Muscle: an independent contributor to the neuromuscular spinal muscular atrophy disease phenotype

Spinal muscular atrophy (SMA) is a pediatric-onset neuromuscular disorder caused by insufficient survival motor neuron (SMN) protein. SMN restorative therapies are now approved for the treatment of SMA; however, they are not curative, likely due to a combination of imperfect treatment timing, inadequate SMN augmentation, and failure to optimally target relevant organs. Here, we consider the implications of imperfect treatment administration, focusing specifically on outcomes for skeletal muscle. We examine the evidence that muscle plays a contributing role in driving neuromuscular dysfunction in SMA. Next, we discuss how SMN might regulate the health of myofibers and their progenitors. Finally, we speculate on therapeutic outcomes of failing to raise muscle SMN to healthful levels and present strategies to restore function to this tissue to ensure better treatment results.

SMA is commonly referred to as a spinal motor neuron disease (16). Consequently, muscle atrophy in the disease is often presumed to derive primarily from spinal motor neuron degeneration and denervation. Yet for clinicians and scientists familiar with the disease, and notwithstanding the neurodegenerative aspect of the condition, there has always been suspicion of a primary muscle defect in SMA. Indeed, such speculation dates as far back as the 1950s, when mild SMA was initially described (5). Patients with this relatively benign form of SMA were initially considered to have muscular dystrophy of the limb-girdle type with, among features associated with the latter condition, hypertrophy of the calf muscles. Other notable reports implicating a primary muscular component in SMA include one published in 1980 by Dutch scientists describing hypertrophy of calf muscles in patients with mild SMA that was accompanied by abnormally high levels of serum creatine kinase (CK) (17) — a marker of muscle breakdown. A study in the 1970s identified muscle pathology, including disorganization of the myofibrils, sarcomeres, and filaments, in patients with severe as well as mild SMA (18). More recently, rare instances of individuals with homozygous SMN1 deletions have been reported to exhibit myopathic phenotypes rather than the classical neurogenic abnormalities associated with SMA (19, 20). While these more recent studies benefited from genetic confirmation of a diagnosis of SMA, the older studies described above were unable to rely on tests of SMN1 integrity. Instead, SMA diagnoses were based on biopsies, overt phenotypes, and clinical presentation.

With the identification of the genetic cause of SMA in 1995, studies investigating a role for skeletal muscle in driving the neuromuscular SMA phenotype have become more refined. Nevertheless, those that relied on human muscle biopsies obtained prior to the identification of the SMN1 gene may not, in every instance, have genotyped patient samples before utilizing them for investigation. Still, the experimental outcomes of these studies are thought-provoking and largely support a primary muscle defect in SMA. Notable among these are several that employed nerve and muscle cocultures to ascertain the disease-triggering effects of SMA muscle. These experiments revealed that when cultured with rat embryonic spinal cord explants, muscle cells from type I and type II, but not type III, SMA patients triggered rapid degeneration of the cocultures (21, 22). Interestingly, such degeneration was not observed when muscle cells from patients with distinct neurodegenerative diseases (ALS) or myopathies (DMD, nemaline myopathy, mitochondrial myopathies) were employed and, furthermore, did not appear to involve a soluble neurotrophic or neurotoxic factor (23).

Follow-up cell culture studies from the same group detected intrinsic defects of myogenesis in type I SMA. These defects, characterized by impaired fusion of type I SMA myoblasts into myotubes, were not observed with cells from patients with intermediate or mild SMA (24). Moreover, myogenesis defects occurred despite normal myoblast proliferation but were accompanied by reduced levels of nicotinic acetylcholine receptors in myotubes.

Notions of a discernible disease-causing effect of low SMN in muscle have been bolstered, reiterated, and further refined in the last two decades with the use of cultured cells, invertebrate models, and numerous independent lines of model mice. Several studies are worth highlighting. For instance, consistent with the existence of myoblast fusion defects in type I SMA patients, two studies — one employing C2C12 myoblast cells and one using satellite cells from a mouse model of severe SMA — reported perturbed myogenesis as assessed by quantification of multinucleate myotube formation (25, 26). In each instance, the muscle cells were purified and cultured without CNS tissue, diminishing any confounding influence of SMA motor neurons. These studies were accompanied by evaluation of myogenic factor expression in purified myoblasts and intact muscle from SMA model mice (26–28). The studies concluded unanimously that myogenic factor expression is perturbed under conditions of low SMN. However, the pattern of disrupted myogenic factor expression differed somewhat among the various studies. Whereas one (28) reported reduced Pax7, MyoD, and myogenin expression in muscle from symptomatic mutants of a commonly employed SMA mouse model that expresses two SMN2 copies in an Smn-null background (29), another employing SV40 large T antigen–transformed myoblasts from the same line of mice demonstrated reduced Pax7 expression accompanied by an increase in MyoD and myogenin (27). Interestingly, muscle tissue extracted from a second, symptomatic intermediate SMA mouse model exhibited markedly elevated expression of all three myogenic proteins (28).

Notwithstanding these somewhat disparate findings, investigators explored how low SMN disrupts myogenesis. In the first of two elegant studies, it was shown that the myogenic program is activated prematurely in the most severe form of SMA but subsequently stalls as myofiber formation is initiated (26). In the second, the molecular basis of myoblast fusion defects was investigated and led to the discovery that two fusogenic factors — myomaker and myomixer — are reduced in severe SMA model mice and in C2C12 cells exhibiting modest (~55%) knockdown of SMN (30). Restoration of SMN in C2C12 myoblasts raised myomaker levels. Intriguingly, however, myomixer levels remained unchanged, and myogenic programming was not fully rescued. Still, AAV9-mediated overexpression of myomixer mitigated disease in a mouse model of SMA, suggesting that this factor links SMN to muscle pathology in SMA.

Additional evidence in support of a primary myopathy in SMA stems from investigations in invertebrate and mammalian models of the disease. In a fly model of SMA, morphology of the thoracic muscles was profoundly altered, and the authors of this study went on to show that SMN interacts and colocalizes with myofibrillar α-actinin (31). Consistent with the observation in flies, SMN was also shown to appear in perfect register with mouse myofibrillar α-actinin, suggesting a muscle-specific function for SMN protein that is conserved across species. In another study, mice with selective ablation of muscle SMN were reported to develop a severe dystrophic phenotype and died from the disease by approximately 1 month of age (32). While complete SMN ablation is incompatible with the survival of any cell type and does not accurately model human SMA, a more recent study (33) in which SMN was reduced to disease-relevant levels in skeletal muscle also concluded that low protein in this tissue is sufficient to trigger disease. Congruent with these findings, selective restoration of SMN to skeletal muscle of SMA mice restored myofiber size, increased animal weight, improved motor performance, and significantly enhanced life span relative to that of mutants ubiquitously depleted of SMN. Importantly, mice with skeletal muscle SMN restoration continued to exhibit neurodegeneration and loss of synaptic integrity due to persistently low SMN expression in the CNS (34). While these studies support an important role for skeletal muscle in driving the overall neuromuscular SMA phenotype, one report concluded otherwise (35). In this study, a Myf-Cre driver was used to deplete SMN in muscle but did not adversely affect the health of the resulting mice when assessed as young adults. Moreover, in contrast to the above-mentioned study by Martinez et al. (34), a study by Gavrilina et al. reported that selective restoration of SMN to myofibers of severely affected SMA mice failed to mitigate disease (36). The most salient of the various studies cited here are listed along with their main conclusions in Table 1. The chief pathologies associated with low SMN in skeletal muscle are further summarized and rendered as a schematic (Figure 1).

Figure 1

Schematic of the chief cell- and non-cell-autonomous defects arising from low SMN in skeletal muscle tissue. The loss of SMN1 specifically in muscle results in both cell- and non-cell-autonomous effects. In muscle, reduced SMN levels leads to central nucleation of myofibers, altered regeneration, dysfunctional myogenesis, myofiber dystrophy, and impaired acetylcholine receptor (AChR) clustering. In addition, loss of SMN in muscle results in fragmented endplates, neurofilament (NF) accumulation at nerve terminals, increased circulating levels of creatine kinase (CK), altered function of neuromuscular junctions (NMJs), and nerve and muscle degeneration. Figure panels were constructed by the authors from material either generated in the laboratory or created using BioRender.com.

Table 1

Principal studies substantiating an independent role for skeletal muscle in SMA

Considering the evidence for a cell-autonomous role for skeletal muscle in driving SMA pathology, an obvious question centers on whether the myopathy originates in mature myofibers, muscle progenitors, or independently in the two cell types. In historical studies involving human SMA muscle biopsies, defects of the myofibers were the focus of attention and markedly easier to appreciate than possible defects of muscle satellite cells (18). On the other hand, outcomes of in vitro studies that employed myoblasts to detect defects of myogenesis imply defects originating in muscle progenitors (24, 27, 37). Attempts to discern the precise contribution of myofibers versus muscle progenitors to muscle defects in SMA were most directly initiated early this century (32, 38). In these studies, muscle-specific ablation of SMN, in mutants harboring two intact inducible Smn-knockout alleles floxed at exon 7 (Smn^F7/F7), resulted in a discernibly milder phenotype than that observed in related mutants heterozygous for the intact allele (Smn^F7/Δ7). The milder phenotypes of the Smn^F7/F7 mutants, which also harbored a human skeletal actin–Cre (HSA-Cre) driver to inactivate the floxed allele specifically in myofibers, were suggested to be the result of healthy satellite cells, which should have two intact alleles and therefore express WT levels of the SMN protein (38). Satellite cells expressing half the WT level of SMN were presumed to be the reason for the severe phenotype exhibited by Smn^F7/Δ7 mice.

However, a more prosaic explanation for the disparate severities that was not considered is the inherent inefficiency of Cre-mediated inactivation of floxed alleles, and the possibility that mutants with two Smn^F7 alleles merely ended up harboring greater numbers of myonuclei with incomplete allele inactivation. As a result, such mutants would express incrementally higher levels of muscle SMN than Smn^F7/Δ7 mutants, in which inactivation of just one intact floxed allele is required for total muscle SMN ablation.

Notwithstanding the caveat identified in the above study — and the debatable strategy of addressing the tissue-specific requirements for SMN by completely ablating rather than reducing SMN, as in human SMA — subsequent investigations reaffirmed the idea that defective satellite cells contribute to muscle pathology in SMA. Thus, in a mouse model that more accurately mimicked the genetics of human SMA, neonatal mutants had an average number of satellite cells exhibiting normal proliferative potential. However, differentiation of the SMA satellite cells was abnormal, based on the premature expression of myogenic markers, and these cells failed to generate myotubes efficiently (26).

Novel lines of model mice were generated to cement notions of a disease-triggering role for low SMN in skeletal muscle and address questions of the origin of muscle cell–autonomous defects in SMA more accurately (33). Two distinguishing characteristics made these mutants especially useful. First, SMN depletion was specifically targeted to skeletal muscle; and second, low SMN in muscle was nevertheless maintained, as in human SMA, by expression of one or two copies of SMN2. Accordingly, the resulting mutants not only expressed disease-relevant levels of SMN in muscle but also enabled investigation of the effects of depleting the protein in both muscle progenitors and mature myofibers. Several inferences may be drawn from these analyses. First, depletion of SMN in skeletal muscle progenitors is sufficient to trigger disease (Figure 2A). Moreover, the severity of the myopathies and overall phenotypes closely correlated with absolute SMN levels (Figure 2B). Second, examinations of mutants that derived residual muscle SMN from two SMN2 copies hinted at a role for this protein in satellite cells, implying that myopathy in SMA must originate, at least in part, in these progenitors. For instance, it was shown that compared with the modest muscle degeneration observed in mutant animals, a disproportionately large number of myofibers had centrally located nuclei (Figure 2C). Such nuclei are generally associated with regenerating fibers but may also be a consequence of untimely activation — consistent with premature differentiation of these cells (26). Finally, restricting SMN depletion to mature myofibers instead of inducing it in muscle progenitors resulted in milder phenotypes. Still, the eventual appearance of muscle pathology in this last set of mutant mice (HSA-Cre;SMN2;Smn^F7/–) unequivocally assigned a role to SMN in sustaining the health of mature myofibers. Collectively, the various studies cited here suggest that low SMN is damaging to both muscle progenitors and the myofibers that arise from them.

Figure 2

Depletion of SMN specifically in skeletal muscle is sufficient to cause pathology. (A) Confocal transverse section image of the calf muscle of a mutant mouse selectively depleted of the SMN protein in skeletal muscle tissue. Muscle cell–autonomous pathology is observed in the form of degenerating fibers penetrated by circulating IgG (asterisks), infiltrating microglia (arrows) and numerous myofibers containing abnormal, centrally positioned nuclei (arrowheads). Muscle was dual stained with antibodies against Iba-1 and mouse IgG to visualize microglia and damaged myofibers, respectively. Scale bar: 50 μm. (B) Kaplan-Meier survival curves depicting the correlation between SMN2 copies, and thus absolute SMN levels, in muscle and life span of the SMA mutants. P < 0.0001, log-rank test, n ≥ 16 mice of each cohort. (C) Enumeration of degenerating myofibers and cells harboring central nuclei in the gastrocnemius of mutants selectively depleted of SMN in skeletal muscle. Roughly nine times as many SMA fibers were found to display central nuclei compared with those that were degenerating (IgG-positive). *P < 0.05, ** P < 0.01, t tests, n ≥ 300 fibers from n ≥ 3 mice of each cohort. Panels adapted from Kim et al. (33).

If the evidence cited here, which in our view is compelling, is truly reflective of a cell-autonomous role for muscle in driving SMA pathology, it prompts several questions. First, how does SMN maintain the health of myofibers and their progenitors? Second, what are the minimum levels of SMN required by muscle to sustain viability? Third, is the level of SMN necessary to ensure early postnatal muscle growth identical to that required for muscle maintenance? Fourth, do muscles, akin to motor neurons, exhibit differential vulnerabilities to low SMN? Finally, what are the potential repercussions of failing to restore muscle SMN to healthy levels and how can muscle SMN deficiency despite treatment with SMN augmenting agents be overcome? In the remainder of the Review, we attempt to address these biologically and clinically relevant questions.

SMN has been implicated in a multitude of functions (39) and new ones continue to be revealed (40). Yet there is no obvious function, pathway, or set of factor(s) that connect SMN uniquely to the health of muscle. One intuitive means of establishing links between SMN and muscle health involves a careful multiomic analysis of SMA and healthy muscle. In this regard, the newer lines of SMA model mice — which not only vary in severity, but are engineered to preclude the confounding influence of low SMN in neighboring tissue — could be especially useful (33). For example, comparisons of perturbations in mice with muscle-specific SMN depletion to those in mutants with systemically low SMN ought to be instructive and distinguish primary muscle defects from those downstream of motor neuron dysfunction. We advocate for such investigations to be carried out not only on different muscles from the model mice but also — to the extent possible — on patient tissue. Specifically examining muscle progenitors in this manner is also certain to cast novel light on myopathy in SMA. The evidence for a role for SMN in satellite cells is persuasive, yet not unequivocal. These cells are critical not only during early postnatal muscle growth (41) but also as a means of replenishing injured or aged muscle. Indeed, the importance of these cells has been described in several muscle diseases, notably DMD, limb-girdle muscular dystrophy, myopathy associated with dystroglycan dysfunction, Fukuyama congenital muscular dystrophy, and sarcopenia (42–46). Moreover, investigations of the cells in the context of these conditions have cast important light on their characteristics, how they respond to build or replenish lost muscle, whether their activities ameliorate or worsen disease, and the mechanisms that operate within them to balance self-renewal with differentiation (47). These studies have revealed that only a small (10%–20%) proportion of satellite cells, defined by their expression of the transcription factor Pax7, are truly quiescent and able to self-renew. Such self-renewal is critically dependent on the capacity for asymmetric cell division — into a quiescent satellite cell and a second activated myoblast that initiates the myogenic program. The second cell, which continues to express Pax7 in the short term and is therefore frequently attributed the status of a muscle stem cell, engages in symmetric cell division to expand the number of activated myoblasts and, consequently, grow or replace lost muscle. A fine balance exists between asymmetric and symmetric satellite cell division, and one that is regulated by a complex network of factors including signaling from Notch, Jak2/Stat3, Wnt7a, and p38MAPK (48–56). Disruption of this balance exacerbates muscle loss in a variety of myopathies (57). Indeed, gradual satellite cell exhaustion and eventual senescence, owing to protracted cycles of myofiber degeneration and regeneration, in DMD (58, 59) are aggravated by loss of muscle protein in these cells. Dystrophin, for instance, establishes satellite stem cell polarity and thus regulates asymmetric division into quiescent and activated daughter cells (60). Could SMN also influence such pathways, either through orchestration of the splicing cascade and subsequent expression of factors important to muscle function or in some hitherto undescribed manner? Investigation of such questions, particularly in light of reports of premature satellite cell activation in SMA, is a biological and clinical imperative.

Despite a meager understanding of pathways underlying a specific role for SMN in muscle satellite cells, there are some aspects of SMA myopathy that are not only well recognized but also explained by phenomena revealed in other muscular dystrophies. For instance, studies of model mice suggest that levels of SMN typical of type I SMA will likely have a profound and direct effect on muscle (33). In what resembles an all-or-none effect, an incremental increase in the protein significantly mitigates disease severity and muscle pathology.

Thus, model mice expressing SMN from two rather than one SMN2 copy are relatively mildly affected. Early postnatal muscle development is restored, and life span is extended from roughly 2½ weeks in the former mutants to approximately 13 months in the latter. Additionally, there is little evidence of perturbed myogenic factor expression in the mutants with the mild phenotype, and onset of muscle pathology in these mice is considerably delayed. This phenotype suggests, at least in model mice, that two SMN2 copies are sufficient to ensure muscle growth through prepubertal life but fail to maintain myofiber health during adulthood. Moreover, certain muscles, such as the flexor digitorum brevis (FDB), are more vulnerable to low SMN than others (33), with myopathy, in general, likely to be accelerated by sustained muscle activity. Elevated CK values, especially in patients with milder SMA, are suggestive of delayed-onset, activity-driven myopathy (61). Still, studies to better explain how low SMN triggers muscle pathology are urgently needed and not only promise to shed new light on SMN biology but could also prove useful in developing and optimizing novel SMA therapies for patients. Such studies will also be instructive for understanding the differential vulnerability of muscles to low SMN (62, 63) and the extent to which the vulnerability arises in the pre- versus postsynaptic compartments of neuromuscular synapses. Low SMN in muscle appears to have a retrograde effect on nerve terminals, as suggested by evidence of neurofilament accumulation in the terminals of severely affected mutants with skeletal muscle–specific depletion of SMN (33). Whether this pathology eventually results in denervation of the muscle remains to be determined and will have to be assessed in milder mutants with life spans long enough for the process of axonal retraction to run its course. Nevertheless, it is clear that neuromuscular junction (NMJ) function is altered in mice selectively depleted of SMN in muscle, as quantal content, a measure of neurotransmission is perturbed in these mutants (33).

Although low SMN in skeletal muscle is sufficient to damage the tissue and very likely triggers defects not just in myofibers but in muscle stem cells as well, the molecular mechanisms underlying the pathology remain poorly understood. Defining these mechanisms is important not only because SMN, like other muscle proteins such as dystrophin, sustains muscle health, but also because doing so informs treatments for SMA. Nusinersen, the antisense oligonucleotide approved for treatment of the disease, is administered intrathecally and does not increase levels of intact SMN transcript from SMN2 in muscle of treated patients (64).

Onasemnogene abeparvovec, an AAV9-mediated gene replacement therapy for SMA, is delivered systemically and therefore expected to target skeletal muscle. However, treatment with this agent is not without caveats. For instance, it is currently only approved for patients under two years of age. Moreover, as it is a one-time treatment, muscle turnover in treated patients would eventually result in loss of the episomal therapeutic molecule and reversion of transduced cells to an SMA state. Finally, it is unclear whether AAV9 delivers the replacement gene efficiently to nondividing muscle stem cells. AAV6, a related capsid that exhibits robust tropism for muscle, fails to target satellite cells (65), and it is possible that AAV9 displays similar characteristics. Risdiplam — a small-molecule splice-switcher and the most recently approved SMN-augmenting agent to be added to the arsenal of SMA therapies — is reported to modulate SMN2 splicing systemically but has only relatively modest effects on boosting the protein, raising plasma SMN levels in treated types I and II patients by approximately 2.5-fold (66, 67) and muscle SMN of mutant mice by 210% (68). Given the profoundly low (10%–15% of WT) baseline levels of SMN in severely affected patients (64, 69), treatment using currently approved doses of an agent such as risdiplam is unlikely to raise SMN to what are considered the “safe” levels found in asymptomatic carriers. Rather, protein may only be increased to levels (30%–40%) approximating those in patients with type III or type IV SMA (70, 71), which is insufficient to prevent disease. These observations, and empirical data about the requirements for SMN in muscle (33), raise the question: What sort of therapeutic outcome might one expect, particularly in the long term, from current SMA therapies? One potential outcome, notwithstanding an immediate benefit following treatment of the presymptomatic patient, is the evolution of a delayed-onset, chronic myopathy. Onset and severity of such pathology will almost certainly vary and be subject to SMA type, treatment timing, and targeting efficiency.

While the outcome described above is not inevitable, clinicians and families ought to be prepared; therefore, it is prudent to consider adjunct treatments to ensure that the health of muscle in treated patients is sustained. There are several strategies worth considering. One strategy that may be considered targets the TNF-like weak inducer of apoptosis (TWEAK)/TNF receptor superfamily member FGF-inducible 14 (Fn14) pathway. This pathway has been implicated in myriad cellular activities, including muscle protein degradation, atrophy, muscle oxidative metabolism, and myofiber regeneration (72). Upregulation of the pathway following muscle damage suggests that it may be involved in tissue repair. Indeed, in two mouse models of SMA, expression of TWEAK and Fn14 was reduced. Moreover, stimulating the pathway with an Fc-TWEAK agonist modestly mitigated disease in the two models (73). Selective androgen receptor modulators (SARMs) might also be considered in combating muscle dysfunction in SMA. These agents, which mimic the muscle-building anabolic effects of androgens without their pronounced adverse effects — including virilization in women and cardiac and prostatic hypertrophy — were initially developed for the treatment of sarcopenia but have also produced benefit in a rodent DMD model (74–76). The adjunct therapy that has perhaps garnered the greatest interest as a means to combat primary muscle disease involves interfering with myostatin signaling. Myostatin, a member of the TGF-β superfamily, is expressed predominantly in skeletal muscle, where it negatively regulates the growth of the muscle by binding to the type IIB activin receptor (ActRIIB) (77). Blocking myostatin activity is known to increase skeletal muscle mass (78, 79) and was reported to mitigate disease in the mdx mouse model of DMD (80). Accordingly, a number of studies have investigated the effects of suppressing myostatin activity for SMA (81–87). These preclinical studies have produced disparate data, with some claiming benefit and others failing to mitigate disease. Nevertheless, the positive outcomes reported have appeared promising enough to prompt a number of clinical trials (88–90). One of these, which employed apitegromab — a human monoclonal antibody that binds the pro-forms of myostatin, thereby inhibiting its activation — concluded that the agent, when used in combination with nusinersen, synergized with the SMN-augmenting therapeutic molecule to ameliorate disease (91). Despite the range of options presented here, it is unclear to what extent any of the associated pathways truly restore SMN activities or functions to muscle. Without restoring such activities, the various adjunct treatments considered here will be of incremental value. This sobering thought justifies renewed efforts to define precisely how SMN sustains muscle health. Identifying the molecular pathways and the various mediators that govern this process remains the most assured means by which any myopathy in SMA will eventually be addressed in the clinic.

We thank members of the Monani laboratory for comments and suggestions. SMA projects in the laboratory are supported by funding from AFM-France, Cure SMA, the Hope for Children Research Foundation, the NIH (R01NS123292 and R01NS104218 to URM), and a Muscular Dystrophy Association Development Grant to NNJ. Authorship order was decided by the alphabetical order of last names.

Address correspondence to: Umrao R. Monani, P&S, Room 5-422, 630 W. 168th St., New York, New York 10032, USA. Phone: 212.342.5132; Email: um2105@columbia.edu.

Conflict of interest: The Monani laboratory receives research funding from Sarepta Inc.

Copyright: © 2023, Jha et al. This is an open access article published under the terms of the Creative Commons Attribution 4.0 International License.

Reference information: JCI Insight. 2023;8(18):e171878. https://doi.org/10.1172/jci.insight.171878.

1. Werdnig G. Two early infantile hereditary cases of progressive muscular atrophy simulating dystrophy, but on a neural basis. 1891. Arch Neurol. 1971;25(3):276–278.
   View this article via: CrossRef PubMed Google Scholar
2. Hoffmann J. Ueber chronische spinale Muskelatrophie im Kindesalter, auf familiärer Basis. Deutsch Z Nervenheilk. 1893;3:427–470.
   View this article via: CrossRef Google Scholar
3. Hoffmann J. Weiterer Beitrag zur Lehre von der hereditären progressiven spinalen Muskelatrophie im Kindesalter nebst Bemerkungen über den fortschreitenden Muskelschwund im Allgemeinen. Deutsch Z Nervenheilk. 1897;10:292–320.
   View this article via: CrossRef Google Scholar
4. Hoffmann J. Dritter Beitrag zur Lehre von der hereditären progressiven spinalen Muskelatrophie im Kindesalter. Deutsch Z Nervenheilk. 1900;18:217–224.
   View this article via: CrossRef Google Scholar
5. Kugelberg E, Welander L. Heredofamilial juvenile muscular atrophy simulating muscular dystrophy. AMA Arch Neurol Psychiatry. 1956;75(5):500–509.
   View this article via: CrossRef PubMed Google Scholar
6. Thompson J, Bruce A. Progressive muscular atrophy in a child with a spinal lesion. Edinb Hosp Rep. 1893;1:372.
7. Dubowitz V. Infantile muscular atrophy. A prospective study with particular reference to a slowly progressive variety. Brain. 1964;87:707–718.
   View this article via: CrossRef PubMed Google Scholar
8. Clermont O, et al. SMN gene deletions in adult-onset spinal muscular atrophy. Lancet. 1995;346(8991–8992):1712–1713.
   View this article via: CrossRef PubMed Google Scholar
9. Gilliam TC, et al. Genetic homogeneity between acute and chronic forms of spinal muscular atrophy. Nature. 1990;345(6278):823–825.
   View this article via: CrossRef PubMed Google Scholar
10. Lefebvre S, et al. Identification and characterization of a spinal muscular atrophy-determining gene. Cell. 1995;80(1):155–165.
    View this article via: CrossRef PubMed Google Scholar
11. Aartsma-Rus A. FDA approval of nusinersen for spinal muscular atrophy makes 2016 the year of splice modulating oligonucleotides. Nucleic Acid Ther. 2017;27(2):67–69.
    View this article via: CrossRef PubMed Google Scholar
12. Gittings LM, Sattler R. Recent advances in understanding amyotrophic lateral sclerosis and emerging therapies. Fac Rev. 2020;9:12.
    View this article via: CrossRef PubMed Google Scholar
13. Babbs A, et al. From diagnosis to therapy in Duchenne muscular dystrophy. Biochem Soc Trans. 2020;48(3):813–821.
    View this article via: CrossRef PubMed Google Scholar
14. Mongiovi P, et al. Patient reported impact of symptoms in spinal muscular atrophy (PRISM-SMA). Neurology. 2018;91(13):e1206–e1214.
    View this article via: CrossRef PubMed Google Scholar
15. Mix L, et al. Quality of life in SMA patients under treatment with nusinersen. Front Neurol. 2021;12:626787.
    View this article via: CrossRef PubMed Google Scholar
16. Sumner CJ, Fischbeck KH. Spinal muscular atrophy. In: Gilman S, ed. Neurobiology of Disease. Academic Press; 2007:501–511.
17. Bouwsma G, Van Wijngaarden GK. Spinal muscular atrophy and hypertrophy of the calves. J Neurol Sci. 1980;44(2–3):275–279.
    View this article via: CrossRef PubMed Google Scholar
18. Szliwowski HB, Drochmans P. Ultrastructural aspects of muscle and nerve in Werdnig-Hoffmann disease. Acta Neuropathol. 1975;31(4):281–296.
    View this article via: CrossRef PubMed Google Scholar
19. Muqit MM, et al. Phenotypic variability in siblings with type III spinal muscular atrophy. J Neurol Neurosurg Psychiatry. 2004;75(12):1762–1764.
    View this article via: CrossRef PubMed Google Scholar
20. Vajsar J, et al. Congenital cytoplasmic body myopathy with survival motor neuron gene deletion or Werdnig-Hoffmann disease. Neurology. 1998;51(3):873–875.
    View this article via: CrossRef PubMed Google Scholar
21. Braun S, et al. Constitutive muscular abnormalities in culture in spinal muscular atrophy. Lancet. 1995;345(8951):694–695.
    View this article via: CrossRef PubMed Google Scholar
22. Guettier-Sigrist S, et al. Muscle could be the therapeutic target in SMA treatment. J Neurosci Res. 1998;53(6):663–669.
    View this article via: CrossRef PubMed Google Scholar
23. Braun S, et al. Degeneration of cocultures of spinal muscular atrophy muscle cells and rat spinal cord explants is not due to secreted factors and cannot be prevented by neurotrophins. Muscle Nerve. 1997;20(8):953–960.
    View this article via: CrossRef PubMed Google Scholar
24. Arnold AS, et al. Reduced expression of nicotinic AChRs in myotubes from spinal muscular atrophy I patients. Lab Invest. 2004;84(10):1271–1278.
    View this article via: CrossRef PubMed Google Scholar
25. Shafey D, et al. Hypomorphic Smn knockdown C2C12 myoblasts reveal intrinsic defects in myoblast fusion and myotube morphology. Exp Cell Res. 2005;311(1):49–61.
    View this article via: CrossRef PubMed Google Scholar
26. Hayhurst M, et al. A cell-autonomous defect in skeletal muscle satellite cells expressing low levels of survival of motor neuron protein. Dev Biol. 2012;368(2):323–334.
    View this article via: CrossRef PubMed Google Scholar
27. Bricceno KV, et al. Survival motor neuron protein deficiency impairs myotube formation by altering myogenic gene expression and focal adhesion dynamics. Hum Mol Genet. 2014;23(18):4745–4757.
    View this article via: CrossRef PubMed Google Scholar
28. Boyer JG, et al. Myogenic program dysregulation is contributory to disease pathogenesis in spinal muscular atrophy. Hum Mol Genet. 2014;23(16):4249–4259.
    View this article via: CrossRef PubMed Google Scholar
29. Monani UR, et al. The human centromeric survival motor neuron gene (SMN2) rescues embryonic lethality in Smn(-/-) mice and results in a mouse with spinal muscular atrophy. Hum Mol Genet. 2000;9(3):333–339.
    View this article via: CrossRef PubMed Google Scholar
30. McCormack NM, et al. Survival motor neuron deficiency slows myoblast fusion through reduced myomaker and myomixer expression. J Cachexia Sarcopenia Muscle. 2021;12(4):1098–1116.
    View this article via: CrossRef PubMed Google Scholar
31. Rajendra TK, et al. A Drosophila melanogaster model of spinal muscular atrophy reveals a function for SMN in striated muscle. J Cell Biol. 2007;176(6):831–841.
    View this article via: CrossRef PubMed Google Scholar
32. Cifuentes-Diaz C, et al. Deletion of murine SMN exon 7 directed to skeletal muscle leads to severe muscular dystrophy. J Cell Biol. 2001;152(5):1107–1114.
    View this article via: CrossRef PubMed Google Scholar
33. Kim JK, et al. Muscle-specific SMN reduction reveals motor neuron-independent disease in spinal muscular atrophy models. J Clin Invest. 2020;130(3):1271–1287.
    View this article via: JCI CrossRef PubMed Google Scholar
34. Martinez TL, et al. Survival motor neuron protein in motor neurons determines synaptic integrity in spinal muscular atrophy. J Neurosci. 2012;32(25):8703–8715.
    View this article via: CrossRef PubMed Google Scholar
35. Iyer CC, et al. Low levels of survival motor neuron protein are sufficient for normal muscle function in the SMNΔ7 mouse model of SMA. Hum Mol Genet. 2015;24(21):6160–6173.
    View this article via: CrossRef PubMed Google Scholar
36. Gavrilina TO, et al. Neuronal SMN expression corrects spinal muscular atrophy in severe SMA mice while muscle-specific SMN expression has no phenotypic effect. Hum Mol Genet. 2008;17(8):1063–1075.
    View this article via: CrossRef PubMed Google Scholar
37. Guettier-Sigrist S, et al. Possible pathogenic role of muscle cell dysfunction in motor neuron death in spinal muscular atrophy. Muscle Nerve. 2002;25(5):700–708.
    View this article via: CrossRef PubMed Google Scholar
38. Nicole S, et al. Intact satellite cells lead to remarkable protection against Smn gene defect in differentiated skeletal muscle. J Cell Biol. 2003;161(3):571–582.
    View this article via: CrossRef PubMed Google Scholar
39. Monani UR. Spinal muscular atrophy: a deficiency in a ubiquitous protein; a motor neuron-specific disease. Neuron. 2005;48(6):885–896.
    View this article via: CrossRef PubMed Google Scholar
40. Kim JK, et al. A spinal muscular atrophy modifier implicates the SMN protein in SNARE complex assembly at neuromuscular synapses. Neuron. 2023;111(9):1423–1439.
    View this article via: CrossRef PubMed Google Scholar
41. Bachman JF, et al. Prepubertal skeletal muscle growth requires Pax7-expressing satellite cell-derived myonuclear contribution. Development. 2018;145(20):dev167197.
    View this article via: CrossRef PubMed Google Scholar
42. Boyer JG, et al. Depletion of skeletal muscle satellite cells attenuates pathology in muscular dystrophy. Nat Commun. 2022;13(1):2940.
    View this article via: CrossRef PubMed Google Scholar
43. Kudryashova E, et al. Satellite cell senescence underlies myopathy in a mouse model of limb-girdle muscular dystrophy 2H. J Clin Invest. 2012;122(5):1764–1776.
    View this article via: JCI CrossRef PubMed Google Scholar
44. Cohn RD, et al. Disruption of DAG1 in differentiated skeletal muscle reveals a role for dystroglycan in muscle regeneration. Cell. 2002;110(5):639–648.
    View this article via: CrossRef PubMed Google Scholar
45. Kanagawa M, et al. Impaired viability of muscle precursor cells in muscular dystrophy with glycosylation defects and amelioration of its severe phenotype by limited gene expression. Hum Mol Genet. 2013;22(15):3003–3015.
    View this article via: CrossRef PubMed Google Scholar
46. Fry CS, et al. Inducible depletion of satellite cells in adult, sedentary mice impairs muscle regenerative capacity without affecting sarcopenia. Nat Med. 2015;21(1):76–80.
    View this article via: CrossRef PubMed Google Scholar
47. Feige P, et al. Orienting muscle stem cells for regeneration in homeostasis, aging, and disease. Cell Stem Cell. 2018;23(5):653–664.
    View this article via: CrossRef PubMed Google Scholar
48. Conboy IM, Rando TA. The regulation of Notch signaling controls satellite cell activation and cell fate determination in postnatal myogenesis. Dev Cell. 2002;3(3):397–409.
    View this article via: CrossRef PubMed Google Scholar
49. Price FD, et al. Inhibition of JAK-STAT signaling stimulates adult satellite cell function. Nat Med. 2014;20(10):1174–1181.
    View this article via: CrossRef PubMed Google Scholar
50. Tierney MT, et al. STAT3 signaling controls satellite cell expansion and skeletal muscle repair. Nat Med. 2014;20(10):1182–1186.
    View this article via: CrossRef PubMed Google Scholar
51. Troy A, et al. Coordination of satellite cell activation and self-renewal by Par-complex-dependent asymmetric activation of p38α/β MAPK. Cell Stem Cell. 2012;11(4):541–553.
    View this article via: CrossRef PubMed Google Scholar
52. Bernet JD, et al. p38 MAPK signaling underlies a cell-autonomous loss of stem cell self-renewal in skeletal muscle of aged mice. Nat Med. 2014;20(3):265–271.
    View this article via: CrossRef PubMed Google Scholar
53. Nakamura A, et al. Up-regulation of mitogen activated protein kinases in mdx skeletal muscle following chronic treadmill exercise. Biochim Biophys Acta. 2005;1740(3):326–331.
    View this article via: CrossRef PubMed Google Scholar
54. Wissing ER, et al. P38α MAPK underlies muscular dystrophy and myofiber death through a Bax-dependent mechanism. Hum Mol Genet. 2014;23(20):5452–5463.
    View this article via: CrossRef PubMed Google Scholar
55. Le Grand F, et al. Wnt7a activates the planar cell polarity pathway to drive the symmetric expansion of satellite stem cells. Cell Stem Cell. 2009;4(6):535–547.
    View this article via: CrossRef PubMed Google Scholar
56. Bentzinger CF, et al. Fibronectin regulates Wnt7a signaling and satellite cell expansion. Cell Stem Cell. 2013;12(1):75–87.
    View this article via: CrossRef PubMed Google Scholar
57. Dumont NA, Rudnicki MA. Targeting muscle stem cell intrinsic defects to treat Duchenne muscular dystrophy. NPJ Regen Med. 2016;1:16006.
    View this article via: CrossRef PubMed Google Scholar
58. Blau HM, et al. Defective myoblasts identified in Duchenne muscular dystrophy. Proc Natl Acad Sci U S A. 1983;80(15):4856–4860.
    View this article via: CrossRef PubMed Google Scholar
59. Sacco A, et al. Short telomeres and stem cell exhaustion model Duchenne muscular dystrophy in mdx/mTR mice. Cell. 2010;143(7):1059–1071.
    View this article via: CrossRef PubMed Google Scholar
60. Kuang S, et al. Asymmetric self-renewal and commitment of satellite stem cells in muscle. Cell. 2007;129(5):999–1010.
    View this article via: CrossRef PubMed Google Scholar
61. Rudnik-Schöneborn S, et al. Analysis of creatine kinase activity in 504 patients with proximal spinal muscular atrophy types I-III from the point of view of progression and severity. Eur Neurol. 1998;39(3):154–162.
    View this article via: CrossRef PubMed Google Scholar
62. Murray LM, et al. Selective vulnerability of motor neurons and dissociation of pre- and post-synaptic pathology at the neuromuscular junction in mouse models of spinal muscular atrophy. Hum Mol Genet. 2008;17(7):949–962.
    View this article via: CrossRef PubMed Google Scholar
63. Woschitz V, et al. Mouse models of SMA show divergent patterns of neuronal vulnerability and resilience. Skelet Muscle. 2022;12(1):22.
    View this article via: CrossRef PubMed Google Scholar
64. Ramos DM, et al. Age-dependent SMN expression in disease-relevant tissue and implications for SMA treatment. J Clin Invest. 2019;129(11):4817–4831.
    View this article via: JCI CrossRef PubMed Google Scholar
65. Arnett AL, et al. Adeno-associated viral (AAV) vectors do not efficiently target muscle satellite cells. Mol Ther Methods Clin Dev. 2014;1:14038.
    View this article via: CrossRef PubMed Google Scholar
66. Baranello G, et al. Risdiplam in type 1 spinal muscular atrophy. N Engl J Med. 2021;384(10):915–923.
    View this article via: CrossRef PubMed Google Scholar
67. Mercuri E, et al. Clinical studies of RG7916 in patients with spinal muscular atrophy: SUNFISH part 1 study update. Neuromuscul Disord. 2017;27(suppl 2):S209.
68. Poirier A, et al. Risdiplam distributes and increases SMN protein in both the central nervous system and peripheral organs. Pharmacol Res Perspect. 2018;6(6):e00447.
    View this article via: CrossRef PubMed Google Scholar
69. Kobayashi DT, et al. Utility of survival motor neuron ELISA for spinal muscular atrophy clinical and preclinical analyses. PLoS One. 2011;6(8):e24269.
    View this article via: CrossRef PubMed Google Scholar
70. Coovert DD, et al. The survival motor neuron protein in spinal muscular atrophy. Hum Mol Genet. 1997;6(8):1205–1214.
    View this article via: CrossRef PubMed Google Scholar
71. Lefebvre S, et al. Correlation between severity and SMN protein level in spinal muscular atrophy. Nat Genet. 1997;16(3):265–269.
    View this article via: CrossRef PubMed Google Scholar
72. Winkles JA. The TWEAK-Fn14 cytokine-receptor axis: discovery, biology and therapeutic targeting. Nat Rev Drug Discov. 2008;7(5):411–425.
    View this article via: CrossRef PubMed Google Scholar
73. Meijboom KE, et al. Dysregulation of tweak and Fn14 in skeletal muscle of spinal muscular atrophy mice. Skelet Muscle. 2022;12(1):18.
    View this article via: CrossRef PubMed Google Scholar
74. Narayanan R, et al. Development of selective androgen receptor modulators (SARMs). Mol Cell Endocrinol. 2018;465:134–142.
    View this article via: CrossRef PubMed Google Scholar
75. Dalton JT, et al. The selective androgen receptor modulator GTx-024 (enobosarm) improves lean body mass and physical function in healthy elderly men and postmenopausal women: results of a double-blind, placebo-controlled phase II trial. J Cachexia Sarcopenia Muscle. 2011;2(3):153–161.
    View this article via: CrossRef PubMed Google Scholar
76. Cozzoli A, et al. GLPG0492, a novel selective androgen receptor modulator, improves muscle performance in the exercised-mdx mouse model of muscular dystrophy. Pharmacol Res. 2013;72:9–24.
    View this article via: CrossRef PubMed Google Scholar
77. McPherron AC, et al. Regulation of skeletal muscle mass in mice by a new TGF-beta superfamily member. Nature. 1997;387(6628):83–90.
    View this article via: CrossRef PubMed Google Scholar
78. Lee SJ, McPherron AC. Regulation of myostatin activity and muscle growth. Proc Natl Acad Sci U S A. 2001;98(16):9306–9311.
    View this article via: CrossRef PubMed Google Scholar
79. Zhu X, et al. Dominant negative myostatin produces hypertrophy without hyperplasia in muscle. FEBS Lett. 2000;474(1):71–75.
    View this article via: CrossRef PubMed Google Scholar
80. Qiao C, et al. Myostatin propeptide gene delivery by adeno-associated virus serotype 8 vectors enhances muscle growth and ameliorates dystrophic phenotypes in mdx mice. Hum Gene Ther. 2008;19(3):241–254.
    View this article via: CrossRef PubMed Google Scholar
81. Rose FF Jr. , et al. Delivery of recombinant follistatin lessens disease severity in a mouse model of spinal muscular atrophy. Hum Mol Genet. 2009;18(6):997–1005.
    View this article via: CrossRef PubMed Google Scholar
82. Sumner CJ, et al. Inhibition of myostatin does not ameliorate disease features of severe spinal muscular atrophy mice. Hum Mol Genet. 2009;18(17):3145–3152.
    View this article via: CrossRef PubMed Google Scholar
83. Rindt H, et al. Transgenic inactivation of murine myostatin does not decrease the severity of disease in a model of spinal muscular atrophy. Neuromuscul Disord. 2012;22(3):277–285.
    View this article via: CrossRef PubMed Google Scholar
84. Liu M, et al. Activin receptor type IIB inhibition improves muscle phenotype and function in a mouse model of spinal muscular atrophy. PLoS One. 2016;11(11):e0166803.
    View this article via: CrossRef PubMed Google Scholar
85. Long KK, et al. Specific inhibition of myostatin activation is beneficial in mouse models of SMA therapy. Hum Mol Genet. 2019;28(7):1076–1089.
    View this article via: CrossRef PubMed Google Scholar
86. Feng Z, et al. Pharmacologically induced mouse model of adult spinal muscular atrophy to evaluate effectiveness of therapeutics after disease onset. Hum Mol Genet. 2016;25(5):964–975.
    View this article via: CrossRef PubMed Google Scholar
87. Zhou H, et al. Myostatin inhibition in combination with antisense oligonucleotide therapy improves outcomes in spinal muscular atrophy. J Cachexia Sarcopenia Muscle. 2020;11(3):768–782.
    View this article via: CrossRef PubMed Google Scholar
88. ClinicalTrials. A Study to Investigate the Safety and Efficacy of RO7204239 in Combination With Risdiplam (RO7034067) in Participants With Spinal Muscular Atrophy (MANATEE). https://classic.clinicaltrials.gov/ct2/show/NCT05115110 Updated August 3, 2023. Accessed August 21, 2023.
89. ClinicalTrials. A Study to Evaluate the Efficacy and Safety of Taldefgrobep Alfa in Participants With Spinal Muscular Atrophy (RESILIENT). https://classic.clinicaltrials.gov/ct2/show/NCT05337553 Updated August 21, 2023. Accessed August 21, 2023.
90. ClinicalTrials. Efficacy and Safety of Apitegromab in Patients With Later-Onset Spinal Muscular Atrophy Treated With Nusinersen or Risdiplam (SAPPHIRE). https://classic.clinicaltrials.gov/ct2/show/NCT05156320 Updated August 4, 2023. Accessed August 21, 2023.
91. Barrett D, et al. A randomized phase 1 safety, pharmacokinetic and pharmacodynamic study of the novel myostatin inhibitor apitegromab (SRK-015): a potential treatment for spinal muscular atrophy. Adv Ther. 2021;38(6):3203–3222.
    View this article via: CrossRef PubMed Google Scholar