The rise of rat models for Duchenne muscular dystrophy and therapeutic evaluations

Abstract

Duchenne muscular dystrophy (DMD) is a devastating X-linked neuromuscular disorder characterized by the absence of a functional dystrophin, leading to progressive muscle loss responsible for cardiorespiratory failure and premature death. While mouse, dog, and pig models have long supported DMD preclinical research, each has limitations in terms of phenotype severity, translational relevance, cost, or ethical acceptance. The emergence of genetically engineered DMD rat models marks a major advancement, offering an intermediate platform that combines practical handling, robust disease features, and disease trajectory accuracy with human patients. Rat models exhibit early, progressive and severe skeletal and cardiac pathology, including impaired muscle regeneration due to satellite cell senescence, all of which closely mirrors patient pathology. In vivo single-nucleus transcriptomics has further highlighted the complexity of fibrotic, inflammatory, and stem cell dysfunction across affected tissues. Importantly, DMD rat models have proven valuable for preclinical therapeutic studies, including gene and exon-skipping therapies, small compounds or cell-based interventions, and senescence-targeting strategies. They have also supported functional, histological, and molecular endpoints aligned with clinical practice. Importantly, DMD rat lines are not phenotypically uniform. Variations in mutation type, involvement of specific dystrophin isoforms, spontaneous exon skipping, and genetic background lead to differences in disease onset, severity, organ involvement, and survival. These distinctions influence the suitability of each model for precision therapeutic strategies. DMD preclinical rat models therefore provide a powerful complementary tool that fits into a continuum of modeling to advance understanding of pathogenic mechanisms, biomarker discovery, and translational research. Their progressive adoption will be accelerating the development of more effective and clinically relevant therapies for patients affected by dystrophin deficiency.

Background

DMD is a severe form of inherited neuromuscular disorder with a prevalence of approximately 1 in 5,000 live male births worldwide [1]. It is characterized by progressive muscle wasting that leads to loss of ambulation, respiratory insufficiency, and premature death, making it one of the most debilitating genetic diseases of childhood. DMD is caused by mutations in the X-linked DMD gene, which encodes dystrophin, a large cytoskeletal protein critical for the stability and function of skeletal and cardiac muscle. The absence of dystrophin renders skeletal muscle fibers highly vulnerable to mechanical stress during contraction, leading to necrosis that triggers cycles of degeneration, regeneration, chronic inflammation, and ultimately, leads to progressive loss of muscle tissue that are gradually replaced by fibrotic and adipose tissue. In addition, cardiomyopathy progresses alongside skeletal muscle degeneration, leading to premature death due to cardiorespiratory failure before the third decade of life. DMD is one of the largest genes in the human genome, spanning 79 exons and encoding several isoforms expressed in skeletal and cardiac muscle, but also in regions of the nervous system and other organs [2]. Despite decades of research, current therapies have aimed to manage symptoms and slow disease progression, but there is still no efficient curative treatment [3, 4]. DMD is part of the dystrophinopathies spectrum, which also includes Becker muscular dystrophy (BMD), a milder form caused by in-frame mutations in the DMD gene that lead to a truncated, partially functional dystrophin protein.

Main text

DMD animal modeling

Animal models play a critical role in understanding DMD pathogenesis, and in testing therapeutic strategies. The mdx mouse, bearing a Dmd gene mutation, remains the most used model. It has been instrumental for studying basic disease mechanisms, dystrophin function, and evaluating gene and cell therapies. However, mdx develops a mild, slowly-progressive phenotype, with near-normal lifespan and limited fibrosis and muscle weakness compared to human patients [5]. In addition, differences in growth dynamics and metabolic or energetic requirements between rodents and humans may further contribute to the disparity in disease severity and progression observed between mdx mice and DMD patients.

To better recapitulate human disease, other species such as the dog offer physiological and clinical similarity to human DMD, but with practical challenges and limitations [6]. The advent of genome editing technologies has enabled the development of DMD models aiming to bridge the gap between the mouse and existing larger mammalian models. These include rabbits [7], pigs [8], non-human primates [9] and rat models (Table 1). Functionally, DMD rats exhibit a progressive and severe DMD pathology associated with significant reduction in muscle mass and strength, early cardiac and diaphragm involvement, all associated with precocious lethality. These phenotypes are more severe and human-like than those observed in mice, while preserving the practical advantages of rodents over larger mammals.

Table 1.

Dystrophin-deficient rat models, pathophysiology, and therapeutic evaluations

Name Mutation Genetic background (Initial report) Survival	Skeletal muscle phenotype	Heart phenotype	Other impairments	Therapeutic evaluations	References
DUCHENNE MUSCULAR DYSTROPHY MODELS
DMD mdx Deletion of 11 bp in exon 23 Sprague Dawley (10) Median survival 13 months	Histology: No dystrophin ↑ Fibrosis ↑ Adipose infiltration ↑ Fiber degeneration ↑ Inflammation ↑ Regeneration Functional defects: ↓ Muscle mass ↓ Strength ↓ Endurance Molecular defects: ↑ Fibrosis/inflammation signatures. Molecular biomarkers: ↑ CK (serum) ↑ Inflammatory cytokine (muscle) ↑ Immune cell markers (muscle) ↑ Urinary Titin.	Histology: ↑ Cardiomyopathy ↑ Fibrosis ↑ Interstitial collagen. Functional defects: ↓ Cardiac output and contractility ↑ Heart rate variability and arrhythmias. ↑ Diastolic dysfunction in aged animals. Molecular defects: ↑ Cardiac stress markers, ↑ Fibrotic and inflammatory signatures. Molecular biomarkers: ↑ Cardiac troponins (serum) ↑ Brain natriuretic peptide with cardiac dysfunction.	↓ Body mass. Behavioral defects: ↑ stress sensitivity ↑ cognitive and emotional impairments, ↓ spontaneous activity. Neuronal changes: ↑ brain inflammation markers (S100β, Tau) ↑ GFAP. Brain Imaging: ↑ structural and metabolic abnormalities. Neurochemistry: ↑ GABA and NAA (cortex, hippocampus).	Gene Therapy: AAV9-micro-dystrophin → restored muscle strength and cardiac function. Immunomodulation: Anti-CD45RC monoclonal antibodies → immune tolerance. Metabolic Modulators: Taurine supplementation → ineffective. Heart Rate Modulation: Ivabradine → reduces heart rate and improves cardiac calcium handling and function. Calcium modulators: Prevention of calcium homeostasis alterations → improved muscle phenotype.	10, 25, 26, 28, 29, 32–34, 37–39, 44–46
W-Dmdem1Kykn (cDMR) F0: deletions of 9 to 577 bp in exon 3 ± flanking regions, or of 1 to 186 bp in exon 16 ± flanking regions; insertion of 2 to 4 bp in exon 3. F1: deletion of 10 bp in exon 16, or insertion of 1 bp and substitution of 2 bp in exon 16 Wistar-Imamichi (11)	Histology: No dystrophin ↑ Fibrosis ↑ Adipose infiltration ↑ Necrosis ↑ Inflammation ↑ Stem cell dysfunction (senescence) Tongue muscle less affected than masseter. Functional defects: ↓ Muscle mass ↓ Strength ↓ Endurance	Histology: ↑ Cardiomyopathy ↑ Fibrosis. Functional defects: ↓ Cardiac output and contractility ↑ Heart rate variability and arrhythmias.	↓ body mass. Skeletal: ↓ trabecular bone volume with altered microstructure.	Ivabradine: → Improved cardiac function and calcium homeostasis. Metabolic supplementation: Ketogenic diet → improves skeletal muscle but exacerbated cardiomyopathy. Senolytic: ABT263 → improves muscle strength and regeneration. Genetic: p16 deletion → reduces senescence and improves repair, but development of rhabdomyosarcoma. Cell therapy: Grafting of human myoblasts in immunodeficient rats.	11, 15, 22, 24, 31, 35, 40–43, 47
DMD-KO Deletion of 14 bp in exon 3 Wistar-Imamichi (15)	Histology: No dystrophin ↑ Fibrosis ↑ Inflammation Functional defect: ↓ Strength	N/A	N/A	Selective androgen receptor modulator TEI-SARM2: → improved muscle strength	15
Dmd-KO Deletion of exons 22 to 26 Wistar-Imamichi (12)	Histology: No dystrophin ↑ Inflammation ↑ Regeneration Functional defect: ↓ Strength Imaging (MRI): ↑ Inflammation ↑ Regeneration	N/A	Brain: ↓ brain volume Neurochemistry: ↑ GABA and NAA (prefrontal cortex) ↑ GABA (hippocampus)	N/A	12, 27
R-DMDdel52 Deletion of exon 52 (118 bp) + 70 bp of flanking regions Sprague Dawley (13) Median survival 11–12 months	Histology: No dystrophin ↑ Fibrosis ↑ Adipose infiltration ↑ Fiber degeneration ↑ Inflammation ↑ Regeneration ↑ Stem cell dysfunction (senescence) Functional defects: ↓ Muscle mass ↓ Strength ↓ Endurance Molecular analyses: nNOS deficiency snRNA-seq. Biomarkers: ↑ CK (serum) ↑ COMP (muscle)	Histology: ↑ Cardiomyopathy (severe) ↑ Fibrosis ↑ Inflammation Functional defects: ↓ Cardiac output and contractility ↑ Heart rate variability and arrhythmias. ↑ Hypertrophic remodeling ↓ Ejection fraction Molecular analyses: ↑ Stress response (snRNA-seq) ↓ Tmem65 mislocalized Cx43.	↓ body mass.	Forskolin: → Improves muscle regeneration and reduces senescence, long-term cardiac impairment.	13, 16, 20, 36
R-DMDdup10-17 Duplication of a ~ 125 kb sequence encompassing exons 10 to 17 Sprague Dawley (14) Median survival 11–12 months	Histology: No dystrophin ↑ Fibrosis ↑ Adipose infiltration Functional defects: ↓ Muscle mass ↓ Strength ↓ Endurance ↓ Spontaneous locomotion ↓ Ventilatory function. Molecular biomarkers: ↑ CK/MYOM3 (serum)	Histology: ↑ Cardiomyopathy (severe) ↑ Fibrosis Functional defects: ↓ Cardiac output and contractility ↑ Heart rate variability and arrhythmias. Molecular biomarker: ↑ hs-cTnT.	↓ body mass.	N/A	14
BECKER MUSCULAR DYSTROPHY MODELS
DMD-IF Deletion of exon 3 to 16 Wistar–Imamichi rats. (18)	Histology: ↓ Dystrophin (reduced, truncated) ↑ Fibrosis (mild) ↑ Adipose infiltration (mild) Functional biomarkers: ↓ muscle mass (delayed) ↓ (Late and mild) strength Molecular biomarkers: ↑ CK (mild, serum) ↑ Titin (mild, urines)	Histology: ↑ Fibrosis ↑ Inflammation Function: ↑ Cardiac dysfunction (delayed and mild)	N/A	N/A	18
R-BMDdel45-47 Deletion of 36,854 bp including exons 45 to 47 Sprague Dawley (16)	Histology: ↓ Dystrophin (levels, truncated) ↑ Fibrosis (mild) = Repair (not affected). Functional defect: ↓ (Late and mild) strength Molecular defect: ↓ nNOS. Molecular biomarker: ↑ (mild) CK (serum)	Histology: ↑ Cardiomyopathy (severe and delayed) ↑ Fibrosis (delayed) Functional defects: ↓ Cardiac output and contractility ↑ Heart rate variability and arrhythmias. ↑ Hypertrophic remodeling ↓ Ejection fraction Molecular analyses: ↑ Stress response (snRNA-seq) ↓ Tmem65 mislocalized Cx43.	N/A	N/A	16
Dmd Δ45 Deletion of exon 45 Sprague Dawley (19) Moderately reduced lifespan.	Histology: ↑ Fibrosis (variable) ↑ Regeneration (transient) ↑ Inflammation (transient) Functional defects: ↓ Muscle mass ↓ Strength ↓ Endurance ↓ Neuromuscular activity. Molecular biomarkers: ↑ CK (serum, transient) ↑ MYOM3 (serum, dynamic) ↓ Dystrophin (reduced levels, transient, spontaneous exon skipping and revertant fibers).	Histology: ↑ Fibrosis (delayed) ↑ Inflammation (late and moderate) Functional defects: ↑ Pulmonary hypertension (late) ↑ Diastolic dysfunction in aged animals ↓ Ejection fraction Molecular analysis: ↑ immune-related pathways	Moderately reduced lifespan. Cognitive defects: ↓ exploratory behavior ↓ working memory. ↑ anxiety	N/A	19

Overview of DMD and BMD rat models

A variety of DMD rat models have been developed (Table 1) to more accurately recapitulate the spectrum of mutations observed in dystrophin deficient patients [2]. In 2014, two landmark studies independently established the first DMD rat models: The Dmd^mdx rat line targeted the exon 23 [10], while the W-Dmd^em1Kykn (cDMDR) lines carried out-of-frame alleles in exons 3 or 16 [11]. Later, a distinct DMD rat model, the Dmd-KO rat line, was generated by introducing a targeted deletion of exons 22 through 26 [12], and the R-DMDdel52 rat model featured a deletion of exon 52, a region within the major human mutation hotspot [13]. More recently, the R-DMDdup10-17 model was engineered with a duplication of exons 10 to 17, thereby modelling pathogenic genomic duplications found in DMD patients [14]. Finally, Nakamura et al. also reported a DMD-KO line with a 14 bp deletion in exon 3 leading to complete absence of dystrophin [15].

Rat models mimicking BMD have also been generated. While the R-BMDdel45–47 model [16], encoding an internally truncated dystrophin protein, was designed to delete exons 45–47 to mimic the most common BMD mutations seen in patients [17], the del3–16 rat was selected among the rats generated by targeting exons 3 and 16 as it expresses an in-frame truncated dystrophin [18]. In addition, a recently generated Dmd^Δ45 rat line, carrying a deletion of exon 45 that is frequently mutated in DMD patients, exhibits a milder muscle phenotype than other DMD rat models, due to spontaneous exon 44 skipping, which partially restores the reading frame and results in an age-dependent increase in revertant dystrophin-positive fibers, and a longer survival [19].

Skeletal muscle damage and wasting

DMD rat models consistently replicate the progressive skeletal muscle pathology seen in DMD patients (Table 1). Genetic ablation of dystrophin reproducibly leads to a near-complete loss of protein and fewer than 5% –albeit variable between the models– of dystrophin-positive revertant fibers [10, 11, 13, 14]. Histological analyses revealed early, widespread myofiber necrosis with declining regenerative capacity, progressing to severe muscle degeneration accompanied by fibrosis, fiber size variation, inflammation, fat infiltration, and fiber-type remodeling — features consistent with advanced human DMD pathology (Table 1).

While these alterations affect most muscle groups, the extraocular muscles are spared similarly to DMD patients [20], and the tongue is also less affected than the masseter [21]. The extensive necrosis in skeletal muscles leads to release of abundant proteins that can be detected in the blood (creatine kinase, CK, or myomesin-3, MYOM3), and in the urine (i.e. titin fragments) (Table 1), providing minimally invasive and reliable biomarkers. Functionally, muscle wasting correlates with reduced voluntary activity, muscle force, endurance and impaired in vivo contractility, while neuromuscular junction disruption and early denervation contribute to functional decline [9, 11, 13, 21].

In vivo single-cell and single-nucleus transcriptomics have further revealed profound microenvironmental remodeling, including an expansion of fibro-adipogenic progenitors with human-like pro-fibrotic signatures such as elevated COMP [13]. Immune infiltrates display mixed M1/M2 phenotypes, and extracellular matrix genes are upregulated in stromal and endothelial cells [13]. Despite their preserved numbers, muscle stem (satellite) cells exhibit hallmarks of senescence, such as elevated p16^Ink4a, impaired proliferation/differentiation, and cell cycle arrest, leading to precocious regenerative failure [20, 22]. These senescent traits in muscle stem cells and the resulting loss of regenerative capacity —identified in R-DMDdel52, cDMDR rats and DMD patient biopsies— were not observed in mdx mice due to their milder pathology, highlighting the relevance of DMD rat models for uncovering early, patient-relevant disease mechanisms [23].

Cardiac muscle impairment

Dystrophin-deficient rats show early and extensive myocardial remodeling (Table 1), widespread inflammatory infiltration and fibrosis [24]. Lesions have been detected as early as 3 months and progressively worsen with age, closely resembling human DMD pathology. Functionally, echocardiographic and hemodynamic studies have revealed ventricular chamber dilation, early systolic and diastolic dysfunction, and a progressive heart failure with reduced ejection fraction phenotype, along with conduction defects such as QT interval changes and arrhythmias [16, 25].

At the molecular level, transcriptomic analyses have highlighted the upregulation of fibrosis- and stress-related genes, activation of the renin–angiotensin–aldosterone pathway, and impaired calcium handling [25, 26]. Single-nucleus RNA-seq of cardiac muscle has further revealed widespread transcriptional dysregulation across cardiomyocytes, fibroblasts, endothelial, and immune cells, implicating non-myocyte populations in adverse remodeling [13].

Neurological alterations

DMD rats show reduced brain volume and elevated GABA and N-acetylaspartate in the prefrontal cortex, with increased hippocampal GABA, indicating disrupted inhibitory neurotransmission and osmotic imbalance paralleling cognitive deficits in DMD patients [27]. Behavioral studies have reported mild neuromotor deficits and heightened stress responses, potentially linked to loss of the full-length Dp427 dystrophin isoform in cerebellar and limbic regions [28]. Molecular analyses have highlighted early neuronal alterations, including S100β and microtubule-associated protein tauupregulation in peripheral nerves, suggesting ongoing denervation [29]. At the neuromuscular junction, rats show pronounced structural disruption and impaired transmission, contributing to reduced excitability and greater injury-induced force loss [12]. Neurobehavioral assessments have further demonstrated stress sensitivity and deficits in complex motor and cognitive tasks, mirroring neuropsychiatric features seen in some DMD patients [30].

Bone defects

In a study examining bone health in the W-Dmd^em1Kykn rat model, bone mineral density was significantly reduced at both 15 and 30 weeks of age compared to controls. While trabecular bone volume and number were similar at 15 weeks, both were significantly lower in DMD rats by 30 weeks, accompanied by deteriorated microstructural parameters such as reduced connectivity density and altered structure model index, indicating an age-related trabecular bone loss and microstructural deterioration, reflecting bone fragility associated with disease progression [31].

BMD rat models

R-BMDdel45-47 rats express reduced but detectable levels of dystrophin, resulting in milder muscle pathology, less pronounced fibrosis, and preserved ambulation and strength, closely paralleling the clinical course of Becker patients [16]. Similarly, the DMD-IF rat expresses a proximally truncated dystrophin (exon 3–16 deletion) that retains key functional domains necessary for partial sarcolemma stability, but with moderate increases in muscle damage, modest fibrosis, and delayed onset of functional deficits compared to DMD models. Cardiac involvement in these models is delayed but progresses into a severe phenotype at a later stage (6 to 12 months) of the disease [18].

The preservation of dystrophin expression in these models (Table 1) allows for the study of genotype-phenotype relationships and for the evaluation of exon-skipping and gene-editing therapeutic strategies generating in-frame deletions. Together, these Becker-like rat models provide essential comparative platforms for unraveling the molecular basis of dystrophin function, disease progression, and therapeutic response in the spectrum of dystrophinopathies.

Therapeutic evaluations in DMD rat models

Therapeutic evaluation in DMD rat models has rapidly progressed, leveraging the rat’s closer physiological resemblance to human disease and enabling a translational platform for viral, pharmacological and regenerative interventions (Table 1).

Vector-based microdystrophin and gene delivery

Gene replacement therapy with microdystrophin vectors has been assessed in DMD^mdx rats, confirming restoration of the dystrophin-associated protein complex in both skeletal and cardiac muscle, and demonstrating improved muscle force and reduced pathology [32–34]. Notably, transgene expression is maintained long-term and significantly extends the average lifespan of DMD^mdx rats from ~ 15 months to over 25 months [34], a critical outcome toward the successful translation of gene therapy for DMD.

Cell therapy

Immunodeficient DMD rat lines have been created by combining Dmd-null and “nude” background, enabling long-term engraftment and functional evaluation of human myogenic cell transplantation. These rats have been used to demonstrate the successful engraftment and differentiation of human myoblasts [35].

Pharmacological interventions

Targeted pharmacological interventions, including senolytics (e.g., ABT263) to eliminate senescent satellite cells [22] and small molecules such as forskolin to rescue stem cell regenerative capacity [20, 36], have demonstrated substantial improvements in muscle histology, strength, and delay of disease progression. Unfortunately, long term forskolin treatment leads to cardiac impairments [36]. Beyond muscle regeneration, immunomodulatory strategies have been investigated: anti-CD45RC monoclonal antibodies promote immune tolerance and reduced muscle inflammation in DMD rats, pointing to potential immunotherapy approaches [37]. In addition, TEI-SARM2, a selective androgen receptor modulator shows slightly improved muscle function (grip, tetanic force) without side effects [15]. Stabilizing calcium homeostasis improves the muscle phenotype, with even greater benefits observed when combined with AAV-microdystrophin therapy [38]. Cardiac-targeted therapies such as ivabradine, a heart rate–lowering agent, offers cardioprotective effects, improves cardiac output and reduces heart rate in DMD rats [25, 26, 36].

Metabolic supplementation

While giving positive outcome in mdx mice, taurine fails to improve muscle phenotype or strength in DMD rats [39] highlighting the limitations of dietary interventions. In addition, a medium‑chain triglyceride containing ketogenic diet improves skeletal muscle [40, 41] but exacerbates cardiomyopathy [42].

Genetic modification

Genetic ablation of p16 (a tumor suppressor gene) improves the dystrophic phenotype of a DMD rat, but adverse effects in dystrophic rats are evident by about 9 months as these animals develop rhabdomyosarcoma [43].

Heterogeneity across DMD rat models and implications for model selection

Direct comparison between DMD rat models is challenging, as methodologies and disease stage at analysis vary between studies. Although all DMD rat models recapitulate the core features of severe dystrophinopathy — complete or near-complete loss of dystrophin, myofiber necrosis associated with elevated blood CK/MYOM3, inflammation and fibrosis, reduced muscle force, and cardiomyopathy — disease severity, progression rate, and organ involvement differ between lines. Nonetheless, several key factors account for phenotypic differences: (i) mutation type and location — point mutations, deletions or duplications leading to out-of-frame deletions generally abolish dystrophin expression and cause a severe DMD pathology in rats, whereas in-frame mutations may retain partial function (i.e. BMD models). Importantly, some mutations also affect specific dystrophin isoforms encoded by internal promoters, while others spare them, contributing to additional phenotypes such as specific cognitive impairments. This aspect is particularly relevant when selecting models for therapies that must target both muscle and non-muscle isoforms; (ii) spontaneous exon skipping or revertant fibers — for example, the Dmd^Δ45 line exhibit partial exon skipping that mitigates disease severity; (iii) genetic background/strain effects, which can modulate inflammation, regenerative capacity, and cardiac susceptibility; and (iv) method of model generation, as CRISPR guide choice, mosaicism, or off-target events can alter disease onset and severity.

For example, R-DMDdel52 and R-DMDdup10–17 rats show early, rapidly progressive, and severe skeletal and cardiac pathology with a median survival of about 11–12 months, whereas the original DMD^mdx line exhibits severe but slightly delayed skeletal and cardiac involvement with slightly longer median survival (~ 13 months). In contrast, DmdΔ45 and Becker-like models such as R-BMDdel45–47 and DMD-IF display a milder, later-onset phenotype due to expression of a truncated dystrophin.

These differences, reflecting mutation class, isoform involvement, spontaneous exon skipping or revertant fiber formation, and genetic background, directly influence muscle, cardiac, and even cognitive pathology. Recognizing this phenotypic spectrum, together with the genetic basis of each model, is essential for selecting the most appropriate system for a given therapeutic evaluation.

Conclusions: rat DMD models provide a robust preclinical platform

DMD rat models occupy a valuable intermediate position between traditional mouse models and larger animals such as dogs or pigs. While canine and porcine models offer high physiological relevance and more easily share environmental factors, their use is constrained by high costs, reduced ethical acceptance, and logistical complexity, limiting them primarily to late-stage therapeutic validation.

A key advantage of rat models is their size, approximately ten to twenty times larger than mice depending on their age, which enables more precise physiological, histological, behavioral, and imaging analyses, while remaining significantly more accessible and cost-effective than large animal models. Notably, DMD rats exhibit a more severe and progressive phenotype than mdx mice, with pathological features that more closely resemble the human disease, enhancing their value for both mechanistic investigations and therapeutic testing.

Although mouse models remain indispensable for early-stage research due to their genetic tractability, extensive transgenic resources, and low costs, rats offer distinct advantages. Their broader behavioral repertoire and more complex social interactions allow for refined assessments of motor coordination, cognition, and emotional responses, all features that are challenging to evaluate in mice. Combining manageable handling, moderate costs, and progressive disease phenotypes, DMD rat models represent a compelling platform for translational research. Their use has already led to important discoveries, including the identification of a progressive decline in muscle stem cell regenerative capacity [20], the demonstration that cAMP/PKA pathway stimulation can restore muscle repair [36] or AAV-microdystrophin preclinical evaluations [32, 34].

In summary, DMD rats should be considered complementary to —rather than replacements for—mouse and large animal models. The optimal choice of a model should be tailored to the research question, the mechanism of interest, and the stage of preclinical development. No single animal model fully recapitulates the complexity of human DMD, and model selection must be guided by specific study objectives, disease severity, experimental scale, and translational endpoints. Integrating data from multiple models across the research pipeline offers the most robust foundation for clinical translation. Used strategically, DMD rat models are poised to accelerate the path from discovery to therapy evaluation in dystrophinopathies.

Abbreviations

AAV

Adeno-associated virus

CK

Creatine kinase

COMP

Cartilage oligomeric matrix protein

DMD

Duchenne muscular dystrophy

DMD-KO

DMD knockout (model)

DMD-IF

In-frame Dmd mutation model

GFAP

Glial fibrillary acidic protein

GABA

Gamma-aminobutyric acid

hs-cTnT

High-sensitivity cardiac troponin T

MRI

Magnetic resonance imaging

MYOM3

Myomesin-3

NAA

N-acetylaspartate

nNOS

Neuronal nitric oxide synthase (NOS1)

SARM

Selective androgen receptor modulator

TRPC1 / TRPC3

Transient receptor potential canonical

Boxed insert in the review

Engaging with World Duchenne Awareness Day and Advancing Global Health Priorities

Despite remarkable scientific progress, there is still no cure for DMD. This review, which offers the first comprehensive synthesis of DMD rat models, aims to support ongoing international efforts to accelerate therapeutic discovery and deepen our understanding of the disease. It is aligned with the objectives of World Duchenne Awareness Day, recognized annually on September 7 by the United Nations (https://www.un.org/en/observances/duchenne-awareness-day). By aligning with this important observance, this review contributes to a global call for greater awareness, coordinated research efforts, and sustained investment in translational science that can meaningfully improve patient outcomes.

In addition, this review contributes to the realization of the United Nations Sustainable Development Goal 3 — Ensure healthy lives and promote well-being for all at all ages — by providing a timely and comprehensive analysis of novel preclinical models that offer significant improvements over existing systems. In particular, DMD rat models reproduce the complexity and severity of human pathology more accurately than traditional mouse models. Their adoption will be key to testing next-generation therapies, improving translational validity, and ultimately accelerating the development of effective treatments.

Finally, this review supports Sustainable Development Goal 8 — Promote sustained, inclusive and sustainable economic growth, full and productive employment and decent work for all — by acknowledging that chronic childhood diseases like DMD significantly reduce workforce participation, not only for patients but also for families and caregivers. Therapeutic innovations that delay or mitigate disease progression can have direct economic benefits by extending the productive years of individuals with DMD, supporting their access to employment, and reducing long-term caregiving burdens. By aligning with global awareness efforts and sustainable development goals, this review aims not only to consolidate scientific progress but also to emphasize the broader societal implications of advancing care and research for Duchenne muscular dystrophy.

Author contributions

FR wrote the first draft of the review; PL generated the first draft of the table; PL and LT formatted the bibliography; PL, VT and LT substantially revised the text and table content. All co-authors revised the text.

Funding

Financial support was obtained from the “Association Française contre les Myopathies” (AFM) via TRANSLAMUSCLE I and II programs (projects 19507 and 22946).

Data availability

No datasets were generated or analysed during the current study.

Declarations

Ethics approval and consent to participate

Not applicable.

Consent for publication

All authors have read and approved the final manuscript and consent to its submission and publication.

Competing interests

The authors declare no competing interests.