# Lysosomal damage is a therapeutic target in Duchenne muscular dystrophy

Abbass Jaber, Laura Palmieri, Rania Bakour, Nathalie Bourg, Ai Vu Hong, Elise Lachiver, Carinne Roudaut, Jérôme Poupiot, Sonia Albini, Daniel Stockholm, et al.

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## ► To cite this version:

Abbass Jaber, Laura Palmieri, Rania Bakour, Nathalie Bourg, Ai Vu Hong, et al., Lysosomal damage is a therapeutic target in Duchenne muscular dystrophy. Science Advances, 2025, 11 (43), pp.eadv6805. 10.1126/sciadv.adv6805. hal-05386319

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DISEASES AND DISORDERS

# Lysosomal damage is a therapeutic target in Duchenne muscular dystrophy

Abbass Jaber<fnref n="1" />,², Laura Palmieri<fnref n="1" />,², Rania Bakour<fnref n="1" />,², Nathalie Bourg<fnref n="1" />,², Ai Vu Hong<fnref n="1" />,², Elise Lachiver<fnref n="1" />,², Carinne Roudaut<fnref n="1" />,², Jérôme Poupiot<fnref n="1" />,², Sonia Albini<fnref n="1" />,², Daniel Stockholm<fnref n="1" />,<fnref n="3" />, Laetitia Van Wittenberghe<fnref n="1" />, Adeline Miranda<fnref n="1" />, Guillaume Tanniou<fnref n="1" />, Nathalie Danièle<fnref n="1" />, Inès Barthélémy⁴, Stephane Blot⁴, Mai Thao Bui<fnref n="5" />, Bornale Das⁶, Edoardo Malfatti⁶,⁷, Teresinha Evangelista<fnref n="5" />, Isabelle Richard<fnref n="1" />,²,⁸, David Israeli<fnref n="1" />,²*

Duchenne muscular dystrophy (DMD), a muscle degenerative disease affecting young boys, arises from the loss of dystrophin. Current gene therapy approaches aim to restore a shortened form of dystrophin (microdystrophin) via adeno-associated vector delivery. While recent clinical studies show promise, therapeutic efficacy remains incomplete, emphasizing the need for improved approaches. Here, we identified lysosomal perturbations in myofibers of patients with DMD and animal models, an overlooked mechanism of cellular damage in muscular dystrophies. These were notably marked by the up-regulation and recruitment of Galectin-3, a biomarker of lysosomal membrane permeabilization, to lysosomes, alongside alterations in lysosome number, morphology, and function. Microdystrophin therapy in *Dmd*^mdx mice fails to fully correct these damages. However, combining it with trehalose, a lysosome-protective disaccharide, substantially improves the outcome, enhancing muscle function, myopathology, and transcriptome. These findings highlight lysosomal damage as an important pathomechanism in DMD and suggest that combining trehalose with gene therapy could enhance therapeutic efficacy.

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INTRODUCTION

Duchenne muscular dystrophy (DMD) is an X-linked recessive muscle disease as a result of the absence of dystrophin, the protein encoded by the *DMD* gene. It affects roughly 1 of 5000 boys and is the most common form of childhood muscular dystrophy. The loss of dystrophin in the skeletal myofiber leads to the destabilization of the dystrophin-associated glycoprotein complex (DAGC), which connects the cytoskeleton to the extracellular matrix (1). This disruption triggers a cascade of pathological events including perturbations of calcium homeostasis, oxidative stress, mitochondrial dysfunction, chronic inflammation, fibroadipocytic transformation, and consequently, the loss of muscle mechanical function (2). Current clinical management of DMD involves a multidisciplinary approach to address both muscular and extramuscular manifestations of the disease (3). Approved treatments include mainly glucocorticoids, which are recommended as early as possible (4) and throughout the patient's life (5). Although glucocorticoids have shown a significant improvement in motor function, they remain merely a symptomatic treatment delaying disease progression (6). Besides, several therapeutic approaches under investigation aim at restoring a functional form of dystrophin. One of the most advanced and promising approaches is microdystrophin [µDystrophin (µDys)] gene therapy, which involves delivering a shortened but functional variant of the dystrophin gene

using a recombinant adeno-associated vector (rAAV) (7). A major
challenge in this therapeutic approach is optimizing the dosage.
High doses may trigger severe immune responses and organs' toxic-
ity (8), while low doses may fail to achieve the desired therapeutic
effect. Although one µDys gene therapy (delandistrogene moxeparv-
ovec) obtained Food and Drug Administration (FDA) approval for
ambulatory patients, the functional improvements for patients have
fallen short of preclinical expectations (9, 10). This limited clinical
benefit could be attributed to several factors, including the lack of
certain functional domains found in full-length dystrophin (11).
Therefore, µDys gene transfer may not be able to reverse all the path-
ological mechanisms in the muscle. Identifying specific pathological
mechanisms that remain uncorrected by µDys gene therapy and ex-
ploring combinatorial approaches with complementary treatments
targeting these mechanisms are crucial.

It is increasingly appreciated that the lack of dystrophin results not only in the mechanical instability of the skeletal myofiber but also in a vast spectrum of metabolic perturbations (12). Among these, plasma dyslipidemia and perturbation of lipid metabolism were identified (13), whereas in an investigation of expression of circulating microRNA in a large DMD cohort, we have recently showed the perturbation of cholesterol metabolism in the dystrophic muscle in DMD (14, 15). Excessive cholesterol levels have frequently been associated with an impaired lysosomal function, particularly in neurodegenerative diseases (16, 17). We therefore hypothesized that cholesterol accumulation in the dystrophic muscle could be associated with the impaired lysosomal function. To assess the lysosomal function in DMD, we first evaluated the expression and localization of Gal-3 (or LGALS3), a known biomarker of lysosomal membrane permeabilization (LMP). We found that Gal-3 is significantly upregulated in the skeletal myofibers of patients with DMD and various animal models, localizing to the lysosomes. LMP was also detected in a limb-girdle muscular dystrophy type R5 (LGMDR5) mouse model and patient with LGMDR5, suggesting a common perturbation

<footnote n="1">¹Généthon, 91000 Evry, France. ²Université Paris-Saclay, Univ. Evry, INSERM, Généthon, INTEGRARE research unit UMR_S951, 91000 Evry-Courcouronnes, France.</footnote>

<footnote n="3">³Ecole Pratique des Hautes Etudes, PSL Research University, Sorbonne Université, INSERM, CRSA, 75012 Paris, France. ⁴Univ. Paris-Est Créteil, INSERM U955 IMRB, Ecole Nationale Vétérinaire d'Alfort, EFS, IMRB, Team BNMS, Maisons-Alfort, France.</footnote>

<footnote n="5">⁵Muscle Pathology Unit, Institut de Myologie and Neuropathology Department, Sorbonne Université, Pitié-Salpetrière Hospital, AP-HP, Paris, France. ⁶Mondor Institute of Biomedical Research, University of Paris Est Créteil, INSERM U955, Créteil, France. ⁷AP-HP, North-East Ile-de-France Neuromuscular Pathology Reference Center, Henri Mondor Hospital, Créteil, France. ⁸Atamyo Therapeutics, 1, bis rue de l'internationale, Evry, France.</footnote>

<footnote>*Corresponding author. Email: israeli@genethon.fr</footnote>
across different types of muscular dystrophy. Using $Dmd^{mdx-4Cv}$ mice, we detected and characterized various lysosomal perturbations in the dystrophic muscle, such as an enlargement and increased number of lysosomes and decreased acidification and proteolytic activity. In addition, we observed the activation of the endolysosomal damage response, which includes lysosomal replacement, repair and removal, and the perturbation of autophagy. LMP was exacerbated in dystrophic mice fed a cholesterol-rich diet and was not fully corrected by µDys gene therapy. In contrast, gene therapy showed better correction in the LGMDR5 mouse model when delivering the complete sequence of the missing γ-sarcoglycan coding sequence, suggesting that the corresponding lysosomal dysfunction is reversible in other muscular dystrophies, but not by the sole µDys in DMD. Subsequently, we selected trehalose (TR), an FDA-approved compound known to restore the lysosomal function, for use in combination with a suboptimal dose of AAV-µDys gene therapy. The combined treatment resulted in the improved correction of dystrophic parameters, including motor function, muscle histology, and transcriptome signature, compared to µDys suboptimal gene therapy alone.

## RESULTS

### Lysosome membrane permeabilization impairs lysosome function in muscle cells and can be detected with LGALS3 puncta assay

To investigate whether cholesterol accumulation in dystrophic muscle is associated with the impaired lysosomal function, we sought a method to quantify lysosomal damage. Several studies have recently focused on lysosomal dysfunction, in which LMP plays a central role (18). However, very few studies have investigated lysosome damage in muscle. One identified marker of LMP is Gal-3 (or LGALS3), which is part of the galectins, a family of lectins that bind specifically to carbohydrates (19). These lectins are present in the cytosol, nucleus, and extracellular matrix and are translocated to the membrane of damaged lysosomes before their removal by the autophagy machinery (20). To analyze LGALS3 expression in a myogenic environment and its capacity to detect LMP, we differentiated healthy human immortalized myoblasts into elongated myotubes for 7 days and then performed immunostaining for LGALS3 and lysosome-associated membrane protein 2 (LAMP2), which is commonly used as a lysosome marker (21). A diffuse expression of LGALS3 was observed in the cells (fig. S1A). Treatment with L-leucyl-L-leucine methyl ester (LLOMe), a lysomotropic agent that induces lysosome-specific membrane damage (22), for 30 min at 2.5 mM triggered the formation of LGALS3-positive puncta, which colocalized with LAMP2, indicating typical damaged lysosomes. An up-regulation of LGALS3 was also detected by Western blotting after 30 min to 1 hour of LLOMe treatment (fig. S1, B and C). LGALS3 puncta were markedly reduced 3 hours after treatment, and LGALS3 expression was restored to the control level, demonstrating efficient lysosomal repair by the cells. This pattern correlated inversely with the amount of LGALS3 released in the media, detected by an enzyme-linked immunosorbent assay (ELISA) assay (fig. S1D), where a significant increase was detected in the media only after 3 hours of LLOMe treatment. We then aimed to check the effect of LMP on the lysosomal function after LLOMe treatment. For that, we performed pH probing using a combination of two dextran conjugates [molecular weight (MW) = 10 kDa], which are endocytosed by the cells and accumulate selectively in the lysosomes (fig. S1E). One dextran is conjugated with Oregon Green, whose

emission is quenched under acidic environments such as the lysosomal lumen, and the other is conjugated with tetramethylrhodamine, whose emission in the red spectrum is pH independent (23). Monitoring the ratio between the green and red dextran-coupled fluorophores demonstrated a decrease in acidification after treatment with LLOMe for 30 min (higher green/red ratio) (fig. S1F), indicating an impact of LMP on the lysosomal function in myotubes. Collectively, these data indicate that LGALS3 is expressed in muscle cells and is recruited specifically to damaged lysosomes. Immunostaining for LGALS3 could be therefore applied to detect LMP in skeletal muscle and to provide insight into the lysosomal function.

### LGALS3 is up-regulated in dystrophic muscle and correlates with LMP

After validating LGALS3 as a biomarker for LMP in muscle cells, we aimed at studying its expression and localization in DMD. First, we evaluated Gal-3 expression at the mRNA and protein levels in young 7-week-old $Dmd^{mdx-4Cv}$ mice. A significant up-regulation was observed in both mRNA transcripts and protein in the gastrocnemius (GA) [mRNA: fold change (FC) = 32; protein: FC = 14.4], tibialis anterior (TA) (mRNA: FC = 21; protein: FC = 22.6), and diaphragm (mRNA: FC = 10; protein: FC = 41.2) muscles of dystrophic mice compared to wild-type (WT) mice (Fig. 1, A to C). In addition, a significant increase in LGALS3 was also detected in the serum of $Dmd^{mdx-4Cv}$ mice compared to control mice (FC = 1.8) (Fig. 1D), suggesting a release of LGALS3 from the muscle. To evaluate whether LGALS3 is recruited to the damaged lysosomes in the context of LMP in the dystrophic muscle, cross sections of TA muscles were immunostained for LGALS3 and LAMP2 (Fig. 1E). Confocal immunofluorescence images showed a colocalization of LGALS3 puncta with LAMP2 inside the myofibers of young $Dmd^{mdx-4Cv}$ mice, indicating a substantial amount of LMP. Notably, LGALS3- and LAMP2-positive areas were also observed outside the myofibers in regions consistent with inflammatory cellular infiltrates characteristic of dystrophic muscle. This aligns with prior reports attributing Gal-3 up-regulation in DMD animal models primarily to LGALS3-positive infiltrating immune cells such as macrophages (24–26). To confirm the inflammatory origin of these regions, coimmunostaining for LGALS3 and CD11b was performed on serial TA muscle cross sections (Fig. 1F). An overlay of both staining (CD11b and LGALS3) was observed between the myofibers, confirming the presence of infiltrating macrophages, while LGALS3 staining was also evident within myofibers. This indicates that the up-regulation of Gal-3 in muscle lysates reflects contributions from both immune infiltrates and myogenic sources.

To specifically quantify LMP in myofibers, we measured the number of LAMP2+LGALS3+ puncta within myofibers, excluding inflammatory areas using a segmentation method (fig. S2A). This analysis revealed a significant increase in LAMP2+LGALS3+ puncta in dystrophic muscle compared to WT controls (mean puncta = 9 for $Dmd^{mdx-4Cv}$ versus 1 in WT) (Fig. 1G), confirming that LGALS3 up-regulation in $Dmd^{mdx-4Cv}$ muscles is not solely due to macrophage infiltration but also reflects myogenic up-regulation and recruitment to the lysosome, suggesting substantial LMP in dystrophic myofibers.

To validate these findings in human and large animal models, we analyzed muscle biopsies from patients with DMD and golden retriever muscular dystrophy (GRMD) dogs. LGALS3 up-regulation within myofibers was consistently observed in biopsies of patients with DMD from children aged 2 and nearly 4 years (Fig. 1H and
<figure><img src="image_3.png" /><figcaption>**Fig. 1.** LGALS3 up-regulation in dystrophic muscle correlates with LMP. (A) Western blot analysis of TA, GA, and diaphragm muscles and serum samples of 7-week-old WT and *Dmd<sup>mdx-4Cv</sup>* mice (n = 4 to 6) for LGALS3. Total protein staining was used as the loading control. (B) Relative *Lgals3* mRNA expression in the TA, GA, and diaphragm, normalized to *Rplp0* (FC shown, average ± SD, n = 6). (C) Band intensity quantification of LGALS3 relative to total protein staining in TA, GA, and diaphragm muscles (FC shown, average ± SD, n = 4 to 6). (D) Band intensity quantification of LGALS3 relative to total protein staining in serum (FC shown, average ± SD, n = 6). (E) Representative confocal images of TA muscle cross sections immunostained for LAMP2 (green) and LGALS3 (red). Scale bars, 100 or 25 µm (zoomed-in images). (F) Confocal images of serial transversal sections of TA muscles from a *Dmd<sup>mdx-4Cv</sup>* mouse immunostained for LAMP2/LGALS3 and LAMININ/CD11b. Zoomed-in images show the inflammatory area of dystrophic muscle. Scale bars, 200 or 50 µm (zoomed-in images). (G) Quantification of double-positive puncta (LAMP2+LGALS3+) in the myofibers of TA muscles of WT and *Dmd<sup>mdx-4Cv</sup>* mice (average ± SD, >400 myofibers analyzed from n = 6). (H) Representative confocal images of transversal sections from muscles of a patient with DMD and a healthy control (2-year-old) and a GRMD dog and a WT control (6-month-old), immunostained for LAMP2 and LGALS3. Left panels show H&E labeling from the same muscle biopsies. Scale bars, 25 or 10 µm (zoomed-in images) and 50 µm (H&E images). An unpaired two-tailed *t* test was used for statistical comparisons of WT and *Dmd<sup>mdx-4Cv</sup>* groups. *t* test: \*P < 0.05, \*\*P < 0.01, and \*\*\*P < 0.0001.</figcaption></figure>
fig. S2B) and in 6-month-old GRMD dogs (Fig. 1H). In all cases, LGALS3 colocalized with LAMP2 in most myofibers, further supporting the presence of LMP and myofiber-intrinsic Gal-3 up-regulation across species.

Alteration of the lysosome structure and function in the dystrophic muscle

After confirmation of LMP incidence in the dystrophic muscle, we sought to evaluate parameters of lysosomal stress. First, we evaluated the lysosome number with a Western blot for LAMP2 in WT and *Dmd<sup>mdx-4Cv</sup>* muscles (Fig. 2A and fig. S3A). An up-regulation of LAMP2 was observed in both GA (FC<sub>GA</sub> = 3.9) (Fig. 2B) and TA (FC<sub>TA</sub> = 4.1) (fig. S3A). Moreover, a quantification of LAMP2 spots on transversal sections from TA muscles immunolabeled with LAMP2 showed a significant increase in spot number (mean = 14.3 and 12.2 for *Dmd<sup>mdx-4Cv</sup>* and WT, respectively) (fig. S3, B and C) and spot size (FC = 1.7) (fig. S3D), two main criteria of lysosomal stress. To further analyze the lysosome morphology and number at the single myofiber level, flexor digitorum brevis (FDB) muscle fibers were isolated from *Dmd<sup>mdx-4Cv</sup>* and WT mice and immunostained for LAMP2 and LGALS3. Three-dimensional (3D) reconstruction of confocal stack images showed a significant accumulation of lysosomes in the *Dmd<sup>mdx-4Cv</sup>* myofibers (Fig. 2C and movie S1), which colocalized with LGALS3, especially around the nuclei, indicating a substantial amount of LMP. However, not all lysosomes (LAMP2+ spots) were positive for LGALS3, indicating the existence of a population of nondamaged lysosomes. 3D modeling of LAMP2 spots (Fig. 2D and movie S1) showed a significant increase in lysosome number (FC<sub>number</sub> = 3.4) (Fig. 2E) and a denser distribution in the *Dmd<sup>mdx-4Cv</sup>* myofibers, as indicated by the distance to the five nearest spots (FC<sub>5 nearest spots</sub> = -1.65) (Fig. 2F). An increase in the average spot diameter (FC<sub>Diameter</sub> = 1.2) was observed in the *Dmd<sup>mdx-4Cv</sup>* myofibers (Fig. 2G). When plotting the spot diameter on a density plot, three populations of LAMP2 spots were distinguished in the *Dmd<sup>mdx-4Cv</sup>* myofibers versus one main population in the WT myofibers (Fig. 2H). Both WT and *Dmd<sup>mdx-4Cv</sup>* myofibers displayed a population of average size spots. *Dmd<sup>mdx-4Cv</sup>* myofibers showed additionally a population of enlarged spots, corresponding to typical damaged enlarged lysosomes, indicative of lysosomal stress and accumulation of undegraded materials. An additional population of smaller spots was detected in *Dmd<sup>mdx-4Cv</sup>* myofibers. These smaller spots may correspond to newly generated lysosomes and/or to protolysosomes, which are fragments of recycled lysosomal membranes following the elimination of damaged lysosomes (27).

To investigate the lysosomal ultrastructure in greater detail, we performed transmission electron microscopy on TA muscles from 7-week-old WT and *Dmd<sup>mdx-4Cv</sup>* mice. Although the sarcomere architecture appeared largely preserved in dystrophic muscle at this stage, we observed a higher number of enlarged and electron-dense lysosomes in *Dmd<sup>mdx-4Cv</sup>* muscles compared to WT (highlighted by red circles) (Fig. 2I). Quantitative analysis confirmed a significant increase in lysosome size, with an average diameter of 383 nm in dystrophic muscle versus 220 nm in WT (Fig. 2J and fig. S3E). In WT samples, lysosomes (blue circles) were smaller, maintained a round morphology, and exhibited a mix of electron-pale and electron-dense appearances (fig. S3E). In contrast, lysosomes in *Dmd<sup>mdx-4Cv</sup>* muscle were more osmiophilic, consistent with the accumulation of undegraded material. Morphologically, dystrophic lysosomes were heterogeneous, ranging from moderately enlarged vesicles to abnormally

large structures (>500 nm), with some resembling lipofuscin granules, indicative of impaired lysosomal clearance (fig. S3E).

To determine whether these structural abnormalities were associated with the impaired lysosomal function, we isolated primary myoblasts from WT and *Dmd<sup>mdx-4Cv</sup>* limb muscles and assessed lysosomal proteolytic activity using polymeric bovine serum albumin (BSA) labeled with red BODIPY (4,4-difluoro-4-bora-3a,4a-diaza-s-indacene), a self-quenched substrate that fluoresces upon proteolytic cleavage (Fig. 2K). This assay is based on the principle that the densely labeled Dye-Quenched (DQ) Red BSA exhibits quenched fluorescence until it is cleaved by lysosomal proteases, at which point fluorescent monomers accumulate and fluorescence increases (23). Myoblasts were incubated with DQ Red BSA, fixed, and imaged after 4 hours. *Dmd<sup>mdx-4Cv</sup>* myoblasts exhibited a significant reduction in lysosomal proteolytic activity, as reflected by the decreased mean fluorescence intensity (MFI) of DQ Red BSA (Fig. 2L), indicating a compromised lysosomal function in dystrophic muscle cells.

Given that glycogen can accumulate in dysfunctional lysosomes, we also performed periodic acid-Schiff (PAS) staining on TA muscles from WT and 7-week-old *Dmd<sup>mdx-4Cv</sup>* mice (fig. S3F). Quantification of muscle cross sections revealed a slight, although nonsignificant, increase in PAS staining in dystrophic muscle, suggesting the early stages of glycogen accumulation. This is consistent with a previous report describing glycogen buildup in older *Dmd<sup>mdx</sup>* mice (28).

Collectively, these data demonstrate that LMP occurs in the dystrophic muscle of *Dmd<sup>mdx-4Cv</sup>* mice, as evidenced by the increased presence of LAMP2+LGALS3+ puncta within myofibers. In addition, multiple indicators of lysosomal stress were observed, including lysosome enlargement, increased number and electron density, and accumulation of undegraded material, all which are hallmarks of impaired lysosomal clearance. These structural and morphological abnormalities are accompanied by a significant reduction in lysosomal proteolytic activity, further supporting the presence of lysosomal dysfunction in dystrophic muscle. Together, these findings suggest that both lysosomal membrane integrity and degradative capacity are compromised in *Dmd<sup>mdx-4Cv</sup>* muscle fibers.

Lysosome membrane permeabilization persists with age and is a common feature across different muscular dystrophies

To address whether LMP is only an early process in dystrophic pathology, given that only young mice were evaluated above, or a persistent feature with disease evolution, 1-year-old *Dmd<sup>mdx-4Cv</sup>* and control mice were also studied. An up-regulation of *Lgals3* transcripts (FC = 29 for TA and 6.5 for GA) (fig. S4A), an up-regulation of LGALS3 protein expression (FC = 52 for TA and 23 for GA) (fig. S4, B and C), and a high number of LAMP2+LGALS3+ puncta (fig. S3, D to G) were detected in both TA and GA muscles (mean = 10.7 and 9.1 in *Dmd<sup>mdx-4Cv</sup>* versus 3.9 and 3.1 in WT for TA and GA, respectively). Older WT mice displayed more LMP compared to young WT mice, indicating a possible degradation of lysosomal function over time.

To evaluate whether lysosomal degradation was a specific DMD feature or whether it might be a common feature in muscular dystrophies, transversal GA muscle sections from an LGMDR5 (or γ-sarcoglycanopathy) mouse model (*Sgcg<sup>-/-</sup>*) were immunostained for LAMP2 and LGALS3 (fig. S5A). A significant increase in LAMP2+LGALS3+ puncta was observed in *Sgcg<sup>-/-</sup>* muscle compared to the WT control (mean = 8.3 versus 1.9 for control) (fig. S4B),
<figure><img src="image_4.png" /><figcaption>**Fig. 2.** Lysosomal stress in dystrophic muscle. (A) Western blot analysis of GA muscle for LAMP2 (n = 6). (B) Band density quantification of LAMP2 relative to glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (average ± SD, n = 6). (C) Reconstructed 3D images of isolated myofibers from the FDB muscle of *Dmd*<sup>mdx-4Cv</sup> and WT mice, immunostained for LAMP2 and LGALS3. Scale bars, 10 or 20 µm. (D) 3D modeling of LAMP2 spots. Green spots show average sized lysosomes (diameter <1.5 µm), and purple spots show enlarged lysosomes (diameter >1.5 µm). Scale bars, 10 µm. (E) FC of the number of lysosomes per myofiber (average ± SD, n = 4). (F) Analysis of five nearest spots to each spot, represented as a density plot (with a nonlinear fit Gaussian curve). (G) Quantification of LAMP2 spot diameter (min to max points). (H) Density plot showing the distribution of LAMP2 spot diameter. (I) Ultrastructural transmission electron micrograph of *Dmd*<sup>mdx-4Cv</sup> and WT TA muscles. The left panel shows low-magnification images, and red circles indicate larger and denser lysosomes in *Dmd*<sup>mdx-4Cv</sup> muscles compared to WT muscles (lysosome indicated by a blue circle). The right panel shows a high-magnification image highlighting a large (390 nm) lysosome found in the *Dmd*<sup>mdx-4Cv</sup> muscle. Scale bars, 3 µm (upper left panel), 1 µm (lower left panel), and 300 nm (zoomed-in image). (J) Quantification of lysosome diameter from electron micrographs (nm; min to max values shown) (n = 2). (K) Representative confocal images show the fluorescence of DQ-Red BSA after an uptake of 4 hours in primary myoblasts isolated from WT and *Dmd*<sup>mdx-4Cv</sup> muscles. Bright fluorescence puncta of DQ-Red BSA indicate delivery to acidic lysosomes of the cells. Scale bars, 50 and 10 µm (zoomed-in images). (L) Quantification of MFI per cell in each condition (n > 158 cells). An unpaired two-tailed t test was used for statistical comparisons of WT and *Dmd*<sup>mdx-4Cv</sup> groups. t test: \*P < 0.05, \*\*P < 0.001, and \*\*\*P < 0.0001.</figcaption></figure>
indicating a substantial amount of lysosomal damage. In addition, a muscle biopsy from a patient with LGMDR5 was analyzed. A positive pattern of LAMP2⁺LGALS3⁺ staining was detected inside the myofibers (fig. S5C), although to a lesser extent than DMD muscle biopsies, possibly correlating the level of lysosomal damage and the severity of the dystrophic phenotype. Together, these findings show that LMP, which can be detected with LGALS3 recruitment to the lysosomes, might be a common cellular perturbation in muscular dystrophies.

## Defects in lysosomal recovery and reacidification following membrane injury

We next investigated the ability of lysosomes to recover from membrane damage in dystrophic cells. For that, we performed a classical lysosomal injury assay on primary myoblasts isolated from WT and *Dmd<sup>mdx-4Cv</sup>* mice. Cells were treated with LLOMe to induce LMP and then either fixed immediately or allowed to recover in fresh media for 1 or 3 hours before analysis (Fig. 3A).

First, we performed immunostaining of LAMP2 and LGALS33 to check membrane integrity (Fig. 3B). At the baseline, *Dmd<sup>mdx-4Cv</sup>* myoblasts exhibited significantly more LGALS3⁺ puncta than WT cells (Fig. 3C), suggesting substantial LMP even in the absence of external stress. As expected, LLOMe treatment induced a marked increase in LGALS3 puncta in WT myoblasts (Fig. 3D). However, *Dmd<sup>mdx-4Cv</sup>* myoblasts showed no further detectable increase, likely due to already elevated basal levels (Fig. 3E). Following a 1-hour washout, LGALS3 puncta in WT cells returned to near-baseline levels and were further reduced after 3 hours, indicating efficient lysosomal recovery. In contrast, *Dmd<sup>mdx-4Cv</sup>* myoblasts showed a significant increase in LGALS3 puncta after both 1- and 3-hour washouts, indicating persistent membrane damage and impaired recovery.

We also analyzed lysosomal acidification using LysoTracker Red, which accumulates in lysosomes under normal acidic conditions (Fig. 3A). Disruption of lysosomal integrity and pH elevation lead to decreased LysoTracker fluorescence. Under basal conditions, *Dmd<sup>mdx-4Cv</sup>* myoblasts displayed significantly fewer LysoTracker-positive dots than WT cells (Fig. 3, F and G), consistent with reduced acidic lysosomes. LLOMe treatment further decreased LysoTracker dots in both genotypes, reflecting acute lysosomal dysfunction (Fig. 3F). In WT cells, LysoTracker dots partially recovered after 1 hour and were fully restored after 3 hours (Fig. 3H). However, in *Dmd<sup>mdx-4Cv</sup>* myoblasts, recovery was minimal, with only a slight increase in LysoTracker dots at 1- and 3-hour postwashout, and levels remained significantly lower than in WT (Fig. 3I). We further quantified the MFI of LysoTracker-positive dots as a proxy for lysosomal acidity. At the baseline, *Dmd<sup>mdx-4Cv</sup>* cells had significantly lower MFI compared to WT (P = 0.0048; Fig. 3J). In WT myoblasts, LLOMe treatment led to a shift toward lower MFI on density plots, which progressively normalized after 1 and 3 hours of recovery, with a complete overlap of the MFI distribution with control at 3 hours (Fig. 3K). In contrast, *Dmd<sup>mdx-4Cv</sup>* cells showed a similar drop in MFI after LLOMe exposure, but this defect persisted even after 3 hours, with no significant recovery (Fig. 3L), suggesting a defective reacidification of lysosomes.

Together, these results reveal an intrinsic defect in lysosomal recovery and acidification in *Dmd<sup>mdx-4Cv</sup>* myoblasts, independent of the complex inflammatory and fibrotic environment of dystrophic muscle. While WT cells can restore lysosomal integrity and acidity after acute injury, *Dmd<sup>mdx-4Cv</sup>* myoblasts failed to do so, further supporting the hypothesis of a primary cellular lysosomal defect.

## Activation of the endolysosomal damage response in *Dmd<sup>mdx-4Cv</sup>* muscles

Lysosomal stress, characterized by LMP and increased lysosome size, is a known trigger of the endolysosomal damage response. This response, coordinated by key mediators such as LGALS3, promotes lysosomal repair, removal, or replacement under physiological conditions (29). To investigate whether and to what extent this damage response is activated in dystrophic muscle, we analyzed markers of lysosomal stress and compensatory pathways.

We first examined the lysosomal biogenesis pathway, mediated by the lysosomal master transcription factor EB (TFEB). Immunostaining of muscle cross sections from 7-week-old *Dmd<sup>mdx-4Cv</sup>* and WT mice revealed a higher overall expression of TFEB in dystrophic muscle, with pronounced localization in nuclei but also in the cytoplasm and on the plasma membrane (Fig. 4A). Although nuclear TFEB accumulation was significantly higher in *Dmd<sup>mdx-4Cv</sup>* muscles compared to WT (Fig. 4B), only about 37% of nuclei were positive for TFEB, indicating partial TFEB activation with incomplete nuclear translocation. To assess downstream TFEB activity, we performed bulk RNA sequencing of GA muscles from 7-week-old WT and *Dmd<sup>mdx-4Cv</sup>* mice. Differential gene expression analysis revealed the up-regulation of several canonical TFEB targets in *Dmd<sup>mdx-4Cv</sup>* muscles, including genes encoding lysosomal enzymes (e.g., cathepsins), proton pumps, and lysosomal membrane glycoproteins (Fig. 4C), confirming the transcriptional activation of the lysosomal pathway. Among these, cathepsins B and D (CTSB and CTSD, respectively) were further validated by quantitative real-time fluorescence PCR (qPCR) and immunostaining in TA muscles, confirming their increased expression in *Dmd<sup>mdx-4Cv</sup>* muscle (fig. S6, A and B). Although cathepsin up-regulation in the dystrophic muscle had been previously reported (30–32), the mechanistic basis remained unclear. Our results support the hypothesis that this up-regulation is a consequence of lysosomal stress and TFEB pathway activation (29, 33). Notably, in WT muscle, CTSB and CTSD showed punctate localization that colocalized with LAMP2, consistent with their confinement within lysosomes. In contrast, *Dmd<sup>mdx-4Cv</sup>* muscle showed a more diffuse expression pattern, especially for CTSB. This shift suggests the leakage of cathepsins into the cytoplasm, a hallmark of LMP, which can contribute to tissue damage by promoting proteolysis, inflammation, and cell death (34).

To determine whether similar stress responses occur in a pure myogenic context, we analyzed TFEB expression in primary myoblasts. While TFEB was undetectable by immunofluorescence in WT myoblasts (Fig. 4D), *Dmd<sup>mdx-4Cv</sup>* myoblasts displayed a stronger TFEB signal, particularly in the nucleus. These results were confirmed by the Western blot analysis of nuclear and cytosolic fractions (Fig. 4E), showing increased TFEB levels in both compartments in *Dmd<sup>mdx-4Cv</sup>* myoblasts (Fig. 4F), consistent with observations in muscle tissue and indicating TFEB up-regulation and partial activation.

In parallel, we evaluated the endosomal sorting complex required for transport (ESCRT)-mediated lysosome repair pathway, which is responsible for the quick repair of acute lysosomal leakages (35). Upon damage, ESCRT subunits such as ALIX (ALG-2-interacting protein X or programmed cell death 6-interacting protein) and CHMP4B (charged multivesicular body protein 4B) are recruited to the lysosomal membrane in a $Ca^{2+}$-dependent manner (35, 36). Immunostaining for ALIX and LAMP2 revealed a significant accumulation of ALIX puncta colocalizing with LAMP2 in *Dmd<sup>mdx-4Cv</sup>* muscles compared to WT (mean = 13.9 versus 6.8 puncta, respectively)
<figure><img src="image_5.png" /><figcaption>**Fig. 3.** The lysosomal damage assay in primary myoblasts reveals early defects. (A) Schematic representation of the experimental setup. Cells were treated with LLOMe (2.5 mM) for 30 min to induce LMP and deacidification, resulting in decreased LysoTracker fluorescence. Washout steps indicate LLOMe removal and incubation with fresh media, enabling lysosomal repair and reacidification, reflected by the recovery of LysoTracker red fluorescence. (B) Representative confocal images of WT and *Dmd<sup>mdx-4Cv</sup>* primary myoblasts immunostained for LAMP2 and LGALS3. Scale bars, 30 or 10 µm. (C to E) Violin plots showing the number of LGALS3-positive puncta per cell (normalized to cell surface area). At least 65 cells per condition were quantified from two independent experiments. (C) Comparison between untreated WT and *Dmd<sup>mdx-4Cv</sup>* cells. LLOMe treatment and washout results are shown for (D) WT and (E) *Dmd<sup>mdx-4Cv</sup>* cells. (F) Representative confocal images of WT and *Dmd<sup>mdx-4Cv</sup>* primary myoblasts stained with LysoTracker Red DND-99 after 30 min of incubation. Scale bars, 10 µm. (G to I) Violin plots showing the number of LysoTracker-positive puncta per cell (normalized to cell surface area). (G) Comparison between untreated WT and *Dmd<sup>mdx-4Cv</sup>* cells. LLOMe treatment effects are shown in (H) WT and (I) *Dmd<sup>mdx-4Cv</sup>* cells. At least 70 cells per condition were analyzed. (J to L) MFI per LysoTracker dot, represented as density plots with nonlinear Gaussian curve fitting. (J) Baseline comparison between WT and *Dmd<sup>mdx-4Cv</sup>* cells (unpaired two-tailed *t* test, \*P = 0.0049). LLOMe treatment effects are shown in (K) WT and (L) *Dmd<sup>mdx-4Cv</sup>* cells. For statistical analysis, unpaired two-tailed *t* tests were used for comparisons between WT and *Dmd<sup>mdx-4Cv</sup>*. \*\*\*\*P < 0.0001. A one-way ANOVA with Tukey's multiple comparisons test was used to assess differences across treatment conditions within the same genotype. *P* values: \*\*\*\*P < 0.05 and \*\*\*\*P < 0.0001; ns, nonsignificant.</figcaption></figure>
<figure><img src="image_6.png" /><figcaption>**Fig. 4.** Evaluation of lysosomal biogenesis and repair pathways in *Dmd<sup>mdx-4Cv</sup>* mice. (A) Confocal images of serial transversal sections of GA muscle immunostained for TFEB and laminin. Scale bars, 100 or 25 µm. (B) Quantification of TFEB-positive nuclei (average ± SD, *n* = 5). (C) RNA sequencing analysis of GA muscle from 7-week-old *Dmd<sup>mdx-4Cv</sup>* and control mice, displaying the expression of certain genes under the control of TFEB (average FC shown, *n* = 5). (D) Representative confocal images of WT and *Dmd<sup>mdx-4Cv</sup>* primary myoblasts immunostained for TFEB. Scale bars, 40 or 10 µm. (E) Western blot analysis of nuclear and cytosolic fractions from WT and *Dmd<sup>mdx-4Cv</sup>* primary myoblasts for TFEB. A-tubulin and lamin B were used as loading controls for cytosolic and nuclear fractions, respectively. (F) Band intensity quantification of TFEB relative to A-tubulin (for cytosolic fractions) and lamin B (for nuclear fractions) (average ± SD). (G) Representative confocal images of TA transversal sections immunostained for ALIX and LAMP2. Scale bars, 25 µm. (H) Quantification of LAMP2<sup>+</sup>ALIX<sup>+</sup> puncta per myofiber (average ± SD, >692 myofibers analyzed from *n* = 6). (I) Western blot of TA muscle for ALIX and (J) band density quantification of ALIX relative to GAPDH (average ± SD, *n* = 6). An unpaired two-tailed *t* test was used for statistical comparisons of WT and *Dmd<sup>mdx-4Cv</sup>* groups. *t* test: \*P < 0.05, \*\*P < 0.01, and \*\*\*P < 0.0001.</figcaption></figure>
(Fig. 4, G and H). Increased ALIX protein levels were also confirmed by Western blotting in TA lysates (Fig. 4I). Similarly, we observed increased colocalization of LAMP2 and CHMP4B in *Dmd<sup>mdx-4Cv</sup>* muscle (fig. S6, C and D), although CHMP4B protein levels were unchanged (fig. S6E). Together, these results indicate the activation of the ESCRT-dependent repair pathway in response to lysosomal injury in dystrophic muscle.

## Increased lysophagy and impaired lysosome-autophagosome fusion in the dystrophic muscle

Given the accumulation of damaged lysosomes, we next evaluated lysophagy, the selective autophagic process that targets damaged lysosomes for degradation via autophagosome sequestration and subsequent fusion with intact lysosomes. Although overall autophagy defects have been previously reported in DMD, mainly due to mechanistic target of rapamycin (mTOR) hyperactivation (37–39), the link of these defects to lysosomal dysfunction remains unclear. First, we performed Western blotting on the TA for the *Microtubule Associated Protein 1 Light Chain 3 Beta* (MAP1LC3B or LC3B). An increase in LC3-I in the *Dmd<sup>mdx-4Cv</sup>* muscle (FC = 1.7) was observed (Fig. 5A). However, the ratio between the lipidated form (LC3-II), generated during the autophagosome formation and representing active autophagy, and the nonlipidated LC3-I was significantly reduced in *Dmd<sup>mdx-4Cv</sup>* muscles (FC = -1.9). This correlated with the increase in *Sequestosome 1* expression (SQSTM1 or p62) detected by Western blotting and puncta quantification (mean = 37.6 for *Dmd<sup>mdx-4Cv</sup>* versus 22.6 for WT) (Fig. 5B and fig. S6, F and G), an autophagy substrate cargo protein known to be integrated into the autophagosomes and degraded during autophagy. These results confirmed a previous report in the mdx model (37) and support the hypothesis of a decrease in the autophagic flux in the dystrophic muscle. To better understand the link of these defects to lysosomes, we performed immunostaining of LAMP2 and LC3B on TA cross sections (Fig. 5C). A colocalization of LAMP2 and LC3B was notably detected around the nuclei in both WT and *Dmd<sup>mdx-4Cv</sup>* muscles, indicating lysosome-autophagosome fusion (40, 41). Analyses of LAMP2 and LC3B particles showed an increase in LAMP2⁺ (mean = 27 for *Dmd<sup>mdx-4Cv</sup>* versus 23.9 for WT) but not of LC3B⁺ particles (mean = 19.3 for *Dmd<sup>mdx-4Cv</sup>* versus 19.7 for WT, nonsignificant difference) in *Dmd<sup>mdx-4Cv</sup>* muscle compared to WT (Fig. 5D). However, the number of LAMP2⁺LC3B⁺ particles, representing autolysosomes, was significantly decreased in *Dmd<sup>mdx-4Cv</sup>* muscle compared to WT (mean = 12.8 for *Dmd<sup>mdx-4Cv</sup>* versus 17.1 for WT). This analysis confirmed an increase in lysosome (LAMP2⁺LC3B⁻ particles) ratio in the *Dmd<sup>mdx-4Cv</sup>* muscle compared to WT control (65% for *Dmd<sup>mdx-4Cv</sup>* versus 46% for WT) (Fig. 5E) and a decrease in autolysosome (LAMP2⁺LC3B⁺ particles) ratio (35% for *Dmd<sup>mdx-4Cv</sup>* versus 54% for WT), suggesting a defect in lysosome-autophagosome fusion in the *Dmd<sup>mdx-4Cv</sup>* muscle. To specifically assess lysophagy, we analyzed the colocalization of LC3B and LGALS3 (Fig. 5F). The proportion of LC3B⁺LGALS3⁺ puncta (relative to total LC3B⁺ particles), indicative of autophagosome recruitment to damaged lysosomes, was significantly increased in *Dmd<sup>mdx-4Cv</sup>* muscle compared to WT (Fig. 5G). This increase was paralleled by a reduction in LC3B⁺LGALS3⁻ puncta, suggesting a shift toward lysophagy over other forms of selective autophagy. Similar trends were observed with SQSTM1: Both the number and proportion of SQSTM1⁺LGALS3⁺ puncta (relative to total SQSTM1) were elevated in *Dmd<sup>mdx-4Cv</sup>* muscle (fig. S6, G and H), further supporting increased targeting of damaged lysosomes for

degradation. Last, triple immunostaining for LAMP2, LGALS3, and LC3B (Fig. 5H) revealed the colocalization of all three markers, consistent with the sequestration of damaged lysosomes (LGALS3⁺LAMP2⁺) within LC3B⁺ autophagosomes—a hallmark of active lysophagy (42, 43). Together, these findings suggest that lysosomal defects may be contributing to an impaired lysosome-autophagosome fusion and an increase in lysophagy rate, all of which may lead to the overall perturbation of autophagy.

## Lipid perturbations in *Dmd<sup>mdx-4Cv</sup>* mice correlate with a damaged lysosomal system

We have reported altered lipid metabolism and cholesterol accumulation in dystrophic myofibers in DMD (14); however, the consequences of these alterations—and their potential contribution to disease progression, particularly in relation to lysosomal dysfunction—remain poorly understood. To further clarify the nature of these lipid perturbations, we evaluated muscle biopsies of young *Dmd<sup>mdx-4Cv</sup>* mice and a young patient with DMD. In agreement with our earlier findings (14), filipin staining revealed a significant accumulation of free cholesterol in dystrophic muscle (fig. S7A). Complementary lipidomic analysis of GA muscles from young *Dmd<sup>mdx-4Cv</sup>* mice confirmed this finding, revealing increased levels of cholesterol, cholesterol esters, and sphingomyelin relative to WT controls (fig. S7, B to E). Moreover, qPCR analysis showed dysregulation of key genes involved in cholesterol homeostasis, such as 3-hydroxy-3-methylglutaryl coenzyme A reductase (*Hmgcr*) involved in cholesterol synthesis or the adenosine 5'-triphosphate-binding cassette transporter (*Abca1*) and apolipoprotein E (*ApoE*) involved in cholesterol efflux (fig. S7F). These results reinforce that cholesterol accumulation is an early and intrinsic feature of DMD pathology, possibly contributing to broader metabolic and cellular dysfunction.

To assess whether cholesterol accumulation exacerbates the dystrophic phenotype and affects the lysosomal function, young WT and *Dmd<sup>mdx-4Cv</sup>* mice were fed either a cholesterol-rich high-fat diet or a standard diet for 4 weeks (Fig. 6A). Oil red O staining confirmed hepatic lipid accumulation in both WT and dystrophic mice under high-cholesterol diet (HCD) (Fig. 6B), although no significant changes in body or organ weights were observed (fig. S8A). Evaluation of disease severity by Sirius Red staining of the diaphragm and GA muscles revealed a significant increase in fibrosis in *Dmd<sup>mdx-4Cv</sup>* mice on HCD compared to those on a standard diet (Fig. 6, C to E). In contrast, WT mice showed no difference in fibrosis between diets. These findings suggest that excess dietary cholesterol exacerbates muscle pathology selectively in dystrophic mice. Nevertheless, no increase in muscle inflammation or sarcolemma permeabilization was caused by the HCD, as shown with CD11b and immunoglobulin (IgG) labeling on GA cross sections (fig. S8, C to E).

Measurement of free cholesterol in GA muscle lysates confirmed increased levels in *Dmd<sup>mdx-4Cv</sup>* mice on standard diet relative to WT controls (FC<sub>mdx-chow/WT-chow</sub> = 2.2) (Fig. 6F). Notably, cholesterol levels increased in both WT and *Dmd<sup>mdx-4Cv</sup>* mice on HCD, suggesting that dietary cholesterol is taken up even by healthy muscle. However, colabeling with filipin and LAMP2 revealed that in WT mice, accumulated cholesterol was not localized within lysosomes, whereas in *Dmd<sup>mdx-4Cv</sup>* mice, filipin-positive cholesterol colocalized with LAMP2, indicating lysosomal sequestration (Fig. 6, G and H). Thus, while HCD increases total muscle cholesterol in both genotypes, lysosomal accumulation of cholesterol occurs only in dystrophic
<figure><img src="image_7.png" /><figcaption>**Fig. 5.** Assessment of autophagy and lysophagy in *Dmd*<sup>mdx-4Cv</sup> muscle. (A) Western blot analysis of TA muscle for LC3, with GAPDH and total protein staining used as loading controls. Graphs show the band density quantification of LC3-I and LC3-II/LC3-I ratio, normalized to GAPDH (average ± SD, *n* = 6). (B) Western blot analysis of TA muscle for SQSTM1, with GAPDH and total protein staining used as loading controls. The graph shows the band density quantification of SQSTM1, normalized to GAPDH (average ± SD, *n* = 6). (C) Representative confocal images of TA transversal sections immunostained for LAMP2 and LC3B. White particles in merged images depict LAMP2<sup>+</sup>LC3B<sup>+</sup> particles representing autolysosomes. Scale bars, 100 or 25 µm. (D) Quantification of LAMP2<sup>+</sup>, LC3B<sup>+</sup>, and LAMP2<sup>+</sup>LC3B<sup>+</sup> particles in the myofibers (average ± SD, >1045 myofibers analyzed from *n* = 6). (E) Lysosome and autolysosome ratios were estimated by the quantification of the percentages of LAMP2<sup>+</sup>LC3B<sup>−</sup> (lysosomes) and LAMP2<sup>+</sup>LC3B<sup>+</sup> (autolysosomes) particles of total LAMP2<sup>+</sup> particles per myofiber (average ± SD). (F) Representative confocal images of TA transversal sections immunostained for LC3B and LGALS3. Scale bars, 25 µm. (G) Scatterplots (average ± SD) showing ratios of active lysophagy and non-lysophagy-related autophagy, estimated from the quantification of the percentages of LC3B<sup>+</sup>LGALS3<sup>+</sup> (lysophagy) and LC3B<sup>+</sup>LGALS3<sup>−</sup> (non-lysophagy-related autophagy) particles of total LC3B<sup>+</sup> particles per myofiber (average ± SD). (H) Representative confocal images of TA transversal sections immunostained for LC3B, LGALS3, and LAMP2. LAMP2<sup>+</sup>LC3B<sup>+</sup>LGALS3<sup>+</sup> puncta represent damaged lysosomes sequestered by autophagosomes. Scale bars, 25 µm. An unpaired two-tailed *t* test was used for statistical comparisons of WT and *Dmd*<sup>mdx-4Cv</sup> groups. *t* test: \* *P* < 0.05, \*\* *P* < 0.01, and \*\*\*\* *P* < 0.0001.</figcaption></figure>
<figure><img src="image_8.png" /><figcaption>**Fig. 6.** HCD exacerbates lysosomal cholesterol accumulation. (A) Schematic representation of the study setup. Three-week-old WT and *Dmd<sup>mdx-4Cv</sup>* mice were fed with a standard chow diet or an HCD/high-fat diet for 4 weeks. (B) Cross sections of livers stained with Oil Red O, labeling nonpolar lipids. (C) Sirius red (SR) staining of diaphragm and GA muscles cross sections. Scale bars, 200 or 100 µm (diaphragm) and 500 or 100 µm (GA). (D and E) Quantification of fibrosis (Sirus red-positive area) (average ± SD, *n* = 6). (F) Free cholesterol quantification in whole GA muscle lysates (average ± SD, *n* = 4). (G) Representative confocal images of serial transversal sections of GA muscle labeled with filipin (free cholesterol) and Syto13 (nuclei) and immunostained with LAMP2. Scale bars, 25 µm. (H) Colocalization analysis of filipin signal overlapping LAMP2, calculated by Manders' overlap coefficient (average ± SD, *n* = 3). An ANOVA test with Tukey's multiple comparisons test was used for statistical comparisons. \*P < 0.05, \*\*P < 0.05, \*\*\*P < 0.001, and \*\*\*\*P < 0.0001.</figcaption></figure>
muscle, pointing to a defect in cholesterol handling and lysosomal processing in *Dmd*<sup>mdx-4Cv</sup> mice.

To test whether cholesterol accumulation contributes to LMP, we analyzed the colocalization of LGALS3 and LAMP2 in GA muscles (Fig. 7A). In *Dmd*<sup>mdx-4Cv</sup> mice, HCD significantly increased the number of LAMP2<sup>+</sup>LGALS3<sup>+</sup> puncta compared to standard diet (FC<sub>mdx-HCD/mdx-chow</sub> = 1.8) (Fig. 7B), along with elevated *Lgals3* mRNA (FC<sub>mdx-HCD/mdx-chow</sub> = 1.75) and protein expression (FC<sub>mdx-HCD/mdx-chow</sub> = 14.25) (Fig. 7, C to E). A smaller but detectable increase in LAMP2<sup>+</sup>LGALS3<sup>+</sup> puncta was also observed in WT-HCD compared to WT-chow mice (mean = 3.2 versus 0.1 in WT-chow), although without significant changes in Gal-3 transcript or protein levels, suggesting minimal lysosomal stress in healthy mice under HCD.

We next assessed autophagy perturbations under these conditions by monitoring SQSTM1 and LC3B (Fig. 7A). HCD led to an increase in SQSTM1 puncta in both WT and *Dmd*<sup>mdx-4Cv</sup> mice (mean: 3.2 → 10.7 in WT; 17.9 → 29.2 in *Dmd*<sup>mdx-4Cv</sup>) (Fig. 7F), suggesting the accumulation of undegraded autophagy substrates. We also evaluated LGALS3/LC3B colocalization (Fig. 7A). The proportion of LC3B<sup>+</sup>LGALS3<sup>+</sup> puncta—indicative of damaged lysosomes being targeted by autophagosomes—increased in both WT and *Dmd*<sup>mdx-4Cv</sup> mice under HCD (from 5.9 to 14% in WT; from 50.9 to 94.8% in *Dmd*<sup>mdx-4Cv</sup>), indicating enhanced lysophagy activity (Fig. 7G). These results suggest that dietary cholesterol promotes lysosomal damage and induces lysophagy, with a more pronounced effect in dystrophic muscle.

Together, these findings indicate that excess cholesterol contributes to lysosomal injury in skeletal muscle, with greater severity in dystrophic muscle because of an underlying susceptibility. While HCD triggers some lysosomal stress in WT muscle, *Dmd*<sup>mdx-4Cv</sup> muscle exhibits cholesterol sequestration within lysosomes and a high increase in lysophagy and accumulation of autophagy substrates, highlighting a pathogenic positive feedback loop between lipid dysregulation and impaired lysosomal function in DMD.

## LMP rescue varies with gene therapy modality in dystrophic models

Following the identification of LMP in both DMD and LGMDR5 mouse models, we sought to evaluate the extent to which current gene therapy strategies can correct these defects. This comparison is particularly relevant given the shared pathological hallmark of both diseases: disruption of the DAGC. Despite this similarity, the therapeutic approaches differ—DMD gene therapy involves the delivery of a truncated version of the large dystrophin gene (µDys), while LGMDR5 therapy enables the delivery of a full-length γ-sarcoglycan (SGCG) gene. To evaluate µDys gene therapy, 6-week-old *Dmd*<sup>mdx</sup> mice were systemically injected with serotype 9 AAV (AAV9) vectors encoding µDys under the control of the muscle-specific spc5.12 promoter. Two doses were tested: a suboptimal (low) dose and a higher (therapeutic) dose known to be effective in this model (Fig. 8A) (44). For LGMDR5, we used the *Sgcg*<sup>-/-</sup> knockout model and administered AAV9-SGCG at an established optimal dose of 2 × 10<sup>13</sup> vg/kg (45).

For µDys gene therapy, a dose effect was noticed in terms of vector transduction assessed by the quantification of viral copy number (VCN) (fig. S9A) and transgene expression at the mRNA (fig. S9B) and protein levels (fig. S9, C and D), with the high dose transducing nearly all GA muscle fibers (~95% of positive myofibers) (fig. S9D).

As for the LGMDR5 gene therapy, the vector was detected in terms of VCN and mRNA transcript (fig. S9, E and F), and an optimal expression of the transgene is achieved (fig. S9G), transducing all muscle fibers with the human SGCG protein (fig. S9H). Assessment of systemic disease markers revealed the partial normalization of circulating creatine kinase (CK) and myomesin-3 (MYOM3) fragments in *Dmd*<sup>mdx</sup> mice treated with µDys, even at the high dose, although values did not reach WT levels (Fig. 8, B and C). In contrast, SGCG gene therapy in *Sgcg*<sup>-/-</sup> mice fully normalized both biomarkers to WT values. Functional rescue, evaluated via an escape test, showed no improvement with the low µDys dose, while the high dose significantly improved muscle strength compared to untreated *Dmd*<sup>mdx</sup> mice, although not fully restoring it to WT levels (Fig. 8D). SGCG gene therapy, however, restored muscle force to levels indistinguishable from WT controls.

We next examined whether gene therapy corrected lysosomal abnormalities. In *Dmd*<sup>mdx</sup> mice, high-dose µDys treatment significantly decreased *Lgals3* mRNA in the GA by 5.5-fold relative to untreated mice (Fig. 8E). SGCG gene therapy also restored *Lgals3* expression to WT levels. However, immunostaining for LGALS3 and LAMP2 revealed persistent LAMP2<sup>+</sup>LGALS3<sup>+</sup> puncta in *Dmd*<sup>mdx</sup> mice treated with µDys (Fig. 8F). No reduction was observed with the low dose, and only a modest decrease was seen with the high dose (median = 6 in untreated versus 5 in µDys-high) (Fig. 8G), indicating incomplete correction of LMP. In stark contrast, SGCG gene therapy in *Sgcg*<sup>-/-</sup> mice resulted in a marked reduction of LAMP2<sup>+</sup>LGALS3<sup>+</sup> puncta (median = 5 in untreated versus 1 in treated), reflecting the effective restoration of lysosomal integrity. These findings suggest that a subtherapeutic dose of µDys is insufficient to ameliorate lysosomal damage, and even high-dose µDys therapy—despite efficient transduction and transgene expression—only partially corrects lysosomal dysfunction. By comparison, gene therapy in the LGMDR5 mouse model achieved full correction of LMP and broader disease hallmarks. This underscores the importance of directly targeting lysosomal dysfunction in DMD. Combining µDys gene therapy with complementary strategies aimed at restoring the lysosomal function may be necessary to achieve full therapeutic benefit in DMD, as demonstrated by the more complete rescue observed in SGCG-treated *Sgcg*<sup>-/-</sup> mice.

## TR-mediated lysosomal correction improves microdystrophin gene therapy at a suboptimal dose in a combined therapy setup

After the identification of lysosomal damage in the dystrophic muscle and the insufficiency of the µDys gene therapy alone to correct these damages, we designed a combined therapy setup where a suboptimal dose of rAAV-µDys would be administered with a drug capable of alleviating lysosomal dysregulations. For that, TR, an FDA-approved dietary disaccharide, was selected, as it has been shown to activate lysosomal biogenesis and alleviate lysosomal damage in various neurodegenerative diseases (46, 47). TR was administered alone (2% in drinking water) or in combination with a suboptimal dose of rAAV-µDys in *Dmd*<sup>mdx-4Cv</sup> mice. To this end, five groups of male mice were included in this study (Fig. 9A). TR was administered from 3 weeks of age until the end of the study 4 weeks later. rAAV-µDys was injected intravenously 1 week after the beginning of treatment with trehalose. First, we assessed whether TR treatment affected muscle transduction or transgene expression. Quantification of VCN and mRNA transcripts in the GA and TA muscles showed no significant differences between the two groups injected with the vector with or without TR
<figure><img src="image_9.png" /><figcaption>**Fig. 7. HCD increases lysosomal damage in skeletal muscle.** (A) Representative confocal images of transversal sections of GA muscle immunostained for LAMP2 and LGALS3 (upper panel), LC3B and LGALS4 (middle panel), and SQSTM1 (lower panel). Scale bars, 25 µm. (B) Quantification of double-positive puncta (LAMP2⁺LGALS3⁺) in the myofibers of GA muscles (scatterplots with median, >784 myofibers analyzed from *n* = 6). (C) Relative *Lgals3* mRNA expression in the GA normalized to *Rplp0* (average ± SD, *n* = 5 or 6). (D) Western blot analysis of GA muscles for LGALS3 (*n* = 6). Total protein staining is used as the loading control. (E) Band intensity quantification of LGALS3 relative to total protein staining (average ± SD, *n* = 6). (F) Quantification of SQSTM1⁺ in the myofibers of GA muscles (*n* > 119 myofibers). (G) Scatterplots (average ± SD) showing ratios of active lysophagy and non-lysophagy-related autophagy, estimated from the quantification of the percentages of LC3B⁺LGALS3⁺ and LC3B⁺LGALS3⁻ particles of total LC3B⁺ particles per myofiber (average ± SD, >539 myofibers). An ANOVA test with Tukey's multiple comparisons test was used for statistical comparisons. \*P < 0.05, \*\*P < 0.05, \*\*\*P < 0.001, and \*\*\*\*P < 0.0001.</figcaption></figure>
<figure><img src="image_10.png" /><figcaption>**Fig. 8.** Evaluation of gene therapy strategies for DMD and LGMDR5 in corresponding mouse models. (A) Schematic representation of the study setup. Six-week-old *Dmd*<sup>mdx</sup> mice were injected intravenously with a rAAV encapsulating a µDys encoding sequence, under the control of an spc512 promoter, at two doses of 5 × 10<sup>12</sup> vg/kg (low dose) and 1 × 10<sup>13</sup> vg/kg (high dose). Four-week-old *Sgcf*<sup>−/−</sup> mice were injected intravenously with a rAAV9 encapsulating the human sequence of SGCG, under the control of a desmin promoter, at a dose of 2 × 10<sup>13</sup> vg/kg. IV, intravenous. (B) Dosage of CK in serum of mice at the end of the study (average ± SD). (C) ELISA quantification of MYOM3 in the serum of the mice before euthanasia. Data are presented as the scatterplot (average ± SD). (D) Global force evaluation by an escape test (force in newton normalized to body weight in grams) before euthanasia of mice (average ± SD). (E) Relative *Lgals3* mRNA expression in the GA normalized to *Rplp0* (average ± SD). (F) Representative confocal images of transversal sections of GA muscle immunostained for LAMP2 and LGALS3. Scale bars, 25 µm. (G) Quantification of double-positive puncta (LAMP2<sup>+</sup>LGALS3<sup>+</sup>) in GA myofibers (median and range shown). An ANOVA test with Tukey's multiple comparisons test was used for statistical comparisons. \*P < 0.05, \*\*P < 0.05, \*\*\*P < 0.001, and \*\*\*\*P < 0.0001. Gray asterisks represent relative comparison to the "WT_Phy_Ser" group, and red asterisks represent comparison to the "mdx_Phy_Ser" group.</figcaption></figure>

(fig. S10, A and B). No significant difference was noticed in the number of dystrophin-positive fibers as well (fig. S10, C and D). Serum was collected at the end of the study before functional evaluation, and different circulating biomarkers were quantified. CK levels decreased significantly compared to control *Dmd*<sup>mdx-4Cv</sup>, reaching nonsignificant difference from WT mice only for mice treated with the combined

therapy (FC<sub>mdx-µDys/mdx</sub> = -2.7; FC<sub>mdx-µDys+TR/mdx</sub> = -5.5) (Fig. 9B). MYOM3 levels were significantly lower with TR treatment alone (FC<sub>mdx-TR/mdx</sub> = -1.3) but still not normalized to the WT level even with the combined therapy (Fig. 9C).

As for the global muscle strength assessed by an escape test, a slight improvement in muscle strength compared to *Dmd*<sup>mdx-4Cv</sup> was
<figure><img src="image_11.png" /></figure>

<figure><img src="image_12.png" /><figcaption>**Fig. 9.** TR partially restores dystrophic phenotypes and potentiates µDys gene therapy in *Dmd<sup>mdx-4Cv</sup>* mice. (A) Study setup. *Dmd<sup>mdx-4Cv</sup>* mice were started on TR treatment (2% dilution in drinking water) at 3 weeks. For groups 4 and 5, mice were injected intravenously with a rAAV9 encapsulating the µDys sequence at a dose of 7 × 10¹² vg/kg. (B) Seric CK analysis before the escape test (average ± SD, n = 5). (C) ELISA quantification of MYOM3 before the escape test (average ± SD, n = 5). (D) Global force evaluation (normalized to mice body weight, N/g) by the escape test (average ± SD, n = 5). (E) Histological characterization of muscles. Top panel: H&E labeling of GA muscle cross sections. Middle panel: Sirius red labeling of diaphragm and GA muscles. Bottom panel: Representative images of GA muscle serial cross sections immunostained for mouse immunoglobulin (IgG), laminin, and CD11b. Scale bars, 200 µm (H&E) and 500 and 100 µm (zoomed-in images). (F) Analysis of myofiber size distribution in GA muscles represented by the variance coefficient calculated as [SD of the muscle fiber size/mean of muscle fiber size] * 1000 (median with min-max values). (G) Centronucleation index in the GA muscle (median with min-max values). (H and I) Fibrosis analysis of GA and diaphragm by the quantification of Sirius red-positive areas on transversal sections (average ± SD). (J) Quantification of IgG uptake by myofibers (positive myofiber number normalized to muscle cross-sectional area). (K) Evaluation of CD11b⁺ cell infiltration in the muscle by the quantification of CD11b area. An ANOVA test with Tukey's multiple comparisons test was used for statistical comparisons (compared to WT control and mdx control). ANOVA: *P* < 0.05, **P* < 0.05, ***P* < 0.001, and ****P* < 0.0001. Black asterisks represent relative comparison to the "WT control" group, and red asterisks represent comparison to the "mdx control" group. An unpaired two-tailed *t* test was used for statistical comparisons between mdx-µDys and mdx_µDys + TR groups. *t* test: *P* < 0.05 and **P* < 0.01.</figcaption></figure>
observed in mice treated with TR alone or µDys alone (Fig. 9D), albeit not statistically significant. Only mice treated with the combined therapy showed a significant improvement in muscle strength compared to the untreated *Dmd<sup>mdx-4Cv</sup>* mice and showed no longer a significantly different strength from the WT group. To confirm and expand on these findings, we replicated the study in an independent cohort and added further functional tests. In the forelimb grip strength test, TR alone led to a modest, nonsignificant increase in peak grip strength, while µDys alone had no detectable effect (fig. S11A). However, the combined treatment significantly increased peak grip strength compared to mdx controls and reached values not significantly different from WT. Normalization to body weight confirmed this effect, with a similar trend showing recovery to WT levels in the combined group (fig. S11B). We then evaluated the contractility of extensor digitorum longus muscles ex vivo. Maximal tetanic force (a measure of sustained contraction) was not significantly improved by TR or µDys alone but was significantly increased by the combined treatment (156 ± 12.8 for mdx versus 184 ± 25.6 nM/mm² for mdx + µDys + TR) (fig. S11, C and D). Twitch force (response to a single stimulus) was reduced in mdx mice compared to WT (42.6 ± 7.9 versus 32.2 ± 10.5 nM/mm²). TR alone did not restore twitch force, while µDys and µDys + TR both returned values to WT levels (µDys: 40.9 ± 7.2; µDys + TR: 38.1 ± 5.7 nM/mm²) (fig. S11, E and F). These additional functional tests suggest that µDys gene therapy alone can restore twitch (short, isolated) contractions but not sustained force or functional grip strength, and TR supplementation substantially boosts sustained force generation and overall muscle function when combined with µDys.

We then performed histological characterization, following the different treatments (Fig. 9E). Hematoxylin and eosin (H&E) staining of the GA transversal section showed an improvement of muscle histology for both TR- and µDys only-treated mice compared to nontreated *Dmd<sup>mdx-4Cv</sup>* mice (Fig. 9E). However, a better correction was achieved in the combined therapy group, where myofiber size distribution was best restored (Fig. 9F), and the centronucleation index decreased (Fig. 9G). Fibrosis was assessed by Sirius Red labeling in the diaphragm and GA, which are among the most fibrotic muscles in this mouse model (Fig. 9E). As for fibrosis in the GA, the TR-treated mice showed a significant reduction, and the combined therapy group had significantly lower fibrosis than the µDys only-treated group (Fig. 9H). In the diaphragm, neither TR nor µDys treatment alone decreased fibrosis (Fig. 9I). Only the combination of treatments reduced fibrosis significantly in this muscle. In addition, myofiber necrosis was evaluated in the GA by IgG uptake (Fig. 9E). A significant reduction of IgG-positive myofibers was observed in both TR- and µDys-treated mice, with a better correction in TR-treated mice versus µDys only-treated mice (nonsignificant difference from WT controls) (Fig. 9J). The best correction was attained in the combined therapy group, where very few IgG-positive myofibers were detected, indicating a better integrity of myofiber sarcolemma. CD11b staining was also performed to assess the inflammation level in the GA muscle (Fig. 9E). A significant decrease in inflammatory areas was observed with both TR and µDys treatments, with the combined therapy reaching the best correction (most significant decrease compared to nontreated *Dmd<sup>mdx-4Cv</sup>*) (Fig. 9K).

To investigate whether TR ameliorates lysosomal function and defects, we quantified LAMP2⁺LGALS3⁺ puncta—indicative of lysosomal damage—in TA muscle sections (Fig. 10, A and B). *Dmd<sup>mdx-4Cv</sup>* muscles exhibited a high number of these puncta (mean = 10.4

compared to WT controls (mean = 0.5). µDys monotherapy partially reduced their number (mean = 4.7), while TR monotherapy produced a more substantial reduction (mean = 2.3). Notably, only the combined treatment normalized the number of LAMP2⁺LGALS3⁺ puncta to WT levels (mean = 1.1), indicating a complete correction of lysosomal damage. Further morphometric analysis of LAMP2-stained sections revealed enlarged lysosomes in *Dmd<sup>mdx-4Cv</sup>* muscles, consistent with lysosomal stress (Fig. 10C). Both µDys and TR reduced lysosome size, but the most robust normalization was observed with the combined treatment. The lysosome number, which was also elevated in *Dmd<sup>mdx-4Cv</sup>* remained unaffected by TR alone but was significantly decreased by µDys and further reduced with the combined therapy (Fig. 10D). Changes of lysosome number mirrored changes in TFEB activity, the key regulator of lysosome biogenesis and subsequently number. Although TFEB was partially activated in dystrophic muscle, its nuclear translocation and lysosome biogenesis were unchanged by TR alone (fig. S12, A and B). µDys decreased TFEB nuclear localization, and the combined treatment fully normalized both TFEB activity and the lysosome number.

To explore effects on autophagy status, we quantified SQSTM1 puncta (Fig. 10E). TR significantly reduced SQSTM1 levels (mean = 23.2) to near-WT levels (mean = 22.5), whereas µDys alone also reduced them, albeit less effectively (mean = 28.4). The combined treatment led to the lowest levels of SQSTM1 puncta (mean = 16.5), suggesting the enhanced clearance of autophagic substrates. Last, we assessed lysophagy via LC3B⁺LGALS3⁺ ratios (Fig. 10F). This ratio was elevated in dystrophic muscle (mean = 56.8% versus 11.9% in WT), significantly reduced by TR (mean = 21.8%) but not by µDys alone (mean = 58.5%). The combined treatment decreased this ratio below WT levels (mean = 1.9%), suggesting a restored lysosomal function and a shift toward other selective autophagy pathways.

To assess the effects of different treatments on the reversion of transcriptomic perturbations in dystrophic muscle, we performed the analysis of bulk RNA sequencing on the GA muscles. On the first two principal components of the principal components analysis (PCA) (Fig. 10G), the group treated with TR clustered around the untreated *Dmd<sup>mdx-4Cv</sup>* group, and the group treated only with µDys was widely distributed on the PCA. Only the group treated with µDys + TR was tightly clustered close to the WT group. Next, we investigated the evolution of all differentially expressed genes (DEGs) found in the *Dmd<sup>mdx-4Cv</sup>* untreated group (comparison: untreated *Dmd<sup>mdx-4Cv</sup>* versus WT, adjusted *P* value <0.05) upon different treatments (Fig. 10, H and I). Similar to PCA, the heatmap of the total 8625 DEGs showed comparable profiles between the TR-treated and untreated *Dmd<sup>mdx-4Cv</sup>* groups, while µDys-alone treatment shifted the heatmap signature closer to WT (Fig. 10H). Notably, the closest profile to WT was achieved with the combined therapy. Among the 4088 down-regulated and 4537 up-regulated DEGs identified in *Dmd<sup>mdx-4Cv</sup>* muscle, only 6 (0.15%) up-regulated and 4 (0.08%) down-regulated DEGs were restored by TR, while µDys alone restored 2394 (58.5%) up-regulated and 3010 (66.3%) down-regulated DEGs (Fig. 10I). The combination of TR and µDys restored 2887 (70%) down-regulated and 3564 (78.5%) up-regulated DEGs. In addition, the expression of more genes was partially or completely normalized by the treatment of µDys + TR compared to µDys alone. A total number of 2009 down-regulated and 2213 up-regulated DEGs were incompletely restored with µDys compared to 1271 down-regulated and 1360 up-regulated DEGs for µDys + TR. We also performed gene set enrichment analysis (GSEA) to evaluate the correction
<figure><img src="image_13.png" /><figcaption>**Fig. 10.** TR alleviates lysosomal damage and enhances transcriptomic correction in µDys-treated *Dmd*<sup>mdx-4Cv</sup> mice. (A) Representative confocal images of TA muscle sections immunostained for LAMP2 and LGALS3 (upper panel), LC3B and LGALS3 (middle panel), and SQSTM1 (lower panel). Scale bars, 25 µm. (B) Quantification of LAMP2<sup>+</sup>LGALS3<sup>+</sup> puncta in TA myofibers (average ± SD, >538 myofibers analyzed from *n* = 5). (C and D) Density plot with a nonlinear fit Gaussian curve, showing the distribution of average spot size (C) and lysosome number per myofiber (D) (>538 myofibers analyzed from *n* = 5). (E) Quantification of SQSTM1+ puncta in TA myofibers (average ± SD, >552 myofibers analyzed from *n* = 5). (F) Ratios of active lysophagy and non-lysophagy-related autophagy, estimated from the quantification of LC3B<sup>+</sup>LGALS3<sup>+</sup> and LC3B<sup>+</sup>LGALS3<sup>-</sup> percentages (average ± SD, >529 myofibers analyzed from *n* = 5). (G to I) Comparison of global transcriptomic changes in GA muscle. (G) PCA plot using the expression level of 10000 genes with the highest variance. Gene expression was first normalized by size factors and transformed by variance-stabilizing transformation using the DESeq2 RStudio package. (H) Heatmap presenting log<sub>2</sub>FC in comparison to WT muscle for all 8625 DEGs found in mdx muscle. The log<sub>2</sub>FC values are illustrated in row z-scores, colored from blue to red, arranged from the lowest to highest. (I) Volcano plots of multiple comparisons illustrating transcriptomic changes before and after different treatments. Down-regulated and up-regulated DEGs are colored in blue and red, respectively, in all volcano plots. The right upper panel shows volcano plots comparing mdx control to treated groups, in which DEGs are correctly restored after treatment. The lower right panel shows treated groups compared to WT control, in which significant DEGs are the genes that are not or incompletely restored after treatment. An ANOVA test with Tukey's multiple comparisons test was used for statistical comparisons in (B) to (E). ANOVA: ****P < 0.0001. Black asterisks represent relative comparison to the "WT control" group, and red asterisks represent comparison to the "mdx control" group.</figcaption></figure>
of known dysregulated pathways in the dystrophic muscle, such as inflammatory response, calcium signaling, nitrogen species metabolic process, and apoptosis (fig. S13, A and B). Metabolic pathways like cholesterol homeostasis, lipid storage, endocytosis, and MTORC1 (mTOR complex 1) signaling (fig. S13C) were also evaluated. Notably, these pathways were all significantly closer to normal with the combined treatment of µDys + TR compared to µDys alone. Together, these data suggest that combining TR and µDys treatments substantially improves pathophysiology, lysosomal damage, and transcriptional dysregulation compared to suboptimal dosed gene therapy standalone treatment.

## DISCUSSION

The classical view of DMD pathophysiology centers on mechanical destabilization of skeletal myofibers as a result of the absence of dystrophin, rendering them susceptible to contraction-induced damage and sarcolemma disruption (1, 48). However, emerging evidence has highlighted additional layers of complexity, particularly metabolic disturbances (12), such as mitochondrial dysfunction (12, 49–51) and lysosomal perturbations (39, 52–54). Perturbations of lipid metabolism were also reported (55). In particular, we and others (14, 56, 57) have recently detected perturbations of cholesterol metabolism. Such disruptions of cholesterol metabolism are often linked with lysosomal damage in both neurodegenerative and lysosomal storage diseases (16, 58–61).

The lysosomal damage biomarker Gal-3 (20) has been reported to be up-regulated in the dystrophic muscle (24). This up-regulation was initially attributed to infiltrating inflammatory cells (25), particularly macrophages, which express high levels of Gal-3 (26). However, recent studies also support Gal-3 expression within muscle fibers themselves, where it plays a role in regeneration (62). In our study, we confirmed Gal-3 up-regulation in dystrophic muscle and demonstrated its localization to enlarged lysosomes within myofibers. Assessing total Gal-3 expression alone is insufficient to evaluate lysosomal damage in dystrophic muscle, as macrophages— which are abundant in this context—contribute substantially to overall Gal-3 levels. Therefore, to more accurately quantify lysosomal damage within myofibers, we focused on the detection of LAMP2<sup>+</sup>LGALS3<sup>+</sup> puncta, providing a more specific marker of lysosomal injury in muscle fibers and enabling the assessment of responses to therapeutic interventions.

We further characterized the lysosomal alterations in dystrophic muscle, observing both an increased number and size of lysosomes, along with the presence of large, electron-dense lysosomes on electron micrographs. Notably, similar lysosomal abnormalities were present in isolated Dmd<sup>mdx-4Cv</sup> primary myoblasts, including evidence of LMP (LAMP2<sup>+</sup>LGALS3<sup>+</sup> puncta), impaired acidification, reduced proteolytic activity, and defective recovery after lysosomal injury. The fact that these defects persist in isolated cells, outside the dystrophic tissue environment (which contains additional stressors such as inflammation, fibrosis, oxidative stress, calcium overload, and lipid accumulation), suggests that lysosomal dysfunction may be at least partly cell-intrinsic. Whether these are truly intrinsic defects or carryover effects from the in vivo environment remains to be determined.

An interesting observation was the impaired recovery of lysosomal integrity and acidity in Dmd<sup>mdx-4Cv</sup> myoblasts, in contrast to the effective, time-dependent recovery observed in WT cells. This

suggests that Gal-3 recruitment alone is insufficient to restore the lysosomal function in DMD cells. In whole muscle, we observed activation of the endolysosomal damage response, including lysosome repair, replacement, and removal pathways. These responses involved both Gal-3 up-regulation (29) and increased nuclear translocation of the lysosomal biogenesis regulator TFEB (33), along with the increased expression of TFEB target genes such as cathepsins (30–32, 53). However, the extent to which these compensatory mechanisms restore the lysosomal function appears limited. TFEB translocation was incomplete, with a substantial portion of TFEB retained in the cytosol. Furthermore, persistent LMP (LAMP2<sup>+</sup>LGALS3<sup>+</sup> puncta) was detected in 1-year-old dystrophic mice, indicating unresolved lysosomal stress despite chronic activation of these compensatory pathways.

We also identified lysosomal defects that may contribute to impaired autophagy. While previous studies have linked defective autophagy in DMD primarily to mTOR hyperactivation (37–39), we observed a reduced rate of autophagosome-lysosome fusion and increased lysophagy. Similar lysosomal dysfunction has been described in glycogen storage disease type II (GSDII/Pompe disease), where impaired autophagosome-lysosome fusion occurs despite suppressed mTOR signaling (63). In this context, overactivation of TFEB can rescue the autophagic block (64), suggesting that defective lysosomal function—rather than mTOR signaling alone—may underlie impaired autophagic flux. In DMD muscle, our findings support a model in which the decreased availability of functional lysosomes limits autophagosome-lysosome fusion, contributing to autophagic dysfunction. This deficiency appears only partially compensated by TFEB-mediated lysosomal biogenesis. Moreover, we found that lysophagy, the selective clearance of damaged lysosomes, is strongly up-regulated in dystrophic muscle. While this process aims to restore lysosomal homeostasis, it may divert autophagy machinery from other selective autophagy pathways (65), further limiting the muscle's ability to maintain proteostasis. However, these conclusions remain preliminary. Additional studies are needed to confirm the causal contribution of lysosomal dysfunction to autophagy impairment in DMD. In particular, more detailed analyses of autophagy flux and lysophagy flux are required, as well as mechanistic studies dissecting the interplay between lysosomal repair, biogenesis, and autophagosome-lysosome fusion. Such work will be essential to fully clarify the mechanisms driving autophagy defects in dystrophic muscle.

To directly assess whether cholesterol accumulation contributes to lysosomal damage, we fed mice an HCD. While increased cholesterol levels were observed in the skeletal muscle of both WT and dystrophic mice, lysosomal cholesterol accumulation, lysosomal damage, and fibrosis were markedly more pronounced in dystrophic muscle. Although previous studies have reported that lipid- and cholesterol-rich diets can worsen the dystrophic phenotype in DMD and other muscular dystrophies (56, 66), here we establish a link between elevated muscle cholesterol, lysosomal dysfunction, and exacerbation of disease severity in the context of muscular dystrophy.

We next investigated whether gene therapy could reverse lysosomal damage in dystrophic muscle. In the Dmd<sup>mdx</sup> mouse model, we observed a dose-dependent reduction in LMP, measured by LAMP2<sup>+</sup>LGALS3<sup>+</sup> puncta, with increasing doses of AAV-µDys. However, despite near-complete myofiber transduction with the highest µDys dose, a significant level of residual lysosomal damage persisted. In the LGMDR5 model, where we also detected substantial LMP, gene therapy using a full-length γ-sarcoglycan transgene
(45) resulted in an almost complete correction of lysosome damage. This discrepancy is particularly notable given the pathophysiological similarities between DMD and LGMDR5, both of which involve the disruption of the DAGC. A key distinction between these two gene therapies lies in the restored protein: full-length γ-sarcoglycan in LGMDR5 versus a truncated μDys lacking much of the spectrin-repeat rod domain in DMD. These findings raise the possibility that the rod domain of dystrophin may play an important role in maintaining lysosomal homeostasis in skeletal muscle. To determine whether the residual lysosomal damage in μDys-treated DMD muscle reflects a limitation of the microdystrophin construct itself or whether it is due to irreversible damage present before treatment, further studies are needed. Future work should assess the therapeutic potential of longer dystrophin constructs containing additional rod domains (67) or full-length dystrophin (68). These investigations will help clarify whether full structural restoration of dystrophin is required for complete lysosomal repair or whether some damage becomes irreversible at advanced disease stages.

After observing that μDys gene therapy alone did not fully correct LMP, we sought to test a combined therapeutic approach using μDys and a pharmacological agent capable of mitigating lysosomal damage. TR was selected on the basis of its reliable safety profile, affordability as a dietary disaccharide (69), and prior clinical testing in neurodegenerative disease (70) and oculopharyngeal muscular dystrophy (71, 72). In addition, TR is known to promote lysosomal biogenesis and autophagy (47), making it a promising candidate for targeting lysosomal dysfunction. Our experimental design compared the effects of TR alone, a suboptimal dose of AAV-μDys alone, and their combination. TR monotherapy produced notable benefits, particularly reducing LMP and other lysosomal stress markers and decreasing myofiber membrane permeabilization and necrosis (IgG uptake). The combined therapy of μDys and TR yielded the most robust therapeutic outcomes. These findings highlight the role of TR in alleviating lysosomal dysfunction and suggest a direct link between lysosomal damage and myofiber necrosis. In the skeletal muscle, we observed the normalization of myofiber size distribution, improved centronucleation index, reduced fibrosis and inflammation, and enhanced muscle function, including global muscle force, grip strength, and isolated tetanic force, compared to untreated dystrophic mice. Transcriptomic analysis further supported the additive effect of the combined treatment. While TR alone had a limited impact on transcriptomic correction, its combination with μDys amplified the normalization of key pathways and biological processes, suggesting a synergistic interaction between μDys and TR. It is also worth noting that TR has been shown in neurodegenerative models to modulate the intestine microbiota, oxidative stress, and inflammation (73), all which are disrupted in DMD. Therefore, the therapeutic benefits observed here may involve not only direct effects on the endolysosomal system but also modulation of broader pathophysiological processes. One limitation of our study is that we tested only a single dose of TR and μDys. Future work should explore dose escalation and long-term treatment effects, as well as investigate whether initiating therapy at later disease stages can reverse established lysosomal damage. Such studies will be crucial to define the therapeutic window and optimal treatment strategies for addressing lysosomal dysfunction in DMD.

In summary, our study reveals a previously underappreciated and critical role for lysosomal damage in the pathophysiology of DMD. We demonstrated LMP in muscle from patients with DMD

and GRMD dogs and extensively in the *Dmd*<sup>mdx-4Cv</sup> mouse model and primary muscle cells. Whether lysosomal dysfunction is a primary driver of disease progression or a secondary consequence of other pathological events remains an important question for future research. This issue is especially relevant given that lysosomal damage is only partially corrected by μDys gene therapy in mice. Our proof of concept that combining lysosomal repair strategies with μDys gene therapy improves outcomes paves the way for novel combinatorial treatments for DMD.

## MATERIALS AND METHODS

### Patients' biopsies

Open muscle biopsies were obtained for diagnostic purposes from the quadriceps (patient 1) or the deltoid muscles (patients 2, 3, 4, and 5) (see biopsies' details in table S1). The muscle location was not available for patient 6. All patients provided informed consent for the use of their biological samples for research purposes, in accordance with French regulations. The specimens were snap-frozen in isopentane cooled in liquid nitrogen for histoenzymology and immunohistochemistry. Transversal cryosections were stained using standard histological and histoenzymological methods. Histologically normal muscles served as controls. Genetic confirmation was done by Sanger sequencing.

### Animal care and use

WT (C57BL/10J and C57BL/6J), *Dmd*<sup>mdx</sup> (C57BL/10ScSn-Dmdmdx/J), and *Dmd*<sup>mdx-4Cv</sup> (B6Ros.Cg-Dmdmdx-4Cv/J) mice were supplied by Charles Rivers Laboratories, Miserey, France. Animals were housed in cages with regular 12-hour day/12-hour night cycles in a specific pathogen-free barrier facility and provided with water and food ad libitum. All animal experimentation procedures were conducted according to French and European guidelines for the care and use of animals for experimental purposes. All procedures were approved by ethical committee CEEA-51 and the French Ministry of Higher Education, Research, and Innovation under APAFIS numbers 19736 (DAP2018-024-B) and 35896 (DAP2022-004-C). Only male mice were used for our studies. The high-fat cholesterol-rich diet was supplied by SAFE (260HF diet supplemented with 20 g cholesterol/kg). TR was supplied by Sigma-Aldrich (ref. T9449) in powder form and dissolved in the mice's drinking water at a 2% concentration, which was changed twice a week. For rAAV injections, 100 µl of viral vectors diluted in saline solution was injected intravenously into the tails of the mice. For blood sampling, mice were anesthetized with isoflurane or a mixture of ketamine (100 mg/kg)/xylazine (10 mg/kg), and blood was collected from the retro-orbital vein on dry capillaries and centrifuged at 10,000 rpm for 10 min. Serum was then collected and stored at -80°C until analysis. Mice were euthanized by cervical dislocation at the end of the studies, and tissues were either dissected, mounted transversely on corks, and frozen in cooled isopentane or frozen directly in liquid nitrogen. Samples were stored at -80°C until molecular and histological analyses.

The dogs included in this study were housed and cared for according to current European Union regulations. Muscle biopsies were sampled in the context of a project authorized by the ethical committee ComEth ANSES-EnvA-UPEC (national registration number 16) under the approval number 11-01-11/06. Muscle biopsies from GRMD and WT littermates were surgically sampled from the tibialis cranialis muscle under general anesthesia (intravenous propofol induction,
maintenance isoflurane in 100% O₂) with an adapted analgesia (intravenous morphine). The muscle biopsies were immediately snap-frozen in isopentane cooled in liquid nitrogen and subsequently stored at -80°C until cryosectioning.

## Muscle function assessment

An escape test was performed to measure the overall muscular force of mice according to standard procedures. Briefly, mice were placed in a 30-cm-long tube with both ends open, and their tails, which were connected to a force transducer, were gently pinched to induce escape in the opposite direction. The force exerted was then measured by the detector. The procedure was performed around 15 times, and the average of the five highest measured forces was normalized to the mouse's weight.

Forelimb muscle strength was assessed by a grip test using a T-shaped bar connected to a force transducer (Bioseb). Briefly, mice were allowed to grasp the bar with their forelimbs, after which they were gently pulled backward by the tail in a steady and continuous motion until they released their grip. Three consecutive trials were performed per mouse, and the peak force exerted before release was recorded as the maximum grip strength (in grams). Both the absolute maximum grip strength and strength normalized to body weight (g/g) were calculated.

For in vitro mechanical force measurements, extensor digitorum longus muscles were dissected and placed in an oxygenated Tyrode buffer bath (Aurora Scientific System) at room temperature (RT) to maintain physiological conditions. Each muscle was mounted between bipolar electrodes and attached to a force transducer. Tetanic force was elicited by delivering 600-mA stimulations (400 ms, 125 Hz). For twitch force measurements, a single 600-mA pulse (equivalent to one action potential) was applied. Maximal forces recorded were normalized either to the muscle's cross-sectional area (calculated using the optimal muscle length $L_o$) or to the muscle weight.

## Transgene constructs and viral production

The µDys construct has been previously described (44). The µDys coding sequence was inserted into a transcription cassette under the control of the muscle promoter spc5.12, flanked by our optimized sequences of the inverted-terminal repeats (ITRs) (referred to as ITR-GNT), and encapsulated in an AAV9. The γ-sarcoglycan (SGCG) construct has been previously described (45). The sequence of the human copy of SGCG was inserted into a transcription cassette under the control of the desmin muscle promoter, flanked by the WT ITR sequences (ITR-2), and encapsulated in an AAV9 vector.

All rAAVs in our studies were produced in HEK 293-T cells [human embryonic kidney (HEK) 293-T cells] using the triple transfection protocol. For that, HEK 293-T cells were grown in suspension (seeded at 0.25 × 10⁶ cells/ml) under agitation (37°C, 5% CO₂) in Freestyle F17 medium (Life Technologies, ref. A13835-01), supplemented with GlutaMAX-I at 4 nM (Life Technologies, ref. 35050-038), Pluronic F-68 at 0.08% (Life Technologies, ref. 24040-032), and anticlumping agent (Life Technologies, ref. 01-0057AE). Three days after seeding, cells were transfected with three plasmids encoding adenovirus helper proteins (pAd Helper), AAV Rep and Cap proteins, and the transgene expression cassette flanked by ITRs. Three days after transfection, cells were harvested, chemically lysed, treated with benzonase (Millipore), and filtered. Viral capsids were purified by affinity chromatography, formulated in sterile phosphate-buffered saline (PBS), and stored at -80°C.

Titers of rAAV vector stocks were determined using qPCR. Viral DNA was extracted using MagNA Pure 96 DNA and the NA small volume viral kit (Roche Diagnostics) according to the manufacturer's instructions. PCR was performed with Absolute ROX mix (TaqMan, Thermo Fisher Scientific) using ITR-specific primers (list in table S2).

## Cell culture

Immortalized human myoblasts were generated from a healthy donor (C25 cell line) at the Myoline platform (Institute of Myology, Paris). Immortalized myoblasts were grown in skeletal muscle growth medium (Promocell, ref. C-23060), supplemented with a supplement mix (Promocell, ref. C-39365), 15% fetal bovine serum (Gibco, 10270-106), 1% GlutaMAX (Thermo Fisher Scientific, ref. 35050061), and 1% gentamycin (Life Technologies, ref. 15750-037). For myotube generation, myoblasts were grown until confluency, growth medium was then replaced by a differentiation medium (Promocell, ref. C-23061), and cells were differentiated for 6 to 7 days before analysis.

Primary myoblasts were isolated from 6- to 7-week-old WT and *Dmd*<sup>mdx-4Cv</sup> mice euthanized by cervical dislocation. Skeletal muscles from the triceps, the gluteus, and hindlimbs—including TA, GA, and quadriceps—were dissected, rinsed, and transferred into sterile PBS. Under a hood, connective tissue, aponeurosis blood vessels, and nerves were carefully removed. Muscles were minced into small fragments using sterile scissors and digested in 10 ml of enzymatic solution containing collagenase A (5 U/ml; Sigma-Aldrich, 11088793001) and dispase II (60 U/ml; Thermo Fisher Scientific, 11510536) in Dulbecco's modified Eagle's medium (DMEM)/Ham F-12 (1:1) (Life Technologies; DMEM: 31966047; Ham F-12: 31765035). The tissue suspension was incubated at 37°C with periodic mechanical dissociation every 10 min by pipetting sequentially with 10-ml, 5-ml, and lastly, 2-ml pipettes until fragments could easily pass through a 2-ml pipette (typically after 50 to 60 min). A final step included passing the solution through an 18G needle with a 10-ml syringe using spotted ebb and flow. The resulting cell suspension was sequentially filtered through 100-, 70-, and 40-µm cell strainers to remove debris. Each filter was rinsed with 10 ml of complete culture medium: DMEM/Ham F-12 (1:1), 20% fetal bovine serum (Gibco), 2% Ultroser G (Sartorius, 15950-017), and gentamicin (20 µg/ml; Life Technologies, 15750037). After the last stainer step, the cells are centrifuged at 300g for 10 min, and the cell pellet was recovered in complete medium. The cellular solution was subjected to two rounds of preplating (20 min each) to reduce fibroblast contamination, after which the nonadherent cells were seeded onto collagen-coated flasks in complete medium. The culture medium was changed every 2 to 3 days. Cells were passed no more than twice before use in experiments.

For lysosomal injury assays, cells were plated on a clear-bottom microscopy dish (Ibidi, ref. 81158) (~60,000 cells/cm²) or in six-well plates. Cells were then treated with a solution of LLOMe (Santa Cruz, ref. sc-285992) at a final concentration of 2.5 mM in the medium and incubated for the indicated time at 37°C, after which the medium was collected, and the cells were either pelleted and frozen at -80°C for Western blotting or fixed for immunofluorescence assays. Lysotracker Red DND (Invitrogen, L7528) was loaded in the media at a final concentration of 75 nM for 30 min at 37°C before cell fixation and imagination.

For immunofluorescence, cells were briefly washed two times with Hanks' balanced salt solution (HBSS) buffer (calcium and magnesium, Gibco), fixed with a 4% ice-cold paraformaldehyde (PFA)
solution (Thermo Fisher Scientific, ref. 28906) for 15 min, then washed three times with PBS, and permeabilized for 10 min with 0.25% Triton X-100 (Sigma-Aldrich). Immunofluorescence was performed according to the protocol described below.

For LGALS3 measurement in the media, media were collected after LLOMe treatment and frozen at -80°C. An ELISA for human LGALS3 (Invitrogen, kit BMS279-4) was performed according to the manufacturer's instructions.

Dextran pH probing was performed as described before (23). Briefly, myotubes were loaded with a mix of Dextran Tetramethylrhodamine (10,000 MW; Invitrogen, D1817) and Dextran Oregon Green (10,000 MW; Invitrogen, D7171) at a final concentration of 100 µg/ml and incubated for 6 hours at 37°C. Cells were then washed with HBSS and incubated overnight in fresh media (chasing of the dextran into the lysosomes). Later, LLOMe was diluted in the media at a 2.5 mM concentration for 30 min, after which the cells were washed twice with HBSS, fixed in 4% ice-cold PFA, and mounted in DAPI (4',6-diamidino-2-phenylindole)-Fluoromount-G (Southern Biotech, ref. 0100-20).

The DQ-Red BSA assay was performed as previously described (23). Briefly, cells were seeded onto clear-bottom microscopy dishes (Ibidi, ref. 81158) and cultured to 80 to 90% confluence. Cells were then incubated for 4 hours at 37°C in 5% CO₂ with prewarmed trafficking medium consisting of DMEM KnockOut (Gibco, 10829018) supplemented with 1% fetal bovine serum, 1% minimum essential medium nonessential amino acids (Gibco, 11140050), 1% GlutaMAX (Gibco, 11574466), and 1% Hepes buffer. DQ-Red BSA (Molecular Probes, D-12051) was added to the medium at a final concentration of 10 µg/ml. Following incubation, cells were fixed with 4% PFA and subsequently imaged.

## Molecular analysis

### Viral genome copy number quantification

Genomic DNA was extracted from frozen muscle and organ samples or sections. Briefly, samples (frozen muscle and tissue sections) were lysed using the Bead Mill (Fisherbrand) machine with radioimmunoprecipitation assay solution (Thermo Fisher Scientific, ref. 89901) for 40 s (speed, 6 m/s) and then centrifuged for 10 min (2000g, 4°C). DNA was later extracted from the recovered supernatant with the NucleoMag Pathogen extraction kit (Macherey-Nagel, ref. 744210.4) using the KingFisher machine (Thermo Fisher Scientific, ref. 5400640). DNA concentrations were quantified by NanoDrop (NanoDrop 8000) and stored at -20°C. For VCN analysis, multiplex channel reactions were performed with the digital droplet PCR probe system. Extracted DNA (50 ng) was used with the following PCR amplification conditions: initial denaturation at 95°C for 10 min, followed by 40 cycles of denaturation at 94°C for 10 s, annealing at 60°C for 10 s, and transcript extension at 72°C for 1 min and 30 s. Following this, fluorescent dye stabilization was performed at 98°C for 10 min. VCN values were calculated using the ratio with the *Rplp0* gene amplification. For this analysis, multiplex channels were analyzed depending on the type/concentration of probe. Primers and probes that were used are listed in table S2. The analysis was performed using Quantasoft Analysis Pro software.

### RNA isolation and reverse transcription qPCR

Total RNA was extracted from frozen muscle samples or sections. A 400-µl Nucleozol solution (Macherey-Nagel, ref.740404) was added per sample, and the tissues were lysed using the Bead Mill 24 (Fisherbrand) (6 m/s for 40 s). Next, 160 µl of deoxyribonuclease (DNase)/

ribonuclease free water was added, and the tubes were mixed vigorously and incubated at RT for 10 min. Samples were then centrifuged (1000g, 15 min, 20°C), and total RNA was extracted from the supernatant using magnetic beads (Macherey-Nagel, ref. 744503.12) with the Ideal-32 system (ID solutions). RNA samples were then treated with DNase (30 min, 37°C) (TURBO DNA-free kit, Thermo Fisher Scientific), and 1 µg of RNA per sample was used for reverse transcription reaction using the RevertAid H Minus First Strand cDNA synthesis kit (Thermo Fisher Scientific, K1632) according to the manufacturer's instructions. The cDNA samples were then diluted (1:9) and analyzed by qPCR using a LightCycler 480° (Roche), with either SYBR Green PCR assays (Thermo Fisher Scientific, ref.4309155) or TaqMan gene expression assays (Thermo Fisher Scientific). The primer sequences used are shown in table S2. Each reaction was performed in duplicate, and the C~t~ (cycle threshold) values for *Rplp0* were used to normalize gene expression using Livak's secondary derivation method (74).

### Protein isolation and Western blotting

Frozen tissues were cut using a cryostat into 10-µm sections and lysed with radioimmunoprecipitation assay lysis buffer (Thermo Fisher Scientific, ref. 89901) using the Bead Mill 24 (Fisherbrand) for 40 s at 6 m/s. Samples were then centrifuged for 10 min (2000g, 4°C), after which the supernatant was collected and treated with DNase (1:1000, Millipore) and a protease inhibitor cocktail (Roche) for 45 min. For nuclear and cytoplasmic protein isolation, the NE-PER kit (Thermo Fisher Scientific, 78333) was used according to the manufacturer's instructions. The total protein concentration was measured using the Pierce BCA Protein Assay kit (Thermo Fisher Scientific, ref. 23225). Equal amounts of protein (between 20 and 30 µg) were loaded and separated on NuPAGE precast gels (Thermo Fisher Scientific). Proteins were transferred to nitrocellulose membranes using the iBlot 2 Dry Blotting system (Thermo Fisher Scientific), and total protein quantification was performed using the Revert Total Protein Stain kit (LI-COR). Membranes were later incubated with a blocking solution (Odyssey Blocking Buffer, LI-COR) for 1 hour at RT. Primary antibody incubation was done overnight at 4°C (dilutions and details of primary antibodies are presented in table S3). Membranes are then washed three times in a Tween-tris-buffered saline (TTBS) solution and incubated with secondary antibodies (1:10 000, listed in table S5). Membranes are washed again with TTBS and scanned using the Odyssey CLx scanner (LI-COR). Images were analyzed using Image Studio Lite (version 5.2), pixels of selected bands were quantified, and the signal was normalized to either mentioned reference protein or to total protein staining.

### Lipidomic analysis

Muscle sections were weighted and lysed in PBS and then diluted to 5 mg/ml. Shotgun lipidomic analysis was performed by Lipotype (Dresden, Germany). This technique involves automated sample extraction, automated direct sample perfusion, and high-resolution mass spectrometry to capture lipid class-specific internal standards. Briefly, lipid extraction was performed using chloroform and methanol (75). Samples were spiked with lipid class-specific internal standards before extraction. After drying and resuspension in the mass spectrometry acquisition mixture, the lipid extracts were subjected to mass spectrometry analysis. Mass spectra were acquired on a hybrid quadrupole/Orbitrap mass spectrometer equipped with an automated nanoflow electrospray ion source in positive and negative ion modes. LipotypeXplorer software is used for lipid identification on mass spectra. Data processing and analysis are carried out using
the Lipotype information and management system and Lipotype-Zoom web software.

## Quantification of free muscle cholesterol

Muscle sections were weighed (between 10 and 20 mg) and then lysed using a mix of chloroform/isopropanol/NP-40 (7:11:0.1) for 1 min with the Bead Mill 24 (Fisherbrand). Samples were then centrifuged at 15,000g for 10 min, and supernatants were transferred to new tubes, dried at 50°C, and centrifuged under vacuum for 30 min. The pellet was then resuspended in cholesterol quantification kit buffer (Invitrogen, ref. A12216) at a concentration of 20 µg/µl and vortexed vigorously. Free cholesterol quantification was performed in duplicate using the Amplex Red Cholesterol kit (Invitrogen, ref. A12216), following the manufacturer's instructions. Cholesterol concentrations were normalized to tissue weight for each sample.

## RNA sequencing

Essentially as described previously (76), RNA was extracted from muscle sections, and purified RNA samples were first checked for RNA quality using the Bioanalyzer 2100 (Agilent). Samples with an RNA integrity index greater than 8 were then used for RNA sequencing (Genewiz). The Stranded Total RNA Library Prep Kit (Illumina) was used to create sequencing libraries, which were sequenced according to the Illumina protocol on the NovaSeq instrument (Illumina), yielding around 20 million paired-end reads per library. Paired reads were filtered and subjected to quality control using fastp. They were then mapped onto the GRCm38/mm10 genome using HISAT2, and count tables were generated using htseq-count. DEGs were identified using the DESeq2 R package. Pathway analysis was carried out in R-Studio (version 4.0.4) using either overrepresentation methods with Gene Ontology or functional class scoring with GSEA. GSEA was performed using the clusterprofile R package.

## Serum biomarker analyses

Quantification of CK in serum was performed using a FUJI DRI-Chem nx500 analyzer (DMV Imaging). Ten microliters of diluted serum was deposited on FUJI DRI-CHEM CPK-PIII plates (DMV Imaging, ref. 15809463), and CK concentrations were automatically generated in international units per liter of serum (U/liter). Fragments of the myofibrillar structural protein MYOM3 have been identified as being abnormally present in the sera of dystrophic animal models (77). To quantify these fragments in mouse serum, a Meso Scale Discovery (MSD) ELISA was performed. An anti-MYOM3 polyclonal capture antibody (Proteintech Lab, ref. 17692-1-AP) was coated to the wells of a multiarray plate containing electrodes (MSD, ref. L15XA-3) (overnight, 4°C). The plate was then washed three times with a washing solution (PBS + 0.05% Tween 20), and the wells were saturated with a 3% BSA solution (1 hour, RT). After three washes, diluted sera were added (1:50 dilution in a 1% BSA solution), and the plate was incubated for 2 hours at RT under rigorous agitation. A standard range of MYOM3 peptide was also used to calculate concentrations. Revelation was carried out using a mouse monoclonal antibody (Anti-51r20, developed by Proteogenix) coupled to a sulfo-tag (Mesoscale, ref. R31AA-2). The plate was incubated for a further 2 hours under agitation and then washed three times. Last, 150 µl of MSD-Gold reading buffer (MSD, ref. R92TG-2) was added, and the electrochemiluminescence was read using the MESO QuickPlex SQ 120 (MSD). MYOM3 concentrations were calculated using the MYOM3 standard range and expressed in pg/µl.

## Single myofiber isolation and analysis

The two FDB muscles were surgically isolated from euthanized *Dmd<sup>mdx-4Cv</sup>* and C57BL/6J male mice (6 weeks old), rinsed in PBS,

and placed in a digestion solution [collagenase (2.5 U/ml) in DMEM, Sigma-Aldrich, United States, 11088793001] for 1 hour and 30 min at 37°C on a rotated wheel. The collagenase solution was removed by fiber sedimentation (10 min, RT), and the isolated muscle fibers were transferred in a 2-ml tube and fixed with an ice-cold 4% PFA solution for 20 min. Fixed myofibers were then washed three times with PBS (10 min, RT) and permeabilized with 0.5% Triton X-100 (Sigma-Aldrich) for 10 min. Afterward, 10% goat serum (GS) (Agilent, ref. X090710) was used for blocking (30 min, RT) before a second blocking with a "mouse-on-mouse" blocking solution (Invitrogen, ref. R37621). Primary antibodies (listed in table S4) were incubated overnight at 4°C on an inclined agitator. Myofibers were then washed three times with a TTBS solution (10 min, RT) before the incubation with secondary antibodies (listed in table S5, 1:200 dilution). Last, myofibers were washed three times with PBS and mounted on glass slides with a DAPI-Fluoromount-G mounting medium (Southern Biotech, ref. 0100-20).

Stack images were acquired with a LEICA TCS SP8 spectral confocal microscope (Leica, Germany) running LASX software (Leica, Germany) using an HC PL APO CS2 20×, 0.75-numerical aperture (NA) dry objective. A stack image was taken every *z* = 0.3 µm, and 3D myofibers images were reconstructed using Imaris software (version 9.9.1). All myofiber images were treated similarly, and LAMP2 spot analyses were performed using the spot modeling function (average spot diameter, 1 µm).

## Ultrastructural electron microscopy analyses

TA muscle samples were fixed, embedded, sectioned, and analyzed using a transmission electron microscope as described in (78). Lysosomes were observed in longitudinal muscle sections (10 sections per animal) from *n* = 2 WT and *n* = 2 *Dmd<sup>mdx-4Cv</sup>* mice.

## Histological and image analysis

For all histological studies, muscles were dissected, fixed on corks, frozen in isopentane cooled in liquid nitrogen, and then stored at -80°C. Transverse cryosections (8 µm thick) were prepared from frozen muscles, air dried, and stored at -80°C.

## Histochemistry

H&E staining was carried out in accordance with standard procedures. This staining combines nuclear staining with hematoxylin and cytoplasmic staining with eosin. Analysis of myofiber size distribution and centronucleation index was performed as previously described in (79).

PAS staining was performed on muscle cryosections to detect glycogen content. Sections were fixed in Carnoy's solution (60% ethanol, 30% chloroform, and 10% glacial acetic acid) for 10 min, rinsed, and incubated in 0.5% periodic acid for 10 min. After additional rinsing, sections were stained with Schiff reagent for 5 min, washed, dehydrated, and mounted.

Oil Red O staining identifies hydrophobic lipids in the liquid phase. The colored body, called lysochrome, moves from the solvent in which it is found to the tissue lipid, and the lipid droplets are colored red and the nuclei blue.

Sirius Red staining was used to assess tissue fibrosis, combining red staining for collagen fibers with yellow staining for cytoplasm. Briefly, slides were treated with acetone for 1 hour, fixed in formalin for 5 min, and then incubated in Bouin's solution for 10 min. Slides were then rinsed twice with water and stained in a Sirius Red solution (Sigma-Aldrich, ref. 365548) (diluted in 0.1% picric acid) for
1 hour, washed with water and dehydrated with ethanol (70% for 1 min, 95% for 1 min, and 100% for 2 min), and then cleared in a xylene bath (two washes for 1 min). Slides were mounted in mounting medium and air dried.

All bright-field histological labeling images were digitized using an Axioscan Z1 slide scanner (Zeiss, Germany) under a Zeiss Plan-Apochromat 10×/0.45 M27 dry lens (Zeiss, Germany). Tiled scanner images were reconstructed using ZEN software (Zeiss, Germany) (pixel size, 0.45 µm).

QuPath (version 0.4.3) software was used for muscle fibrosis quantification from Sirius Red staining. For each muscle scan, a small artificial neural network was trained to classify positive and negative fibrotic pixels and subsequently used to quantify fibrotic areas.

### Immunofluorescence

**Galectin-3 and all coimmunostainings of LAMP2.** Slides were fixed in ice-cold 4% PFA (Thermo Fisher Scientific, ref. 28906) for 3 min, washed two times in PBS, and then permeabilized in an ice-cold methanol gradient (10 min, 70%; 10 min, 95%; 20 min, 100%). After three washes with PBS, a 10% GS solution (Agilent, ref. X090710) was used for blocking (RT, 30 min). A second "mouse-on-mouse" blocking solution (Invitrogen, ref. R37621) was also used to block endogenous binding to mouse immunoglobulins (RT, 30 min) for all immunostainings with mouse primary antibodies on mouse muscles. Slides were then incubated with primary antibodies at 4°C overnight (diluted in 1% GS; see table S3 for specific dilutions). Next, slides were washed three times in a TTBS solution before incubation with secondary antibodies for 2 hours at 4°C (Molecular Probes Alexa Fluor secondary antibodies were used at 1:200 dilution; see table S5). Last, the slides were washed three times in a PBS solution for 5 min, rinsed in Milli-Q water, and mounted with DAPI-Fluoromount-G mounting medium (Southern Biotech, ref. 0100-20).

**Other immunostainings.** Slides were fixed for 10 min in a 4% methanol-free ice-cold PFA solution (Thermo Fisher Scientific, ref. 28906), permeabilized in 0.1% Triton for 5 min, and blocked for 45 min in a 10% GS solution (Agilent, ref. X090710). For staining using primary mouse antibodies, an additional 30-min blocking was done with a "mouse-on-mouse" solution (Invitrogen, R37621). Next, slides were incubated with the primary antibody solution at 4°C overnight (diluted in 1% GS; references and dilutions are listed in table S4). Slides were then washed three times in PBS and incubated for 2 hours at 4°C with a secondary antibody solution (Molecular Probes Alexa Fluor secondary antibodies were used at 1:200 dilution; listed in table S5). Last, the slides were washed three times in a PBS solution for 5 min, rinsed in Milli-Q water, and mounted with DAPI-Fluoromount-G mounting medium (Southern Biotech, ref. 0100-20) or Fluoromount-G medium (Southern Biotech, ref. 0100-01).

**Detection of free cholesterol by filipin staining.** Free cholesterol was detected using the fluorophore filipin (Sigma-Aldrich, ref. SAE008). Filipin was incubated with the secondary antibody solution at a concentration of 100 µg/ml, along with Syto13 (Invitrogen, S7575) for nuclear staining.

### Image acquisition

Once the slides have been dried, they were imaged with a LEICA TCS SP8 spectral confocal microscope (Leica, Germany) running LASX software (Leica, Germany) using an HC PL APO CS2 20×, 0.75-NA dry objective, 40×, 1.3-NA oil immersion, or 63×, 1.4-NA oil immersion objective. Acquired images were processed uniformly with Fiji ImageJ for final figures.

For dystrophin, IgG, and CD11B staining, images were digitized using the Axioscan Z1 slide scanner (Zeiss, Germany) under a Zeiss

Plan-Apochromat 10×/0.45 M27 dry lens (Zeiss, Germany) and using an ORCA-Flash4.0 complementary metal-oxide semiconductor digital camera (Hamamatsu, Japan). Tile scanner images were reconstructed with ZEN software (Zeiss, Germany) (pixel size, 0.65 µm).

### Image analyses

**Myofiber segmentation.** For images with laminin or wheat germ agglutinin labeling for the membranes, the Cellpose2 cyto2 model (80) was fine-tuned on manually myofiber-labeled images (hyperparameters: n_epochs = 200, learning_rate = 0.05, and weight_decay = 0.0001). The labeled dataset used in fine-tuning was prepared in such a way that the model can simultaneously segment myofibers and ignore low-quality staining areas. Fine-tuned models were then used to extract myofiber masks. Reconstruction of myofiber masks of whole-scan images was done using the cellpose package (80). Reconstructed masks were then converted into regions of interest (ROIs) for subsequent quantification (each ROI corresponds to an individual myofiber) using the Labels>To_Rois.py Fiji plug-in (81). For all images acquired with confocal microscopy, the cellpose2 cyto2 model was fine-tuned on manually labeled images on the basis of the background noise of the A488 channel.

**Puncta quantification.** All images from the same experiment or panel were processed uniformly using Fiji ImageJ. Median filters were applied to reduce noise, and thresholding was performed to select the puncta to be analyzed. For overlay quantification, the image calculator function "AND" was used. Various parameters of puncta were quantified within each individual ROI, where one ROI corresponds to one myofiber.

**Overlay analysis.** ImageJ's JaCOP plug-in was used to analyze colorization by calculating Manders coefficients. These coefficients indicate the fraction of the fluorescence signal from one channel localized in the fluorescence pixels of a second channel.

### Statistical analysis

Statistical analysis was performed using GraphPad Prism 10 (GraphPad Software Inc., La Jolla, CA), R 4.2.2., or Python 3.10. Variances were assumed to be equal; a Shapiro-Wilk test was used to check normality. Student's t test was used to compare the means of two groups. A one-way analysis of variance (ANOVA) test was used to compare more than two groups when values followed a normal distribution, accompanied by Tukey's multiple comparisons test; otherwise, a nonparametric Kruskal-Wallis test with Dunn's multiple comparisons was used. Error bars on graphs represent standard deviation (SD), unless specified otherwise. Results were considered significant when *P* values or adjusted *P* values were less than 0.05.

## Supplementary Materials

The PDF file includes:

Figs. S1 to S13

Tables S1 to S5

Legend for movie S1

Legends for supplementary Excel files S1 to S3

Other Supplementary Material for this manuscript includes the following:

Movie S1

Supplementary Excel Files S1 to S3

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**Acknowledgments:** We are grateful to the "Association Française contre les Myopathies" (AFM) and "Institut National de la Santé Et de la Recherche Médicale" (INSERM) for support. We thank the "Imaging and Cytometry Core Facility" and the "Preclinical Evaluation Platform" at Généthon for technical support. We are members of Genopole, the first French biocluster dedicated to genetics, biotechnology, and biotherapies. We are also grateful to Ile-de-France Region, the Conseil Départemental de l'Essonne (ASTRE), INSERM, and GIP Genopole (Evry) for contributions to equipment acquisition. **Funding:** D.I. and I.R. were funded by INSERM (FranceRelance no. 221030A10). **Author contributions:** Conceptualization: A.J. and D.I. Methodology: A.J. and D.I. Investigation: A.J., L.P., R.B., N.B., E.L., C.R., J.P., S.A., L.V.W., A.M., G.T., N.D., B.D., and E.M. Resources: I.B., S.B., M.T.B., T.E., I.R., and D.I. Data curation: A.J. and A.V.H. Software: A.J., A.V.H., and D.S. Formal analysis: A.J., A.V.H., and D.S. Validation: A.J. Visualization: A.J. Supervision: A.J., I.R., and D.I. Project administration: A.J., I.R., and D.I. Funding acquisition: I.R., and D.I. Writing—original draft: A.J. Writing—review and editing: A.J., T.E., I.R., and D.I. A.J., L.P., R.B., N.B., A.V.H., E.L., C.R., J.P., S.A., D.S., L.V.W., A.M., G.T., N.D., I.B., S.B., M.T.B., B.D., E.M., T.E., I.R., and D.I. discussed the results and approved the manuscript. **Competing interests:** A.J. and D.I. are inventors on patent applications (EP2319441.4 and EP2319461.2) submitted by Généthon and related to the use Galectin-3 as a biomarker of lysosomal damage in muscular dystrophy and TR-based combined therapy for lysosomal damage. I.R. is a cofounder and chief scientific officer of Atamyo Therapeutics. The other authors declare that they have no competing interests. **Data and materials availability:** All data needed to evaluate the conclusions in the paper are present in the paper and/or the Supplementary Materials. The transcriptomic data generated in this study have been deposited in the NCBI BioProject database under accession code PRJNA1281841: www.ncbi.nlm.nih.gov/bioproject/?term=PRJNA1281841.

Submitted 2 January 2025
Accepted 22 September 2025
Published 22 October 2025
10.1126/sciadv.adv6805