PPMO-based exon skipping therapy improves respiratory function in the mdx mouse model of Duchenne muscular dystrophy

Abstract

Duchenne muscular dystrophy (DMD) is a severe X-linked recessive disorder caused by a deficiency of dystrophin, leading to progressive muscle degeneration and eventually cardiorespiratory failure. Exon-skipping therapies using cell-penetrating peptide-conjugated phosphorodiamidate morpholino oligomers (PPMOs) restore production of a shortened but functional dystrophin protein. Since respiratory insufficiency is the leading cause of morbidity and mortality in DMD, we sought to examine the impact of PPMO on respiratory pathology. We evaluated the effects of RC-1001, a PPMO targeting a dystrophin mutation, in mdx mice, a preclinical model of DMD. These mice received monthly intravenous doses of RC-1001 (30, 50, or 100 mg/kg), starting at 2 months of age, and were monitored until the study endpoint at 12 months of age. Respiratory function was evaluated using whole-body plethysmography and the forced oscillometry technique, followed by histological and molecular analysis of respiratory muscles. PPMO-treated mice showed dose-independent improvements in respiratory function, with postmortem studies revealing significant dystrophin restoration, reduced inflammation, and decreased fibrosis in respiratory muscles. Additionally, dystrophin restoration and strength improvements were observed in limb muscles. Overall, PPMO-mediated exon skipping effectively targets respiratory pathology and is a promising therapy for respiratory insufficiency in patients with DMD.

Graphical abstract

Duchenne muscular dystrophy results in respiratory muscle decline due to the absence of dystrophin. RC-1001 exon-skipping therapy in mdx mice restored dystrophin, reduced inflammation and fibrosis, and improved respiratory function. These findings suggest that PPMO-based exon therapy effectively mitigates respiratory impairment in DMD.

Introduction

Duchenne muscular dystrophy (DMD) is an X-linked recessive disease caused by mutations in the DMD gene that lead to dystrophin deficiency.¹ Dystrophin connects the intracellular cytoskeleton to the extracellular matrix and plays a crucial role in muscle cell membrane stabilization during muscle contraction.^2,3,4,5 The lack of dystrophin results in contraction-induced muscle damage, which leads to chronic inflammation, fibrosis, and progressive muscle degeneration.^4,6,7 Patients with DMD typically begin to experience muscle weakness at 2–3 years of age, which progresses to a loss of ambulation by 10–12 years of age. At that time, these patients also develop progressive respiratory muscle weakness, which often leads to respiratory failure within the second or third decade of life.^8,9,10,11

Exon-skipping therapy using antisense oligonucleotides (AOs) (synthetic nucleic acid analogs) targets the exon containing a nonsense mutation. This approach alters the mRNA structure to restore the correct reading frame and enable the production of a functional, but shortened, dystrophin protein.^12,13,14 The Food and Drug Administration (FDA) has approved several phosphorodiamidate morpholino oligomer (PMO) exon-skipping therapies for DMD. These PMOs can be used in patients with exon 51-, 53-, and 45-eligible skipping mutations.^{15,16,17,18,19} Treatment with PMO increases the production of an internally truncated but functional dystrophin in skeletal muscles.^{20,21,22,23,24} However, PMOs require weekly infusion, and their cell penetration is suboptimal.²⁵

Peptide-conjugated PMOs (PPMOs) are cell-penetrating peptides linked to the PMO backbone, which enhance cellular uptake of AOs, thereby improving exon-skipping efficiency and dystrophin restoration. PPMOs allow for a decreased frequency of dosing compared to PMOs and result in improved pathology.^26,27,28,29 However, the impact of PPMO therapies on respiratory muscle structure and function is unknown. In this study, we investigated the efficacy of RC-1001, a DMD exon 23-skipping PPMO, in improving respiratory pathology in the mdx mouse model of DMD.³⁰ Mdx mice carry a nonsense mutation in exon 23 of the dystrophin gene and are the most commonly used mouse model for preclinical studies of DMD.^31,32 These mice exhibit muscle fibrosis and inflammation, as well as respiratory function deficits, rendering them an ideal model to examine the impact of PPMO on respiratory muscle structure and function.^33,34,35,36 The primary objective of this study was to evaluate whether RC-1001 restores dystrophin expression and reduces respiratory muscle pathology. In addition, we examined the impact of RC-1001 on respiratory insufficiency and pulmonary mechanics in mdx mice.

Results

RC-1001, a proprietary PPMO, targets mouse dystrophin exon 23 to restore the reading frame and allow for production of a truncated but functional dystrophin protein.³⁰ mdx mice received monthly intravenous (i.v.) RC-1001 (via tail-vein injection) from 8 to 48 weeks of age. The treated mice were divided into three groups based on the dose they received throughout the study: a low dose of 30 mg/kg (n = 8), an intermediate dose of 50 mg/kg (n = 8), and a high dose of 100 mg/kg (n = 8). As controls, C57BL/10ScSnJ wild-type (hereafter referred to as “WT”) mice (n = 8) and saline-treated mdx mice (n = 8) served as disease controls (Figure 1A). Mice underwent whole-body plethysmography (WBP) and behavioral testing every month until they were 52 weeks old. Monthly assessments were conducted 1 day prior to treatment with RC-1001. At 52 weeks of age, mice were evaluated for pulmonary mechanics using the forced oscillometry technique and euthanized for postmortem analysis (Figure 1B).

Figure 1.

Experimental design of the study

(A) Schematic representation of PPMO and saline treatment in WT and mdx mice. mdx mice were treated with three doses of PPMO—30 mg/kg (low dose), 50 mg/kg (intermediate dose), and 100 mg/kg (high dose). As controls, WT and mdx mice were treated with saline. PPMO was injected monthly via intravenous tail vein injections.

(B) Schematic representation of the study timeline. Mice were injected with either saline or PPMO at 8 weeks of age and then monitored monthly for breathing using WBP and for behavioral analysis until the end of the study. At the study endpoint, FlexiVent was used to assess respiratory mechanics prior to euthanizing the mice at 52 weeks of age (1 year) for postmortem analysis.

Over the course of the study, four mice died: a WT mouse (n = 1; died at 48 weeks of age), an mdx mouse receiving 30 mg/kg PPMO (n = 1; died at 15 weeks of age), one mdx mouse receiving 50 mg/kg PPMO (n = 1; died at 52 weeks of age), and one mdx mouse receiving 100 mg/kg PPMO (n = 1; died at 40 weeks of age). Therefore, although 8 mice were initially assigned to each treated and untreated cohort, data are presented from n = 7 for each group.

RC-1001 restores the expression of dystrophin in mdx mice

We assessed the dose-dependent effects of RC-1001 by evaluating exon 23-skipping efficiency and dystrophin protein restoration across multiple tissues. These included muscles regulating breathing (diaphragm, tongue, intercostal, and parasternal muscles) (Figure 2), limb muscles (tibialis anterior [TA] and quadriceps), and cardiac muscles (Figure S1). Treatment with RC-1001 resulted in a significant induction of exon 23 skipping in respiratory muscles. mdx mice treated with RC-1001 at doses of 30, 50, and 100 mg/kg showed 28%, 30%, and 50% exon-skipping levels in the diaphragm, (Figure 2A), 23.5%, 31%, and 44.5% in the tongue (Figure 2B), 49.7%, 53.7%, and 73.9% in the intercostal muscles (Figure 2C), and 52.8%, 49%, and 80% in the parasternal muscles, respectively (Figure 2D) (Figures 2A–2D; p < 0.05, PPMO-treated vs. saline-treated mdx mice).

Figure 2.

Restoration of dystrophin in muscles

(A–C) Exon skipping analysis of diaphragm (A), tongue (B), parasternal (C), and intercostal (D) by qPCR.

(E–H) Dystrophin protein quantification in diaphragm (E), tongue (F), parasternal (G), and intercostal (H) by western blot. Statistical significance was determined using one-way ANOVA, followed by an uncorrected Fisher’s LSD test (∗p < 0.05; ∗∗p < 0.01; ∗∗∗p < 0.001; ∗∗∗p < 0.001).

(I–L) Representative western blots showing dystrophin expression in diaphragm (I), tongue (J), parasternal (K), and intercostal (L).

Concordantly, treatment with RC-1001 significantly increases dystrophin expression in the respiratory muscles of mdx mice (Figures 2E–2H). The levels of dystrophin expressed in the mdx mice treated with RC-1001 at doses of 30, 50, and 100 mg/kg are 42.6%, 41.0%, and 51.3% in the diaphragm (Figure 2E); 29.0%, 29.0%, and 52.2% in the tongue (Figure 2F); 47.3%, 52.5%, and 71.7% in the intercostals (Figure 2G); and 55.5%, 49.8%, and 69.0% in the parasternal muscles (Figure 2H), respectively. (Figures 2E–2H; p < 0.001 RC-1001 treated vs. saline-treated mdx). While treatment with PPMO significantly increases dystrophin expression in treated mdx mice compared to saline-treated mdx controls (p < 0.05, PPMO-treated vs. saline-treated mdx mice), the expression of dystrophin in tissues from PPMO-treated mice is still significantly lower than in tissues from saline-treated WT mice (p < 0.05, saline-treated WT vs. PPMO-treated mdx mice). This is consistent with a partial restoration of dystrophin in mdx mice with PPMO treatment. Representative western blots for the respiratory muscles—diaphragm (Figure 2I), tongue (Figure 2J), intercostal (Figure 2K), and parasternal muscles (Figure 2L)—show robust expression of dystrophin.

We also examined the limb muscles, including TA, quadriceps, and cardiac muscle, by quantifying exon-skipping efficiency (Figures S1A–S1C) and performing western blot analysis for dystrophin expression (Figures S1D–S1H). Similar to respiratory muscles, limb muscles exhibited a dose-dependent increase in exon skipping, including the TA and quadriceps (Figures S1A and S1B). The extent of exon-skipping in mdx mice treated with RC-1001 at doses of 30, 50, and 100 mg/kg is 44.3%, 51%, and 79.7% in the TA (Figure S1A) and 55%, 57.1%, and 79.5% in the quadriceps (Figure S1B), respectively. The levels of dystrophin expression in mdx mice treated with RC-1001 at doses of 30, 50, and 100 mg/kg are 43.7%, 41.8%, and 56.3% in TA (Figures S1D and S1G) and 40.4%, 44.9%, and 81.3% in quadriceps (Figures S1E and S1H). respectively. Limb muscles also demonstrate efficient restoration of dystrophin in RC-1001-treated mdx mice (Figures S1D, S1E, S1G, and S1H). In contrast, modest but significant levels of exon skipping (Figure S1C) and dystrophin expression (Figures S1F and S1I) are detected in the cardiac muscle of RC-1001-treated mdx mice compared to saline-treated mdx mice. Exon skipping and dystrophin expression in cardiac tissue are minimal compared to skeletal and respiratory muscles. Specifically, PPMO-treated mdx mice show only 2.5%, 4.2%, and 12.8% exon skipping and 0.9%, 2.2%, and 9% dystrophin expression with doses of 30, 50, and 100 mg/kg of RC-1001, respectively.

RC-1001 improves muscle histopathology and morphology in mdx mice

Next, we investigated the effects of RC-1001 on histopathological changes in key respiratory muscles, including the diaphragm, a major inspiratory muscle (Figure 3), and the tongue, which is important for upper airway patency (Figure 4).^37,38,39,40 Immunohistochemistry (IHC) of dystrophin expression shows restoration of dystrophin in the diaphragm (Figure 3A) and the tongue (Figure S2A) of mdx mice. Histological analysis using hematoxylin and eosin (H&E) staining reveals significantly elevated inflammation in saline-treated mdx mice, reflected by the area infiltrated by immune cells in the diaphragm (21.4%) (Figures 3B, 3E) and tongue (9%) (Figures S2B and S2E) compared to WT diaphragm (1.6%) and tongue (3.4%) (p < 0.0001 in the diaphragm, p < 0.01 in tongue muscles of saline-treated mdx vs. WT). PPMO treatment of mdx mice with 30, 50, and 100 mg/kg of RC-1001 reduces the area of immune cell infiltration to 10.5%, 6%, and 4.5% in the diaphragm (Figure 3B, 3E), and 4.7%, 3.2%, and 3.6% in the tongue (Figures S2B, S2E), respectively (p < 0.05, RC-1001-treated vs. saline-treated mdx mice).

Figure 3.

Enhanced expression of dystrophin and attenuation of pathology in the diaphragm of PPMO-treated mdx mice

(A–D) Representative images of dystrophin (A), hematoxylin and eosin (B), Picosirius red (C), and laminin (D) staining of the diaphragm from saline- and RC1001-treated WT and mdx mice. Scale bars, 90 μm. The white arrow indicates immune cell infiltration, and the asterisk indicates centralized nuclei.

(E–H) Quantification of the percentage area infiltrated by immune cells (E) and myofibers with centralized nuclei (F) from H&E-stained sections, intensity of Picrosirius red staining (G), and muscle fiber cross-sectional size (minimal Ferret diameter) (H) from laminin-stained images of the diaphragm. Data are presented as mean ± SEM. Statistical significance was determined using a mixed-model two-way ANOVA, followed by Fisher’s LSD test (∗p < 0.05; ∗∗p < 0.01; ∗∗∗p < 0.001; ∗∗∗p < 0.001).

Figure 4.

Improved pulmonary mechanics in PPMO-treated mdx mice

(A–H) Pulmonary mechanics in saline-treated WT and mdx mice and PPMO-treated mdx mice were measured using FlexiVent for overall resistance (Rrs; A), compliance (Crs; B), elastance (Ers; C), Newtonian resistance (Rn; D), tissue dampening (G; E), and tissue elastance (H; F).

Furthermore, mdx mice exhibit a higher proportion of myofibers with centralized nuclei—a hallmark of muscle regeneration and pathology^41,42—in the diaphragm (21.4%) (Figures 3B, 3F) and tongue (30%) (Figures S2B, S2F) compared to WT diaphragm (4.3%) and tongue muscles (2.6%) (p < 0.05, saline-treated-mdx vs. WT). Although RC-1001-treated-mdx mice demonstrate reduced inflammation, the number of myofibers with centralized nuclei remains unchanged in both the diaphragm and tongue. (p > 0.05, RC-1001-treated vs. saline-treated-mdx mice).

Next, using picrosirius red staining, we investigated the accumulation of collagen in the extracellular matrix to assess fibrosis. Compared to WT diaphragm (7%) and tongue (7.6%), collagen deposition is significantly elevated in saline-treated mdx diaphragm (48%) (Figures 3C and 3G) and tongue (17.6%) (Figures S2C, S2G) (p < 0.0001 in diaphragm; p < 0.01 in tongue muscles of saline-treated mdx vs. saline-treated WT). A partial reduction of fibrosis is evident in the diaphragm (20.4%, 19.6%, 13.2%) of mdx mice treated with 30, 50, and 100 mg/kg of RC-1001 (Figures 3C, 3G; p > 0.0001, RC-1001-treated vs. saline-treated mdx mice). However, no impact is observed on fibrosis in the tongue of RC-1001-treated-mdx mice (Figures S2C, S2G; p > 0.05, RC-1001-treated vs. saline-treated-mdx mice).

In addition, morphometric analysis of the diaphragm shows a significant reduction in myofiber size (mean minimal Feret’s diameter) in mdx diaphragms (38.5 μm) (Figure 3D, 3H) and tongue (76 μm) relative to WT diaphragm (66 μm) and tongue (114 μm) (Figures S2D, S2H; p < 0.0001 in the diaphragm and p < 0.01 in the tongue of saline-treated mdx vs. saline-treated WT). RC-1001 increases myofiber diameter in the diaphragm of mdx mice, indicating improved muscle atrophy (Figure 3D, 3H; p < 0.001, RC-1001-treated vs. saline-treated mdx mice). However, myofiber size in the tongue remained reduced regardless of RC-1001 treatment (Figures S2D, S2H; p > 0.05, RC-1001-treated vs. saline-treated mdx mice).

RC-1001 ameliorates respiratory insufficiency in mdx mice

To evaluate the functional impact of exon-skipping therapy on respiratory performance, we assessed awake spontaneous breathing (WBP) and pulmonary mechanics (FlexiVent). Forced oscillation measurements via FlexiVent from the saline-treated mdx mice exhibit significantly impaired pulmonary mechanics compared to saline-treated WT mice. Specifically, saline-treated mdx mice have increased respiratory system resistance (Rrs) (Figure 4A), decreased compliance (Crs) (Figure 4B), and increased elastance (Ers) (Figure 4C), with decreased Newtonian resistance (Rn) (Figure 4D), increased tissue damping (G) (Figure 4E), and increased tissue elastance (H) (Figure 4F) compared to saline-treated WT mice (p < 0.001, saline-treated mdx vs. WT).

RC-1001 treatment significantly improves these mechanical parameters. Compared to saline-treated mdx mice, RC-1001-treated mdx mice have reduced Rrs, increased Crs, and decreased Ers (Figures 4A–4C). Improvements are also observed in Rn, G, and H (Figures 4D–4F) (p < 0.05; RC-1001-treated vs. saline-treated mdx mice). Notably, respiratory mechanics in PPMO-treated mdx mice approach WT values, indicating substantial improvement of pulmonary mechanics in mdx mice (p > 0.05; RC-1001-treated vs. saline-treated WT mice). No significant differences are observed between the three PPMO dose groups in any of the pulmonary mechanical parameters.

To assess the breathing response to a respiratory challenge, we measured respiratory parameters at “baseline” (normoxic) conditions and at “challenge” (hypoxic-hypercapnic) conditions using WBP at 52 weeks (Figure 5). At baseline (FiO₂: 0.21, N₂ balance), no significant differences are observed in minute ventilation (MV), respiratory frequency (F), tidal volume (TV), peak inspiratory flow (PIF), or peak expiratory flow (PEF) between saline-treated WT and mdx and RC-1001-treated-mdx mice (p > 0.05 across all groups). However, during a respiratory challenge of hypoxia and hypercapnia (FiO₂: 0.10, FiCO₂: 0.07, N₂ balance), saline-treated mdx mice had significantly reduced MV (Figure 5A), F (Figure 5B), PIF (Figure 5D), and PEF (Figure 5E) compared to WT controls (p < 0.05; saline-treated mdx mice vs. WT). Although not significant, a downward trend in TV was also observed in mdx mice (Figure 5C).

Figure 5.

PPMO ameliorates respiratory deficits in mdx mice

(A–F) Whole-body plethysmography measurements of respiratory function in 52-week-old saline-treated WT and mdx mice and PPMO-treated mdx mice were performed to evaluate minute ventilation (A), frequency (B), tidal volume (C), peak inspiratory flow (D), peak expiratory flow (E), and enhanced pause (penH) (F) at room air (“baseline”) and during a maximal respiratory challenge with hypoxia and hypercapnia. Data are presented as mean ± SEM. Statistical significance was determined using a mixed-model two-way ANOVA, followed by Fisher’s LSD test. Statistical significance for PenH was determined using one-way ANOVA. (∗p < 0.05; ∗∗p < 0.01; ∗∗∗p < 0.001; ∗∗∗∗p < 0.0001).

PPMO treatment led to significant improvements in all measured ventilatory parameters under challenge conditions. RC-1001-treated mdx mice had increased MV, F, TV, PIF, and PEF relative to saline-treated mdx controls (p < 0.05; RC-1001-treated vs. saline-treated mdx mice). The lowest dose group (30 mg/kg) showed the most pronounced improvement, while higher doses (50 and 100 mg/kg) exhibited greater variability (Table 1).

Table 1.

Ventilation during normoxia and in response to hypoxia-hypercapnic challenge in mice

	Saline treated		RC-1001 treated
	WT	mdx	mdx 30 mg/kg	mdx 50 mg/kg	mdx 100 mg/kg
Baseline (21% O2,N2 balance)
MV	30.9 ± 2.3	35.3 ± 1.6	37.8 ± 2.6	36.7 ± 2.8	34.3 ± 1.6
F	166.5 ± 9.8	171.9 ± 2.4	160.6 ± 6.1	152.2 ± 10.1	148.4 ± 5.8
TV	0.19 ± 0.005	0.21 ± 0.007	0.25 ± 0.01	0.25 ± 0.02∗	0.23 ± 0.01
PIF	2.5 ± 0.15	2.6 ± 0.1	3.1 ± 0.2	3.1 ± 0.2	2.9 ± 0.1
PEF	1.8 ± 0.1	2.8 ± 0.2	2.7 ± 0.3	2.6 ± 0.2	2.5 ± 0.1
PenH	1.4 ± 0.2	2.9 ± 0.2∗	1.9 ± 0.3#	2.1 ± 0.3#	2.0 ± 0.2#
Challenge 10% O2,7% CO2,N2 balance
MV	133.4 ± 7.5	112.3 ± 6.6∗	160.8 ± 6.7∗#	140.1 ± 6.9# $	146.5 ± 9.4#
F	306.9 ± 9.1	279.8 ± 6.1∗	305.7 ± 5.9#	274.1 ± 13.9∗ $	290 ± 11.6
TV	0.44 ± 0.16	0.40 ± 0.02	0.53 ± 0.03∗#	0.50 ± 0.03∗#	0.51 ± 0.03∗#
PIF	7.1 ± 0.5	5.6 ± 0.4∗	9.1 ± 0.6∗#	7.8 ± 0.6# $	8.1 ± 0.5#
PEF	7.6 ± 0.5	6.8 ± 0.4	9.9 ± 0.5∗#	8.7 ± 0.7#	9.4 ± 0.8∗#
PenH	1.4 ± 0.06	1.9 ± 0.05∗	1.7 ± 0.2∗	1.7 ± 0.09∗	1.8 ± 0.06∗

Minute ventilation (MV), frequency (F), tidal volume (TV), peak inspiratory flow (PIF), and peak expiratory flow (PEF) were measured in wild-type, mdx, and PPMO-treated mdx mice at baseline (normoxia; room air) and during a respiratory challenge using WBP. Data are presented as mean ± standard error.

‘∗’p < 0.05 between saline-treated WT vs. saline-treated and RC-1001-treated mdx mice; “#” p < 0.05 between saline-treated mdx vs. RC-1001-treated mdx mice; “$” p < 0.05 between 30 mg/kg RC-1001-treated mdx mice vs. 50 and 100 mg/kg RC-1001-treated mdx mice.

Although we had no a priori expectations of changes in enhanced pause (PenH), a non-invasive surrogate measure of airflow resistance assessed using WBP in a sealed chamber,⁴³ RC-1001 treatment led to a modest but statistically significant reduction in PenH. PenH was significantly elevated in saline-treated mdx mice at baseline compared to WT controls (Figure 5F; p < 0.001). RC-1001 treatment significantly reduced PenH values in mdx mice, bringing them closer to levels observed in WT mice (Figure 5F; p < 0.01 for PPMO-treated vs. saline-treated mdx mice).

RC-1001 increases limb muscle strength in mdx mice

To compare with prior PPMO or PMO animal studies, we examined the impact of RC-1001 on the TA as a representative muscle to evaluate both the histological effects of long-term treatment (Figure S3) and its functional impact on muscle strength (Figure 6). Treatment with RC-1001 resulted in dystrophin expression in the TA muscles of mdx mice (Figure S3A). A significantly increased area of immune cell infiltration was observed in mdx mice 6.5%) compared to saline-treated WT TA muscles (0.7%). However, RC-1001 treatment at doses of 30, 50, and 100 mg/kg in mdx mice markedly reduced inflammation to 0.7%, 0.8%, and 0.7%, compared to WT muscles (Figures S3B, S3E). Similar to the diaphragm and tongue muscles, there is an increase of centralized nuclei in the myofibers of mdx TA (45.9%) compared to WT (4.9%). No differences are observed in myofibers with centralized nuclei between saline- and RC-1001-treated mdx mice (60.5%, 68.5%, and 59.6%) (Figures S3B, S3F). When evaluated for fibrosis, collagen accumulation was increased in mdx mice (9.1%) compared to WT (4.5%) but was substantially reduced in RC-1001-treated mdx mice (4.9%, 4.1%, and 5.5%) (Figures S3C, S3G). Similarly, TA fiber size was significantly decreased in mdx mice (71.5 μ) compared to WT (99 μ) and increased with all doses of RC-1001 treatment (106, 101, and 110 μ). (Figures S3D, S3H). At baseline (8 weeks of age), prior to RC-1001 treatment, mdx mice had a significant decrease in forelimb grip strength compared to WT (p < 0.01), and this persisted until the study endpoint (52 weeks). Although differences in strength were negligible between RC-1001-treated and saline-treated mdx mice (p > 0.05) at younger ages, an increase in strength was evident in the RC-1001-treated groups at 32 weeks of age and continued until the study endpoint (Figure 6) (p < 0.05, saline vs. RC-1001-treated mdx mice). The mdx mice treated with the highest dose of RC-1001 (100 mg/kg) had the maximum gain in strength (p > 0.05, WT vs. 100 mg/kg PPMO-treated mice), while lower (30 mg/kg) and intermittent doses (50 mg/kg) showed a trend toward improved strength (p < 0.05, WT and untreated mdx vs. lower and intermittent dose PPMO-treated mdx mice) (Figure 6A).

Figure 6.

PPMO improves muscle strength in mdx mice

(A–B) Measurements of grip strength (A) and wire hang test (B) in 52-week-old saline-treated WT and mdx mice and PPMO-treated mdx mice. Data are presented as mean ± SEM. Statistical significance was determined using a mixed-model two-way ANOVA, followed by Fisher’s LSD test. Statistical significance for PenH was determined using one-way ANOVA. (∗p < 0.05; ∗∗p < 0.01; ∗∗∗p < 0.001; ∗∗∗∗p < 0.0001).

Similar results were observed with the wire hang test. Irrespective of treatment, at a younger age, mdx mice were unable to hang for long and tended to have a shorter latency to fall compared to saline-treated WT mice. However, starting at 16 weeks of age, saline-treated mdx mice continued to fall much earlier than WT and RC-1001-treated mdx mice, and this trend persisted until 52 weeks of age (p < 0.05, saline-treated vs. PPMO-treated mdx mice) (Figure 6B). In summary, long-term treatment with RC-1001 significantly improved muscle strength and provided sustained motor endurance in mdx mice.

Discussion

The main outcome of this study is that RC-1001, a proprietary PPMO, improves respiratory insufficiency in the DMD mdx mouse. Long-term administration of RC-1001 at three different doses results in exon skipping and robust expression of a truncated yet functional dystrophin protein in the skeletal muscles studied, including the muscles of respiration and the TA. Further, this treatment significantly decreased inflammation and fibrosis in these muscles. Ultimately, all three tested doses of RC-1001 alleviate respiratory histopathology, improve respiratory function, and enhance pulmonary mechanics in mdx mice.

Exon skipping in DMD

Dystrophin maintains the structural integrity of muscle fibers by anchoring the cytoskeleton to the extracellular matrix through the dystrophin-associated glycoprotein complex (DGC). This connection is important for distributing mechanical stress during muscle contraction.^2,4,44,45,46 In DMD, the absence of functional dystrophin makes muscle fibers highly vulnerable to contraction-induced damage, triggering a pathological cascade of muscle degeneration, chronic inflammation, and fibrosis. The sustained loss of functional myofibers, extensive fibrotic remodeling, and the diminished regenerative capacity of muscles result in progressive muscle weakness.^{35,47,48,49,50}

Exon-skipping therapies are FDA-approved treatments that use AOs to modulate pre-mRNA splicing and produce a partially functional dystrophin transcript.^51,52,53 Currently, four different PMO therapies are FDA approved for patients with DMD: eteplirsen (Exondys 51), golodirsen (Vyondys 53), viltolarsen (Viltepso), and casimersen (Amondys 45). These therapies target mutations amenable to exon 51, 53, and 45 skipping, respectively. However, a weekly dose of PMO is required to maintain effective exon-skipping levels. A previous longitudinal clinical study of patients treated with eteplirsen—an FDA-approved PMO therapy—reported slowing of the rate of decline in respiratory parameters but an inability to reverse respiratory pathology.²⁰ PPMO is the next generation of PMO, conjugated with a cell-penetrating peptide that enhances cellular uptake and is designed to overcome the delivery limitations of traditional PMOs.^16,17,18,19

Previous studies demonstrated that repeated monthly dosing of RC-1001 (PPMO) significantly outperformed AVI-4225 (PMO) treatment in mdx mice.³⁰ After three doses of 40 mg/kg, RC-1001 produced exon-skipping rates of 98.8% in quadriceps, 87.4% in diaphragm, and 43.2% in heart, with corresponding dystrophin levels of 38.9%, 50.9%, and 8.6%. In comparison, PMO-treated (AVI-4225) mdx mice showed much lower exon skipping (53.4%, 0.5%, 0%) and dystrophin expression (8.5%, 1.1%, 0%).³⁰ Based on these findings, we focused this study on PPMO-based treatment to address respiratory dysfunction in DMD.

In this study, we evaluated the impact of the PPMO RC-1001 administered over a year. RC1001 is designed to induce exon 23 skipping in the mdx dystrophin gene and to improve respiratory function in the DMD mdx mouse model. A previous study using RC-1001 demonstrated that monthly administration at 40 mg/kg significantly increased exon skipping in the diaphragm—reaching 87.4% after 3 months—compared to 49.1% following a single injection. This repeated dosing also led to markedly higher dystrophin expression in the diaphragm.³⁰ Building on these findings, we administered RC-1001 monthly at three different doses (30, 50, and 100 mg/kg) in mdx mice, starting at 8 weeks of age. We observed significant exon skipping with noteworthy dystrophin expression in the respiratory muscles at all three doses. However, exon skipping and dystrophin expression were dose dependent. Notably, muscles from mdx mice treated with 100 mg/kg RC-1001 showed significantly higher exon skipping and dystrophin levels compared to the lower dose groups.

Although exon skipping in the diaphragm peaked at the highest dose, dystrophin expression was similar across all doses. We also observed significant functional recovery in limb muscles, as measured by both grip strength and latency to fall in the wire hang test. Gan et al. reported improvements in grip strength following RC-1001 treatment, which aligns with our findings.³⁰ Overall, RC-1001 treatment in mdx mice resulted in dose-dependent exon 23 skipping and partial restoration of dystrophin expression, particularly in respiratory and limb muscles, with limited efficacy in cardiac tissue.

Analysis of muscle structure revealed that PPMO-based treatment reduces inflammation in tissues, including the diaphragm, tongue, and TA. However, in contrast to the diaphragm and TA, tongue muscles exhibited fibrosis and smaller-sized fibers at all treatment doses. The tongue also showed decreased exon skipping and dystrophin expression at the lower doses. The tongue comprises interwoven intrinsic and extrinsic muscles oriented in multiple directions (longitudinal, transverse, and vertical), which may hinder uniform therapeutic delivery and lead to uneven distribution across muscle layers. Additionally, unlike limb muscles, the tongue is innervated by multiple cranial nerves responsible for motor, sensory, and taste functions. This complex innervation could influence local immune responses, therapeutic uptake, and functional outcomes. The tongue contains a distinct mix of slow- and fast-twitch muscle fibers arranged in a complex spatial pattern. This heterogeneity may affect how the muscle responds to dystrophin restoration and remodeling stimuli.

Notably, despite reduced inflammation following PPMO treatment, the number of centralized nuclei remains significantly elevated in the diaphragm, tongue, and TA muscles after chronic administration of RC-1001. Centralized nuclei are characteristic of regenerating myofibers and are frequently observed in dystrophic muscle. Compared to WT mice, mdx myofibers exhibit a higher number of centralized nuclei, which may result from disrupted microtubular organization and impaired nuclear anchoring mechanisms.^54,55 Similarly, a previous study by Gushchina et al. reported that centralized nuclei persisted despite PPMO therapy. Specifically, they reported that a single i.v. injection of PPMO (31-mer PPMO targeting exon 2, PPMO-A) in the Dup2 mouse model of DMD (mice with exon 2 duplication representing single-exon duplication in patients with DMD) induced robust exon 2 skipping and dystrophin restoration in skeletal muscles, including the diaphragm and TA. However, they found that Dup2 mice receiving 80 mg/kg of the 31-mer PPMO did show a significant but modest decrease in myofibers with centralized nuclei at 60 days post-treatment in the TA (56.2%) and diaphragm (∼40%) compared to saline-treated TA (∼66%) and diaphragm (∼45%).⁵⁶ These findings indicate that PPMO treatment does not fully reverse nuclear centralization, suggesting that persistent deficits in myofiber maturation may remain. Further investigation is warranted to elucidate why centralized nuclei persist despite dystrophin expression and decreased inflammation. Strategies to address these limitations could include optimizing dosing or repeated administration to enhance structural remodeling. Additional approaches, such as satellite cell activation or gene-editing technologies (example: CRISPR-Cas9), may provide more durable correction of dystrophin deficiency and support proper nuclear positioning.

Quantitative analysis of myofiber cross-sectional area revealed substantial structural remodeling in the skeletal muscles of mdx mice. Consistent with previous studies, we observed a significant increase in the proportion of small-diameter fibers and a reduction in large-diameter fibers, reflecting ongoing cycles of muscle degeneration and regeneration.^57,58 The mdx muscles also have a decreased proportion of type IIx fibers,⁵⁸ which are large, fast-twitch, high-force-generating fibers, and this may explain the reduced muscle strength and functional impairments associated with the dystrophic phenotype. We observed that treatment with RC-1001 significantly restored myofiber size in the diaphragm and TA muscles. Additional functional assays are needed to assess the potential physiological impact of this treatment.

Consistent with prior findings, RC-1001 showed lower efficacy in cardiac tissue than in skeletal muscle. Limited dystrophin restoration in the heart is a concern for elderly patients or those with cardiac comorbidities. Future studies combining exon-skipping approaches with AAV-based gene therapy may provide better cardiac dystrophin restoration and AAV functional cardiac benefit.

Impact of exon-skipping therapy on respiratory function in DMD

In DMD, dystrophin deficiency leads to progressive weakening of the respiratory muscles, including the diaphragm, intercostals, and accessory muscles of respiration. This respiratory muscle weakness leads to restrictive lung disease characterized by reduced lung volumes and impaired ventilation. Respiratory compromise is further exacerbated by scoliosis and thoracic deformities.^8,59,60 In addition, dystrophic oropharyngeal and tongue muscles impair swallowing, decrease airway protection, and increase the risk of aspiration pneumonia.^39,40 The risk of pneumonia is further increased by ineffective cough and poor airway clearance.^9,61 In the absence of therapeutic intervention, respiratory dysfunction in males with DMD culminates in respiratory failure, leading to death during the second to third decade of life.^11,62

Respiratory dysfunction in DMD has been extensively characterized in a range of preclinical models. Among these, the mdx mouse remains the most widely used model and has been well studied for respiratory deficits. Several studies have documented reductions in resting ventilation at a younger age (2–3 months),^58,63,64,65 while others have reported significant differences in ventilatory parameters between mdx and WT mice at older ages (10–16 months).^66,67,68,69 In line with the latter group of studies, our current findings did not reveal significant differences in respiratory parameters while breathing room air (FiO₂: 0.21) between mdx and WT mice. Despite some discrepancies in the literature, there is growing consensus that respiratory function in mdx mice progressively declines with age, coinciding with the progression of pathology in respiratory muscles, including fibrosis, fiber atrophy, and neuromuscular junction (NMJ) disruption.⁷⁰ In younger mdx mice, compensatory mechanisms may help maintain relatively normal respiratory function at baseline, thereby masking subclinical deficits under resting conditions.⁷¹ Respiratory impairments become more apparent when the mice are exposed to respiratory stressors. In our study, exposure to combined hypoxic and hypercapnic conditions (FiO₂: 0.10; FiCO₂: 0.07) revealed significant impairments in mdx mice across multiple ventilatory parameters, including F, TV, MV, PIF, and PEF. The inability to mount an appropriate ventilatory response to hypoxia and hypercapnia is consistent with our previous data in mdx mice (6 and 12 months of age).³⁵ In addition, a similar pattern was observed in mdx;utrn^−/− double knockout mice, which exhibit a more severe phenotype due to the absence of both dystrophin and utrophin. While these mice displayed subtle differences from WT controls during eupneic breathing, their respiratory deficits under challenge were striking,⁷² indicating that physiological reserve and the capacity to mount an appropriate ventilatory response during stress are significantly compromised in dystrophic animals. In this study, RC-1001 improved respiratory parameters in awake, spontaneously breathing mice during a hypoxic and hypercapnic challenge. The lowest dose of RC-1001 (30 mg/kg) showed a better respiratory response under maximal chemostimulation by hypoxia and hypercapnia compared to higher doses. This indicates that a low dose may achieve sufficient dystrophin restoration while avoiding potential dose-related adverse effects.

In addition, this study demonstrates that RC-1001 improves respiratory mechanics in mdx mice. Untreated mdx mice exhibit significantly increased respiratory system resistance and elastance, along with reduced compliance—findings consistent with restrictive lung disease. Notably, all tested doses of RC-1001 therapy significantly improved pulmonary mechanics and ameliorated the restrictive respiratory pattern in 52-week-old mdx mice, suggesting that pulmonary function is achieved once a therapeutic threshold of dystrophin expression is achieved, beyond which additional drug offers no added benefit. Moreover, the forced oscillation technique may lack sensitivity to detect subtle dose effects. These findings highlight the potential of low-dose exon-skipping therapy as an effective strategy for restoring pulmonary function.

RC-1001 specifically induces exon 23 skipping in the mdx mouse model, and this study demonstrated the therapeutic potential of PPMO chemistry on respiratory outcomes. Translation to patients would require designing new PPMOs tailored to each clinically relevant exon mutation for direct clinical application. Vesleteplirsen (SRP-5051) is the first PPMO targeting exon 51 to enter clinical trials and generated foundational data on the efficiency, safety, and pharmacokinetics of the treatment. Although this program was discontinued due to issues with long-term safety and tolerability, the increased dystrophin expression observed supports the development of next-generation PPMOs with improved safety profiles.

Conclusions

This study demonstrates that PPMO-mediated exon skipping improves respiratory muscle histopathology, increases dystrophin expression, and reduces inflammation. While dystrophin restoration and exon-skipping efficiency are dose-dependent, inflammation is reduced across all dose levels, and centralized nuclei persist despite treatment. Further, PPMO-treated mdx mice show significant improvements in respiratory functions, including awake spontaneous breathing and pulmonary mechanics. In conclusion, PPMO therapy enables a decreased dosing frequency compared to PMO and effectively ameliorates respiratory muscle histopathology while enhancing respiratory function in the mdx mouse model of DMD.

Materials and methods

Mice

The mdx (C57BL/10ScSn-Dmd^mdx/J) and age-matched WT (C57BL/10ScSnJ) male mice were used in this study. These mice were purchased from the Jackson Laboratory (Strain # 001801 and 000476) at 6 weeks of age, and all experiments began at 8 weeks of age. Animals were bred and housed at the Duke University Division of Laboratory Animal Resources on a 12-h light/dark cycle with ad libitum access to food and water. Mice were euthanized at 52 weeks of age. All animal studies were conducted under protocols approved by the Duke University Institutional Animal Care and Use Committee (IACUC).

PPMO treatment

RC-1001 was manufactured by Sarepta Therapeutics, Inc. (Cambridge, MA, USA).³⁰ PPMO was injected systemically via the tail vein of 8-week-old mdx mice at final doses of 30, 50, and 100 mg/kg monthly (every 4 weeks) until the mice reached 48 weeks of age and were euthanized at 52 weeks of age. WT and mdx control groups were injected with saline. Each mouse was injected with either saline or PPMO in a total volume of 200 μL.

Droplet digital PCR

Tissue samples were homogenized in FastPrep Matrix S tubes (MP Biomedicals, Cat#: 1169250-CF) using a Spex SamplePrep GenoGrinder tissue homogenizer for 45 s at a speed of 1,600, followed by 15 s of rest, for 3 cycles. Samples were then centrifuged at 12,000 RCF for 5 min at room temperature. For more fibrous tissues, samples were subjected to an additional 2 cycles in the GenoGrinder and centrifuged again under the above-specified settings. Total RNA was isolated from homogenized tissues using a Zymo Quick-RNA MiniPrep Kit (Cat#: R1055). RNA concentrations were measured using a Lunatic UV/Vis spectrophotometer. RNA was converted into cDNA using a High-Capacity cDNA Reverse Transcription Kit with RNase inhibitor (Thermo Fisher Scientific, Cat#: 4374966).

Assay primers and probes for measuring exon 23 skipping were ordered from IDT using the sequences described previously,⁷³ except for the skip assay probe. Probes were designed with Zen/Iowa Black FQ quenchers and FAM or HEX dyes. Separate skip and nonskip reactions were set up using 900 nM each of forward and reverse primers, 250 nM probe, cDNA reverse transcribed from 15 ng of RNA, ddPCR Supermix for probes (no dUTP) (Bio-Rad, Cat#: 186–3024), and nuclease-free water. Droplets were generated using the Bio-Rad Automated Droplet Generator, followed by thermocycling according to the guidelines given by the ddPCR Supermix, except for the annealing temperature, which was 61°C for the skip assay and 53°C for the nonskip assay. Droplets were read using the Bio-Rad QX200 Droplet Reader, and droplet reads were analyzed using QuantaSoft software.

Percent exon skipping was calculated as (skip copies/(skip + non-skip copies)) ∗ 100.

Primers and probes for the exon 23 skipping ddPCR assay

Skip assay

Forward primer: AGC AGT CAG AAA GCA AAC TCT CTG.

Reverse primer: TTC AGC CAT CCA TTT CTG TAA GGT.

Probe: AGA CTC GGG AAA TTA CAG AAT CAC.

Non-skip assay

Forward primer: ACT GAA TAT GAA ATA ATG GAG AGA CT.

Reverse primer: GCC ATT TTG CTC TTT CAA A.

Probe: AAA TTA CAG GCT CTG CAA AG.

Automated western blot method

Dystrophin quantification in tissues was measured by ProteinSimple Jess. Tissues were cut to weigh approximately 20 mg and homogenized in 300 μL of homogenization buffer in Fast Prep Matrix S tubes. The homogenization buffer was prepared as follows: 5 mL of 8 M urea (Thermo Fisher Scientific, Cat#: NC9554009), 2.5 mL of 0.5 M Tris (Thermo Fisher Scientific, Cat#: AAJ63735AP), 2 mL of SDS (Thermo Fisher Scientific, Cat#: BP1311-200), 0.5 mL of DI water, and one protease inhibitor tablet (Thermo Fisher Scientific, Cat#: PIA32953). Samples were homogenized using the Spex SamplePrep GenoGrinder with pre-chilled blocks, using the same homogenization settings as described in the droplet digital PCR method, except that centrifugation of the Jess samples was done at 4°C. The supernatant was then transferred to Eppendorf tubes and placed at −80°C for an hour before thawing on ice. Samples were centrifuged at 12,000 RCF for 10 min at 4°C. The samples were then transferred to Eppendorf tubes, and protein concentrations were measured using the Pierce BCA assay (Thermo Fisher Scientific, Cat#: 23225) per the manufacturer’s instructions for the microplate method. Samples were then diluted to 0.2 μg/μL protein concentrations in 0.1× sample buffer provided with the Jess kit (Protein Simple, Cat#: SM-W008). A dystrophin standard curve was prepared using pooled saline-treated C57BL/10ScSn WT and mdx muscle lysates at a protein concentration of 0.2 μg/μL.

Samples were diluted in the fluorescent 5× master mix (Protein Simple, Cat#: PS-FL03-8) before incubating at 95°C for 5 min with gentle shaking. The plate was then centrifuged at 2,500 RCF for 5 min and placed on ice. The rest of the plate was prepared following the manufacturer’s instructions for the Total Protein module (Protein Simple, Cat#: DM-TP01) with Replex (Protein Simple, Cat#: RP-001) and milk-free antibody diluent (Protein Simple, Cat#: 043–524). The primary antibody for dystrophin (Abcam, Cat#: ab154168) was prepared at a 1:1000 dilution in the milk-free antibody diluent. Samples were analyzed on a 66–440 kDa, 25-capillary cartridge using the Total Protein and Replex assays with the chemiluminescent Anti-Rabbit Detection Module (Protein Simple, Cat#: DM-001).

The dystrophin signal was normalized to the total protein signal in each capillary, and the percent WT value was interpolated from the dystrophin standard curve.

Tissue preparation for immunohistology

Muscles were harvested upon euthanization of the mice and embedded in optimum cutting temperature (OCT) compound (Tissue-Tek, Cat#: 4583) for cryoprotection. The tissues were then frozen in liquid nitrogen-chilled 2-methylbutane and stored at −80°C. Serial transverse muscle sections (10 μm) were cut using a cryostat at −20°C and mounted onto polylysine-coated glass slides (VWR, Cat#: 48311-703). These slides with tissues were stored at −80°C until staining was performed.

Histological analysis

Histological analysis included H&E staining to examine inflammatory cell infiltration and centralized nuclei in muscle, Picrosirus red staining to analyze collagen deposition, and laminin staining to quantify the minimal ferret diameter of muscle fibers. The protocols for each staining and data analysis are described below.

H&E

To examine putative inflammatory cell infiltration of muscle and centralized nuclei, tissue sections were stained with H&E. Slides containing tissues were rinsed in distilled water, followed by staining with hematoxylin (Sigma Aldrich, Cat#: 65067-M) for 5 min. Sections were then treated with differentiating solution (Epredia, Cat#: 74211), blued in Scott’s tap water substitute (Electron Microscopy Sciences, Cat #: 26070-07), and counterstained with Eosin Y (Sigma-Aldrich, Cat#: HT110116) for 1–2 min. Slides were dehydrated in ethanol, cleared in xylene, and mounted with Epredia Cytoseal Mountant 60 (Cat#: 23–244257) before imaging.

Picrosirus red staining

Picrosirus red staining was performed to analyze the percentage of collagen deposition. The Picrosirus red solution was prepared by dissolving 0.1 g of Direct Red (Sigma-Aldrich, Cat#: 365548) in 100 mL of 1.3% picric acid (Sigma-Aldrich, Cat#: P6744). Slides were rehydrated through graded ethanol, stained with 0.1% Sirius Red solution in saturated picric acid for 60 min, and then washed briefly in 0.5% acetic acid. Slides were dehydrated in ethanol, cleared in xylene, and mounted with Epredia Cytoseal Mountant 60 (Thermo Fisher Scientific, Cat#: 22-050-262) before imaging.

Laminin staining

Sections were blocked with 1% BSA (VWR, Cat#: 0332) for 20 min, followed by blocking with 5% goat serum for an hour, then incubated overnight at 4°C with a primary anti-laminin antibody at a 1:100 dilution (Sigma-Aldrich, Cat#: L9393). After washing, slides were incubated with a secondary antibody, goat anti-rabbit IgG-fluorescein isothiocyanate (FITC) antibody (Millipore Sigma, Cat#: F9887) at a 1:100 dilution in 1% BSA for 1 h in the dark and mounted with Vectashield (Vector Laboratories, Cat#: H1200).

Dystrophin staining of muscles

Sections were washed in PBS, blocked with M.O.M. (mouse-on-mouse) blocking reagent (Cat#: MKB-2213-1) for 2 h, and incubated overnight at 4°C with an anti-dystrophin antibody (Abcam, Cat#: ab15277) at a 1:250 dilution in 1% BSA. After washing with PBS, sections were incubated with an Alexa Fluor 488 goat anti-rabbit secondary antibody at a 1:500 dilution (Cat#: A11008, Thermo Fisher Scientific) in 1% BSA for 1 h in the dark, washed, and mounted with DAPI Fluoromount-G (Cat#: 100-20, Southern Biotech).

Data analysis

Images of the tissues were acquired using an ECHO Revolve Microscope (Echo, San Diego, CA) at 20× magnification in brightfield. A total of five tissue sections per animal were examined and scored using ImageJ software. Images were acquired from multiple regions of interest per tissue per animal. Quantification was performed using ImageJ by two observers blinded to the genotype and treatments administered to the mice. Putative inflammatory cell infiltration was quantified and expressed as a percentage of the total muscle area. For Picrosirus red-stained tissues, images were analyzed using a color threshold, and the collagen area was expressed as a percentage of the total muscle area. Muscle fiber minimal Feret diameter—the shortest distance between two parallel lines tangent to the fiber boundary—was measured using ImageJ software. Multiple fibers per section were analyzed, and average Feret diameters were computed per sample.

WBP

WBP was performed as described previously.^74,75 Unanesthetized and unrestrained mice were placed in a Plexiglas chamber (DSI, St. Paul, MN), and data were collected and analyzed using FinePointe Software. Ventilation was monitored during eupneic breathing (normoxia: FiO₂: 0.21; N₂ balance) and during a respiratory challenge with hypercapnia and hypoxia (FiCO₂: 0.07, FiO₂: 0.10; N₂ balance).

Baseline: Mice were acclimatized to the chamber for the first 30 min in room air and later analyzed for approximately 1.5 h. Within this period, 5 min of regular breathing were selected as the “baseline.”

Challenge: Following the period of normoxia, mice were exposed to a respiratory challenge with hypercapnia and hypoxia for 10 min. The final 5 min of the challenge period were selected for analysis.

Mice were then returned to normoxia. WBP was performed monthly as the mice aged to detect the progression of respiratory dysfunction.

Forced oscillometry (FlexiVent system)

Airway mechanics were measured using forced oscillometry (FlexiVent, Scireq), as previously described.⁷⁶ In brief, mice were anesthetized with sodium pentobarbital (65 mg·kg⁻¹, administered via intraperitoneal injection). A tracheal cannula was inserted and secured with sutures (Lundbeck, Inc., Deerfield, IL, USA). Mice were paralyzed with pancuronium bromide (0.25 mg·kg⁻¹) and ventilated with 100% oxygen (Sigma, St. Louis, MO, USA) at a constant volume of 7 mL·kg⁻¹ and a frequency of 180 breaths·min⁻¹. Measurements were obtained by analyzing pressure and volume signals acquired in reaction to predefined, small-amplitude oscillatory airflow waveforms (perturbations) applied to the airways. Rrs, Ers, and Crs were obtained by assessing the mouse’s response to a single-frequency forced oscillation maneuver. Resistance of the central airways (Rn) and small airways, G, and H are parameters of the constant phase model (CPM) measured using broadband low-frequency forced oscillation maneuvers over a range of frequencies (Quick Prime-3).

Muscle strength analysis

Neurobehavioral assessments were performed as described previously.^72,77

Wire hang test: Mice were placed on an elevated wire grid. The grid was inverted and suspended above the cage for 120 s. The latency to fall was recorded. This test was performed with two trials per session, and the average of the two trials is reported.

Forelimb grip test: Forelimb grip strength was measured to assess maximal peak force (g), as previously described.⁷⁸ The Alemno digital grip strength meter (Ahlborn, Holzkirchen, Germany) equipped with a mesh screen was used for this test. Each mouse was placed on the metal grid and allowed to grip the mesh. The mouse was then gently pulled by the tail until it released its grip. The average of three consecutive measurements was calculated and normalized to the weight of each mouse.

Statistics

The effects of RC-1001 treatment on exon skipping, dystrophin protein, FlexiVent measurements, and quantification of IHC images for area of infiltration, centralized nuclei, fibrosis, and minimal Ferret diameter were analyzed using one-way analysis of variance (ANOVA). Fisher’s least significant difference (LSD) was used for post hoc analysis. Two-way ANOVA with Fisher’s LSD post hoc analysis were used to analyze WBP at baseline and challenge, as well as behavioral assessments at multiple time points. All data and statistical analyses were performed using GraphPad Prism 10.1.

Data and code availability

The datasets generated in the current study are available from the corresponding author upon reasonable request.

Acknowledgments

We appreciate the assistance of Victoria McQuade and the Rodent Inhalation Core Facility for FlexiVent measurements, as well as Alec Wright (Sarepta Therapeutics) for assistance with ddPCR assay design. This study was funded by Sarepta Therapeutics, Inc. and NIH RO1 HL171282-01 (MKE). We would also like to acknowledge BioRender for assistance with the creation of the graphical abstract.

Author contributions

D.D.B., M.Y.H.R., G.H., E.C., S.M., J.J., and S.L. were involved in methodology, data acquisition, and analysis. L.E.H., M.A., and A.S. were involved in methodology. O.C., A.D.P., and E.L. were involved in data analysis. D.D.B. and M.K.E. drafted the original manuscript, and M.K.E. supervised the project. All authors reviewed and edited the manuscript.

Declaration of interests

This study was sponsored by Sarepta Therapeutics, Inc. G.H. and E.C. were employees of Sarepta Therapeutics, Inc., at the time of this research.