Cell therapy for Duchenne muscular dystrophy: promises, challenges, and controversies

Abstract

Despite extensive studies, Duchenne muscular dystrophy, a neuromuscular disorder caused by the lack of dystrophin, a key muscle structural protein, remains an incurable disease. One of the potential treatment options currently being investigated is cell therapy, although it has not yet been clinically established. Several strategies, including muscle satellite cells, mesoangioblasts (vessel-associated multipotent stem cells), and induced pluripotent stem cell (iPSC)-derived muscle cells, have emerged as tools for restoring dystrophin expression and regenerating damaged muscle tissue. Nevertheless, each of these approaches faces significant limitations, including poor cell engraftment, low delivery efficiency, and the risk of immune rejection. Furthermore, long-term safety, the possibility of tumorigenicity, and off-target effects must be rigorously evaluated. Importantly, the latter technology, utilizing cardiomyocytes differentiated from iPSC, holds the potential for addressing cardiomyopathy, the major cause of death of DMD patients. At the same time, several interventions using cells with claimed stem cell potential have emerged, raising both scientific and ethical concerns. This review summarizes recent advancements in the development of cell therapies for DMD, highlighting promising progress while critically analysing questionable approaches.

Introduction

Duchenne muscular dystrophy (DMD) represents one of the rare genetic disorders, affecting approximately 1 in 5,000–6,000 boys. The primary cause of the disease, which drives its rapid and irreversible progression, is the absence of functional dystrophin, a 427 kDa actin-binding protein and a crucial component of the dystrophin-glycoprotein complex (DGC). More than 7,000 patient-specific mutations have been identified in the DMD gene, the largest human gene, which contains 79 exons and approximately 2.4 million base pairs [1].

The predominant muscle phenotype of DMD is attributed to the absence of dystrophin, which connects the intracellular cytoskeleton to the extracellular matrix by linking F-actin and the DGC complex, functioning as the ‘shock absorber’ and force transducer in muscle and cardiomyocytes [2, 3]. DGC maintains the integrity of muscle fibers and regulates several cellular pathways, including nitric oxide (NO) synthesis and calcium entry [4, 5]. Therefore, a lack of dystrophin results in increased sarcolemmal damage caused by contractions and disrupts the homeostasis of NO and Ca²⁺. However, dystrophin appears to be also expressed in other cells, including cardiomyocytes, smooth muscle cells, and many non-muscle cells [6–9], although the presence and significance in fibroblasts and endothelial cells are disputable. Dystrophin was also claimed to serve roles beyond its structural function, e.g., it is presumed to be a tumor suppressor gene [10], though this hypothesis requires further validation. Accordingly, mutations in DMD patients, which either result in a complete loss of dystrophin or, in rarer cases, the production of dysfunctional protein, significantly affect the function of skeletal and respiratory muscles, the heart, the nervous system, and other organs (for a review, see e.g [11]).

DMD patients suffer from progressive muscle dystrophy with a plethora of clinical signs. The first symptoms of the disease, identified mostly by parents, are frequently noticeable in 1-3-year-old boys and manifest, among others, as a struggle to sit down, stand up, walk, jump, climb stairs, and run. In young boys, symptoms often include calf muscle enlargement, poor head control, muscle pain, and general clumsiness. A characteristic feature is the Gowers’ sign, where weakness in the proximal muscles forces the child to use their hands to rise from a lying position.

Between the ages of 8 and 14, patients lose ambulation due to gradual muscle atrophy and become dependent on wheelchairs. Scoliosis and joint contractures develop as muscular weakness increases, which in turn worsens restrictive lung disease [12]. Therefore, with disease progression, respiratory failure occurs, which can be alleviated through the use of nocturnal non-invasive ventilation (NIV), diurnal non-invasive ventilation via a mouthpiece, or, finally, ventilation via tracheostomy [13]. Furthermore, immobility increases the risk of fracture and accelerates the loss of bone density [14]. Of note, cardiorespiratory complications have become one of the primary causes of mortality from the second decade of life in DMD men. Importantly, female carriers are also at risk of cardiomyopathy [15–18]. Unfortunately, heart problems in DMD patients may be undetected without detailed examination [19, 20]. Many teenage individuals have no classic symptoms of heart failure, or these symptoms are not recognized properly, as they mostly use wheelchairs and do not undergo increased cardiac workload. This results in a significant delay in their proper assessment and the late start of pharmacological therapies [21]. Therefore, to prevent the early onset of heart failure, initiating cardiac-focused interventions before ventricular dysfunction is detected may be crucial. Nevertheless, so far there is no effective treatment for dystrophin-dependent cardiomyopathy, which is ultimately fatal in DMD patients (who, due to their very severe overall condition, typically do not qualify for heart transplantation), highlighting the need to identify novel therapeutic avenues and mechanistic insights [22–24]. In addition to well-known skeletal and cardiac muscle symptoms, DMD is associated with cognitive disorders, language, and learning disabilities, failure to thrive, and metabolic disorders (obesity, insulin resistance, glucose intolerance) [22, 25]. A summary of the disease progression and its key hallmarks is presented in Fig. 1.

Fig. 1.

DMD progression. Mutations in the DMD gene, resulting in the absence of dystrophin, lead to muscle degeneration and heart complications. These are exacerbated by chronic inflammation, accelerated fibrosis, increased oxidative stress, disturbed regeneration, and impaired angiogenesis. Symptoms typically begin in early childhood (2–3 years), with weakness in the lower limbs, difficulty walking, and gait abnormalities. In adolescence (~ 12 years), most individuals are wheelchair-bound, and respiratory and cardiac issues become more severe. In adulthood, ventilatory support is often required, and life expectancy is significantly reduced due to respiratory and heart failure

Satellite cell dysfunction in Duchenne muscular dystrophy

In DMD, impaired muscle regeneration is linked to disturbed production of myogenic progenitor cells. Importantly, dystrophin is not only a structural protein in muscle fibers but is also present in muscle stem cells, known as satellite cells (mSCs), characterized as CD45⁻CD31⁻α7-integrin⁺Sca-1⁻ cells that express Pax7, a key marker essential for their survival and proper function [26]. Dumont et al. [27] demonstrated that dystrophin plays a critical role in regulating the polarity and asymmetric division of mSCs. In regenerating healthy muscle, activated mSCs undergo both symmetric cell division to maintain the muscle stem cell pool and asymmetric cell division to give rise to myogenic progenitor cells that expand and contribute to efficient muscle repair and regeneration. In dystrophic skeletal muscle cells, symmetric expansion dominates over asymmetric cell division, resulting in a reduced pool of myogenic progenitor cells and impaired muscle regeneration. Interestingly, although altered, both increased [28–31] and decreased [32, 33] SC numbers have been reported in various animal dystrophic models; their functional exhaustion due to telomere shortening, induction of cellular senescence, and suppression of proliferation has been consistently observed [34, 35]. Our studies performed in mdx mice (the commonly used murine DMD model with a point mutation in exon 23 of the Dmd gene) suggest that the compromised regenerative potential is not primarily due to a decreased abundance of mSCs, as we found a higher number of Pax7⁺ cells in dystrophic muscles [36–38]. However, in D2-mdx mice, which demonstrate more severe muscle damage, the number of satellite cells is lower than in healthy DBA counterparts [32]. Moreover, several studies, including ours, indicated that murine mSCs isolated from mdx mice [39] or human embryonic stem cell (hESC)-derived mSCs, as reported by Chal et al. [40], present disturbed differentiation. Additional pathological features of dystrophic mSCs include mitochondrial abnormalities evidenced by atypical morphology [41, 42], altered mitochondrial gene expression, and significantly diminished respiratory function [43]. Granet et al. [32] found reduced expression of autophagy-related genes in mSCs isolated from mdx and D2-mdx mice, and demonstrated that treatment with an autophagy-promoting peptide enhanced the differentiation capacity of DMD progenitor cells. Notably, analysis of single-nuclei human DMD transcriptomic data also revealed dysregulated autophagy pathways in diseased mSCs [32], underscoring the universal role of impaired autophagy in their dysfunction. All of these defects may result in elevated oxidative stress, higher production of reactive oxygen species (ROS), reduced ATP generation, and changes in overall cellular metabolism. Notably, dystrophin restoration using CRISPR/Cas9 editing has been shown to improve the energetic properties of dystrophic progenitor cells [44]. In addition, DMD pathology is associated with the premature onset of senescence of mSCs, which may contribute to their impaired regenerative capacity [45, 46]. Moreover, dystrophic mSCs, due to elevated transforming growth factor beta (TGFβ) signaling, may display enhanced fibrogenic activity at the expense of myogenic potential [47]. Such misguided differentiation reflects changes in cellular plasticity, resulting in excessive connective tissue formation and a more pro-fibrotic environment in affected muscles. Finally, as mSCs rely on tightly regulated epigenetic mechanisms and signaling pathways to maintain optimal function, the lack of dystrophin may disrupt these regulatory networks, resulting in regenerative failure of dystrophic muscle. It should be emphasized, however, that the primary cause of the disease is the absence of dystrophin in muscles, which leads to fiber degeneration. Satellite cell dysfunction, described here briefly, summarized in Fig. 2, and comprehensively discussed in several recent review papers [48, 49], may contribute to disease progression over time.

Fig. 2.

Satellite cell dysfunction in DMD. Dystrophic satellite cells exhibit a range of dysfunctions, including reduced asymmetric division, increased senescence, disrupted autophagy, impaired epigenetic regulation of gene expression, altered cell signaling and metabolism, and mitochondrial dysfunction. Dystrophin deficiency also leads to satellite cells’ impaired differentiation and dysregulated plasticity, ultimately promoting a fibrotic rather than myogenic lineage commitment

Taking all factors into consideration, restoring the proper function of mSCs appears to be crucial. Some studies suggest that treatment with various pharmacological compounds that modify signaling pathways may, for example, correct impaired mSC polarity (for references, see [48, 49]). However, a key unresolved technical question regarding potential DMD treatments remains the effectiveness of gene therapy, specifically, whether mSCs can be successfully transduced, and if so, what the long-term impact of such transduction is on their function.

Treatment of Duchenne muscular dystrophy

Despite enormous efforts in basic research and a better understanding of the molecular mechanisms, effective therapies for DMD remain elusive. Current therapeutic strategies can be divided into three approaches (Fig. 3). The commonly used pharmacological treatment (Table 1, with detailed references) only mitigates the downstream effects of dystrophin deficiency, like inflammation, fibrosis, and oxidative stress. The gold standard drugs, used for more than 30 years, glucocorticoids (prednisone, prednisolone, and deflazacort), through reducing inflammatory response by inhibition of the NK-κB pathway, can prolong the ambulation of patients, increase muscle strength, slow the progression of scoliosis, and improve cardiovascular functions [50–53]. Despite these undeniable benefits on cardiac and skeletal muscle performance, their long-term daily use is associated with a high risk of several side effects, including weight gain, delayed growth, osteoporosis, adrenal insufficiency, behavioral changes, and gastrointestinal complications [54]. Some studies suggest that administering glucocorticoids less frequently, such as twice a week instead of daily, may be equally effective while causing fewer adverse effects [52, 55]. More recently, vamorolone (VBP-15), a synthetic anti-inflammatory steroid and mineralocorticoid receptor antagonist, has been shown to offer similar benefits with a more favorable side-effect profile. In vivo studies on mdx dystrophic mice, along with clinical trials, have demonstrated a positive effect from vamorolone administration [54, 56]. Given the encouraging results from Phase 1 and 2 trials, the Food and Drug Administration (FDA) registered this drug in October 2023, followed by approval from the European Medicines Agency (EMA) in December 2023. The Phase 3 VISION-DMD trial (NCT03439670, ClinicalTrials.gov) further confirmed the drug’s efficacy, as well as its favorable safety and tolerability profile over a 48-week treatment period. Moreover, cardiac abnormalities are, at least to some extent, delayed by treatment with angiotensin-converting enzyme inhibitors (ACEi), aldosterone receptor blockers (ARBs), β-adrenergic receptor (β-AR) blockers, and mineralocorticoid receptor antagonists (MRAs). Also, modulators of utrophin (the paralogue of dystrophin) or inhibitors of histone deacetylases are being tested (reviewed in [11]). The ongoing research also explores the therapeutic potential of gasotransmitters, including hydrogen sulfide (H₂S) [57–61], in mitigating disease progression (see Table 1 for additional information and references).

Fig. 3.

Treatment of DMD. The key treatment strategies employed to manage the disease include pharmacological therapies, mostly glucocorticosteroid treatment to slow muscle degeneration; genetic therapies to restore micro-, mini-, or full-length dystrophin and microutrophin; and cell therapies with muscle satellite cells, myoblasts, and mesoangioblasts to regenerate muscle tissue. Among these strategies, future therapies are focused on the use of therapeutic gasotransmitters, like hydrogen sulfide, the CRISPR/Cas9 system for gene editing, and induced pluripotent stem cell-derived muscle cells or cardiomyocytes for cell-based approaches

Table 1.

Examples of Pharmacological DMD therapies, used in patients or tested in animal models and/or in humans. Drugs officially approved and registered for the treatment of DMD are indicated in bold

Type of drug/compound	Exemplary drug/compound	Mechanism of action	References
glucocorticoids	prednisone prednisolone deflazacort	inhibition of the NK-κB pathway and reducing the inflammatory response	[50–53]
synthetic steroids	vamorolone	anti-inflammatory; fewer side effects due to selective mineralocorticoid receptor antagonism	[54, 56]
PDE5 inhibitors	sildenafil tadalafil	inhibiting of cGMP phosphodiesterase (PDE5) activity and prolongation of the biological half-life of cGMP	[52, 170]
ACEi angiotensin-converting enzyme inhibitors	perindopril enalapril captopril	inhibition of Ang II formation and metabolism	[21, 171–173]
ARBs angiotensin receptor blockers	losartan	competitive inhibition of angiotensin II (Ang II); binding to the angiotensin 1 receptor	[21, 174, 175]
β-AR beta-adrenergic receptor	bisoprolol metoprolol carvedilol	inhibition of β-adrenergic receptor (selective or nonselective)	[176–178]
MRAs mineralocorticoid receptor antagonists	eplerenone spironolactone	blocking the endogenous mineralocorticoid, aldosterone, at its receptors	[179–181]
utrophin upregulation	ezutromid	an aryl hydrocarbon receptor (AhR) antagonist; failed in clinical trials	[182, 183]
HDACi histone deacetylase inhibitors	givinostat suberoylanilide hydroxamic acid (SAHA)	inhibition of histone deacetylase activity	[184]
H 2S donors	sodium hydrosulfide (NaHS) sodium sulfide (Na2S) GYY4137	an increase in hydrogen sulfide levels results in anti-inflammatory, antioxidant, anti-fibrotic, and proangiogenic effects (tested in animal DMD models)	[57, 61, 185–187]

Although pharmacological therapies can improve the quality of life of DMD patients and slow disease progression, they do not target the underlying cause of the disorder. In contrast, this could be achieved with the use of genetic therapies, such as read-through therapy, an antisense oligonucleotide (ASO)-mediated exon skipping, vector-mediated gene therapy to restore expression of a shorter form of dystrophin (micro- or mini-dystrophin), or microutrophin and utrophin, the paralogue of dystrophin, expressed during fetal development, and then largely replaced by dystrophin in mature muscle fibers [62, 63], and CRISPR/Cas9-based gene editing (Fig. 3) [64, 65]. It has to be underlined that delivery of the full-length dystrophin gene is challenging because of the large size of its cDNA (~ 11.2 kb protein-coding sequence [66, 67]), which prevents its packaging to adeno-associated viral (AAV) vectors commonly used for muscle cell transduction, able to carry only ~ 4.7 kb transgene [68]. However, innovative methods based on the split-intein technique have recently offered a new way to overcome such limitations and produce full-length dystrophin [69–71].

Finally, cell-based therapies may present a promising avenue for treating DMD (Fig. 3). This approach involves delivering cells capable of expressing dystrophin and could be considered a form of gene therapy, where the cell itself serves as the vector. However, expectations for cell therapy extend beyond gene replacement, as these cells are anticipated not only to express dystrophin but also to differentiate into functional muscle fibers, thereby fully restoring muscle integrity.

Cell therapies for Duchenne muscular dystrophy

Over the years, various cell types have been considered for DMD cell therapy, including some that lack muscle-regenerative potential. However, given the progressive muscle degeneration characteristic of DMD, only cells capable of differentiating into skeletal muscle have the potential to provide meaningful therapeutic benefits.

As mentioned above, the CD45⁻CD31⁻α7-integrin⁺Sca-1⁻ mSCs are bona fide muscle stem cells that differentiate into myoblasts and ultimately form muscle fibers (for a review and references, see [72]). These cells, characterized by the expression of the transcription factor Pax7 and other markers, depending on the state of myogenic activity, are naturally present in skeletal muscles and the diaphragm, making them an obvious candidate for DMD cell therapy [73]. However, dystrophic satellite cells already have impaired functions ([27]; for references see [48, 49]), and when differentiated into muscle, they would not express dystrophin. Accordingly, satellite cell-based therapy requires either the use of allogeneic cells or genetic modification of the patient’s satellite cells to re-express dystrophin.

Cell therapy for DMD must overcome several barriers (Fig. 4). First, isolating and expanding a sufficient number of undifferentiated muscle progenitors capable of repopulating all damaged muscles remains difficult. Second, in the case of the allogeneic approach, overcoming genetic differences between the donor cells and the patient requires immunosuppression. Third, for autologous transplantation, the restoration of dystrophin expression in mSCs of DMD patients would be necessary. However, fourth, the potential immune response against dystrophin has to be carefully considered. Fifth, efficient cell therapy must ensure the high and long-term engraftment of cells across multiple muscles, enabling not only fiber regeneration but also the establishment of a functional satellite cell pool to support future muscle repair. To achieve widespread cell engraftment, systemic delivery appears to be the most logical strategy. Nevertheless, a major limitation is that myoblasts are unable to cross the blood vessel barrier (for a review, see [72]). While local injection into the muscle was tested and demonstrated to be feasible, it only allows for the correction of limited areas within small, superficial muscles [74].

Fig. 4.

Challenges and promises of cell therapies for DMD. Although cell therapies hold considerable promise, they are associated with numerous challenges (as indicated in the upper part of the figure). Among the various well-justified approaches, the use of muscle satellite cells, myoblasts, mesoangioblasts, and pluripotent stem cell-derived muscle cells and cardiomyocytes is accompanied by serious limitations (depicted in the lower part of the figure and discussed in the text)

Satellite cell-based approaches

Transplantation of healthy mSCs has been tested in animal models of muscle injury, demonstrating the feasibility of this approach [75–80]. However, these experiments have also highlighted several limitations. First, the isolation of human mSCs requires an invasive muscle biopsy. Second, the long-term culture of such mSCs results in the loss of their regenerative potential, as the cells undergo senescence [77, 78, 81, 82]. Third, the mSCs isolated from healthy donors are immunologically mismatched with potential recipients, raising concerns about immune rejection. Nevertheless, animal studies have demonstrated the feasibility of their intramuscular application for muscle progenitor/stem cell therapy [83]. The fusion of healthy myoblasts with dystrophin-deficient fibers in mdx mice has been demonstrated [84], generating hope for a similar effect in patients [85]. While the initial, nonrandomized trial of myoblast transfer for DMD therapy claimed some positive effects [86, 87], the overall results were negative [88], indicating a lack of long-term myoblast engraftment and minimal dystrophin expression in the injected muscles [89–91]. Despite being administered in large numbers (exceeding 100 million cells) and through multiple injections, myoblasts failed to engraft effectively due to immune rejection (in the absence of immunosuppression), limited migration beyond the injection site, and their differentiated state.

Intravenous delivery of various stem cells, including mSCs/myoblasts, is ineffective, as the cells become trapped in the lungs and fail to cross the blood vessel wall to reach the muscles [92]. The intra-arterial injection has thus been tested as an alternative; however, its efficacy remains low, with only about 3.2% of injected cells detected in the analyzed muscle Sect. [93]. In the latter study performed in macaques, intra-arterial delivery of β-galactosidase-labeled myoblasts resulted in cell retention in vessels and inefficient muscle penetration. Moreover, even when cells are delivered intramuscularly, they do not survive well (for a review, see [94]). Recently, Saleh et al. [95] demonstrated that human skeletal muscle progenitor cells (isolated from 18-week human fetal muscle) cannot be efficiently systemically delivered into the muscle of immunodeficient dystrophic mice. Importantly, the intravenous delivery was ineffective, while intra-arterial administration allowed limited myoblast extravasation into muscle tissue; however, the efficacy was too low to achieve robust long-term engraftment. The cells have been mainly detected inside blood vessels, and the prominent clotting of these cells was noted in the dystrophic mdxD2-NSG (immunodeficient, to prevent rejection) mice [95]. This again indicates that the systemic delivery of myoblasts, as proposed by some approaches, is unjustified, in light of this and many previous studies.

Mesoangioblasts and similar cells

Pericytes associated with vascular walls in the human skeletal muscle have been demonstrated to differentiate into muscle fibers when injected into murine skeletal muscles. These vessel-associated progenitors are referred to as mesoangioblasts [96–98] and have been the focus of preclinical and initial clinical studies. The potential benefit of these and similar cells [99] has also been linked to their claimed ability to leave the blood vessels after systemic injection. Their potential has been initially tested in mouse [98] and dog [97] models of DMD. In mice, mesoangioblasts derived from mdx animals were corrected using human artificial chromosome-based restoration of the full dystrophin gene [98], while in golden retriever muscular dystrophy (GRMD) dogs, both corrected mesoangioblasts and wild-type cells were injected [97]. Interestingly, autologous mesoangioblasts genetically modified to express dystrophin were less efficient in treating dystrophic dogs than allogeneic ones [97]. Based on these studies, a clinical trial was conducted in five DMD patients who received four intra-arterial injections of HLA-matched mesoangioblasts at 2-month intervals (isolated from the muscles of healthy siblings) [100]. Tacrolimus was administered for immunosuppression, and muscle biopsies analyzed two months after the last injection revealed only low Levels of donor DNA in 4 out of 5 patients. Dystrophin expression was detected in only one case. These results suggest that the efficacy of systemic injection of these cells is quite low and requires substantial improvement. A more detailed review of the mesoangioblast-based approaches has been recently published [101].

The potential of other cell types to enter the muscles after systemic injection was also demonstrated in canine models. Allogeneic muscle stem cells (MuStem cells), distinct from Pax3/Pax7 satellite cells and exhibiting a heterogeneous phenotype (predominantly CD56⁺), were injected systemically (intra-arterially) into immunosuppressed GRMD dogs, resulting in myofiber regeneration, satellite cell replenishment, and improved physical properties of the treated animals [102]. The long-term dystrophin expression was evident, but it was lower than after intramuscular injection. In another study, the same group showed that vascular delivery of allogeneic stem cells in dystrophic dogs requires only short-term immunosuppression, which they ascribed to the low expression of MHC class I antigen [103] as well as the cells’ ability to repress the T-cell proliferation and cytotoxicity [104], a finding observed in human MuStem cells in vitro as well.

Finally, a similar potential in promoting muscle regeneration has been demonstrated for human CD56⁺ MuStem cells transplanted into injured mouse muscles [99]. However, the origin of these cells and their relation to satellite cells is unclear, as they do not express Pax7. If these human myogenic cells represent the same population as canine MuStem cells, which were shown to be effective after intra-arterial delivery [102], they may correspond to mesoangioblast-like cells. Recently, human MuStem cells have been demonstrated to fuse with the nonhuman primate myofibers [105]; however, clinical trials with these cells have not yet been performed.

Pluripotent stem cell-based therapies

In light of the various characteristics of different and numerous cell types used in experimental therapies for DMD (see table S1 in [99]) and the challenges of translating such studies to humans, it may be reasonable to rely on cell types that can be clearly and unequivocally characterized. The application of pluripotent stem cells (PSCs), either embryonic stem cells (ESCs) or induced pluripotent stem cells (iPSCs), represents an attractive option, as they can be efficiently differentiated into muscle cells through genetic overexpression of Pax3/Pax7 [106–108] or small molecule-driven protocols [40, 109, 110] (Table 2). However, this will require overcoming the technical and medical issues associated with these cells.

Table 2.

Examples of pluripotent cell-based approaches to correct dystrophin deficiency in animal models of Duchenne muscular dystrophy

Pluripotent cell	Mode of differentiation	Animal model of DMD	Number and phenotype of cells applied for therapy	Effect after transplantation	References
hESC	Embryoid body differentiation & DOX-induced Pax3 expression	mdx mice; tacrolimus for immunosuppression	1.5 × 106 cells, sorted for PDGF-αR+/Flk-1−; intramuscular (TA) and systemic (i.v. and i.a) injection	Engraftment, also after systemic injection, although to a lesser extent (note: non-sorted Pax3-induced cells formed teratomas)	[106]
hESC	Culture in myogenic medium followed by transduction with adenoviral MyoD vector	Rag/mdx mice	5 × 105 MyoD+ cells into the muscles	Myofibers expressed dystrophin	[188]
hESC hiPSC	DOX-induced Pax7 expression	NSG-mdx4Cv mice	5 × 105 Pax7+ myogenic progenitors	Persistent engraftment for 11 months after transplantations and repopulation of the satellite cell compartment	[189]
mESC	Serum-free protocol (GSK3β inhibitor: CHIR99021, BMP inhibitor: LDN193189, IGF-1, HGF, FGF-2)	Rag1−/−Dmdmdx−5Cv mice	Pax7-GFP cells (artificial chromosome); 1 × 105 into the TA muscle	Foci of dystrophin+ and MyHC+fibers visible one month after injection; Pax7+ satellite-like cells detected under the basal lamina of GFP+ fibers	[40]
hiPSC	Transgene-free or MyoD expression and correction of DMD mutation (CRISPR/Cas9 knockout of exon 45–55)	NSG-mdx mice	1 × 106 NCAM + HNK1− cells injected intramuscularly	Few donor-derived dystrophin-positive myoblasts demonstrated engraftment; efficacy improved in the subsequent study of this group [190]	[191]
hiPSC	CHIR99021-DAPT culture: GSK3β inhibition followed by Notch signalling inhibition; Genetic restoration of dystrophin by artificial chromosome	NSG-mdx4Cv mice NOD-SCID-IL2rγnull-mdx4Cv mice (immunodeficient mice lacking dystrophin)	1–3 × 106 CD57+ (NCAM+) HNK1−cells	Contribution to myofibers	[192]
hESC hiPSC	DOX-induced Pax7 expression (as in [189])	NSG-mdx4Cv mice	5 × 105 CD54, α9β1 integrin, and syndecan 2-purified Pax7+ progenitors	Efficient engraftment	[193]
hESC	Serum-free protocol	NSG-mdx4Cv mice	3 × 105 CD10+CD24− myogenic progenitors into CTX-injured muscles	12% donor-derived satellite cell engraftment	[117]
hPSC	Serum-free protocols (two modes compared)	NSG-mdx mice	1 × 106 ERRB3+ or NGFR+ myogenic progenitors (dystrophin expression restored by CRISPR/Cas9 correction)	Enrichment for ERBB3 and NGFR enriched for Pax7+ fetal and PSC-derived cells and improved engraftment of PSC; PCS-derived muscle progenitors less effective than CD45−CD31−NCAM+ fetal satellite cells (PSC-muscle progenitors did not fuse with the host fibers)	[190]
hiPSC	Transgene-free differentiation	NSG-mdx mice	1–2 × 106 MYF5+ cells; muscles were not CTX-treated	Engraftment; Functional recovery of DMD muscle at 6 weeks after injection	[113]
hiPSC	Transgene-free differentiation	NOG-mdx mice	1 × 105 NGFR+ cells	Better engraftment than negative cells	[118]
hiPSC	Serum-free protocol	NSG-mdx4Cv mice	1 × 106 Pax7:GFP cells	Long-term regenerative potential and maturation of the cells upon in vivo engraftment; In the non-mdx model, the regenerative potential in the secondary recipient demonstrated	[110]
hESC hiPSC	DOX-induced Pax7 expression, integration into safe harbor locus	NSG-mdx4Cv mice	1 × 106 CD54 and syndecan 2 -purified Pax7+ progenitors	Efficient engraftment (2 months)	[194]
hiPSC	DOX-induced Pax7 expression (as in [194])	NSG-mdx mice	1.65 × 106 iPax7+ cells injected into the TA and quadriceps	Long-term (2 months) engraftment; transcriptional maturation	[120]
hiPSC	Pax7 lentiviral overexpression; correction of mutation by CRISPR/Cas9 knock-in to connect exon 44 to exon 58	NSG-mdx4Cv mice	1 × 106 Pax7+ myogenic progenitors transplanted into irradiated TA muscle	Donor-derived myofibers produced	[119]
Teratoma-derived myogenic progenitors	Formation of teratoma in the TA muscle for 4 weeks; isolation of myogenic progenitors	NSG-mdx4Cv mice	2–4 × 105 α7+ VCAM+ myogenic progenitors isolated from teratomas (for teratoma formation, 2.5 × 105 ESCs or 1 × 106 were injected into the TA muscle)	Very efficient engraftment (~ 80%) of the regenerating TA muscle; Matured after transplantation into quiescent muscle stem cells	[115]
Teratoma-derived myogenic progenitors	In vitro expanded progenitors obtained as in [115]; nine orders of magnitude expansion for 37 days	NSG-mdx4Cv mice	4 × 105 α7+ VCAM+ myogenic progenitors isolated from teratomas (for teratoma formation, 2.5 × 105 ESCs were injected into the TA muscle)	Efficient engraftment (40%); functional improvement of the transplanted muscles	[116]

Abbreviations: BMP bone morphogenetic protein, CTX cardiotoxin, DOX doxycycline, FGF-2 fibroblast growth factor-2, GSK3β glycogen synthase kinase-3 beta, hESC human embryonic stem cells, HGF hepatocyte growth factor, iPAX7 doxycycline (dox) inducible expression of PAX7, hiPSC human induced pluripotent stem cells, i.a. intra-arterial, IGF-1 insulin-like growth factor-1, i.v. intravenous, mESC murine embryonic stem cells, NGFR nerve growth factor receptor, PDGF-αRplatelet-derived growth factor α receptor, TA tibialis anterior, VCAM-1 vascular cell adhesion protein 1, α7 α7-integrin

In their initial study, Perlingeiro’s group demonstrated that ESC-derived platelet-derived growth factor alpha receptor-positive and Flk-1-negative cells (PDGF-αR⁺/Flk-1⁻) (hence not endothelial type) possess muscle regenerative potential [106]. Such cells, when injected intramuscularly or systemically (intravenously or intra-arterially) into dystrophic mice, engrafted into the muscles and improved their contractile potential. Subsequently, in 2013, the same group conducted a feasibility study demonstrating the muscle-regenerating potential of iPSC-derived cells in a murine DMD model. In that work, Filareto et al. [111] generated iPSCs from fibroblasts of mice lacking both dystrophin and utrophin, and then they restored the microutrophin (µUtrn) expression, a short version of utrophin, using the Sleeping Beauty transposon system. Modified iPSCs were differentiated into PDGF-αR⁺ skeletal muscle progenitors, which, when transplanted into dystrophic mice, engrafted into the muscle, generating a large number of utrophin-positive myofibers and improving contractile strength. The authors also reported the presence of utrophin-positive fibers in the skeletal muscles and occasionally in the diaphragm after systemic (intravenous) injection of the PSC-derived muscle progenitors, but it was undetected in the hearts of the injected mice [111]. This might indicate that such progenitors can leave blood vessels and enter the muscle. However, the efficacy of that has not been evaluated. Importantly, before cell administration, the mice were also injected with the cardiotoxin to induce muscle injury, a condition that may have facilitated engraftment, as shown previously [102].

The application of PSC-derived muscle progenitors has also been demonstrated in several other studies [40, 109, 112–117], (see also Table 2 for additional references). However, the methods of differentiation show various efficacies, and some of the proposed modalities rely on gene transfer-based overexpression of myogenic factors. Similar to the use of primary satellite cells (described above), the long-term culture of muscle progenitors may limit their therapeutic efficacy. Therefore, future application of PSC-derived myogenic progenitors may require the selection of the proper progenitor cell population to ensure an efficient method for propagating a sufficient number of cells for the therapy. This may be particularly crucial for human progenitor cells, as it has been demonstrated that mSCs lose engraftment capacity after long-term culture. A recent study suggested that CDH13 (T-cadherin or H-cadherin) and fibroblast growth factor receptor 4 (FGFR4) could serve as markers for human myogenic cell purification, with these cells contributing to muscle regeneration in mdx mice [118]. Additionally, Zhao et al. [113] found that human iPSC-derived Myf5⁺ cells, when injected intramuscularly into immunodeficient mdx mice, recovered dystrophin expression and improved muscular function.

A recent proof-of-principle study by Dhoke et al. [119] demonstrated the therapeutic potential of patient-specific iPSCs. The authors generated iPSCs from two individuals carrying mutations in exons 45 and 51 of the DMD gene and successfully corrected these mutations using the CRISPR/Cas9 system. The gene-corrected iPSCs were then differentiated into myogenic progenitors and transplanted into the muscles of immunodeficient dystrophic NSG-mdx^4cv mice, resulting in successful engraftment and dystrophin expression [119]. However, functional improvements in muscle performance were not reported. Notably, iPSC-derived Pax7⁺ progenitors have been shown to undergo transcriptional maturation following transplantation into both dystrophic and non-dystrophic muscle environments [120]. Importantly, the human muscle progenitor Pax7-expressing cells, generated by chemical-driven differentiation, efficiently engrafted into the damaged muscles of NSG-mdx^4cv mice [110], improving the exercise capacity of the mice in the treadmill assay and contributing to the quiescent satellite cell population. Such in vivo matured satellite cells demonstrated the ability to regenerate muscle upon reinjury and to repopulate the muscle niche in secondary recipient mice [110].

Importantly, this study [110] provides proof of principle for the therapeutic potential of iPSC-derived mSCs, which can mature efficiently upon in vivo engraftment, in contrast to in vitro conditions. However, challenges remain in generating a large number of these cells and achieving effective long-term engraftment, particularly after systemic delivery, which remains inefficient for this type of muscle progenitor. Recently, the FDA approved the investigational use of iPSC-derived myogenic progenitors in non-ambulant DMD patients [121], using cells generated through a newly published protocol [108].

Interestingly, Kyba’s team [115] proposed an alternative strategy for generating large numbers of progenitor cells. The authors injected PSCs into the mouse muscle, which resulted in the formation of teratomas. Then, the α7integrin⁺VCAM-1⁺ myogenic progenitors have been FACS-purified from such teratomas, and following injection, these cells demonstrated high engraftment in injured muscle, ameliorating muscular dystrophy [115]. Importantly, such progenitors contributed to the generation of the quiescent Pax7⁺ satellite cells, which responded to subsequent injuries by regenerating muscles. In their follow-up study, the authors successfully propagated these progenitors in vitro, thereby expanding the therapeutic potential of the proposed method while preserving the regenerative capacity of the Pax7⁺-teratoma-derived cells [116].

Recently, the same approach has been tested for the human teratoma-derived progenitors [122], achieving successful in vitro expansion of the CD82⁺ERBB3⁺NGFR⁺ skeletal myogenic progenitor population. Of note, the expression of CD82 (tetraspanin) could serve as a prospective marker for isolating human satellite cells [123]. The results proved the feasibility of testing iPSC-derived muscle progenitors in DMD patients; however, clinical studies have not yet been initiated due to technical constraints and safety concerns. Progress could be expected if the efficient differentiation of progenitors with stable regenerative properties is achieved with minimal genetic manipulation, ideally with transgene-free protocols. In recent years, several such protocols have been proposed and tested [124, 125]. However, the potential of these cells remains to be determined, particularly regarding their stability and survival [126]. A crucial issue will be achieving therapeutic success after systemic delivery, as intramuscular injections can provide only localized improvement, if any. It remains to be established whether PSC-derived muscle progenitors can effectively exit the bloodstream and engraft into the muscle tissue. As noted, such properties are absent for human myoblasts but have been suggested for mesoangioblasts ([97]; for a review, see [101, 127]).

The studies showing the successful engraftment of PSC-derived muscle progenitors were preceded and likely facilitated by muscle irradiation and cardiotoxin injury [122]. Such an approach would be unacceptable in humans, and it is uncertain to what extent the endogenous injury caused by the lack of dystrophin would facilitate cell homing in DMD patients. PSCs may also enable the investigation of the systemic delivery of the myogenic precursors. Through appropriate selection, treatment, or genetic modification, it may be possible to identify cells expressing specific markers that promote or improve their passage through blood vessels into the muscle. In line with such a possibility, Choi et al. [128] recently showed that treatment of CD56⁺ myogenic progenitors with DLL4 and PDGF-BB improves the migration of hiPSC-derived myogenic progenitors.

Other cell types

In addition to satellite cells, vessel-associated mesangioblasts, and PSC-derived myoblasts, several other cell types have been proposed as potential therapies for DMD [127]. These applications are often based on claims that these cells can differentiate into muscle. However, it is crucial to critically evaluate the scientific rationale behind the studies suggesting the therapeutic efficacy of so-called mesenchymal stem/stromal cells (MSCs) and chimeric cells to determine their validity.

Mesenchymal “stem” cells

A widespread misconception has arisen from mesenchymal stromal cell transplantations, commonly but inaccurately referred to as mesenchymal stem cells (hence both abbreviated as MSCs). These cells have been asserted to possess multilineage differentiation potential, enabling them to repair muscle, heart, and other tissues across various diseases. This claim is based on the belief that MSCs undergo differentiation into numerous cell types, including skeletal muscle cells or cardiomyocytes, thereby offering therapeutic avenues for muscular diseases. However, while bone marrow-derived MSCs are capable of differentiating into adipocytes, chondrocytes, and osteoblasts, this does not extend automatically to differentiation into other cell types. Various stromal cells from different organs demonstrate tissue-specific differentiation capacity (for references see: [129, 130]). Their claimed wide differentiation capabilities have been questioned and often attributed to experimental artifacts [129–131]. Some studies have even claimed that allogeneic, adipose tissue-derived MSCs were therapeutically effective in immunocompetent mdx mice [132] and that these cells could persist for several months in non-immunosuppressed GRMD dogs after local or systemic administration [133]. However, these findings have not been reproduced, and their translation to human patients remains highly questionable. The variability in disease progression among DMD dogs, along with concerns about the quality and controls of these studies, raises doubts about the effectiveness of this approach. It has to be stressed that the perpetuated claims on muscle-differentiation properties of various MSCs have been disproven in a landmark study by Paulo Bianco’s group. This research demonstrated that CD146⁺ stromal cells isolated from human muscle could differentiate into myoblasts in a rigorous in vivo assay, while this was not the case for bone marrow, periosteum, or cord blood-derived CD146⁺ cells [134]. This study highlighted that tissue-specific stem/progenitor cells (generally referred to as MSCs) are restricted to differentiating into cell types native to their tissue of origin.

Analysis of outcomes following the use of other “popular” cells, such as umbilical cord Wharton’s jelly-derived cells, also named MSCs, indicates their ineffectiveness in treating conditions like DMD, as discussed in a recent review [135]. Surprisingly, studies on this approach are continued, and it is being promoted as a potential pathway for conditional drug approval.

In sum, the so-called MSCs isolated from several tissues (bone marrow, adipose, Wharton’s jelly of the umbilical cord, teeth, etc.) represent different stromal-cell types with various origins. Their stem cell properties are at best disputable, as such claims are based on assays prone to artifacts (for references, see [129, 134, 136–138]). Unfortunately, these cells are commonly used and even offered commercially as an unregulated and unproven treatment for numerous conditions (for references, see [136–138]).

Chimeric cells

An even more unjustified example of a questionable strategy is the use of so-called chimeric cells. This approach will be examined in greater detail, particularly because it recently received orphan drug designation from the FDA despite unclear justification and well-founded criticism [139].

The concept of using chimeric cells is presented in several papers from one research group, and a clinical trial has also been initiated [140]. According to this strategy, the patient’s myoblasts (lacking dystrophin) are combined (“fused”) in the laboratory with the father’s dystrophin-expressing myoblasts, forming so-called dystrophin-expressing chimeric (DEC) cells, which are then administered intraosseously into the patient’s bones. Proponents of this approach claim that this unusual, uncommon, and invasive route of cell delivery is designed to induce immunological tolerance to the father’s antigens, potentially including dystrophin. However, a detailed review of the published experimental work in animal models, which, according to the authors, proved the feasibility of the approach, fails to substantiate any of their proposed hypotheses.

The concept of using chimeric cells is presented in several papers from one research group, and a clinical trial has also been initiated [140]. According to this strategy, the patient’s myoblasts (lacking dystrophin) are combined (“fused”) in the laboratory with the father’s dystrophin-expressing myoblasts, forming so-called dystrophin-expressing chimeric (DEC) cells, which are then administered intraosseously into the patient’s bones. Proponents of this approach claim that this unusual, uncommon, and invasive route of cell delivery is designed to induce immunological tolerance to the father’s antigens, potentially including dystrophin. However, a detailed review of the published experimental work in animal models, which, according to the authors, proved the feasibility of the approach, fails to substantiate any of their proposed hypotheses.

First, the initial studies were conducted in systems that inherently prevent cell rejection. For example, mdx mice received myoblasts from genetically identical healthy strains, or immunodeficient SCID/mdx mice were transplanted with human myoblasts, which are not rejected in these hosts. Second, and more importantly, these studies did not show the presence of injected cells in the circulation following intrabone administration, an essential step if subsequent muscle engraftment were to occur [140–142]. Third, even if myoblasts could enter the circulation from the bone marrow (which has not been demonstrated), numerous studies have evidenced that they do not migrate from the vasculature into muscle tissue (as discussed above).

The presence of such chimeric myoblasts in the muscles or heart, alongside the reported functional improvements, raises serious concerns. Recently, the administration of chimeric cells to three patients - two six-year-old walking boys and one 15-year-old boy using a wheelchair was reported [140]. The authors claimed that six months after the intraosseous administration of chimeric myoblasts, some symptoms showed signs of improvement. However, the open, non-randomized nature of the studies, which does not account for any accidental changes that may occur in patients, does not allow these results to be considered conclusive evidence of a therapeutic effect.

Moreover, the unproven validity of the hypothesis of administering myoblasts intraosseously, i.e., injecting cells into a site where they do not naturally reside, raises serious methodological and safety concerns. The concept of such invasive treatments, which may create the illusion of potential effectiveness, also brings up ethical issues. Both experimental and clinical applications of this approach have recently been scrutinized by experts in the field. A comprehensive analysis, published on the website of the DMD patients organization, World Duchenne, warns patients about this scientifically, medically, and ethically questionable strategy (see recent comment by Aartsma-Rus [139]).

Unfortunately, some researchers continue to pursue and promote the discredited effects of these cells. This raises ethical concerns about such claims, especially given their lack of scientific validity, methodological rigor, and demonstrable benefit to patients. Generating hype and offering false hope in the absence of a credible therapeutic effect can ultimately be more harmful than providing no intervention at all.

Cell therapies for the treatment of DMD-related cardiomyopathy

The heart does not contain cardiac stem cells. Although various cell-based therapies have been proposed and even clinically tested in patients with heart failure, in light of the negative results (described above), the rationale and justification currently rely on the application of PSCs, which are differentiated into cardiomyocytes before being injected into the heart. However, some other modalities have been suggested in the pursuit of improving heart function. As in skeletal muscles, it has been proposed that the small vessels of the juvenile mouse ventricle contain mesoangioblasts [143]. Aorta-derived mesoangioblasts, characterized by the expression of Nkx2.4, cardiac troponin I, tropomyosin, and α-actin, have been injected into the hearts of young mdx mice [144]. The authors reported that these cells appeared to differentiate into dystrophin-expressing cardiomyocytes and could enhance blood vessel formation, as evidenced by the higher expression of CD31. However, the approach was inefficient in the fibrotic hearts of aged mdx mice, resulting in no functional improvement [144]. Nevertheless, the properties of mesoangioblasts and their contribution to heart repair raise concerns, particularly in light of other studies indicating the lack of progenitor cells in the adult heart.

Cardiosphere cells, isolated from the heart, are considered a type of stromal progenitor cell with claimed immunomodulatory, antifibrotic, and regenerative actions in DMD (for references, see [145]). The clinical studies were based on preclinical work, where mdx mice, systemically injected with these cells, demonstrated improvements in cardiac and skeletal muscle structure and function [145]. These beneficial effects were ascribed to the exosomes [146], which, when secreted from cardiospheres, are claimed to reprogram macrophages and fibroblasts and to affect endogenous cardiac progenitors. A recent study demonstrated that both mouse cardiosphere-derived cells (mCDCs) and extracellular vesicles (EVs) secreted by human cardiosphere-derived cells (hCDC-EVs) exerted a long-term effect on DMD pathology in mdx mice [147]. When young (8-week-old) mice were injected with mCDCs once monthly over a one-year study period, muscle function and structure were preserved, protecting against functional decline even at 12 months of age. In contrast, the efficacy of hCDC-EVs declined after six months of treatment, likely due to the activation of an immune response that resulted in their immunological clearance and a corresponding reduction in therapeutic benefit [147].

These findings prompted the initiation of clinical trials. In the HOPE-Duchenne trial (NCT02485938, ClinicalTrials.gov), 25 non-ambulant patients with a mean age of 17.8 years were randomized to receive an intracoronary infusion of CDCs (CAP-1002) (13 patients) or serve as controls (12 patients) [148]. MRI performed 12 months after treatment revealed significant scar reduction, improvements in inferior wall systolic thickening, and enhanced mid-distal upper limb performance [148]. In the HOPE-2 multicenter Phase 2 trial (NCT03406780, ClinicalTrials.gov), mostly non-ambulant patients were randomized. Eight patients (seven non-ambulant and one in the late ambulant stage) were administered intravenous injection of allogeneic CDCs, derived from donor hearts. Each patient received 1.5 × 108 cells, once every three months, for a total of four infusions, with a 12-month follow-up.

A total of twelve patients were randomized to receive a placebo; however, six of them were excluded from the final analysis due to screening failures. The study concluded that patients treated with CAP-1002 showed stabilization of upper limb function and improvements in cardiac function and structure [149]. However, the study was subsequently discontinued by the sponsoring company, Capricor Therapeutics, with the claims of financial issues. Despite reporting positive outcomes, the approach was associated with serious adverse events, such as strong hypersensitivity reactions in three patients within the treatment group. This led to the protocol modification, requiring all patients to receive glucocorticoids, as well as H1 and H2 blockers [150]. The authors stated, based on previous preclinical studies, that CDCs do not express MHC class II surface antigens or the costimulatory molecules CD80 and CD86. However, the occurrence of hypersensitivity reactions raises concerns about the safety of this approach. Furthermore, the mechanisms underlying the observed cardiac benefits remain unclear, aside from the acclaimed paracrine effect, which has not, however, been conclusively proven. The proposed exosome-mediated profits are still speculative and poorly understood, leaving their true mechanisms of action uncertain. Despite these challenges, recent reports from Capricor Therapeutics suggest that a follow-up study indicates improvements in cardiac function in cardiosphere-treated patients. Based on these findings, the company planned to seek FDA approval [151], however, in July, the FDA rejected the application and did not grant registration [152]. A subsequent trial is currently ongoing (HOPE-3, NCT05126758; ClinicalTrials.org), evaluating the safety and efficacy of allogeneic hCDCs, now referred to as deramiocel, in patients with advanced-stage DMD.

It can be presumed that the PSC-based approaches developed for treating heart failure, which have also been recently tested in humans [153], could potentially be applied in DMD patients. PSCs are the only cells that can be effectively differentiated into cardiomyocytes. The transplantation of either ESC- or iPSC-derived cardiomyocytes has been tested in various animal models of heart failure, including studies in non-human primates (for reviews, see [154–157]). However, applying cardiac cell therapy to DMD patients may be particularly challenging in the advanced stages of the disease. A critical barrier is the overall health status of DMD patients, who are generally considered ineligible for heart transplantations. The same challenges could restrict the application of cell therapy in DMD hearts.

An additional issue limiting the efficacy of cell therapies in DMD is the persistence of the damaged cells in the muscles and heart, where the therapeutic cells are intended to be injected. It is important to note, that to date, the only effective cell therapies require ablation of diseased cells, as is the case in hematopoietic stem cell (HSC) transplantation for blood diseases, or removal of damaged cells, as in burns or treatment of damaged corneal epithelium to provide a favorable environment for the therapeutic cells to take hold and regenerate tissue effectively (for a review, see [138]). The application of cell therapies to severely damaged skeletal and cardiac muscles exposes transplanted cells to unfavorable environments, limiting their survival and therapeutic efficacy. Therefore, combining cell-based approaches with anti-inflammatory and anti-fibrotic treatments emerges as a rational strategy.

Cell therapies for other forms of muscular therapies

DMD represents the most common and remains the most studied form of muscular dystrophy, yet other genetically inherited muscle disorders, such as Becker muscular dystrophy (BMD), limb-girdle muscular dystrophies (LGMD), facioscapulohumeral dystrophy (FSHD), oculopharyngeal muscular dystrophy (OPMD) and congenital forms like LAMA2-related muscular dystrophy, also present significant unmet medical needs. While these diseases share core features like progressive muscle weakness, yet differ markedly in their genetic origins (with the exception of BMD and DMD, which are both caused by mutations in the DMD gene), age of onset, severity, and progression. This diversity presents a considerable challenge in developing universally applicable therapeutic strategies. Among emerging treatments, cell therapy holds promise across multiple dystrophy types. However, most efforts remain at the preclinical stage.

In the experimental approaches (with exemplary strategies for LGMD and FSHD summarized in Table 3), various cell types were used to replace the damaged muscles. Each disease represents specific and different challenges from DMD (except for BMD, which is, in fact, usually a milder form of DMD), and it is beyond the scope of this paper to discuss them in detail. Recent reviews highlighted emerging therapies in various muscular dystrophies have been published [158, 159]. Readers seeking more in-depth information are encouraged to consult these review papers and the original studies referenced in Table 3.

Table 3.

Examples of cell replacement therapies for muscular dystrophies other than DMD in experimental animal models

Dystrophy type	Subtype/ abbreviation*	Mutation (gene)	Cell type used for therapy	Model	Therapeutic effect	References
Limb girdle muscular dystrophy	LGMDR9 FKRP-related (LGMD2I)	FKRP	FKRP L276IKI-mouse derived satellite cells; lentiviral expression of WT human FKRP gene	FKRP L276IKImice	Recovery of α-DG glycosylation; improvement of muscle strength	[195]
	LGMDR9 FKRP-related (LGMD2I)	FKRP	mESC- and hiPSC-derived myogenic progenitors	FKRP P448L-NSG mice	Enhanced specific force in transplanted muscle	[196]
	LGMDR9 FKRP-related (LGMD2I)	FKRP	CRISPR-Cas9 corrected LGMDR9 iPSC-derived progenitors	FKRP P448L-NSG mice	Myofiber and satellite cell engraftment, restoration of α-DG functional glycosylation	[197]
	LGMDR3 α-sarcoglycan-related (LGMD2D)	SGCA	Patient-iPSC-derived-mesoangioblasts; lentiviral restoration of α-sarcoglycan	Sgca-null/scid/beige mice (dystrophic, immune-deficient triple mutant)	Functional amelioration of dystrophic phenotype, restoration of depleted progenitors	[198]
	LGMDR1 calpain3-related (LGMD2A)	CAPN3	CRISPR-Cas9 corrected LGMDR1 iPSC-derived progenitors	Immunodeficient CAPN3-KO mice	Muscle engraftment, rescue of CAPN3 mRNA	[160]
	LGMDR1 calpain3-related (LGMD2A)	CAPN3	CRISPR/Cas9 corrected LGMDR1 iPSC-derived progenitors	Immunodeficient CAPN3-KO mice	CAPN3 expression evident after cell transplantation	[161]
	LGMDR21 POGLUT-1-related (LGMD2U)	POGLUT1	CRISPR-Cas9 corrected LGMDR21 iPSC-derived progenitors	NSG-immunodeficient WT mice; cardiotoxin injury of muscle	Rescue of impaired engraftment exhibited by mutated cells	[199]
	LGMDR2 dysferlin-related (LGMD2B)	DYSF	CRISPR-edited murine primary muscle stem cells	hEx44mut mice (humanized mouse model with Dysf exon 44 with the founder frameshift mutation; autologous therapy)	Regeneration of muscles; repopulation of the muscle stem cell niche	[200]
	LGMD R2 dysferlin-related (LGMD2B)	DYSF	Healthy human and mouse myoblasts	SCID and SJL mice (dysferlinopathy model)	Dysferlin-positive fibers detected in muscle sections; the therapeutic effect limited by poor cell migration	[201]
Facioscapulohumeral muscular dystrophy	FSHD	DUX4 overexpression due to defective epigenetic repression	Mouse ESC-derived muscle progenitors with an induced low level of DUX4	iDUX4pA-HSA mice (doxycycline-induced DUX4 expression)	Reduced fibrosis and improved contractile force with myogenic progenitors expressing a low level of DUX4	[163]
	FSHD	DUX4 overexpression due to defective epigenetic repression	Mouse ESC-derived myogenic progenitors	FRG1 transgenic mice	Functional improvement in muscles of treated male mice (more severely affected than females)	[202]

Abbreviations:CAPN3 calpain 3, DYSF dysferlin,FRG1 FSHD Region Gene 1, FRKP fukutin-related protein, hiPSC human induced pluripotent stem cells, mESC murine embryonic stem cells, POGLUT1 O-glucosyltransferase 1, SGCA  α-sarcoglycan

New and old (in brackets) LGMD subtype nomenclature was given according to Straub et al. [203], and subsequent reports

Based on the experimental studies, a few clinical trials have been recently proposed. The bASKet trial (NCT05588401, ClinicalTrials.gov) is a Phase I/IIa study evaluating the safety of CRISPR/Cas9-corrected autologous satellite cells (GenPHSats) over a 12-month monitoring period in six LGMD patients. The idea is based on the initial works by Selvaraj et al. [160] and Müthel et al. [161], demonstrating the utility of the CRISPR/Cas9 approach to correct mutations in the calpain-3 (CAPN3), the leading cause of the LGMDR1 (Table 3). A similar strategy, VTA-100 by Vita Therapeutics, targeted CAPN3 mutations was cancelled in 2024 due to financial constraints, despite promising preclinical results. The company has since shifted focus to VTA-200, an allogeneic, hypoimmunogenic satellite cell therapy, aiming for broader, off-the-shelf application [162]. It also aims to develop universal iPSC-based therapy for FSHD, targeting double homeobox 4 (DUX4)-driven toxicity. Supporting this strategy, Azzag et al. [163] (Table 3) demonstrated that DUX4-silenced iPSC-derived myogenic cells reduced fibrosis and improved disease features in an FSHD mouse model.

Encouraging clinical results have been reported for OPMD, owing to the localized nature of the disease. In this form of dystrophy, caused by mutations in the PABPN1 gene, the pathology primarily affects a limited set of muscles, including the levator palpebrae, pharyngeal muscles, and proximal limb muscles. This restricted muscle involvement presents a unique therapeutic opportunity: autologous genetically normal myoblasts can be harvested from healthy and functionally intact muscles and locally transplanted into diseased areas without requiring genetic modification or immunosuppressive therapy. This approach was validated by Perié et al. [164], who conducted a phase I/IIa clinical study (NCT00773227, ClinicalTrials.gov) involving 12 OPMD patients. In this study, autologous myoblasts, isolated from unaffected quadriceps or sternocleidomastoid muscles, were injected into the pharyngeal muscles. Over a 2-year follow-up, no adverse effects were reported, while an improvement in the quality of life score was found for all 12 patients, and a cell dose-dependent improvement in swallowing function was observed for 10 patients. However, no recent data are available.

Future directions

Cellular therapies are regarded as potential interventions to address the pathological mechanisms of DMD. However, unlike the well-established application of cell therapy in hematological disorders, their systemic efficacy in muscle diseases remains limited due to the widespread and extensive nature of the affected tissues. Notably, the volume of the tissue requiring treatment is enormous, presenting a significant logistical and biological challenge. Moreover, the success of HSC therapy is partly attributed to the removal (ablation) of diseased cells before transplantation, a step that is not feasible in DMD. In contrast, cell therapies for DMD are introduced into a hostile microenvironment characterized by chronic inflammation, fibrosis, and ongoing muscle fiber necrosis in both skeletal and cardiac tissues. These conditions severely constrain the therapeutic potential of transplanted cells, making them markedly less effective than HSC transplantation following myeloablation.

A promising future strategy to target cardiac fibrosis may involve chimeric antigen receptor-T cell (CAR-T) technology, which has demonstrated preclinical efficacy in models of hypertensive cardiomyopathy and potential utility for cardiac repair [165–167]. CAR-T targeting fibroblast activation protein (FAP), currently considered for treating solid tumors [168], may represent a future direction in DMD therapy. To our knowledge, such a scenario has not been extensively (or almost has not been) explored in the context of DMD-related cardiomyopathy, except for the initial work by Marigny et al. [169] (available only in abstract form), who reported the beneficial effects of FAP-CAR-T cell therapy in lymphodepleted D2.mdx mice. The potential benefits of combining cell therapy with anti-inflammatory and anti-fibrotic approaches in other mouse models and DMD patients remain to be experimentally validated.

Authors’ contributions

A.Ł., J.D. Conceptualization; A.Ł., J.D. Writing and editing; A.Ł. Figure conceptualization and drawing; A.Ł., J.D. Funding acquisition.

Funding

Supported by the grants from the National Science Center: MAESTRO #2018/30/A/NZ3/00412; SHENG2 #2021/40/Q/NZ3/00165; DAINA #2024/52/L/NZ3/00142 to JD, and OPUS #2019/35/B/NZ3/02817 to AŁ.

Data availability

No datasets were generated or analyzed for this manuscript.

Declarations

Competing interests

The authors have no relevant financial or non-financial interests to disclose.