sorted myogenic cells (here called MAB-iPSCs) instead of fibroblast-derived iPSCs (f-iPSCs). The epigenetic memory in MAB-iPSCs provides a more robust myogenic differentiation [25]. I.m. injection of wild-type mouse iPSC-derived MAB-like cells (MIDEMs) resulted in an amelioration of the motor capacity in Sgca-null/scid/beige mice [158]. Although not compared one-on-one, efficacy of (genetically corrected autologous) HIDEMs is likely higher compared to f-iPSCs in vivo.

Survival and engraftment of transplanted cells in vivo

A crucial factor for an effective cell therapy is the early survival of donor cells. The ultimate goal is that the ratio between death and proliferation leads to a good “net” survival post-transplantation, with the whole donor cell population being either stable or growing. Although suggested many times, it seems that myogenic cells are not instantly killed and removed after i.m. injection, but they undergo necrosis or apoptosis in the first few days. Factors that have been implicated in cell death are cellular effectors of an acute inflammatory response, a combination of natural killer cells, T-lymphocytes and complement, and the fact that only a small specific donor cell sub-population with stem cell-like characteristics survives [159].

Various factors are able to improve the survival, migration and engraftment of injected cells. The growth factor, vascular endothelial growth factor (VEGF) reduced hypoxia-induced death of human myoblasts in vitro and in a mouse model [151, 160]. Other growth factors such as insulin-like growth factor-1 (IGF-1) and basic fibroblast growth factor (bFGF) were co-injected and promoted the overall migration of human myoblasts in various mouse models and stimulated cell migration and engraftment of monkey myoblasts in a nonhuman primate model [161]. Treatments with these factors stimulated components of proteolytic systems and thereby enhanced cellular migration. A short term ex vivo treatment using Wnt7a, a member of the Wingless-INT (WNT) family, on satellite stem cells had similar effects and markedly increased cell dispersion and engraftment, which ultimately resulted in improved muscle function of dystrophic muscles [162].

The inhibition of p38 signaling pathway in vitro during SC expansion enhances in vivo engraftment [135]. Identifying additional modifications in the culture protocol that increase the regenerative potential of isolated cells will therefore have large beneficial effects. As an example, improvements have been made by p53 inhibition which can increase survival of edited cells. Nonetheless, the selection of p53-mutated cells raises additional concerns, since p53 inhibition could increase cancer risks [163, 164].

Pre-treatment of receiving skeletal muscle

The harsh environment that cells get exposed to after injection likely plays a role in rapid cell death and low long term viability [165]. MDs cause changes in the skeletal muscle niche leading to a hostile environment for injected cells [166, 167]. The use of immunosuppressive drugs has proven to increase viability of injected cells however it comes with side effects [168]. The encapsulation of cells in polymer-based microcapsules might be effective. The encapsulated cells are protected from the immune system but outward diffusion of factors is still possible [169]. The review of Murua et al. focusses on the cell microencapsulation technology and the possibilities in the clinic [169].

Fibrosis, the progressive replacement of functional tissue by nonfunctional and more rigid connective tissue, occurs in many MDs. In a fibrotic environment cells have been shown to differentiate more towards a fibrogenic fate thereby decreasing their regenerative potential [170, 171]. Improving the quality of receiver muscle by antifibrotic therapies before grafting will be helpful to optimize the fate of implanted cells. The co-injection of myoblasts with other cell types such as pro-inflammatory macrophages increased proliferation and migration, and delayed differentiation [172].

As mentioned before, alterations in Wnt7a and p38 signaling could exert positive effects on myogenic cell migration and engraftment. Besides pretreatment of cells, treating receiving muscle tissues could also potentially be beneficial [135, 173]. Focal treatment of dystrophic muscles using mdx mice showed that Wnt7a treatment efficiently induced satellite cell expansion, myofiber hypertrophy, reduced the level of contractile damage and caused a shift in fiber type toward slow-twitch [174].

The competitive potential of transplanted cells

A crucial factor complicating the efficacy of myogenic cell transplantation is that both mRNA and intracellular proteins in myofibers tend to stay close to the nucleus of origin, in an area called the nuclear domain [175]. Due to this limited spreading, proteins from donor cells remain in the myofiber to which the transplanted cells fused. The size of the nuclear domain differs for each protein, but also depends on the length of the myofiber in which grafted cells are integrated. For DMD, the graft-derived myonuclei can transcribe mRNA and synthesize donor cell-derived dystrophin. In a handful of clinical trials with myoblast transfer, expression of donor-derived dystrophin was indeed confirmed in myofibers of patients [30, 46, 47, 176]. However, the novel proteins remained near the donor myonucleus, the nuclear domain in the myofiber. Muscle sections derived from DMD patients that underwent cell engraftment with healthy myoblasts showed that donor-derived dystrophin expression was restricted to distances of 0.7-2.0 mm in single myofibers [47].

Importantly, in myotonic dystrophy there is no need to ensure distribution of a missing protein along a myofiber, but the removal or reduction of the toxic RNA in a myotube is required for functional improvement. Therefore, once the technical procedures for ex vivo editing and cell transplantation have been established, further studies are required to determine conditions of the most optimal administration and dosage regimen to ensure the regeneration of myofibers and exert a positive effect on muscle function.