Facioscapulohumeral muscular dystrophy: the road to targeted therapies

Mara S Tihaya; Karlien Mul; Judit Balog; Jessica C de Greef; Stephen J Tapscott; Rabi Tawil; Jeffrey M Statland; Silvère M van der Maarel

doi:10.1038/s41582-022-00762-2

. Author manuscript; available in PMC: 2024 Nov 20.

Published in final edited form as: Nat Rev Neurol. 2023 Jan 10;19(2):91–108. doi: 10.1038/s41582-022-00762-2

Facioscapulohumeral muscular dystrophy: the road to targeted therapies

Mara S Tihaya ^1,⁶, Karlien Mul ^2,⁶, Judit Balog ¹, Jessica C de Greef ¹, Stephen J Tapscott ³, Rabi Tawil ⁴, Jeffrey M Statland ⁵, Silvère M van der Maarel ^1,^✉

PMCID: PMC11578282 NIHMSID: NIHMS2032731 PMID: 36627512

Abstract

Advances in the molecular understanding of facioscapulohumeral muscular dystrophy (FSHD) have revealed that FSHD results from epigenetic de-repression of the DUX4 gene in skeletal muscle, which encodes a transcription factor that is active in early embryonic development but is normally silenced in almost all somatic tissues. These advances also led to the identification of targets for disease-altering therapies for FSHD, as well as an improved understanding of the molecular mechanism of the disease and factors that influence its progression. Together, these developments led the FSHD research community to shift its focus towards the development of disease-modifying treatments for FSHD. This Review presents advances in the molecular and clinical understanding of FSHD, discusses the potential targeted therapies that are currently being explored, some of which are already in clinical trials, and describes progress in the development of FSHD-specific outcome measures and assessment tools for use in future clinical trials.

Introduction

Facioscapulohumeral muscular dystrophy (FSHD) is one of the most common inherited muscle disorders of adulthood and, according to the most recent European epidemiological study of FSHD (published in 2014), has a prevalence of 5–12 affected individuals per 100,000 of the population¹. After intensive research, consensus has been reached that FSHD results from epigenetic de-repression of the DUX4 gene in skeletal muscle. This gene encodes the transcription factor double homoebox 4 (DUX4), which is normally silenced in most somatic tissues^2,3. These advances in understanding of the molecular mechanisms underlying FSHD led to new opportunities for the development of disease-altering therapies and prompted a substantial shift in focus for the FSHD research community towards the development of targeted treatments for FSHD. Academic groups and pharmaceutical companies now have active programmes developing targeted therapies for FSHD, and several clinical trials have already been initiated. In anticipation of additional upcoming trials, several challenges remain to be addressed owing to the rarity, clinical variability and slow progression of FSHD⁴.

This Review presents important advances in the molecular and clinical understanding of FSHD. In particular, we discuss the latest disease-altering therapies in various stages of research, ranging from proof-of-concept in vitro studies to clinical trials of a compound with proven DUX4-blocking effects. We propose a model for future clinical trials of disease-altering treatments for FSHD and discuss the outcome measures that are likely to be most clinically meaningful in the clinical trial setting. We also appraise the level of evidence for a proposed molecular biomarker of DUX4 activity in muscle and the utility of MRI neuroimaging biomarkers.

Clinical phenotype

FSHD is named after its distinctive phenotype, which consists of (often asymmetrical) weakness and atrophy of muscles of the face, shoulder girdle and upper arms⁵. Despite its name, the axial and leg muscles can also be affected. Muscle weakness is slowly progressive over the lifetime of a patient, although many individuals report long periods of disease stabilization interrupted by bursts of disease activity that are associated with rapid functional decline^5,6. Although the pattern of muscle involvement is highly characteristic, the degree of muscle involvement is highly variable both between patients and between different muscles within the same patient. Eventually, approximately 20% of the patients with FSHD become wheelchair-dependent⁷.

The pattern of inheritance is most often autosomal dominant; however, in 10–30% of patients, FSHD is caused by de novo mutations, and individuals carrying FSHD mutations show a high frequency of somatic mosaicism⁸. Disease severity varies widely, even within affected families. Genetic studies reveal that up to 30% of family members carry FSHD mutations but have non-penetrant or minimally affected phenotypes that depend on their D4Z4 repeat length^9–12. FSHD penetrance is unevenly distributed among the sexes. Non-penetrance is reported more often in women than in men^9,13, and women with somatic mosaicism of DUX4-permissive, short D4Z4 repeat alleles were found to be non-affected carriers more often than were their male counterparts, even though the percentage of cells with DUX4-permissive, short D4Z4 repeat alleles was higher in mosaic women than in mosaic men¹⁴. Although some studies^9,10,13,15 report higher clinical disease severity scores in male patients, other large cohort studies did not find a difference in disease severity between the sexes^12,16,17. Contrary to the findings of earlier studies, a 2021 report of findings from a longitudinal study in a large FSHD1 cohort in the USA showed that, independent of genetics, female patients were more likely than male patients to progress to wheelchair use¹⁸. A protective effect of oestrogens in women with FSHD has been suggested on the basis of reports of female patients experiencing accelerated disease progression after receiving oestrogen-blocking therapy that triggered early menopause¹⁹. However, a clinical study of lifetime oestrogen exposure did not show any correlation between the (fairly small) variations in oestrogen exposure and FSHD severity²⁰.

Age at symptom onset varies from infancy to late adulthood, but the disease mostly manifests between the age of 15 and 30 years. Disease onset below the age of 10 years is typically associated with a severe disease course²¹. A subgroup of patients with early-onset FSHD and a fairly consistent presentation defined by severe generalized muscle weakness and a high frequency of extramuscular disease manifestations is often referred to as having ‘infantile’ FSHD^22,23. The extramuscular features of FSHD include sensorineural hearing loss, retinal vasculopathy that sometimes progresses to a pathology resembling Coats disease, an increased prevalence of (incomplete) right bundle branch block, restrictive lung disease and the possibility of cognitive impairment and epilepsy²⁴. In patients with typical age at onset of FSHD, systemic features occur rarely and are mostly subclinical; moreover, involvement of the cardiac and respiratory muscles is rare and life expectancy is generally not reduced.

Currently, no curative or pharmacological treatments are available for FSHD. Treatment is focused on optimizing daily functioning, reducing the burden of pain and fatigue and surveillance for extramuscular complications²⁵. Patients might benefit from the use of orthotic devices such as corsets for back support and leg braces for foot drop²⁶. Patients with FSHD should be encouraged to engage in low-intensity aerobic exercise, which has a beneficial effect on chronic fatigue, physical activity and fitness^27–29.

Disease mechanism

Identification of the FSHD-causing gene

In 1990, genetic linkage studies mapped an FSHD-associated locus to the distal end of chromosome 4q^30,31. This work was soon followed by the discovery of de novo genetic rearrangements in the 4q35 locus in patients with FSHD³². These DNA rearrangements were defined as the partial deletion (shortening or contraction) of a tandem macrosatellite repeat called D4Z4 (ref. 33). Because the D4Z4 repeat itself was initially considered to be non-protein coding³⁴, genes in close proximity to the D4Z4 repeat were among the first to be investigated as FSHD candidate genes^35–38. Monosomy of distal 4q was not associated with FSHD, which excluded a loss-of-function mechanism for the observed mutations. Instead, a position-effect variegation model was considered, in which partial deletion of the D4Z4 repeat was thought to lead to altered cis-spreading of the 4q subtelomere chromatin structure resulting in the altered expression of proximal genes^35,39–41. However, numerous follow-up genetic studies in FSHD families challenged this theory by failing to consistently demonstrate any transcriptional dysregulation of candidate proximal genes in FSHD muscle cells^40,42–46. Indeed, some FSHD families have been identified in which the 4q subtelomere disease locus is physically separated from the so-called proximal genes by a 4;10 translocation, which casts further doubt on their involvement in FSHD pathogenesis⁴⁷.

In healthy individuals, the D4Z4 macrosatellite repeat consists of between 8 and ~100 tandem repeat units of 3.3 kb each³² (Fig. 1). Each D4Z4 tandem repeat unit contains a retrogene that includes the full open reading frame of DUX4^30,31,34,48. The discovery of two equally common distal D4Z4 sequence variants, 4qA and 4qB (ref. 49), was followed by the observation that FSHD is exclusively associated with the 4qA variant^50,51. The critical difference between these two genetic variants became apparent when a DUX4 polyadenylation signal (PAS) was found in the pLAM region³³ flanking the most distal D4Z4 unit, which is specific to the 4qA genetic variant^52,53. This PAS is used by DUX4 in FSHD muscle to stabilize its messenger RNAs (mRNAs) and hence facilitate protein translation. Of note, chromosome 10q also harbours a D4Z4 macrosatellite repeat that shows high sequence homology to the 4q D4Z4 macrosatellite repeat^54,55, but size variations in the 10q D4Z4 repeat are typically not associated with FSHD because this chromosome lacks the DUX4 PAS⁵⁶.

The presence of DUX4 mRNA and DUX4 protein in FSHD muscle cell cultures⁵⁷ and their general absence (or occasionally very low abundance) in muscle cell cultures from symptom-free individuals^57,58 confirmed the hypothesis that inappropriate expression of DUX4 in skeletal muscle causes FSHD². This hypothesis is currently known as the unifying genetic model of FSHD.

Repression of the FSHD locus

The D4Z4 macrosatellite tandem repeat is epigenetically silenced in most somatic tissues of healthy individuals. This tandem repeat is CG-rich and has a higher than average frequency of CpG dinucleotides³⁴, making it susceptible to transcriptional repression by DNA methylation⁵⁹. The D4Z4 repeat is also characterized by other hallmarks of constitutive heterochromatin⁶⁰, such as enrichment of the repressive modification trimethylation of histone H3 lysine 9 (H3K9me3) and the presence of repressive chromatin factors such as heterochromatin protein 1 (HP1) isoforms α and γ and cohesin^61,62. Additionally, various chromatin modifiers have been identified that partake in D4Z4 repression, such as structural maintenance of chromosomes flexible hinge domain containing protein 1 (SMCHD1)^3,63, nucleosome remodelling deacetylase complex, chromatin assembly factor 1 complex⁶⁴, Polycomb repressive complex (PRC) 1 (ref. 65) and PRC2. PRC2 mediates the repressive histone mark trimethylation of histone H3 at lysine 27 (H3K27me3)^62,66. The fact that several different transcriptional repressors are associated with D4Z4 suggests a degree of redundancy and highlights the importance of maintaining repression of this locus in somatic cells.

FSHD types 1 and 2

Two types of FSHD can be distinguished on the basis of the underlying epigenetic mechanism that causes D4Z4 chromatin relaxation and results in DUX4 de-repression in skeletal muscle. In autosomal dominant FSHD type 1 (FSHD1), which is the most common form of FSHD and affects 95% of all patients with this disease, de-repression is caused by partial deletion (also termed contraction) of the D4Z4 repeat on one 4qA allele^32,33. Contraction reduces the number of repeat units to 1–10 (ref. 67), which leads to the partial loss of D4Z4 DNA methylation^68,69 and a reduction in repressive H3K9me3 histone modifications on the affected allele⁶¹. Ultimately, these changes result in some degree of chromatin relaxation, which is sufficient to allow DUX4 transcription to occur in skeletal muscle.

In FSHD type 2 (FSHD2), which accounts for most of the remaining 5% of patients with FSHD, even the shortest D4Z4 allele typically still contains between 9 and 20 tandem repeat units^67,70. Although this number of repeats is within the normal range for healthy individuals, in individuals with FSHD2, D4Z4 chromatin relaxation and DUX4 expression are both caused by a mutation in one of the chromatin repressors that normally participate in silencing of D4Z4. The most common cause of FSHD2 is a heterozygous mutation in the SMCHD1 gene, which accounts for >85% of the patients with FSHD2 (refs. 3,71). A rare cause of FSHD2 is a heterozygous mutation in DNMT3B, which encodes DNA methyltransferase 3β (ref. 72). Interestingly, mutations in these two genes can also cause two clinically unrelated conditions, bosma arhinia microphthalmia syndrome (BAMS) and immunodeficiency, centromeric instability and facial anomalies (ICF) syndrome, discussed subsequently. A homozygous mutation in LRIF1 (encoding ligand-dependent nuclear receptor-interacting factor 1) has also been identified as the cause of FSHD2 in one patient⁷³. The roles of these genes in D4Z4 repression are discussed in more detail in the discussion of genotype–phenotype correlations and disease modifiers.

FSHD types 1 and 2 can co-occur in the same individual when an FSHD1 allele size in the upper end of the range (9–10 D4Z4 tandem repeats) is inherited together with an FSHD2-causing mutation, typically in SMCHD1 (ref. 74). These patients are described as having FSHD1 + 2, which is often characterized by an earlier onset of disease and more rapid progression when compared with patients who carry only one FSHD1 allele of an equivalent D4Z4 tandem repeat size⁶⁷. We therefore propose that, instead of considering FSHD1 and FSHD2 as separate disease entities, they rather form a disease continuum in which the D4Z4 tandem repeat copy number mainly dictates the disease characteristics in the lower end of the range (1–6 tandem repeat units), whereas the function of chromatin modifiers such as SMCHD1 takes an increasingly prominent role with increasing D4Z4 tandem repeat copy number⁶⁷.

Rare forms and unexplained FSHD

Rare cases of FSHD have been attributed to unusual DNA alterations that result in D4Z4 chromatin relaxation and permit DUX4 expression^47,75,76. These alterations include deep intronic mutations in SMCHD1 that affect mRNA splicing⁷⁷, small duplications (expansions) of the D4Z4 macrosatellite repeat with or without co-segregation of an SMCHD1 mutation^76,78 and partial deletions of the D4Z4 macrosatellite repeat region that extend proximally into flanking non-D4Z4 sequences^79,80.

For a small number of patients with FSHD who have a normal-sized but transcription-permissive D4Z4 macrosatellite repeat on a 4qA background, the causative mutation has not been identified. Therefore, other FSHD-related genes might exist that perhaps represent other D4Z4 chromatin modifiers⁸¹.

Role of DUX4 in health and disease

To understand the deleterious effects of DUX4 expression in skeletal muscle, it is important to consider the role of this transcription factor in normal development. DUX4 function has been comprehensively reviewed elsewhere⁸²; thus, this section summarizes some of the most important features of DUX4 in healthy individuals and patients with FSHD.

DUX4 is involved in embryonic genome activation, which occurs during early embryonic development at the cleavage stage and acts as a molecular switch that is required to traverse the stages of normal embryonic development^83–85. DUX4 expression peaks at the two-cell stage in mouse embryos and at the four-cell stage in human embryos, and this peak is closely followed by expression of other genes and repetitive elements that are activated by direct binding of DUX4 to their promoters, in a process known as embryonic genome activation or maternal-to-zygotic transition^83,84. DUX4 is capable of altering the chromatin structural landscape to orchestrate the activation of transcription in otherwise silenced regions through recruitment of p300 and CREB binding protein histone acetyltransferases, which induce global reorganization of H3K27 acetylation⁸⁶. DUX4 also induces the expression and incorporation of histone variants H3.X and H3.Y throughout the bodies of DUX4 target genes, which results in increased relaxation of the chromatin structure at those loci and enhanced target gene perdurance⁸⁷.

DUX4 expression is suppressed in most, but not all, adult somatic tissues. Some evidence supports DUX4 expression in human thymus and skin^60,88, which suggests that DUX4 might have roles in these adult somatic tissues, although these studies have not been replicated. DUX4 is also expressed in testes, where its expression originates from both the 4q and 10q loci. An alternative (further downstream) PAS is present on both of these chromosomes, in addition to the one that is active in FSHD skeletal muscle. This downstream PAS results in a unique (germline-specific) splice form of DUX4 (ref. 58). The exact DUX4 gene structure and PAS used during the zygotic cleavage stage are not known.

Experimental induction of DUX4 expression, whether overexpressed or inducibly expressed, is cytotoxic in various types of somatic cells, although it is unclear which cell death pathway is the main contributor to DUX4-induced cytotoxicity. Evidence suggests that both p53-dependent^89–92 and p53-independent^93,94 apoptotic pathways can be effectors of DUX4-induced cell death. Conversely, in induced pluripotent stem cells derived from patients with FSHD, DUX4 is activated via a primate-specific subtelomeric region that contains a p53 binding site⁹⁵. Therefore, p53 is not only induced by DUX4 but also has been identified as a direct activator of DUX4, which creates a DUX4-inducing feedforward loop. Accordingly, p53 is a potential therapeutic target in FSHD⁹⁵. Moreover, p53 is a well-known tumour suppressor and is also the driver of apoptosis in the DNA damage response (DDR) pathway⁹⁶. Further research to elucidate the mechanisms leading to cell death following DUX4 expression could prove beneficial for additional target discovery.

Myoblasts from patients with FSHD are highly susceptible to oxidative stress⁹⁷. Indeed, in a patient-derived induced pluripotent stem cell model, oxidative stress induced by hydrogen peroxide led to DUX4 expression as a result of serine-protein kinase ataxia-telangiectasia mutated-dependent activation of the DDR pathway and subsequent activation of p53 (ref. 98). Of note, in this study, inhibition of mitogen-activated protein kinase (MAPK) family members — among which p38 MAPK can be activated directly by oxidative stress — did not result in decreased DUX4 activation, indicating that DUX4 activation is not a direct effect of oxidative stress itself, but rather is reliant on other factors, such as DDR⁹⁸. Thus, oxidative stress is not only a consequence of DUX4 expression^97,99 but also acts as an inducer of DUX4 expression, possibly creating another p53-mediated feedforward loop. Indeed, given that p53 is the main inducer of apoptosis, it partakes in any cascade that results in DDR activation⁹⁵.

Direct detection of DUX4 protein in FSHD myonuclei is challenging, because this protein is only transiently present and occurs in very few myonuclei. In many studies, therefore, transcriptional upregulation of DUX4 targets in skeletal muscle is considered indirect evidence of the presence of DUX4 protein. Concurrent measures of RNA and protein levels in DUX4⁺ cells showed that expression of DUX4 target genes was highly induced at both the mRNA and protein levels¹⁰⁰. Indeed, a pilot study has provided direct evidence of DUX4 protein in a muscle biopsy sample from a patient with FSHD using a proximity ligation assay¹⁰¹. DUX4 protein abundance in cultured FSHD myocytes increases with myogenic differentiation, but DUX4 positivity remains sporadic, ranging from 1 in 200 myonuclei¹⁰² to 1 in 1,000 (ref. 58) or 1 in 2,000 (ref. 57) myonuclei. DUX4 protein was also detected in differentiated myocytes from some unaffected family members of FSHD patients, albeit with at a much lower abundance, corresponding to a frequency for DUX4 positivity of approximately 1 in 11,000 myonuclei⁵⁷.

In tissue culture, the typical expression pattern comprises a highly DUX4⁺ nucleus flanked by nuclei with somewhat reduced DUX4 positivity, which suggests that DUX4 positivity spreads to neighbouring nuclei after a transcriptional burst^102,103. Expression of DUX4 target genes follows this same pattern of spread and either occurs coincidentally with DUX4 expression or persists after transient DUX4 activation, where it acts as an indicator of an extinguished DUX4 transcriptional peak in the affected cells⁶⁴.

DUX4 has many downstream target loci that are most probably activated in a dose-dependent manner¹⁰⁴. This circumstance would explain why healthy relatives of patients with FSHD can sometimes show low levels of DUX4 expression that do not result in either transcriptional activation of DUX4 target genes or cytotoxicity⁵⁷. Among the genes most robustly upregulated by DUX4, LEUTX, KHDC1L, TRIM43, PRAMEF2, ZSCAN4 and MBD3L2 expression represent a DUX4 signature of potential utility as a molecular biomarker^{64,84,104,105}. Of note, methyl-CpG-binding domain protein 3-like 2 (encoded by MBD3L2) is involved in a positive feedforward loop that amplifies DUX4 expression and potentially facilitates the spread of DUX4 to neighbouring nuclei within individual myofibres⁶⁴.

Single-cell RNA sequencing (RNA-seq) of primary differentiated mononuclear myocytes identified a small subset of cells with an FSHD-specific transcriptional profile, including expression of DUX4 and/or DUX4 target genes. In combination with pseudotime trajectory modelling, early and late DUX4-responding genes were identified. Identification of these genes enabled the reconstruction of an FSHD cellular progression model in which bursts of DUX4 expression leave a long-lasting ‘scar’ of transcriptionally upregulated DUX4 target genes that remain activated long after the peak in DUX4 expression that triggered this transcriptional deregulation has passed¹⁰⁶. Single-nucleus RNA-seq of differentiating primary FSHD2 myoblasts during a time course of 6 days captured native DUX4 expression and identified a subpopulation of cells that express DUX4 signature genes, including the DUX4 paralogue DUXA. DUXA is implicated in the maintenance of DUX4 target gene expression in late stages of myocyte differentiation, suggesting that DUXA can take over the role of DUX4 and create self-sustaining expression of DUX4 signature genes long after DUX4 itself has disappeared from the nuclei¹⁰⁷.

Correlates and modifiers of disease

D4Z4 repeat size

Variations in the age at onset of FSHD, disease severity and penetrance can be partly explained by genetic and epigenetic factors that act as disease modifiers. In FSHD1, disease severity roughly and inversely correlates to the number of D4Z4 repeat units on the contracted allele, especially in patients carrying short repeats (1–6 D4Z4 units). Patients carrying repeats of 1–3 units have a higher risk of belonging to the ‘early onset’ FSHD group, are more severely affected and have faster disease progression than those with longer repeats²¹. However, a few reports have described individuals with unusual non-penetrant repeats of 1–3 D4Z4 units, suggesting that other genetic elements might also play a part in disease presentation¹⁰⁸. Patients with repeats of 7–10 D4Z4 units are more likely than those with shorter repeats to present with increased clinical variability, lower CpG methylation than expected for their D4Z4 repeat size (given that D4Z4 methylation is dependent on repeat size) and generally also have milder disease severity^109,110. Also, repeats of 7–10 D4Z4 units are more often observed in non-penetrant than in penetrant carriers¹². Consistent with their reduced penetrance, D4Z4 repeats of 7–10 units can be observed in the general European population at a frequency of 1–2%^111,112.

DNA methylation affects D4Z4 regulation

DNA methylation is an important epigenetic disease modifier and regulator of D4Z4. Although one study¹¹³ and a review¹¹⁴ questioned the predictive value and diagnostic relevance of DNA methylation in patients with FSHD, possibly because of technical challenges^113,114, substantial evidence supports D4Z4 hypomethylation as an epigenetic hallmark of FSHD^{59,67,68,110,115–123}. Unlike in FSHD1, in which hypomethylation is restricted to the contracted D4Z4 allele and is considered a consequence of the contraction, in FSHD2 all D4Z4 repeats are hypomethylated^116,117. Therefore, considerable reductions in pan-D4Z4 methylation are observed, especially in patients with FSHD2 (refs. 72,73,124). Moreover, specific CpG sites within D4Z4 are differentially methylated in patients with FSHD versus control individuals¹²⁵. The methylation status of diagnostic region 1, which lies 1 kb upstream of the DUX4 open reading frame, can differentiate muscle samples from patients with FSHD from those of healthy control individuals and could also distinguish between muscle samples from patients with FSHD1 and those from patients with FSHD2 (refs. 59,120–123,125). In addition, CpG methylation analysis of the pLAM region can also be used to detect FSHD-associated hypomethylation, with the advantage that this method offers increased specificity for the 4qA disease allele by preventing co-amplification of the 4qB allele (in 4qA/4qB heterozygous carriers) and also by preventing co-amplification of 10q alleles^69,122. D4Z4 CpG methylation status has been associated with variations in clinical disease severity¹²³: among individuals carrying a D4Z4 repeat of 7–10 units, a greater reduction in CpG methylation was found (corrected for D4Z4 repeat size) only in symptomatic individuals with FSHD1 and not in non-penetrant mutation carriers¹²⁶. Moreover, asymptomatic carriers of a shortened D4Z4 repeat were shown to have methylation levels similar to those of healthy control indivduals¹¹⁸.

SMCHD1 acts as an FSHD modifier

SMCHD1 is a chromatin modifier that has been shown to bind to the D4Z4 repeat, and thereby to repress DUX4, in multiple cell lines and in cultured muscle cells from healthy individuals^3,127,128. Heterozygous nonsense or truncating mutations in SMCHD1 and/or SMCHD1 haploinsufficiency lead to decreased SMCHD1 protein levels in fibroblasts from patients with FSHD³ and a decrease in D4Z4-bound SMCHD1^3,128, and both these changes are associated with pan-D4Z4 hypomethylation and DUX4 de-repression in patients with FSHD2 (refs. 3,127,128). Likewise, heterozygous missense mutations in SMCHD1 result in pan-D4Z4 hypomethylation, DUX4 expression in skeletal muscle and manifestation of FSHD2 in individuals with a permissive (4qA) genetic background³. SMCHD1 is not only a causative gene for FSHD2 but has also been shown to function as a disease modifier in FSHD in general. Patients whose FSHD1 allele included a D4Z4 repeat size in the upper end of the range were shown to be more severely affected when they also had a heterozygous SMCHD1 mutation compared with their less severely affected family members who carried the same D4Z4 disease allele but lacked the SMCHD1 mutation^74,119.

Knockdown of SMCHD1 expression in myotubes from patients with FSHD1 increases DUX4 expression^74,127. Moreover, moderate overexpression of SMCHD1 in myotubes from patients with either FSHD1 or FSHD2 resulted in reduced DUX4 expression¹²⁷, as did restoring near-normal SMCHD1 levels by clustered regularly interspaced short palindromic repeat (CRISPR)–Cas9-mediated excision of an intronic cryptic splice site in SMCHD1 in muscle cells from a patient with FSHD2 (ref. 77). These findings not only mark SMCHD1 as an interesting therapeutic target but also demonstrate that DUX4 de-repression in myotubes can be rescued and its effects potentially reversed.

Heterozygous SMCHD1 missense mutations are a cause not only of FSHD2 but also of the rare developmental disorder BAMS. BAMS is characterized by severe craniofacial malformations such as incomplete development of nose and eyes^129,130. Indeed, BAMS can be considered to lie at the severe end of a clinical spectrum that also includes congenital nasal hypoplasia, anosmia, arrhinia and isolated hypogonadotropic hypogonadism, all of which can be associated with SMCHD1 mutations^129–131. Contrary to FSHD2, in which a broad spectrum of SMCHD1 mutations can be found spanning the entire locus, mutations that cause the BAMS phenotype are exclusively of the missense type and are concentrated in an area encoding the extended ATPase domain of SMCHD1^110,132. The nature of these mutations suggest that some, but not all, BAMS mutations are hypermorphic, based on observations of increased ATP hydrolysis by recombinant mutant SMCHD1 in in vitro quantification assays of ATPase function^133,134. Interestingly, two mutations are associated with both FSHD2 and BAMS^130,132. One of these shared mutations has been shown to result in increased ATPase activity of the recombinant protein in vitro¹³³.

Despite this overlap in the genetic causes of FSHD2 and BAMS, patients with FSHD2 seem to be unaffected by any of the dysmorphic features characteristic of BAMS¹³⁵. Conversely, one patient with BAMS who presented with FSHD-like symptoms was found to have a moderately sized DUX4-permissive allele, which suggests that the two disorders can coexist within the same individual¹³⁰. A study that investigated the overlap between BAMS and FSHD phenotypes reported that 3 of 11 patients with BAMS caused by a mutation in SMCHD1 also met epigenetic and genetic criteria for FSHD2: DUX4 expression was found in dermal fibroblasts from these three patients with BAMS after epigenetic de-repression¹³⁶. DUX4 expression was also found in myotubes derived from two of these three patients, who were therefore considered at risk of developing FSHD2. However, DUX4 mRNA levels in the fibroblasts and myotubes of these three patients were much lower than those of patients with FSHD2, and the three patients with BAMS showed no evidence of skeletal muscle involvement on muscle ultrasound or muscle MRI¹³⁶. These data suggest that patients with BAMS can have an (epi)genetic background compatible with FSHD2 yet show no manifestations of this disease and, furthermore, that the BAMS and FSHD2 phenotypes are unrelated¹³⁶. These findings are also in concordance with the incomplete penetrance of FSHD in FSHD families, in which low levels of DUX4 expression in asymptomatic carriers of pathogenic SMCHD1 mutations have also been reported^16,57. However, a molecular explanation for the disparate clinical outcomes of SMCHD1 mutations in BAMS and FSHD remains largely elusive.

DNMT3B and LRIF1: more FSHD modifiers

DNMT3B has also been identified as an FSHD disease-modifying gene. Mutations in this gene resulted in manifest FSHD in carriers of an FSHD1-sized D4Z4 repeat as well as in carriers of a normal-sized D4Z4 repeat⁷². Biallelic DNMT3B mutations also cause autosomal recessive ICF type 1 (ICF1)^137,138. Monoallelic DNMT3B mutations, by contrast, are a cause of FSHD2: in individuals with a monoallelic DNMT3B mutation inherited along with a short (9 repeat units) permissive D4Z4 allele, de-repression of DUX4 has been observed in muscle cell cultures⁷². DNA methyltransferase 3B (DNMT3B) is co-responsible (along with DNMT3A and DNMT3L, which DNMT3B co-regulates) for de novo CpG methylation and plays a crucial part in the establishment of CpG methylation on the inactive X chromosome (among other loci) during early embryonic development^137,139,140. One of the main features of ICF1 is hypomethylation of CpG islands in pericentromeric satellite repeats and other repeat regions in the genome, including D4Z4^141,142. Therefore, DNMT3B acts as a D4Z4 repressor. As in a subset of patients with BAMS, low levels of expression of DUX4 and/or its target genes were observed in dermal fibroblasts from two patients with ICF1 carrying a DUX4 permissive allele, and low levels of DUX4 protein were detected in myotubes derived from a third patient with ICF1 (ref. 72). However, these three patients with ICF1 did not develop any muscle phenotype, possibly owing to their young age and reduced life expectancy⁷². Interestingly, ICF1 mutation carriers (that is, the parents of patients with ICF1, who each carry a monoallelic DNMT3B mutation) are generally unaffected by FSHD2, which suggests that, as is the case for SMCHD1 mutations causing either FSHD2 or BAMS, genotype–phenotype relationships are only partly understood for DNMT3B mutation carriers.

A homozygous mutation in LRIF1 (the gene encoding ligand-dependent nuclear-receptor-interacting factor 1) that abolishes the long isoform of this protein is associated with FSHD2 (ref. 73). LRIF1 is known to interact with SMCHD1 and is responsible for recruitment of SMCHD1 to the inactive X chromosome in a manner dependent on all subtypes¹⁴³ (that is, α, β and γ¹⁴⁴) of HP1. In patient-derived immortalized fibroblasts forced to form myocytes by ectopic expression of myostatin D, lack of the long isoform of LRIF1 not only resulted in a reduction in total LRIF1 binding but also in reduced SMCHD1 binding to D4Z4. As a result, these cells exhibited D4Z4 hypomethylation and DUX4 expression, marking LRIF1 as a D4Z4 chromatin modifier and LRIF1 as a potential FSHD2 gene⁷³.

An integrated model of FSHD progression

We propose an integrated model of FSHD progression on the basis of clinical features, muscle imaging data and molecular findings. According to this model, fatty infiltration of the muscle tissue of single muscles progresses in a nonlinear fashion with phases of muscle inflammation preceding episodes of rapid fatty replacement, consistent with discrete bursts of DUX4 expression^17,150–152. At any given time, different muscles experience different disease stages. This model reflects a slow linear progression of whole body muscle weakness overall, within which individual muscles show a relapsing course.

Even in patients without any clinical symptoms or abnormalities on muscle MRI, mild-to-moderate histological changes in muscle biopsy samples as well as subtle structural changes on muscle ultrasonography can be detected in the leg muscles of patients with FSHD^150,153,154. To date, no studies have been performed to directly link these ultrasonographic and histological changes. The abnormalities seen on muscle ultrasonography might be due to early histological changes, such as intramuscular fibrosis, that cannot be adequately detected with current MRI techniques.

FSHD muscle biopsy samples with no evidence of DUX4 target gene expression still show mild-to-moderate histological changes and elevated expression of genes encoding extracellular matrix components and inflammatory mediators or genes involved in immune responses^150,155. On the basis of these findings, we hypothesize that expression of DUX4 and DUX4 target genes might still occur at levels below the detection threshold before disease progression in an individual muscle. This situation could occur either because DUX4 expression is transient or because DUX4 is expressed only in a very small number of nuclei at any one time, as has been shown in tissue culture⁵⁸. The abnormal histological findings and gene expression patterns detected in FSHD muscles even in the absence of detectable DUX4 mRNA or DUX4 target gene expression might reflect the accumulated damage from intermittent DUX4 expression in nuclei that are too few or are too briefly DUX4⁺ to detect by RNA-seq. These episodes of transient and/or low-level expression of DUX4 might, nonetheless, be sufficient to activate a primary or secondary immune response.

Further muscle deterioration is characterized by progressive fatty infiltration of the muscle tissue, which can be detected and longitudinally tracked by muscle MRI and is associated with advancing muscle weakness^{17,149,156–158}. Evidence is accumulating that this fatty infiltration is often preceded by a phase of muscle inflammation, which can be visualized on muscle MRI as signal hyperintensities on short-tau inversion recovery (STIR) scans (a T2-weighted sequence with nulling of the fat signal)^159,160. Although some muscles initially show inflammation that disappears over time without leaving any macroscopic consequences on muscle MRI, a more common scenario is that muscle inflammation precedes rapid progression of fatty infiltration of the muscle tissue^{17,151,161–163} (Fig. 2).

Fig. 2 | — a, An MRI scan (Dixon sequence) showing mild-to-moderate fatty infiltration in a region of interest (outlined) of the right medial gastrocnemius muscle. b, A corresponding short tau inversion recovery image of the same patient shows a hyperintense signal (arrowhead) indicative of muscle inflammation. c, Follow-up MRI scan (Dixon sequence) of the right medial gastrocnemius muscle 4 years later of the same patient, showing facioscapulohumeral muscular dystrophy progression to near-complete fatty infiltration of the muscle tissue. d, A corresponding short tau inversion recovery image at the same time point shows that the hyperintense signal (muscle inflammation) has disappeared.

A study has investigated the correlation between two MRI characteristics: turbo inversion recovery magnitude (TIRM) hyperintensity (equivalent to the STIR sequence) and quantitative fat fraction; and two FSHD-associated molecular signatures — DUX4 target gene expression signature and PAX7 score — in muscle biopsy samples. TIRM hyperintensities correlated with active disease and were associated with increased expression of the DUX4 target gene signature, whereas increased expression of the DUX4 target gene signature and reduced PAX7 score were both found in muscles with a high fat fraction, which is characteristic of late stages of disease¹⁵². Molecular biomarkers of FSHD are described in more detail subsequently.

At a group level, disease progression is associated with increased expression of DUX4 and DUX4 target genes as well as increased expression of many extracellular matrix, inflammation and immune response genes in muscle biopsy samples, although in individual patients, biopsy samples might show fluctuating (either increased or decreased) expression of these genes over time^104,150,155. In particular, the presence of STIR hyperintense areas seems to be strongly associated with aberrant DUX4 target gene expression, as well as with an active myopathic process marked by ongoing muscle fibre injury and repair in the presence or absence of associated inflammatory changes. Accelerated replacement of muscle by fat has also been associated with an intermediate fat content in muscles^156,162,164. Graphs that arrange muscles in order of their fat content display an hourglass pattern, in which only a few muscles carry an intermediate fat content¹⁶⁴. This fact suggests that once a muscle reaches an intermediate fat content, progression accelerates towards complete fatty replacement.

Once a muscle becomes nearly or completely replaced with fatty infiltrate, it reaches end-stage pathology and inflammation disappears¹⁵¹. At this point, the muscle has no contractile properties left and muscle damage is assumed to be irreversible.

Therapy development

Now that consensus has been reached on the underlying cause of FSHD — namely, the presence of DUX4 in skeletal muscle — the focus of research has shifted to development of disease-altering therapies (Fig. 3). In the past, several non-targeted therapies have been tested on a small scale (reviewed elsewhere¹⁶⁵). In this Review, we focus on targeted therapies that are currently being developed and/or investigated in preclinical and clinical studies.

Fig. 3 | — a, Pathways that are (partly) upregulated by and/or respond to DUX4 protein in skeletal muscle lead to transcriptional deregulation, inflammation and muscle atrophy. Possible interventions targeting either DUX4 function or DUX4 effectors are indicated with red inhibitory arrows. Bold letters refer to detailed figure panels showing the potential mechanisms that can attenuate the cytotoxic effects of DUX4. b, Top left. Bromodomain-containing protein 4 (BRD4) is an epigenetic regulator that activates DUX4 transcription by recruiting additional epigenetic regulators to the D4Z4 repeat. Bromodomain and extraterminal domain (BET) inhibitors block this BRD4-dependent recruitment and reduce *DUX4* transcription. b, Bottom left. Clustered regularly interspaced short palindromic repeat (CRISPR)–dCas9 (‘dead’ caspase 9) inhibition recruits Krüppel-associated box (KRAB) zinc finger proteins and other transcriptional repressors to D4Z4, thereby reducing transcription of *DUX4* and/or DUX4 target genes. b, right. CRISPR–Cas9 editing disrupts the polyadenylation signal (PAS), either through (top) inducing insertion–deletion mutations (indels) via double-stranded DNA breaks (traditional Cas9) or by (bottom) targeted conversion of a single DNA base pair using dCas9 or Cas9 nickase (nCas9) fused to a base editor protein. c, Phosphorodiamidate morpholino oligomers (PMOs), small interfering (si) RNAs or antisense oligonucleotides (AONs) bind to DUX4 transcripts and prevent maturation of pre-messenger RNA (mRNA) or ribosomal binding to mature mRNAs, thereby blocking translation. MicroRNAs that target DUX4 pre-mRNA can induce degradation of the RNA through the endoribonuclease DICER-activated RNA-induced silencing complex. d, At the protein level, DUX4 function can be blocked by the small molecule iP300w, which interferes with DUX4-dependent recruitment of the cyclic adenosine monophosphate response element binding protein (CREB)–p300 lysine acetyltransferase transcriptional activator complex, thereby inducing transcriptional deregulation. Alternatively, introducing competing high-affinity binding sites for DUX4 traps and prevents DUX4 from binding to its target genes. e, Although the exact mechanism of action of the p38 mitogen-activated protein kinase (MAPK) inhibitor losmapimod is unknown, this agent is proposed to reduce inflammation, possibly disrupting a feedforward loop that would otherwise increase *DUX4* expression by increasing intracellular oxidative stress. SMCHD1, structural maintenance of chromosomes flexible hinge domain containing protein 1; UTR, untranslated region.

β2 Adrenergic agonists and p38 MAPK inhibitors

The results of two non-targeted trials of β2-adrenergic agonists in patients with FSHD showed limited beneficial effects on muscle mass and some measures of muscle strength in the short term^166,167 (Fig. 3a). However, neither of these studies met their primary end point of a 52-week change in global strength as assessed by maximum voluntary isometric contraction testing^166,167. More than a decade later, a targeted screen of a library enriched in pharmacological compounds with epigenetic activities revealed the DUX4-repressive action of β2 adrenergic agonists, which marked these compounds as potential DUX4 inhibitors¹⁶⁸. Further investigations using small molecule inhibitors revealed that p38 MAPK is activated by β2 adrenergic signalling¹⁶⁹. Interestingly, in this screen, the p38α and p38β MAPK isoforms were identified as potent inhibitors of DUX4, most probably independently of β2. This effect was confirmed in studies of the p38 MAPK inhibitor losmapimod in a mouse xenograft model, which provided evidence that DUX4-blocking effects can be achieved by targeting p38α and/or p38β MAPKs^169,170. Oral losmapimod has also been tested in a randomized, double-blind phase 2b clinical trial involving 80 patients with FSHD¹⁷¹. Although no change in DUX4-driven gene expression was observed in losmapimod-treated patients, the researchers stated that losmapimod showed a clinically relevant benefit in terms of various functional and patient-reported outcome measures versus placebo. Patients in the treatment arm showed reduced progression of muscle fat infiltration on MRI over the course of the study in intermediately affected muscles when compared with the placebo group, but no difference in the progression of the muscle fat fraction and lean muscle volume. Given the anti-inflammatory properties of losmapimod¹⁶⁹, it cannot be excluded that the benefits of losmapimod in this trial are (at least in part) due to its anti-inflammatory activity, independent of DUX4 reduction (Fig. 3a,e). The drug was well tolerated and no serious adverse effects were reported. These promising results still await peer-reviewed publication^172,173.

BET-bromodomain inhibitors

The small molecule screen that identified the DUX4-repressive qualities of β2-adrenergic agonists also suggested that bromodomain and extra-terminal domain (BET) family members might attenuate DUX4 levels. BET inhibitors act by blocking the activity of bromodomain-containing protein 4 (BRD4), which in turn suppresses transcription of DUX4 mRNA and abrogates DUX4 activity¹⁶⁸ (Fig. 3b, top left). BRD4 binds to acetylated open chromatin where it promotes transcription by recruiting transcription-regulatory complexes. BRD4 has been widely studied for its potential anticancer and anti-inflammatory effects^174,175.

RNA interference

Small interfering RNAs (siRNAs), microRNAs (miRNAs or miRs) and antisense oligonucleotides (AONs) have therapeutic potential in FSHD owing to their capacity to block transcription and/or translation of DUX4 mRNA. Transcribed miRNA and siRNA molecules emerging from the D4Z4 repeat were identified early in the search for FSHD-causing genes⁵³ and thought to be involved in RNA-mediated epigenetic silencing. Transfection with siRNAs bearing identical sequences to D4Z4-derived small siRNAs or miRNAs resulted in a strong reduction in DUX4 mRNA levels, increased H3K9me2 abundance at D4Z4 and increased recruitment of AGO2, an effector of RNA-mediated epigenetic silencing¹⁷⁶ (Fig. 3c, bottom). Furthermore, inhibition of the DICER–AGO2 pathway led to de-repression of DUX4 expression, suggesting that this pathway provides a basal level of suppression¹⁷⁶. Endogenous human miR-675 inhibits DUX4 mRNA in vitro and has muscle-protective effects when used as a gene therapy in an FSHD mouse model. Several small molecules have the potential to increase endogenous production of miR-675, including oestrogen and melatonin, marking miR-675 a candidate drug target¹⁷⁷.

AONs comprise several different classes of molecules with unique chemical properties that can target RNA molecules in a sequence-specific manner. AONs can achieve knockdown of specific transcripts by preventing their translation and/or by inducing transcript degradation through the RNAse H1 pathway, depending on their specific chemistry¹⁷⁸. Multiple studies have demonstrated the effectiveness of AONs for reducing levels of DUX4 protein, mature DUX4 mRNA levels and DUX4 target gene expression, both in vitro (in FSHD primary and immortalized myocytes) and in vivo (in mouse models)^179–183. Systemic delivery of AONs targeting the DUX4 transcript in a transgenic mouse model proved to be successful in reducing both DUX4 mRNA transcript and DUX4 protein levels, as well as reducing DUX4 target gene expression. Compared with control mice that received a non-targeted AON, mice receiving the DUX4-targeted AON exhibited less skeletal muscle pathology and fibrosis, showed less inflammatory dysregulation and performed better on some functional outcome measures, specifically in relation to muscle strength^182–185. AONs are therefore considered a feasible therapeutic strategy for FSHD (Fig. 3c, top).

Vector-based RNA interference offers the advantages of relatively easy in vivo delivery and a high cell transduction efficiency in post-mitotic cells. Adeno-associated virus (AAV) vectors have already been approved for use in human clinical trials, and trials of AAV-mediated therapies with many years of follow-up attest to their excellent safety profile¹⁸⁶. Intramuscular administration of an AAV vector has been used to deliver miR-405 (which targets DUX4 mRNA) in mice co-transduced with AAV-DUX4. This approach successfully reduced DUX4 protein levels and DUX4-induced muscle pathology by directing the DUX4 mRNA transcript to the RNAi degradation pathway^187,188.

Another class of oligonucleotides has demonstrated its potential for DUX4 inhibition. The targeting antisense sequence of the 5ʹ end of U7 small nuclear RNAs, a component of the small nuclear ribonucleoprotein complex involved in 3ʹ end processing of histone pre-mRNAs, was altered to specifically target DUX4 pre-mRNA, thereby interfering with its maturation. Treatment with this agent led to a substantial reduction in DUX4 transcript levels in FSHD muscle cells. The added benefit of this system compared with traditional AONs and RNAi-based therapies is the longevity of the small nuclear ribonucleoprotein complex that is redirected to target DUX4. This complex protects the included small nuclear RNA from degradation and enables intermittent administration of the therapeutic vector (in contrast to traditional AONs, which require continuous dosing)¹⁸⁹. The necessity for lifelong administration (either continuous or intermittent) of RNAi-based therapies can be considered both a disadvantage (compared with single-dose gene-editing therapies, discussed subsequently) and a strength. RNAi-based treatment is generally non-permanent and self-inactivating, which enables dose correction or treatment discontinuation if adverse effects are observed.

DUX4 protein inhibition

DUX4 recruits the histone acetyltransferases p300 and CREB binding protein to its target loci, which then undergo local H3K27 acetylation that in turn leads to target gene expression⁸⁶. Blocking the transactivation capacity of DUX4 could, therefore, be a viable method of interfering with the DUX4 cascade. Treatment with the p300-specific inhibitor iP300w is effective in blocking DUX4 target gene induction, inhibiting DUX4 cytotoxicity and reversing the DUX4-induced global accumulation of acetylated histone H3 (ref. 190) (Fig. 3d, left). Small molecule inhibitors have the added advantage of systemic administration, possibly even as drugs that can be taken orally.

Another method of blocking DUX4 transactivation has been tested in vitro and in vivo using ‘decoy’ DUX4 binding sites, which sequester endogenous DUX4 and prevent it from binding to and activating its target genes¹⁹¹ (Fig. 3d, right). DNA aptamers with optimized DUX4-binding affinity and specificity might further improve this decoy trapping strategy by increasing trapping efficiency¹⁹².

Gene editing

The CRISPR–Cas9 gene-editing system offers a versatile genetic toolbox for the development of DUX4-targeted therapy^193–196. The first use of the CRISPR system in FSHD involved targeting the KRAB transcriptional inhibitor, fused to catalytically inactive, ‘dead’ Cas9 (dCas9), to the DUX4 promoter in FSHD myoblasts (Fig. 3b, bottom left). DUX4 and its target genes were successfully repressed. An increase of known repressive proteins on D4Z4 was also observed, although levels of H3K9me3 and H3K27me3 were not substantially altered¹⁹⁷.

In FSHD2, deep intronic mutations in SMCHD1 that create an aberrant splice site have been targeted using the CRISPR–Cas9 system (Fig. 3b, top right). Excision of the mutation abolished the cryptic splice site and restored the wild-type SMCHD1 transcript, which was followed by a reduction in expression of DUX4 and DUX4 target genes. However, DNA methylation on the D4Z4 locus was not restored. Nonetheless, this study provided a proof-of-concept for the feasibility of this approach⁷⁷.

Subsequent gene-editing strategies have mainly focused on abolishing the DUX4 exon 3 PAS. Targeting of this DUX4 PAS in FSHD myoblasts using CRISPR and TALEN (transcription activator-like effector nuclease) systems successfully eliminated the targeted DUX4 PAS, even though the editing efficiency was low¹⁹⁸. However, this approach did not completely abolish DUX4 transcript levels, even though all three measured DUX4 target genes (MDB3L2, TRIM43 and LEUTX) assessed were downregulated, confirming the specificity of this approach. In fact, the researchers discovered that an alternative PAS, upstream of the targeted PAS, was responsible for the observed residual DUX4 transcript levels¹⁹⁸. The observed redirection to an alternative PAS was identical to that described in earlier AON approaches that targeted DUX4 pre-mRNA¹⁷⁹. Genetic and epigenetic targeting strategies were combined by targeting the DUX4 PAS directly and using the dCas9-KRAB inhibitor to restore the heterochromatin state at the D4Z4 locus. These two approaches led to reductions in DUX4 transcript levels and in DUX4 target gene expression, achieved by successfully deleting the PAS and restoration of the repressive H3K9me3 histone modification on the targeted locus, respectively¹⁹⁹. In a separate study, the adenine base editor system^200,201 was used along with Cas9-nickase²⁰² to induce an AT to CG conversion (Fig. 3b, bottom right) in the DUX4 PAS in FSHD1 and FSHD2 patient-derived immortalized myoblasts, which effectively introduced mutations into the PAS that prevented DUX4 expression²⁰³.

Loci within repeats are challenging and risky to target. Owing to the repetitive nature of these sequences, the presence of multiple CRISPR–Cas9 binding sites can potentially lead to multiple DNA breaks along the locus, which not only creates unwanted off-target effects but also creates multiple danger signals to the cell and undermines editing efficiency at the on-target site by diluting the effect over a large area. Moreover, delivery of gene-editing systems such as CRISPR–Cas9 to muscle cells remains a considerable challenge that needs to be solved before this technique can develop beyond a promising proof-of-concept. One group tackled the delivery problem by using a dCas9 orthologue derived from Staphylococcus aureus (which is compact enough to be packaged into an AAV vector delivery system) rather than the more widely used Streptococcus pyogenes Cas9²⁰⁴. This strategy proved effective in restoring epigenetic repression of DUX4 without notable effects on the D4Z4 repeats adjacent to the targeted site or at the predicted and closest matching off-target genes. Moreover, this approach achieved repression of DUX4 and DUX4 target genes in vivo in an FSHD transgenic mouse model, albeit at a modest level of effect²⁰⁴. Altogether, the different CRISPR-based genetic and epigenetic strategies used so far have risen above a proof-of-principle stage and are viable methods that are worth pursuing towards therapeutic development.

Various approaches have been used with the aim of developing FSHD-specific, targeted therapies for clinical use (Fig. 3b and Table 1). The primary target of these therapies is DUX4, and these agents act by preventing or disrupting DUX4 transcription, preventing translation or inhibiting protein function (Fig. 3). All these potential therapies are in the preclinical stage, with the exception of losmapimod, which is progressing towards a phase 3 clinical trial. Each strategy presents challenges with respect to delivery, efficacy and safety and the need to address the sporadic nature of DUX4 expression. Moreover, the level of DUX4 that can be tolerated in the muscles of patients with FSHD remains unknown. Because DUX4 leaves a transcriptional memory in affected muscle cells, it is also unknown whether abolishing DUX4 expression can fully reverse or stabilize the damage already accumulated in affected muscles.

Table 1 |.

Targeted therapies for facioscapulohumeral muscular dystrophy

Agent	Mechanism	Preclinical and clinical findings	Refs.
Small molecules
β2 Adrenergic receptor agonists	Presumed to inhibit DUX4 transcription by promoting muscle hypertrophy, in part through modulating cAMP levels	↓DUX4 mRNA, ↓DUX4 target gene expression and ↑intracellular cAMP in FSHD patient-derived muscle cells RCTs in 90 and 65 patients with FSHD: ↑lean muscle mass, ↑grip strength; no change or limited improvement in maximum voluntary isometric strength in some muscles	166–168
BET protein BRD4 inhibitor	Inhibition of DUX4 transcription blocks BRD4- dependent recruitment of transcription regulators to D4Z4	↓DUX4 mRNA and ↓ DUX4 target gene expression in FSHD patient-derived muscle cells	168
p38 MAPK inhibitors (losmapimod)	Inhibition of p38 MAPK signalling has anti-inflammatory effects. Decreased DUX4 expression observed through unknown mechanism	↓DUX4 mRNA and ↓DUX4 target gene expression in FSHD patient-derived muscle cells; ↓DUX4 and DUX4 target gene expression in a mouse FSHD xenograft model Phase 2b RCT in 80 patients with FSHD (awaits peer review): improvement in some patient-reported outcomes albeit without DUX4 attenuation (possibly through anti-inflammatory properties); phase 3 trial planned	169–173
P300 histone acetyltransferase inhibitor (iP300w)	Inhibits DUX4-mediated recruitment of p300–CBP to DUX4 target genes by blocking p300	↓DUX4 target gene expression; ↓global acetylated histone H3 abundance (accessible chromatin marker) in FSHD myotubes and in a transgenic FSHD mouse model	190
Oligotherapeutics
PMO and/or AON targeting 3’-UTR of DUX4 pre-mRNA (PAS and/or cleavage sites)	Prevents correct pre-mRNA processing (intron splicing or polyadenylation), inhibiting mRNA stability and promoting transcript degradation	↓DUX4 mRNA and ↓DUX4 target gene expression; cleavage site redirection that rescued DUX4 observed with one PMO; ↓DUX4 protein and ↓atrophic myotubes; ↑muscle mass and strength; ↓myofibre central nucleation and muscle fibrosis; in FSHD patient-derived muscle cells, FSHD patient-derived primary muscle cells and in FSHD xenograft and/or transgenic FSHD mouse models	179–184
AON targeting exon 1 of DUX4 transcript	Inhibits DUX4 mRNA translation, promoting transcript degradation	↓DUX4 mRNA, DUX4 protein and DUX4 target gene expression; ↓skeletal muscle pathology; improved treadmill test results; no improvement in strength test results in a transgenic FSHD mouse model	185
siRNAs bearing sequence homology to DUX4	Exogenous siRNA targeting the 5’-UTR of DUX4 pre-mRNA	↓DUX4 mRNA; ↑H3K9me2 and recruitment of AGO2 (indicating RNA-mediated epigenetic silencing) in FSHD patient-derived muscle cells	176
miR-405	RNA interference through artificial miRNA targeting human DUX4 mRNA open reading frame, inhibiting transcription	↓DUX4 mRNA in luciferase assay screen; ↓skeletal muscle pathology but no notable improvement in grip strength with AAV-miDUX4.405 overexpression in the AAV-DUX4-transgenic mouse model	187,188
miR-675	RNA interference through naturally occurring human miR-675 directly targeting both the 3’-UTR and open reading frame of DUX4 mRNA	↓DUX4 mRNA, ↓DUX4 protein, ↓DUX4 target gene expression, ↓DUX4-induced cell death; ↓transactivation of DUX4-responsive reporter in HEK293T cells transfected with DUX4 and in FSHD patient-derived muscle cells; ↓skeletal muscle pathology in transgenic (AAV-DUX4 induced) FSHD mouse model	177
U7 small nuclear (sn) RNA antisense expression cassettes	Alters target specificity of sn ribonucleoprotein complex to target DUX4 pre-mRNA, inhibiting its maturation	↓DUX4-induced cell death; ↓DUX4 mRNA; ↓DUX4 protein in transfected HEK293 cells overexpressing DUX4; ↓DUX4 mRNA and ↓DUX4 target gene expression in FSHD patient-derived muscle cells	189
Double-stranded (ds) DNA oligonucleotides encoding DUX4 binding sites	Inhibits DUX4 transactivational activity through competition of decoy binding sites with real DUX4 target gene-binding sites	↓DUX4 target gene expression on direct electrotransfection of dsDNA in FSHD patient-derived muscle cells and on systemic delivery of AAV-dsDNA decoy in mice overexpressing DUX4 from transfected plasmid vector	191
DNA aptamers	Bind DUX4 protein with increased affinity and specificity, inhibiting DUX4 target binding and activation	Aptamers enhance affinity and specificity of decoy binding site sequences for DUX4 using recombinant DUX4 (SELEX-derived aptamers have not yet been tested in DUX4-expressing cells)	192
Gene editing
CRISPR–Cas9	Restores SMCHD1 reading frame by removing an intronic mutation causing a cryptic splice site	↑Wild-type SMCHD1; ↓DUX4 mRNA; ↓DUX4 target gene expression in FSHD2 patient-derived muscle cells; no effect on D4Z4 DNA methylation	77
CRISPR–Cas9	CRISPR–Cas9 and TALEN-based elimination of DUX4 PAS (exon 3)	↓DUX4 mRNA and ↓DUX4 target gene expression in double-knockout HCT116 (human colorectal cancer cell) model; use of alternative PAS upstream of targeted PAS in FSHD patient-derived muscle cells	198
CRISPR–Cas9	CRISPR–Cas9 elimination of DUX4 PAS (exon 3)	↓DUX4 mRNA and ↓DUX4 target gene expression in double-knockout HCT116 human colorectal cancer cell model and in FSHD patient-derived muscle cells	199
CRISPR–dCas9 adenine base editor	Induces mutation (AT → CG conversion) in DUX4 PAS	↓DUX4 mRNA and ↓DUX4 target gene expression in FSHD patient-derived muscle cells	203
CRISPR–dCas9 inhibition
CRISPR–dCas9 transcriptional repression	CRISPR–dCas9 orthologue fused to transcriptional repressors of DUX4	↓DUX4 mRNA and ↓DUX4 target gene expression in FSHD primary myocytes; ↓DUX4 and DUX4 target gene expression in an FSHD transgenic mouse model (more modest repression)	204
CRISPR–dCas9–KRAB transcriptional repression	KRAB-repressor targeting the DUX4 gene promoter	↓DUX4 mRNA and ↓DUX4 target gene expression in FSHD primary myocytes; no effect on expression of other genes in the D4Z4 locus (FRG1 and FRG2)	197
CRISPR–dCas9–KRAB transcriptional repression	KRAB-repressor targeting DUX4 transcription activators	↓DUX4 mRNA; ↑H3K9me3 (evidence of altered chromatin) at the targeted D4Z4 locus in FSHD primary myocytes	147
CRISPR–dCas9–KRAB transcriptional repression	KRAB-repressor targeting the DUX4 PAS	↓DUX4 mRNA and DUX4 target gene expression; (partial) restoration of repressive H3K9me3 in FSHD patient-derived muscle cells	199

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AAV, adeno-associated virus; AGO2, argonaute RISC catalytic component 2; AON, antisense oligonucleotide; BET, bromodomain and extra terminal domain; BRD4, bromodomain-containing protein 4; cAMP, cyclic adenosine monophosphate; Cas9, caspase 9; CBP, cyclic adenosine monophosphate response element binding protein (CREB) binding protein; CRISPR, clustered regularly interspaced short palindromic repeat; dCas9, ‘dead’ (DNA cleavage-inactivated) Cas9; DUX4, double homeobox protein 4; FSHD, facioscapulohumeral muscular dystrophy; KRAB, Krüppel-associated box; MAPK, mitogen-activated protein kinase; PAS, polyadenylation signal; PMO, phosphorodiamidate morpholino oligomer; RCT, randomized controlled trial; SELEX, systematic evolution of ligands by exponential enrichment; siRNA, small interfering RNA; TALEN, transcription activator-like effector nucleases; UTR, untranslated region of pre-mRNA; mRNA, messenger RNA; miR, microRNA; SMCHD1, structural maintenance of chromosomes flexible hinge domain containing protein 1.

Clinical trial readiness

Important challenges also remain in the development of highly reliable and responsive biomarkers and clinical outcome measures for use in future clinical trials^4,205. As variability between patients can be large and FSHD progression is generally slow, clinical trials are likely to require high numbers of individuals with this rare disease and a long follow-up time. For example, a natural history study in the 1990s, in which 50 patients with FSHD were followed up for 1 year, calculated that a two-armed clinical trial that used quantitative muscle strength as the primary outcome would require 160 patients per group and 1 year of follow-up to provide 80% power to detect a complete halt in disease progression²⁰⁶. To tackle these challenges, improved clinical trial access, sensitive biomarkers and relevant clinical outcome measures are required.

Clinical trial access

Access to clinical trials is facilitated by the use of patient registries in 14 different countries^207,208. In addition to being an effective means of informing large numbers of patients about new trials, patients who fulfil the (strict) eligibility criteria can be approached selectively through the registry if clinical and genetic data are collected.

Biomarkers

Reliable biomarkers are also required, especially for early (phase 1–2) clinical trials, to accelerate the drug development process. Of most interest are pharmacodynamic biomarkers that can both serve as (secondary) clinical trial end points and provide a ‘proof of mechanism’ that confirms target engagement, thereby improving the chances of a novel therapy moving forward to later-phase and confirmatory clinical trials.

Although DUX4 protein levels might seem at first glance to be an appropriate molecular biomarker for disease activity in patients with FSHD, DUX4 itself is not ideal because this protein can be difficult to detect owing to its low levels and sporadic DUX4 gene expression in skeletal muscle^57,58,209. Instead, the expression of several DUX4 target genes that comprise a DUX4 signature can be measured. This signature is useful for drug screening in patient-derived cells and inducible animal models (that is, models in which DUX4 expression can be switched on specifically in muscle cells) and is also measurable in patient-derived muscle biopsy samples^{104,105,150,210,211}. Knockdown of DUX4 expression substantially decreases the expression of DUX4 target genes^104,180. Various studies^{64,77,128,198} use combinations of 2–4 DUX4-upregulated target genes, among which LEUTX, KHDC1L, PRAMEF2, MDB3L2, ZSCAN4 and TRIM43 comprise a DUX4 signature panel. One study showed that a set of four genes (LEUTX, PRAMEF2, TRIM43 and KHDC1L) showed slightly superior performance in discriminating muscle biopsy samples from patients with FSHD from those of healthy control individuals¹⁰⁴. However, the muscle biopsy samples from healthy control individuals could not be accurately distinguished from those of mildly affected patients with FSHD (mimicking early stage of disease) on the basis of expression of the signature genes alone¹⁵⁵. An RNA-seq study in 36 patients with FSHD suggests that an expanded set of 52 genes, which comprises DUX4 target genes as well as other genes that are differentially expressed in FSHD muscle, encoding complement proteins, inflammatory mediators, lymphocyte markers, immunoglobulins and cell cycle regulators, might have increased discriminative power compared with just the above four DUX4-upregulated target genes for identifying early changes in FSHD muscle¹⁵⁵.

Another candidate biomarker is the PAX7 score²¹². PAX7 is produced in satellite cells and is a marker for these quiescent myogenic progenitors. PAX7 and DUX4 exhibit high sequence similarity in their DNA binding domains and compete for DNA binding to their respective target loci^212,213. DUX4 expression, therefore, competitively inhibits PAX7 target genes in differentiating myogenic cells, which results in an altered ratio between PAX7-induced and PAX7-repressed genes. This ratio is known as the PAX7 score²¹². The PAX7 score correlates with histological and muscle imaging measures of FSHD disease activity in a manner independent of, but partially overlapping with, DUX4 target gene expression^152,214,215. Like DUX4, PAX7 also undergoes sporadic expression, a characteristic that similarly limits its suitability for use as a singular molecular biomarker. In areas where the DUX4 signature was increased in TIRM⁺ muscle biopsy samples, combining both signatures enabled detection of the later stages of disease, which are characterized by fat infiltration. These data suggest that although the PAX7 score and DUX4 signature overlap, they represent different stages of disease¹⁵². A study of FSHD muscle biopsy samples obtained from 36 participants in a different longitudinal study¹⁵⁵ showed an increase in PAX7 target gene repression over 1 year of follow-up, suggesting that this biomarker is associated with FSHD disease progression²¹⁵. Further validation of these potential muscle tissue biomarkers in independent patient cohorts is required, in addition to determining their test–retest reliability, variability between different samples within the same muscle and how large a change would be necessary to see a clinical effect.

Multiple exploratory studies have identified several potential serum biomarkers for FSHD^216–223, the most promising of which are creatine kinase-myocardial band isoforms^216,217, innate immunity mediator protein S100-A8 (ref. 220), IL-6 (ref. 221), complement components²²², miR-206 (ref. 223) and a panel of 19 miRNAs that are upregulated in patients with FSHD²¹⁹. Confirmatory studies need to be conducted to validate these biomarkers.

Several muscle imaging biomarkers are promising and could be especially useful to support the transition from early phase clinical trials to phase 3 trials. Both quantitative muscle MRI and quantitative ultrasonography are able to discriminate between-patient differences in the severity of muscle involvement and, on a cross-sectional level, both correlate strongly with clinical outcome measures^{153,158,164,224–230}. MRI is better than ultrasonography for detecting the late stages of fatty infiltration, but ultrasonography can detect changes in muscles that appear normal on MRI¹⁵³. In the past few years, muscle MRI has been used in two multicentre clinical trials: to assess muscle volume in a phase Ib–II study of recombinant human histidyl-tRNA synthetase and to assess quantitative fat fraction in a phase I and II study of losmapimod. In future clinical trials that include patients in the early disease stages of FSHD, ultrasonography biomarkers might be of particular interest.

As mentioned previously, muscle MRI can identify the muscles with active disease that are at highest risk of accelerated disease progression^17,162,163. MRI-based targeting of such muscles (which are also likely to show high levels of DUX4 target gene expression) for muscle biopsy could be advantageous for assessing early responses of patients to treatment with a DUX4-targeted therapy. As such, the presence of STIR⁺ areas on muscle MRI might serve as an inclusion criterion for clinical trials. This use of imaging biomarkers is especially relevant because finding an appropriate time window for treatment administration might be challenging owing to the variability in disease stages between different muscles within one patient, the variability in disease course between patients and the high likelihood of only transient DUX4 expression. Longitudinal data show that both MRI and ultrasonography can detect progression of fatty infiltration in muscle tissue^{17,149,156,162,230}. Whether the changes in imaging metrics correlate with clinical changes is yet to be established, because in most of these studies no changes occurred in clinical parameters over the course of the study, either because of small cohort sizes (up to 45 patients) or short follow-up periods (a maximum of 2 years). Additional longitudinal imaging studies are also required to define the increase in muscle fatty infiltration that represents a clinically relevant change.

Another MRI metric that could be used to measure disease progression or response to therapy is muscle volume. Although some early studies used muscle computed tomography or dual-energy X-ray absorptiometry to measure changes in muscle SMCHD1 mass, the clinical relevance of this parameter remains questionable because an increase in muscle mass in several clinical trials did not result in increased muscle strength^166,167,231.

Clinical trial outcome measures

Clinical outcome measures that reflect how a patient ‘feels, functions or survives’ are essential for late phase clinical trials and for obtaining treatment approval. Regulatory agencies such as the FDA and EMA favour the use of clinical outcome measures in phase 3 clinical trials^232,233. FSHD-specific outcomes are preferred, because outcome measures designed to assess function in patients with other neuromuscular disorders are often not fully suited to assess change in patients with FSHD, because the rate of progression and distribution of weakness across the body vary across different neuromuscular disorders²³⁴.

In the past few years, multiple FSHD-specific measurement instruments have been developed and are currently undergoing further validation. These include functional outcome measures, most importantly the FSHD Composite Outcome Measure (FSHD-COM)²³⁵ and Reachable Workspace (RWS)^236–238, and two FSHD-specific patient-reported outcomes, namely, the FSHD Health Index (FSHD-HI)²³⁹ and FSHD-Rasch-built Overall Disability Scale (FSHD-RODS)^4,207. A detailed description of these instruments is beyond the scope of this Review. All these outcome measures are currently being tested in longitudinal studies. The results of these studies will be important to determine the sensitivity to change, sample size requirements and minimal clinically relevant changes for each tool. Until these data are available, these outcome measures should be used with caution in clinical trials.

Conclusions

After decades of studying the disease mechanism of FSHD and uncovering many potential therapeutic targets, consensus has been reached that aberrant DUX4 expression is the root cause of FSHD and, as a result, the field is entering a new era focusing on the development of potentially disease-modifying therapy. Many promising molecular pathways and (epi)genetic tools have successfully demonstrated proof-of-concept applications in the treatment of FSHD, and many more are rapidly being uncovered by emerging single-cell and single-molecule approaches, as well as CRISPR–Cas9 genome editing tools. The first DUX4-targeted therapies are either already being investigated or moving towards in vivo studies and making their way towards clinical approval. Of these, losmapimod, a p38 MAPK inhibitor, has been tested in a recent phase 2 trial.

In vivo studies also require reliable efficacy biomarkers. Although the expression of DUX4 signature genes and the PAX7 score seem to be viable biomarkers of active FSHD disease processes in cultured cells and muscle biopsy samples, the search for easily applicable (serum) biomarkers continues. Several muscle imaging modalities are currently being investigated to identify metrics that could be used as non-invasive clinical biomarkers, and these studies have also demonstrated the suitability of quantitative MRI and ultrasonography for measuring FSHD progression.

Various functional and patient-reported outcome measures are currently being tested in longitudinal studies. With more trials on the way, an increasingly important research focus is the further development of clinical outcome measures and investment in worldwide patient registries and trial readiness. At the same time, studies of the basic molecular underpinnings of this disease continue to fuel translational studies that lead to the development of new therapies. This exciting new era is expected to benefit patients and bear witness to the joint efforts of many research groups and industry in making the final leap from bench to bedside.

Key points.

Facioscapulohumeral muscular dystrophy (FSHD), a disorder for which there currently is no cure, is characterized by muscle weakness, predominantly affecting muscles in the face, shoulder girdle and upper arms.
FSHD is associated with epigenetic de-repression of the DUX4 gene, which leads to aberrant expression of the transcription factor DUX4 and cytotoxicity in skeletal muscle cells.
Clinical disease progression occurs in a nonlinear and muscle-by-muscle fashion with phases of muscle inflammation preceding rapid fatty replacement of muscle tissue and muscle wasting.
Consensus has been reached on the pathogenetic mechanism of FSHD, and the field is entering a new era of targeted therapy development.
Disease-altering therapies currently in development range from proof-of-principle gene-editing technologies focusing on reducing DUX4 expression to clinical trials of DUX4-blocking agents.
Clinical trials in FSHD require the development of meaningful patient outcome measures, identification of reliable biomarkers and accurate methods of measuring disease progression, such as MRI and ultrasonography.

Acknowledgements

The authors apologize to those whose work was not cited or insufficiently cited because of size constraints. Our work is supported by the Prinses Beatrix Spierfonds (W.OR19-06, W.OR21-04), public–private partnerships allowances made available by Health-Holland, Top Sector Life Sciences and Health, The Netherlands, the Medical Research Council UK, Stichting Spieren voor Spieren, the National Institute of Neurological Disorders and Stroke (Grant No. P01NS069539) and the National Institute of Arthritis and Musculoskeletal and Skin Diseases (Grant No. P50AR065139). Some of the authors are members of the Radboud University Medical Centre (UMC), Center of Expertise for Neuromuscular Disorders (Radboud-NMD), Netherlands Neuromuscular Center (NL-NMD) and the European Reference Network for Rare Neuromuscular Diseases [ERN EURO-NMD].

Glossary

cis-spreading: Spreading of epigenetic chromatin modifications from a locus to neighbouring loci on the same chromatin strand
FSHD Composite Outcome Measure: (FSHD-COM) An evaluator-administered instrument composed of individually validated functional motor tasks that assess leg, shoulder and arm, trunk and hand function and balance
FSHD Health Index: (FSHD-HI) A patient-reported outcome measures to assess total FSHD health-related quality of life and 14 separate subdomains, each identified by FSHD patients as having the greatest importance to their specific population
FSHD-Rasch-built Overall Disability Scale: (FSHD-RODS) A linear-weighted patient-reported outcome measure for detecting activity and participation restrictions in patients with FSHD
PAX7 score: The ratio of genes induced to genes repressed indicated by the satellite cell marker PAX7, which is closely related to DUX4
Perdurance: Lasting target gene expression followed after a short DUX4 transcriptional burst
pLAM: A DNA region flanking the most distal D4Z4 unit that contains the polyadenylation signal (PAS) of the DUX4 gene, named after the cosmid clone used for its isolation
Proximity ligation assay: An antibody assay based on rolling-circle amplification of a DNA loop formed from ligation of oligonucleotide-labelled probes bound to distinct primary antibodies, used for detection of proteins at single-molecule resolution in unmodified cells and tissues
Pseudotime trajectory modelling: A computational method used in single-cell transcriptomics to establish the pattern of a dynamic cellular process; followed by ordering of cells based on their progression through the process
Reachable Workspace: (RWS) A depth-ranging sensor (Kinect)-based upper extremity motion analysis system that is applied to determine the spectrum of reachable workspace encountered with the arms and shoulders
Retrogene: A segment of DNA reverse-transcribed from mRNA and randomly inserted into a genome

Footnotes

Competing interests

K.M. declares that she has acted as a consultant for Avidity Biosciences. S.J.T. declares that he has acted as a consultant for Avidity Biosciences and is a Board member for Renogenyx. R.T. declares that he has acted as a consultant and/or is a member of the advisory board for Arrowhead Pharma, Avidity Biosciences, Dyne Therapeutics, Fulcrum Therapeutics, Mitsubishi Tanabe Pharma, miRecule Biotech and Roche. J.M.S. declares that he has acted as a consultant and/or is a member of the advisory board for Acceleron, Avidity Biosciences, Dyne Therapeutics, Fulcrum Therapeutics, Ionis, ML Bio Solutions, Mitsubishi Tanabe Pharma, Roche and Sarepta. S.M.v.d.M. declares that he has acted as consultant and/or is a member of the advisory board for Avidity Biosciences, Dyne Therapeutics and Fulcrum Therapeutics and is a Board member for Renogenyx. The other authors declare no competing interests.

Additional information

Peer review information Nature Reviews Neurology thanks the anonymous reviewers for their contribution to the peer review of this work.

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PERMALINK

Facioscapulohumeral muscular dystrophy: the road to targeted therapies

Mara S Tihaya

Karlien Mul

Judit Balog

Jessica C de Greef

Stephen J Tapscott

Rabi Tawil

Jeffrey M Statland

Silvère M van der Maarel

Abstract

Introduction

Clinical phenotype

Disease mechanism

Identification of the FSHD-causing gene

Fig. 1 |. The D4Z4 repeat array on chromosome 4q in healthy individuals and patients with facioscapulohumeral muscular dystrophy.

Repression of the FSHD locus

FSHD types 1 and 2

Rare forms and unexplained FSHD

Role of DUX4 in health and disease

Correlates and modifiers of disease

D4Z4 repeat size

DNA methylation affects D4Z4 regulation

SMCHD1 acts as an FSHD modifier

DNMT3B and LRIF1: more FSHD modifiers

Other factors linked to DUX4 expression

Chromatin remodelling proteins.

Environmental factors.

An integrated model of FSHD progression

Fig. 2 |. Progression of facioscapulohumeral muscular dystrophy can be measured with MRI.

Therapy development

Fig. 3 |. DUX4-mediated pathways and possible methods of therapeutic inhibition.

β2 Adrenergic agonists and p38 MAPK inhibitors

BET-bromodomain inhibitors

RNA interference

DUX4 protein inhibition

Gene editing

Table 1 |.

Clinical trial readiness

Clinical trial access

Biomarkers

Clinical trial outcome measures

Conclusions

Key points.

Acknowledgements

Glossary

Footnotes

References

ACTIONS

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RESOURCES

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