Introduction

DMD is a severe, progressive X-linked recessive disease due to mutations in the dystrophin gene (Dmd) that predominantly disrupts skeletal muscle tissue. It encodes the protein dystrophin, which is highly important for structural integrity in muscle fibers by its subsarcolemmal connection between the actin cytoskeleton and the extracellular matrix. The absence of dystrophin leads to repeated muscle damage, poor regenerative capacity, progressive muscle degeneration, and early mortality1. One of the worst-hit tissues in DMD patients is the skeletal muscle, manifested as muscle wasting, weakness, and fibrosis. Animal models, the mdx mouse models, the DMD dog models, the DMD pig models, and the DMD rat models have thus far been highly instrumental in the study of the pathogenesis of DMD and afford insight into the progress of the disease and possible therapeutic intervention2,3,4.

Of them all, the rat models for DMD are beneficial due to their larger size and physiological similarities to human muscle pathology5,6,7. Such models help in realizing the histopathological changes and the molecular disruptions underlying the disease mechanism in DMD. For instance, dystrophin deficiency in the models closely resembles that in human patients, resulting in degeneration, inflammation, and fibrosis of skeletal muscles, just like in humans8. This again points to a better resemblance of rats to many features of DMD in humans, including more severe muscle damage, more evident cardiomyopathy, more progressive muscle degeneration, and more severe deficits in motor responses5,7. The DMD symptoms are not as severe in mdx mice, with good muscle recovery due to the high activation of satellite cells9,10,11.

A study by Teramoto et al. (2020) described a new rat model harboring in-frame dystrophin gene mutations that mimicked the dystrophic phenotype of Becker muscular dystrophy (BMD). Though this study was mainly associated with BMD, it also gave insight into dystrophin expression patterns and skeletal muscle pathology relevant to the pathogenesis of DMD. Thus, in this model, progressive muscular dystrophy due to the functional absence of the complete dystrophin protein resulted in remarkably distinctive gene expression changes as a starting point in exploring disrupted molecular pathways in DMD12,13. In human models, gene expression profiling has also identified vital information on the biological processes and pathways disrupted by the absence of dystrophin, which includes inflammation, calcium signaling, and metabolic pathways14.

Next-generation sequencing (NGS) technologies aim to perform detailed and large-scale analyses, allowing researchers to study the effects of gene mutations on biological processes and molecular pathways15. A key example is RNA-Seq, which has significantly advanced our understanding of DMD gene expression and related pathway disruptions16. Transcriptomic data analyses allow the identification of differentially expressed genes (DEGs) associated with disease progression and pathway enrichments, highlighting changes in critical signaling processes such as inflammatory response, muscle regeneration, and calcium homeostasis1. This molecular insight is essential for establishing therapeutic targets and testing new treatments to restore dystrophin expression, including gene replacement, antisense oligonucleotides, and gene editing strategies, and/or to compensate for dystrophin deficiency by using pharmacological approaches17.

Despite progress in elucidating the pathology of DMD, critical gaps remain in completing the entire network of molecular pathways affected by dystrophin deficiency. Most previous studies used mdx mouse models and mainly concentrated on selected pathways18. The DMDmdx rat model presents the opportunity to bridge this gap with a much more robust system to pursue pathways for therapeutic implications5.

Accordingly, the present study has developed a comprehensive gene expression and pathway enrichment analysis in the DMDmdx rat model. We implement next-generation sequencing capabilities to probe into the molecular landscape of DMD by identifying essential dysregulated genes and pathways at distinct stages of disease progression. Our research represents another step towards filling the gap in understanding the molecular mechanisms of DMD pathology, within the framework of a new DMDmdx rat model of significant translational importance due to its greater similarity to human DMD. This will provide basic information on the transcriptome profile of skeletal muscles (biceps femoris muscle), which will enable the planning of future studies focused on exploring specific genes and signaling pathways identified in this study, as well as the planning of future therapeutic interventions targeting the molecular factors causing DMD, with an emphasis on therapeutic strategies, gene editing technologies, and personalized medicine approaches.

Materials and methods

Animals and sampling

The Sprague-Dawley (Rattus norvegicus) male dystrophin-deficient rats (strain name: DMDmdx) and wild-type (WT) healthy rats were obtained from Nantes Université and the Nantes Vet School-ONIRIS, Nantes, France5. DMDmdx and WT rats were littermates. All experimental protocols were approved by the 2nd Local Ethical Committee for Animal Experiments, number WAW2/128/2022. All experiments were performed in accordance with relevant guidelines and regulations and reported in accordance with ARRIVE guidelines (https://arriveguidelines.org). We enrolled rats aged 13–14 weeks (around 4 months of age) into two groups: DMDmdx (n = 7), exhibiting a phenotype similar to DMD disease, and WT rats (n = 7). The mean weight of the rats was 470.86 ± 38.22 g and 554.00 ± 49.83 g for DMDmdx and WT, respectively. Rats were housed in the animal house of the Institute of Veterinary Medicine, Warsaw University of Life Sciences, Poland, and kept in plastic cages (Tecniplast S. p. A, 21 020, Italy) in a temperature-controlled room (22 °C ± 1 °C) on a 12 h light/dark cycle, with 50% humidity. Animals were fed the Labofeed B maintenance rat chow (Wytwórnia Pasz Morawski, Poland) ad libitum with free access to water. After 12 h of fasting, animals were anesthetized with an intraperitoneal injection of ketamine/xylazine (90/9 mg/kg) and euthanized by exsanguination via cardiac puncture. Euthanasia was completed by anesthetic overdose, in accordance with Directive 2010/63/EU, Annex IV. The blood was collected by heart puncture using the Sarstedt blood collection system with lithium heparin (Sarstedt AG & Co. KG, Nümbrecht, Germany). Samples were immediately centrifuged to obtain plasma. The right and left hind limb skeletal muscle tissue (biceps femoris) was collected. Samples were quickly cleaned of connective tissue, placed in liquid nitrogen, and stored at − 80 °C until analyzed. Samples for histopathological examination were placed in 10% buffered formalin until analyzed.

Histopathological examination

For histopathological examination, biceps femoris muscle samples (n = 7 per genotype) were fixed in 10% buffered formalin, embedded in paraffin wax, cut in 4 μm sections, and stained with hematoxylin and eosin (H&E). Additional slides were stained with Picrosirius red (PSR) stain for collagen and Koss stain for calcium. Histopathological slides were evaluated with light microscopy using the Leica DM 1000 microscope (Leica Microsystems GmbH, Germany) and KoPa Capture Pro 9.0 software (https://ostec.com.cn/; Guangzhou Ostec Electronic Technology Co., Ltd., China). The size of the fibers was determined in slides stained with Picrosirius red by measuring a minimum of 250 myofibers from each rat in the randomly selected fields using 100x magnification and the above software. Myofibers were measured perpendicular to the axis of the muscle fiber – maximal width and minimal width (max diameter and min diameter, respectively), as was previously described19. Minimal Feret’s diameter was not measured, as maximal and minimal diameters perpendicular to the fiber axis adequately captured fiber size variation. The fibrous connective tissue and skeletal muscle tissue areas were calculated using MicroImage 4.0 software (Olympus Optical CO., GMBH, USA). Fifteen pictures from each rat were taken from representative areas, avoiding superficial areas covered with perimysium from 200x magnification and Picrosirius red staining. Two pathologists independently performed the analyses, each evaluating all samples from all rats.

Biochemical analyses

Biochemical analyses (n = 7 per genotype), including aspartate aminotransferase (AST, U/l), alanine aminotransferase (ALT, U/l), alkaline phosphatase (ALP, U/l), glucose (mg/dl), creatinine (mg/dl), urea (mg/dl), total protein (g/l), albumin (g/l), creatine phosphokinase (CPK, U/l), gamma-glutamyltransferase (GGT, U/l), lactate dehydrogenase (LDH, U/l), total bilirubin (mg/dl), amylase (U/l), uric acid (mg/dl), calcium (Ca, mg/dl), phosphorus (P, mg/dl), magnesium (Mg, mg/dl), potassium (K, mmol/l), and sodium (Na, mmol/l), were determined in Department of Veterinary Laboratory and Clinical Diagnostics, Institute of Veterinary Medicine, Warsaw University of Life Sciences, Poland. The analyses were performed using the Miura One automated biochemistry analyzer (I.S.E. S.r.l., Rome, Italy) and commercially available Pointe Scientific reagent kits (Pointe Scientific Inc., Canton, MI, USA) according to the manufacturer’s protocols (n = 7 per genotype). Blood plasma samples were processed under standardized conditions to ensure the reliability and accuracy of the results.

Oxidative status measurements

Biceps femoris muscle samples (n = 7 per genotype) were homogenized in PBS, vortexed for 15 min, and centrifuged at 500×g for 15 min. The supernatants were used for analyses. Total antioxidant status (TAS) and activities of glutathione reductase (GR), glutathione peroxidase (GPx), and superoxide dismutase (SOD) activities were measured in the homogenates using Randox assay kits (TAS: NX2332, GR: GR2368, GPx: RS504/505/506, SOD: SD125) following the manufacturer’s protocols (Randox Laboratories Ltd., UK). Thiobarbituric acid reactive substances (TBARSs) were analyzed per Aguilar Diaz’s method20.

RNA extraction, validation, and sequencing

According to the manufacturer’s protocol, total RNA was extracted from biceps femoris muscle samples (n = 4 per genotype; individuals most representative of the group average were selected) using the Rneasy Mini Kit (Qiagen, USA). RNase-free DNase was implemented to eliminate any DNA contamination. Quantifying the isolated RNA and libraries was performed on Qubit (Thermo Fisher Scientific, USA), and profiles of RNA and libraries were measured on Tape Station D1000 (Agilent Technologies, USA), generating a quality control (QC) report. The VAHTS Universal v10 RNA-seq Library prep kit for Illumina (Vazyme, China) was used. Sequencing was conducted on Illumina’s NovaSeqX Plus platform with a 25B flowcell (Illumina, USA), generating 100 million PE150 reads. Raw reads were obtained in a FASTQ file format. The files have been checked to assess the sequencing quality according to the Q score. RNA isolation, library preparation, and sequencing were performed at Dante Labs (Dante Labs, L’Aquila, Italy).

RNA-seq data validation and processing

Following RNA-Seq sequencing, the next step was quality control, entailing adapter trimming, removing sequences not from the rat genome, and removing low-quality reads and uncalled bases. Since the data obtained came from Illumina, we utilized FastQC to assess the quality of the files. FastQC is freely available under the URL (http://www.bioinformatics.babraham.ac.uk/projects/fastqc/). Following quality control, we evaluated the mapping of the rat genome. Mapping extensive high-throughput sequencing data against a known genome is essential to RNA-Seq data analysis. In the present work, we used an Ensembl genome version’s Gene Transfer Format (GTF) file for the Rattus norvegicus genome as a reference. Paired-end reads were aligned using Hierarchical Indexing for Spliced Transcripts 2 (HISAT2)21, a software optimized for rapid and efficient mapping. Then, we used the featureCounts package to read and quantify RNA sequencing. FeatureCounts summarizes the aligned reads in SAM format and outputs count data by gene in a matrix.

Differentially expressed genes

Differential gene expression analysis plays a vital role in processing RNA-seq data to investigate gene expression variations across distinct DMDmdx and WT experimental conditions in rat muscle samples. Analysis was performed to find upregulated or downregulated genes, followed by normalized RNA-seq quantification data. The analysis used the DESeq2 pipeline in the R programming language22. All plots were done in R, using functions from ggplot2, Volcano, the principal component analysis (PCA), and pheatmap packages for visualization. Several statistical approaches have been used for detecting DEGs, and the selection criterion was based on log2FC between 0.3 and − 0.3 and P < 0.05.

Functional analysis

Further considerations were made regarding the Gene Ontology (GO) and pathway enrichment studies using the DAVID (https://davidbioinformatics.nih.gov/; Sherman et al., 2022) and KEGG (www.kegg.jp/kegg/kegg1.html) databases. This study divided selections into three categories, including gene sets according to biological processes (BP), cellular components (CC), and molecular functions (MF). Enrichment of specific GO terms and signaling pathways for the used gene set showed significance (P < 0.05) by Fisher’s exact test. Only the statistically significant results were taken into consideration. After this, the findings were visualized using an online tool for further analysis and are represented by charts available at http://www.bioinformatics.com.cn/srplot.

Statistical analysis

Histopathological features, blood plasma biochemical parameters, and antioxidative status were compared using the Student’s t-test with independent variance estimation. Variables with significant differences (p < 0.05) are presented in the graphs. Due to the sample size, normality could not be definitively rejected, justifying the use of the t-test. However, most variables exhibited heteroscedasticity, meaning that the variance increased proportionally with the mean. Therefore, the t-test for unequal variances (Welch’s t-test) was used to evaluate them. Statistical analyses were conducted using STATISTICA 13.3.

Results

Histopathological changes

Histopathological examination of biceps femoris muscle samples collected at 4 months of age from the DMDmdx rats revealed massive multifocal myofiber necrosis, multifocal inflammatory infiltrates with a predominance of mononuclear cells, and marked interstitial hyperplasia of fibrous connective tissue (Fig. 1A-B). None of these changes were found in the WT rats. Moreover, regenerative activity was present in the form of satellite cells arranged at the periphery of the myofibers of DMDmdx rats (Fig. 1C). Marked variation in myofiber size due to large hypertrophic fibers was also noted (Fig. 1D), all compared to normal WT skeletal muscle (Fig. 1E-F). Calcification wasn’t found in either WT or DMDmdx rat biceps femoris samples.

Fig. 1
Fig. 1The alternative text for this image may have been generated using AI.
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Histopathological features in the biceps femoris muscle of DMDmdx and WT rats: A) multifocal necrosis of myofibers (asterisk) and inflammatory infiltrates with a predominance of mononuclear cells (arrows), DMDmdx, H&E stain, 400x; B) satellite cells at the periphery of myofibers (arrows) in DMDmdx rat, indicating myofiber regeneration, H&E stain, 400x; C) marked variation in myofiber size, DMDmdx, H&E stain, 400x; D) interstitial hyperplasia of fibrous connective tissue (arrows) in DMDmdx, PSR stain, 100x; E) normal muscle fibers of WT, H&E stain, 100x, F) normal wild-type muscle of WT, PSR stain, 100x. DMDmdx – Sprague-Dawley DMDmdx rats; WT – Sprague-Dawley wild-type rats; H&E – hematoxylin and eosin; PSR – Picrosirius red.

The biceps femoris muscle fibers diameter analysis in both rat groups showed a statistically significant difference in the max diameter mean value in the DMDmdx group (58.20 ± 4.95 μm) compared to the WT group (64.34 ± 3.48 μm) (p = 0.022). Similarly, the minimum diameter mean value is statistically different, with 25.15 ± 1.72 μm and 36.74 ± 4.54 μm (p = 0.000), for DMDmdx and WT, respectively (Fig. 2A).

Surface area analysis of biceps femoris showed statistically significant differences between connective tissue surface area in DMDmdx versus WT rats, which were 44.29% and 3.97% (p = 0.000), respectively. A similar analysis for muscle tissue in the same groups revealed 55.71% for DMDmdx rat skeletal muscle tissue average area and 96.03% in WT rats (p = 0.000) (Fig. 2B-D).

Fig. 2
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Biceps femoris muscle fiber diameter (µm) (A) and the average fibrous connective tissue and muscle fiber area in WT and DMDmdx rats (percentage) (B). DMDmdx muscle structure, PSR stain, 40x (C) and WT muscle structure, PSR stain, 40x (D). DMDmdx – Sprague-Dawley DMDmdx rats; WT – Sprague-Dawley wild-type rats; PSR – Picrosirius red. The dots represent the mean; the boxes – the standard error; the whiskers – the standard deviation. All comparisons between DMDmdx and WT rats are statistically significant (p < 0.05; n = 7 per genotype).

Blood plasma biochemical parameters

The biochemical analysis of blood plasma showed statistically significantly lower activity in DMDmdx compared to WT rats at 4 months of age in the following enzymes, respectively: mean CPK activity 2864.46 U/l and 1008.83 U/l (p = 0.000); mean LDH activity 1080.43 U/l and 475.03 U/l (p = 0.000); mean ALT activity 102.06 U/l and 45.40 U/l (p = 0.004) (Fig. 3). In addition, a difference in amylase activity was also demonstrated at 578.76 U/l and 948.87 U/l, showing higher activity in WT rats (p = 0.000). The analysis also revealed the electrolytes and minerals level variations between both examined groups: Ca (8.97 mg/l and 9.87 mg/l; p = 0.002) and Na (132.06 mmol/l and 135.76 mmol/l; p = 0.007) levels were higher in WT than in DMDmdx rats. Conversely, P (11.89 mg/l and 6.71 mg/l; p = 0.000), Mg (2.64 mg/l and 2.33 mg/l; p = 0.009), and K (5.77 mmol/l and 4.89 mmol/l; p = 0.003) levels were lower in WT than in DMDmdx rats. Moreover, significant differences in urea and glucose levels were also noticed. Higher levels of glucose (143.43 mg/dl and 197.94 mg/dl; p = 0.000) and lower levels of urea (40.04 mg/dl and 31.84 mg/dl; p = 0.002) were present in WT blood plasma than in DMDmdx rats (Fig. 3). Trends of higher total bilirubin (p = 0.086) and uric acid (p = 0.086) in DMDmdx rats were noticed. A trend of higher AST (p = 0.081) in WT rats was noticed. The remaining parameters, ALP (p = 0.911), creatinine (p = 0.163), GGT (p = 0.591), total protein (p = 0.247), and albumin (p = 0.638), showed no differences between DMDmdx and WT rats.

Fig. 3
Fig. 3The alternative text for this image may have been generated using AI.
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The biochemical analysis of blood plasma. CPK – creatine phosphokinase; LDH – lactate dehydrogenase; ALT – alanine aminotransferase; DMDmdx – Sprague-Dawley DMDmdx rats; WT – Sprague-Dawley wild-type rats. The dots represent the mean; the boxes – the standard error; the whiskers – the standard deviation. All comparisons between DMDmdx and WT rats are statistically significant (p < 0.05; n = 7 per genotype).

Antioxidative status

The analysis of antioxidative status parameters in the biceps femoris muscle homogenate showed significant differences between DMDmdx and WT rats at 4 months of age. The mean TAS levels were 2.05 µmol/l in DMDmdx rats and 3.54 µmol/l in WT rats (p = 0.003) (Fig. 4), indicating a marked reduction in total antioxidant capacity in the dystrophic group. Similarly, GSH reductase activity was significantly lower in DMDmdx rats, with mean values of 2.44 U/l compared to 5.75 U/l in WT rats (p = 0.000). GSH peroxidase activity also showed a significant decrease in DMDmdx rats, with values of 29.51 U/l vs. 52.37 U/l in WT rats (p = 0.000). Additionally, TBARS levels, a marker of lipid peroxidation, were significantly reduced in DMDmdx rats, with mean values of 3.25 nmol/l compared to 5.57 nmol/l in WT rats (0.001). Conversely, the activity of SOD did not differ significantly between the groups, with mean values of 0.61 U/l in DMDmdx rats and 0.57 U/l in WT rats (p = 0.144) (Fig. 4).

Fig. 4
Fig. 4The alternative text for this image may have been generated using AI.
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The antioxidative status parameters. TAS – total antioxidant status; GSH – glutathione; TBARS – thiobarbituric acid reactive substances; SOD – superoxide dismutase; DMDmdx – Sprague-Dawley DMDmdx rats; WT – Sprague-Dawley wild-type rats. The dots represent the mean; the boxes – the standard error; the whiskers – the standard deviation. All comparisons between DMDmdx and WT rats are statistically significant (p < 0.05; n = 7 per genotype).

RNA-sequencing data processing

After removing low-quality reads from raw RNA-Seq data of the transcriptome in the biceps femoris muscle of DMDmdx and WT rats, the remaining reads were matched with the reference genome using HISAT2. Later, > 90–95% mapping rate of the reads to our dataset was obtained. Among those clean reads, more than 80% showed unique mapping in various regions of the rat genome. For further analysis, the non-mapped reads and the reads mapped to more than one position were discarded (Table S1).

Analysis of differential gene expression

According to the DESeq2 pipeline statistical analysis results, the general distribution of DEGs was significantly skewed, with an adjusted p-value less than 0.05. Log2FC greater than 0.3 was the cutoff for fold change, while the tight threshold for the p-value was less than 0.05. The PCA explains 56.2% of the total variance (PC1) and distinguishes DMDmdx and WT groups, while PC2 accounts for 13.9%, highlighting within-group variation (Fig. S1). According to PCA analysis, the clear separation of the DMDmdx and WT groups indicates distinct transcriptional profiles associated with the loss of dystrophin.

The differential gene expression analysis between DMDmdx and WT rat groups revealed 3,615 DEGs. Among them, 1,824 genes were upregulated, while 1,791 were downregulated in the DMDmdx rats compared to the WT group (Fig. 5A; Table S2-S3). The top 10 significantly upregulated and downregulated genes are shown in Fig. 5B, ranked by log2 fold change and p-value.

Fig. 5
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The distribution of DEGs between the DMDmdx and WT rat biceps femoris muscle samples: A) The volcano plot with gene distribution. The log2 fold change (log2FC) is shown on the x-axis, while the -log10 (p-value) is shown on the y-axis. The genes on the left side are downregulated in DMDmdx rats compared to the WT, while genes on the right are significantly upregulated. The vertical dashed lines represent a threshold of differential expression standard value log2FC > |0.3|. The horizontal lines represent the threshold for statistical significance (P < 0.05; n = 4 per genotype). B) Top 10 upregulated and downregulated genes in DMDmdx rats compared to WT. Ranking according to the log2 fold change (log2FC). Significantly upregulated genes are colored green, while significantly downregulated genes are colored red. Each bar displays the level of expression change and the p-value. DMDmdx – Sprague-Dawley DMDmdx rats; WT – Sprague-Dawley wild-type rats; DEGs – differentially expressed genes.

Gene ontology analysis

Gene Ontology enrichment analysis showed that significant terms have emerged in BP, CC, and MF, pointing to significant disturbances in the dystrophic muscle tissue of DMDmdx rats. In BP, the following were the most enriched terms: actin cytoskeleton organization (GO:0030036), extracellular matrix organization (GO:0030198), and actin filament organization (GO:0007015). CC enrichment analysis showed significant GO terms in the cytoskeleton (GO:0005856), extracellular matrix (GO:0031012), and actin cytoskeleton (GO:0015629). Under MF, the enriched terms included ATP binding (GO:0005524), calcium ion binding (GO:0005509), and actin-binding (GO:0003779) (Fig. 6; Table S4).

Fig. 6
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Top 15 enriched Gene Ontology (GO) terms categorized into Biological Processes (BP, red), Cellular Components (CC, blue), and Molecular Functions (MF, green), identified using DEGs in DMDmdx vs. WT rat biceps femoris muscle samples. DMDmdx – Sprague-Dawley DMDmdx rats; WT – Sprague-Dawley wild-type rats; DEGs – differentially expressed genes.

Pathway analysis

Pathway analysis was performed based on two databases: the Kyoto Encyclopedia of Genes and Genomes (KEGG) and Reactome. Both provided complementary information about which pathways are dysregulated in the DMDmdx rat model (when compared to WT rats). KEGG pathway analysis emphasized specific, more essential pathways based on the number of genes. Among the top enriched pathways were the cytoskeleton in muscle cells, MAPK signaling pathway, focal adhesion, regulation of actin cytoskeleton, and the calcium signaling pathway. These pathways will be relevant in maintaining muscle structure, cellular signaling, and contraction. (Fig. 7A; Table S5). The Reactome pathway analysis highlighted immune-related and structural pathways. The top pathways include those related to the immune system, innate immune system, extracellular matrix organization, and muscle contraction (Fig. 7B; Table S6).

Fig. 7
Fig. 7The alternative text for this image may have been generated using AI.
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Pathway analysis of identified DEGs in DMDmdx vs. WT rat biceps femoris muscle samples using A) KEGG and B) Reactome. KEGG pathways are sorted by the number of genes. The left column lists the target pathways, while the right column categorizes them according to their assigned KEGG classifications. Reactome pathways are sorted based on the p-value. DMDmdx – Sprague-Dawley DMDmdx rats; WT – Sprague-Dawley wild-type rats; KEGG – Kyoto Encyclopedia of Genes and Genomes DEGs – differentially expressed genes.

Discussion

Duchenne muscular dystrophy is a severe genetic disorder characterized by progressive muscle degeneration and weakness, resulting from mutations in the Dmd gene that encodes dystrophin, a key protein in maintaining the integrity of muscle cells23. This research work employed the DMDmdx rat model to investigate the molecular changes characteristic of the pathology of DMD in the biceps femoris muscle. Animals were analyzed at 4 months of age when pathology was known to be established, although not as severe as at later time points5. This age was, therefore, relevant for analyzing some pathways that future therapeutics could target.

Histopathological and biochemical assessment

The histopathological picture of the biceps femoris muscle samples was typical for muscular dystrophy, containing extensive myofiber necrosis, inflammatory infiltration dominated by mononuclear cells, and significant interstitial fibrosis in the DMDmdx group, which was not observed in WT rats (Fig. 1). Still, it was similar to the observations of Larcher et al. (2014)5 made on the same animal model, underlining the pathological severity associated with dystrophin deficiency. In biceps femoris of DMDmdx rats, sprouting myoblasts (satellite cells) surround the margins of myofibers, indicating that the regenerative process is still active but insufficient (Fig. 1). Quantitative analysis verified significant differences in muscle morphology of DMDmdx rats compared to WT, including decreased myofiber diameter (Fig. 2A) and increased fibrous connective tissue area (Fig. 2B-D), characteristic of DMD progression24,25. These structural changes lead to the failure of muscle function, as seen in DMD26,27. This study’s biochemical and antioxidative status data provide essential information regarding DMD’s systemic and muscular effects as modeled in Sprague-Dawley DMDmdx rats. Such high levels of muscle-specific enzymes (CPK, LDH, ALT) in DMDmdx rats (Fig. 3) correlate well with the muscle damage and membrane instability characteristic of DMD28 due to the absence of dystrophin29. Moreover, lower amylase activity in DMDmdx rats indicates potential differences in metabolic or pancreatic function that require further exploration.

Several electrolyte disturbances were detected in the DMDmdx rats, including a decrease in calcium and sodium and an increase in phosphorus, magnesium, and potassium (Fig. 3), similar to those observed in other cases of muscular dystrophy and possibly associated with dysregulated muscle homeostasis, including dysregulation of ion channels and muscle degeneration24,29,30. Above and beyond the local intra-/inter-tissue interactions that might occur, global metabolic changes reflect the dystrophic background by producing systemic alterations such as hyperglycemia and lower levels of urea measured in the blood plasma of DMDmdx rats (Fig. 3). These metabolic dysregulations are known to occur in muscle diseases. They may also be related to the alteration of the normal systemic response to exercise or stimulus observed in muscle diseases31.

The antioxidative status analysis reveals a significant oxidative stress burden in DMDmdx rats. Decreased TAS, GSH reductase, and GSH peroxidase activity indicate impaired antioxidant defenses, while elevated TBARS levels (Fig. 4) highlight increased lipid peroxidation, a hallmark of oxidative damage in dystrophic muscle. These findings align with the oxidative stress hypothesis of DMD progression, where excess reactive oxygen species (ROS) amplify muscle damage and inflammation32,33,34. Interestingly, the absence of significant differences in SOD activity between DMDmdx and WT rats suggests selective impacts on specific antioxidant pathways (Fig. 4).

The analysis of 4-month-old DMDmdx rats captures an intermediate stage of DMD pathology, where muscle degeneration, inflammation, and fibrosis are established but not as severe as in older rats5. Our findings likely reflect early-to-mid-stage molecular and histopathological changes compared to aged DMDmdx rats (e.g., 12 months), which exhibit more pronounced fibrosis and muscle loss6. This age was chosen to identify pathways amenable to therapeutic intervention before irreversible damage occurs. Future studies exploring specific signaling pathways with expression changes identified in this study, comparing younger and older DMDmdx rats, will help clarify how these molecular profiles evolve with disease progression. The analyses presented above, together with the earlier study by Larcher et al. (2014)5, conducted using the same DMDmdx rat model, provide valuable insight into the pathophysiology of DMD and form the basis for the primary analysis presented in this study, in which we focused on detailed gene expression profiling using RNA-seq to characterize the skeletal muscle transcriptomic landscape as a foundation for further research involving the DMDmdx model. Differences in histopathological, biochemical, and antioxidative status parameters can provide important context for interpreting molecular mechanisms and identifying new therapeutic targets in DMD.

Gene expression analysis

In our NGS analysis, we identified 3,615 differentially expressed genes (DEGs), with significant enrichment in pathways related to muscle contraction, actin cytoskeleton, and extracellular matrix (ECM) organization (Fig. 5). Identified biological processes are critical for maintaining muscle fiber structure and function35. Their dysregulation reflects core pathological features of DMD, including fiber degeneration and impaired repair36,37. Changes in cytoskeletal components and ECM-related genes underscore the loss of cellular integrity in dystrophic muscle38,39. Moreover, Disruption of actin dynamics contributes to sarcolemmal instability40, while alterations in ATP and calcium-binding functions impair contraction and energy metabolism41,42. Reduced expression of actin-binding proteins further destabilizes muscle structure and regeneration43,44 (Fig. 6). The pathway enrichment performed in our study via KEGG and Reactome showed complementarity: KEGG emphasized cytoskeletal regulation and MAPK signaling, while Reactome highlighted inflammation and ECM remodeling. These findings align with hallmark DMD pathology (Fig. 7)45,46,47,48,49. Moreover, we observed downregulation of key structural genes: Dmd (–1.2), Actc1 (–2.7), Actn1 (–1.0), Col6a1 (–1.7), and Mmp2 (–1.4), indicating impaired contraction, ECM integrity, and cytoskeleton function14,50,51. From a therapeutic perspective, modulating these genes or pathways, for instance by enhancing Mmp2-mediated muscle repair, may aid in limiting muscle damage. Reduced expression of Actb (–0.8), Actg1 (–0.9), and Flna (–0.8) suggests disrupted myogenic differentiation and enhanced fibrosis, as also supported by decreased Col5a1 (-1.39) and Col6a150. Notably, Myh3 (–4.4), Myh8 (–4.3), and Actc1 (-2.7) downregulation further reflect impaired regeneration, compounded by the absence of dystrophin, resulting in muscle weakness, fragility, and susceptibility to necrosis.

Muscle degeneration and regeneration

DMD involves progressive muscle degeneration due to dystrophin deficiency, impairing regeneration. Chronic inflammation and fibrosis further hinder muscle repair, leading to functional decline52. In our study, on the molecular level, the genes Myh9 (-0.8), Myh3, and Actn1 are downregulated, which causes decreased contractility of muscle fibers. These genes are important in providing the mechanical contractions essential in skeletal muscle twitch, and their loss results in poor muscle function. Russell et al. (2023), who investigated the role of myosin inhibition in protecting skeletal muscles in dystrophic mdx mice, found that reducing muscle contraction through selective myosin inhibition protected against muscle stress injury without compromising strength. This suggests modulating myosin activity could be a promising therapeutic approach for improving muscle function in DMD patients53.

The downregulation of Igf1 (-1.3) is significant for muscle regeneration, as it is crucial for satellite cell activation. Research indicates that lower levels of Igf-1 impair satellite cell proliferation and differentiation, which are essential for muscle repair following injury54,55. Conversely, the increased expression of cardiac muscle genes such as Tbx5 (3.03) and Myh6 (1.89) indicates a compensatory response to muscle degeneration. This response, however, is insufficient to counterbalance the loss of skeletal muscle-specific contractile proteins, highlighting a maladaptive shift toward cardiac muscle gene expression in the context of skeletal muscle injury56. Interfering with the signaling pathways that regulate the Igf1 gene could enhance satellite cell activation and muscle formation57.

It is also important to mention that several critical pathways are affected in DMD, such as PI3K/AKT signaling, calcium homeostasis, and cytoskeleton remodeling pathways. In our study, some genes belonging to the abovementioned pathways, including Akt3 (-0.7), Egfr (-0.9), and Rock1 (-1.02), were downregulated. These findings further support the idea of their role in disrupting muscle regeneration and enhancing fibrosis. Specifically, interfering with such molecular pathways, including the PI3K/AKT signaling pathway, which is crucial for muscle regeneration, may create a management approach to the degenerative progression of DMD58. Preventing fibrosis-related pathways, perhaps Rock1 and Pdgfrα (-1.63), may also decrease fibrotic tissue, replacing functional muscles in DMD patients59.

Fibrosis and ECM remodeling

In DMD, fibrosis results from excessive ECM remodeling, driven by chronic inflammation and muscle degeneration, leading to tissue stiffness and impaired muscle function60. Fibrosis, characterized by the deposition of ECM proteins, is another key pathophysiological feature in mdx mice61. In our study, several collagen genes, such as Col1a1 (-2.09), Col1a2 (-2.03), and Col6a1 (-1.07), were found to be downregulated in DMDmdx rats. These findings align with prior research and indicate a reduction in collagen synthesis as fibrosis advances61. However, the early stages of the disease are marked by extensive ECM production, resulting in fibrotic scarring. Interestingly, certain MMPs, such as Mmp2, are downregulated, reflecting decreased ECM turnover. Conversely, Mmp10 is upregulated, highlighting the intricate nature of the remodeling process62.

Fibrosis significantly contributes to the progression and worsening of Duchenne Muscular Dystrophy. Tgfβ1 gene signaling (-1.3) modifies muscle atrophy and non-functional regeneration in DMD. The study by Mázala et al. (2020) aimed to understand Tgfβ-mediated muscle degeneration and the effects of early stages of regenerative failure under the D2-mdx mice model. Tgfβ signaling suppression was demonstrated to prevent muscle wasting by preventing the build-up of fibro-adipogenic progenitor (FAP) in a model study. At the same time, it failed to restore normal myogenesis63. Serrano, Rodriguez, and Muñoz-Cánoves (2017) have discussed this in detail regarding fibrosis development in muscles. According to their review, treatments targeting Tgfβ and other pathways might reduce the extent of fibrosis and positively impact muscle function64.

Oxidative stress

Chronic oxidative stress defines DMD because it substantially drives muscle deterioration and inflammatory processes. Anatomical defects from dystrophin elimination impair muscle cell stability while raising mechanical vulnerability and calcium overload, damaging mitochondrial functions, and generating high ROS levels33. Oxidative stress directly impacts proteins, lipids, and DNA, accelerating muscle degeneration and fibrosis progression and activating inflammatory pathways65,66. In DMD, pathological processes accelerate as antioxidant defenses struggle against oxidative stress, leading to further muscle damage and necrosis34. Research indicates that targeting oxidative stress alongside its related pathways presents the potential to manage DMD67.

The muscle cell generates ROS mainly from mitochondria, while mitochondrial dysfunction substantially leads to oxidative stress that affects DMD patients68. In our research, Actn1 (-1.00) and Myh3 (-4.40) genes exhibit decreased expression levels, which may be the consequence of impaired muscle contractions and distorted energy metabolism properties associated directly with mitochondrial dysfunction. The observed decrease in Akt3 (-0.71) expression levels may suggest that the essential PI3K/AKT pathway cannot activate properly to promote cell survival and oxidative stress responses69. Moreover, an elevated expression level of Keap1 (0.54) can block the signaling of NRF2, the antioxidant system master regulator. In DMD, increased ROS damage occurs due to the cells’ inability to counteract oxidative stress, as the body’s antioxidant system fails to function correctly. The diminished expression of Tgfβ1 (-1.30) identified in our study may interfere with redox balance and inflammatory regulation, causing disturbances in antioxidant responses33. Research indicates that decreased Tgfβr2 (-0.90) receptor levels also limit antioxidant protection against oxidative stress70.

The development of fibrosis in DMD is closely connected to oxidative stress and ECM modifications. Muscle cells can become vulnerable to oxidative damage because Col6a1 (-1.70) and Col5a1 (-1.39) show decreased expression, leading to ECM breakdown in these muscles71. Reduced fibronectin expression, Fn1( -1.13), impairs ECM remodeling, weakens cellular defense against oxidative stress, and contributes to ECM deterioration in dystrophic muscles72. The lower levels of Mmp2 (-1.40) enhance the protection of tissues in the short term but also stimulate fibrotic changes, leading to more oxidative damage73. In addition, DMD leads to prolonged inflammation, which generates ROS primarily through the entry of immune cells and cytokine production. The elevated expression of Tnfrsf19 (3.24) reflects amplified TNF signaling that fuels inflammatory and oxidative damage74. The Dkk1 (4.48) elevates its levels and turns off Wnt pathway operation, leading to increased oxidative stress and disturbances in muscle regeneration75. The decreased expression of Coro1a (-0.84) indicates a weakened immune cell function and promotes chronic inflammation76.

Elevated intracellular calcium plays a central role in DMD pathogenesis because it causes oxidative damage through protease activation and inflammatory pathways77,78. The identified in our study, Atp2b2 (1.91), together with Atp1a2 upregulation, point to a potential compensatory response against calcium overload, but it proves inadequate to stop oxidative damage79. The decreased expression of Pdgfrα (-1.63) in DMD indicates dysfunctional mechanisms for responding to oxidative stress, which can worsen muscle destruction59. The damage pathway of DMD shows complexity because of abnormal calcium and oxidative stress regulation systems.

Inflammation and immune response

In DMD, chronic inflammation exacerbates muscle degeneration, triggering an immune response that further damages tissue, creating a cycle of inflammation and immune-mediated injury80. When inflammation becomes chronic, macrophages and other immune cells sustain inflammation, leading to increased fibrosis. Juban et al. (2018) identified that pro-inflammatory macrophages correlate with fibrosis in DMD. They found that Ly6C-positive macrophages had pro-fibrotic activity through the maintenance of fibroblast collagen I synthesis by integrating high latent Tgfβ1 (-1.3) production related to Ltbp4 expression. However, it was delineated that AMPK activation inhibited the Ltbp4 gene, lowering the fibrosis rate and enhancing the muscle force in the dystrophic mice81. It has been proposed that the pathophysiology of muscular dystrophy involves both regenerative and inflammatory mechanisms82,83. As the disease advances, immune homeostasis is impaired by the downregulation of Lgals3 (-2.3)84 and Cd44 (-1.8)85, related to immune cell recruitment83. Also, Nos2 (1.45) in oxidative stress is increased while the inflammation is still active.

In summary, we constructed a gene interaction network using the STRING plugin in Cytoscape to integrate the key processes implicated in DMD progression. This network highlights significant relationships among the DEGs identified in DMDmdx rats, specifically those associated with the major pathological pathways discussed above. The associated GO terms exhibited high connectivity within the network, underscoring their central role in DMD pathogenesis (Fig. 8).

Fig. 8
Fig. 8The alternative text for this image may have been generated using AI.
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Relevance network of identified DEGs. Nodes in the network represent genes, while the expression status is indicated through color coding, where downregulated genes appear red and upregulated genes show green. Blue squares identify DMD-related biological processes.

Comparison with the m dx mouse model

In the mdx mouse model, a critical animal model for DMD, the primary gene implicated is the Dmd gene itself, characterized by a spontaneous mutation that disrupts dystrophin transcription and translation. Transcriptomic analyses of `mdx` muscle have identified several other genes exhibiting significant expression changes relevant to the disease phenotype. Notably, Myh3 and Myh8, regeneration-linked isoforms of myosin heavy chain genes, show marked alterations. Furthermore, other myosin heavy chain genes, including Myh2, Myh4, and Myh7, alongside the SERCA Ca(2+)-ATPase gene (Atp2a1), demonstrate differential expression patterns contributing to the understanding of muscle pathology in this model86. Another study shows that key pathways include cytoskeleton and ECM organization, mechanotransduction, and Ras, Rho, and Wnt signaling. Genes like Sspn, Rho family GTPases, and “collagen” genes are involved in compensatory remodeling and maintaining muscle integrity87. However, the DMDmdx rat model exhibited more DEGs (3,615 vs. approximately 2,000–3,000 in mdx mice), likely reflecting the more severe muscle pathology observed in rats, as previously reported by Larcher et al.5. Notably, the downregulation of Mmp2 in our study contrasts with its upregulation in some mdx mouse studies, suggesting species-specific differences in ECM remodeling. Previous studies, as well as our own results, highlight the DMDmdx rat model as a complementary model with potentially greater translational relevance due to its closer mimicry of human DMD pathology. Further studies are needed, including a meta-analysis of RNA-seq data from humans, rats, and mice, which should allow for the identification of key differences relevant to selecting the most appropriate experimental model for human DMD.

Advantages of the DMD mdx rat model and alignment with human DMD

The DMDmdx rat model offers significant advantages over the mdx mouse model due to its larger body size, more severe muscle pathology, and closer resemblance to human DMD, including advanced fibrosis and dilated cardiomyopathy5,6. This model more accurately reflects the gradual loss of muscle function and the multisystemic nature of the disease, which in humans affects not only the muscular system but also the cardiovascular and respiratory systems. The larger size of the animals enables the application of more precise experimental and functional procedures, such as muscle strength measurements, biopsies, local injections, and cardiac assessments using echocardiography. Our RNA-seq data align with human DMD studies14, showing dysregulation in muscle contraction (e.g., Actc1, Myh3), ECM organization (e.g., Col6a1), and inflammation (e.g., Nos2), consistent with human muscle biopsy transcriptomes. Moreover, in the rat model, we also observe elevated oxidative stress and gene alterations related to calcium homeostasis and cytoskeletal remodeling, which are likewise reported in human DMD. Unlike mdx mice, which exhibit strong muscle regeneration9, partly due to a large pool of active satellite cells and adequate compensation by utrophin, the limited regenerative potential of DMDmdx rats more accurately reflects the chronic and progressive nature of the disease in humans. Rats do not show the same level of compensatory utrophin expression, resulting in a more pronounced dystrophic phenotype, including persistent muscle damage, increased fibrosis, and reduced tissue regeneration capacity. This makes the DMDmdx rat model significantly more suitable for translational studies, particularly for evaluating the efficacy of therapeutic interventions at early and intermediate stages of the disease (Table 1). However, despite its advantages, the DMDmdx rat model also has certain limitations. These include higher maintenance costs, fewer standardized protocols compared to the well-characterized mdx mouse model, and, as with other models, physiological differences that may affect the translational relevance of some findings.

Table 1 Core and contextual genes dysregulated in the DMDmdx rat model.
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It should be noted, as a study limitation, that the biceps femoris muscle samples used for RNA-seq likely contained a heterogeneous cell population, including myofibers, satellite cells, immune cells (e.g., macrophages), and fibroblasts, particularly in DMDmdx rats, due to inflammation and fibrosis. This cellular diversity may influence the transcriptomic profile, as upregulated immune-related genes (e.g., Nos2, Tnfrsf19) likely reflect immune cell infiltration, while downregulated muscle-specific genes (e.g., Actc1, Myh3) primarily originate from myofibers. Future single-cell RNA-seq studies could help dissect cell-type-specific contributions and provide a more precise interpretation of these molecular changes. Furthermore, as the analysis was performed on only one muscle type, future studies should include additional muscles, such as the diaphragm, a tissue severely affected in DMD and functionally critical in later stages of the disease, to more comprehensively characterize histopathological and molecular changes in the DMDmdx rat model. Our transcriptomic profiling may facilitate this direction of research.

Conclusion

Duchenne muscular dystrophy is a severe X-linked genetic disorder caused by mutations in the Dmd gene, leading to progressive loss of muscular strength. In this study, we further characterize the DMDmdx rat model and have demonstrated that biochemical alterations, such as elevated ALT, LDH, CPK, and other markers, are closely associated with muscle damage and inflammation. Histopathological evaluation of biceps femoris muscle tissue reveals significant fiber attenuation, fibrosis, and mononuclear cell infiltration. Concurrently, quantitative gene analysis indicated the downregulation of genes involved in muscle contraction, ECM structure, and cytoskeletal organization, all contributing to the characteristic muscle phenotype in DMD. Pathways such as actin cytoskeleton organization, calcium signaling, PI3K/AKT, and MAPK were reported to be both enriched and impaired, promoting muscle atrophy and fibrosis. Therapeutic strategies for DMD include antisense therapies restoring dystrophin function, modulation of fibrosis-related genes such as Col6a1 and Mmp2, and targeting signaling pathways like PI3K/AKT to enhance muscle regeneration and function. Additionally, anti-inflammatory approaches, including Nos2 inhibitors and promoting the transition of macrophages from pro-inflammatory (M1) to pro-regenerative (M2) states, hold promise for mitigating chronic inflammation and improving disease outcomes. The studies described above provide a comprehensive transcriptomic characterization of the skeletal muscle of DMDmdx rats, a model for Duchenne muscular dystrophy, whose results may support and guide the development of multifaceted therapeutic approaches targeting the molecular and cellular basis of DMD.