Histological Analysis of Tibialis Anterior Muscle of DMD mdx4Cv Mice from 1 to 24 Months

Abstract

Background:

The mdx-C57/B6 mouse model does not show the clinical signs of Duchenne muscular dystrophy (DMD), although muscles exhibit hallmarks of permanent regeneration and alterations in muscle function. The DMD ^mdx ^4Cv strain exhibits very few revertant dystrophin positive myofibers, making that model suitable for studies on gene and cell therapies.

Objective:

The study appraises the histological evolution of the Tibialis Anterior muscle of WT and DMD ^mdx^4Cv mutant from 1 to 24 months.

Methods:

Histological analysis included a series of immunostainings of muscle sections for assessing tissue features (fibrosis, lipid deposition, necrosis) and cellular characteristics (size of myofibers, number and distribution of myonuclei, number of satellite cells, vessels, macrophages).

Results:

None of the investigated cell types (satellite cells, endothelial cells, macrophages) showed variations in their density within the tissue in both WT and DMD ^mdx^4Cv muscle. However, analyzing their number per myofiber showed that in DMD ^mdx^4Cv, myofiber capillarization was increased from 1 to 6 months as compared with WT muscle, then dropped from 12 months. Macrophage number did not vary in WT muscle and peaked at 6 months in DMD ^mdx^4Cv muscle. The number of satellite cells per myofiber did not vary in WT muscle while it remained high in DMD ^mdx^4Cv muscle, starting to decrease from 12 months and being significantly lower at 24 months of age. Myofiber size was not different in DMD ^mdx^4Cv from WT except at 24 months, when it strongly decreased in DMD ^mdx^4Cv muscle. Necrosis and lipid deposition were rare in DMD ^mdx^4Cv muscle. Fibrosis did not increase with age in DMD ^mdx^4Cv muscle and was higher than in WT at 6 and 12 months of age.

Conclusions:

As a whole, the results show a strong decrease of the myofiber size at 24 months, and an increased capillarization until 6 months of age in DMD ^mdx^4Cv as compared with the WT. Thus, DMD ^mdx^4Cv mice poorly recapitulates histological DMD features, and its use should take into account the age of the animals according to the purpose of the investigation.

Keywords

aging histology duchenne muscular dystrophy

INTRODUCTION

Duchenne muscular dystrophy (DMD) is a progressive X-linked neuromuscular disorder due to mutations in the DMD gene, that encodes for dystrophin. Dystrophin belongs to a large transmembrane complex (dystrophin associated glycoprotein complex) that links the intracellular cytoskeleton of the myofiber to the extracellular matrix. The absence of dystrophin leads to recurrent myofiber damages [1], that trigger the activation of satellite cells, or muscle stem cell (MuSC)s and their entry into the myogenesis program, in attempt to repair the damaged myofibers [2]. Although highly efficient in normal skeletal muscle, the process of regeneration eventually fails in DMD due to chronic myofiber damage [1], leading to the replacement of the muscle parenchyma by fibrosis and lipid deposition. Moreover, a pro-fibrotic environment is associated with a deficit in myogenesis [3]. The most used model for DMD is the mdx mouse, which bears a nonsense mutation in the Dmd gene [4]. On the contrary to human, mdx mice do not show clinical signs of the disease, move and live normally. However, mdx mice are easy to breed and the mdx hindlimb muscle shows signs of permanent regeneration associated with alterations in muscle function when specifically stimulated [5 –7]. Therefore, despite its limitations, the mdx mouse is widely used as a model for understanding the physiopathological mechanisms sustaining DMD and as a preclinical model to test therapeutic strategies.

Most of the longitudinal studies describing mdx skeletal muscle tissue all life long were done with C57/BL10 mice, the original background of the mdx mouse (here after referred as B10-mdx) [8 –11]. More recent studies have been using a C57/B6 background, and the DMD ^mdx^4Cv and DMD ^mdx^5Cv mutants because they show a low frequency of reversion mutations [12].

A crucial parameter to consider when evaluating mdx muscle phenotype or function is the age of the animal. Of most importance, B10-mdx mice show an episode of acute myofiber degeneration at 3-4 weeks of age (at time of weaning) that leads to an important process of regeneration [13 –15]. Thereafter, continuous cycles of damage and regeneration lead to more than 80–90% of myofibers exhibiting hallmarks of regeneration at 3 months of age [15, 16]. Second, from several pioneers studies that investigated B10-mdx muscle at various ages (from 10 to 24 months), it was admitted that time accelerates the dystrophic process, although high variations were observed between the studies [8 , 17–19]. As a result, the literature in the field encompasses studies using a variety of time ranges using the mdx model, animals being considered as “old” from 10 to 22 months.

One of the most commonly used readouts of muscle homeostasis are histological parameters, such as the number and size of myofibers, the number of satellite cells, of vessels, etc. Having established a series of samples that were used for previous studies, we took the opportunity to perform a histological comparative analysis of these samples, which were Tibialis Anterior (TA) muscles of the C57/B6 DMD ^mdx^4Cv mouse strain, from 1 to 24 months of age. TA muscle is one of the mostly used for investigation of MuSC biology. The B6 background is also very popular because of the use of many transgenic strains for the study of muscle biology. In the DMD ^mdx ^4Cv model, a C to T transition in exon 53 at position 7916 creates a premature ochre stop codon (CAA to TAA) [20]. DMD ^mdx ^4Cv mouse strain exhibits low frequency of reversion mutations, rendering that model suitable for gene and cell therapy in preclinical investigations [12].

MATERIALS AND METHODS

Mice experiments and histology

WT and DMD ^mdx^4Cv [20] on C57BL/6J background males from 1 to 24 months of age were used according to the French legislation (Approval from local Animal Care and Use Committee was obtained under ref CEE A34.BC-RM.053.12). 5 mice were used at each time point. The TA muscle was recovered and was frozen in liquid nitrogen-precooled isopentane, and stored at –80°C until use. Ten micrometer-thick cryosections were made and used for stainings and immunostainings.

Histological stainings

Hematoxylin-eosin (HE) was used for morphological observation of the muscle tissue. To label lipids, sections were stained with Sudan Black solution (199664, Sigma) for 2 h and were counterstained with Hemalun (MHS80, Sigma) for 1 min. Whole muscle sections were reconstituted with ImageJ software after recording at ×10 objective using a Zeiss axioskop microscope and an Axiocam ICC5 zeiss camera. The area of black staining (Sudan black staining) was measured using ImageJ software using the AutoThreshold Yen and Analyze/histogram function.

Immunolabeling for satellite cells

Muscle sections were fixed with paraformaldehyde (4%) for 10 min at room temperature and permeabilized with Triton X-100 (0.5%) before acidic antigen retrieval was performed (Citrate buffer 10 mM at 90°C for 5 min). Slides were incubated with primary antibodies against Pax7 (1/50, Developmental Studies Hybridoma Bank, DSHB) overnight at 4°C, then washed with PBS and further incubated with FITC-conjugated secondary antibodies (1/200, Jackson ImmunoResearch Laboratories). A biotin-conjugated secondary antibody (1/200 Vector laboratory, BA-2000) revealed by a DTAF-conjugated streptavidin (1/1000, Beckman Coulter, PN IM0307) was used to amplify the signal as previously described [21]. Muscle sections were then incubated with anti-laminin antibodies (1/100, L9393 sigma) that stains all basal membranes, for 2 h at 37°C, washed and further incubated with Cy3-conjugated secondary antibodies (1/200, Jackson ImmunoResearch Laboratories) for 45 min at 37°C. Sections were washed with PBS, incubated in Hoechst solution for 10 sec, and then mounted with Fluoromount (FP483331, Interchim). About 12 pictures covering all areas of the muscle section were taken at x20 objective using Zeiss Z1 imager microscope and a Photometrics CoolSnap camera. The number of muscle stem cells (Pax7^pos) was manually counted using ImageJ software as well as the number of myofibers (thanks to laminin immunostaining). Results are given in number of cells/myofiber or in number of cells/mm² of muscle section.

Immunolabeling for macrophages

Muscle sections were directly incubated with primary antibodies against F4/80 (1/200 ab6640 Abcam) overnight at 4°C, then washed with PBS and further incubated with Cy3-conjugated secondary antibodies (1/200 Jackson ImmunoResearch Laboratories). Muscle sections were then fixed with paraformaldehyde (4%) for 10 min at room temperature and permeabilized with Triton X-100 (0.5%) before immunolabeling for laminin as described above for satellite cells. About 12 pictures covering all areas of the muscle section were taken at x20 objective using Zeiss Z1 imager microscope and a Photometrics CoolSnap camera. The number of macrophages (F4/80^pos) was manually counted using ImageJ software as well as the number of myofibers (thanks to laminin immunostaining). Results are given in number of cells/myofiber or in number of cells/mm² of muscle section.

Immunolabeling for endothelial cells

Muscle sections were fixed with paraformaldehyde (4%) for 10 min at room temperature and permeabilized with Triton X-100 (0.5%) before incubation with primary antibodies against CD31 (1/200, ab7388, Abcam) overnight at 4°C, then washed with PBS and further incubated with Cy3-conjugated secondary antibodies (1/200, Jackson ImmunoResearch Laboratories). Muscle sections were then treated for the detection of laminin as described above for satellite cells. About 12 pictures covering all areas of the muscle section were taken at x20 objective using Zeiss Z1 imager microscope and a Photometrics CoolSnap camera. The number of capillaries (CD31^pos) was manually counted using ImageJ software as well as the number of myofibers (thanks to laminin immunostaining). Results are given in number of cells/myofiber or in number of cells/mm² of muscle section.

Immunolabeling for collagen 1

Muscle sections were fixed with paraformaldehyde (4%) for 10 min at room temperature and permeabilized with Triton X-100 (0.5%) before incubation with primary antibodies against collagen 1 (1310-01, Southern Biotech) overnight at 4°C, then washed with PBS and further incubated with Cy3-conjugated secondary antibodies (1/200, Jackson ImmunoResearch Laboratories) and mounted with Fluoromount (FP483331, Interchim). About 12 pictures covering all areas of the muscle section were taken at x20 objective using Zeiss Z1 imager microscope and a Photometrics CoolSnap camera. Fibrosis was quantified after collagen I immunolabelling as in [22]. Whole muscle sections were automatically scanned at ×10 of magnification using an Axio Observer.Z1 (Zeiss) connected to a CoolSNAP HQ2 CCD Camera (photometrics).

Immunolabeling for damaged myofibers

Muscle sections were fixed with paraformaldehyde (4%) for 10 min at room temperature and permeabilized with Triton X-100 (0.5%) before incubation with donkey anti-mouse FITC-conjugated IgGs (Jackson ImmunoResearch Laboratories) overnight at 4°C, then washed with PBS and mounted with Fluoromount (FP483331, Interchim). Whole muscle sections were automatically scanned at ×10 of magnification using an Axio Observer.Z1 (Zeiss) connected to a CoolSNAP HQ2 CCD Camera (photometrics). The area of IgG positive myofibers was manually delineated. Results are given in % of the total muscle section area.

Analysis of myofiber CSA

Whole muscle sections were automatically scanned at ×10 of magnification using an Axio Observer.Z1 (Zeiss) connected to a CoolSNAP HQ2 CCD Camera (photometrics). Myofiber cross-section area (CSA) was calculated on whole muscle sections using Open-CSAM software based on laminin staining as previously described [23].

Analysis of myonuclei

12 pictures covering all areas of the muscle section immunolabelled for laminin were taken at x20 objective using Zeiss Z1 imager microscope and a Photometrics CoolSnap camera. The number of myonuclei per myofiber (distinguishing the myofibers with peripheral versus central nuclei) was manually counted using ImageJ software.

Statistics

For each time point, 5 mice were analyzed in a non-blinded way by two independent experimenters. Statistical analyses included One-way Anova after checking normality or Man-Whitney test for non-parametric data. P < 0.05 was considered significant.

RESULTS AND DISCUSSION

TA muscles from WT and DMD ^mdx ^4Cv from 1 to 24 months were proceeded for histological stainings and immunostainings. While DMD ^mdx ^4Cv muscle showed several signs of necrosis, inflammation, heterogeneity in fiber size and signs of regeneration as compared with WT, there was no obvious macroscopic changes in the muscles with age (HE staining examples are given in Figure Supplement 1).

Myofibers

Laminin immunolabeling (Fig. 1A) was used to analyze various parameters of myofibers. After the huge regeneration process observed at 3 weeks, the number of regenerating myofibers increases in mdx muscles, reaching about 80% after a few weeks and remaining high until 2 years of age [19, 24]. We evaluated the number of myonuclei/myofiber in both non-regenerating myofibers (that present a peripheral location of their nuclei) and regenerating/regenerated myofibers (exhibiting myonuclei in a central location) in about 12 pictures taken randomly in the whole section. In WT muscle, myonuclei were mainly present at the periphery of myofibers (Fig. 1B). Rare myonuclei were present in a central position, reflecting isolated fusion events all life long, with no increase with age (Fig. 1B). In DMD ^mdx ^4Cv muscles, the number of myonuclei dramatically increased in myofibers with central myonuclei from 3 months of age (+ 272% vs 1 month), likely reflecting the transition from an acute to a chronic regenerating state of the muscle (Fig. 1B). Concomitantly, the number of nuclei in myofibers exhibiting only peripheral nuclei declined (Fig. 1B). It was previously reported that myofibers isolated from Soleus, Extensor Digitorum Longus and Flexor Digitorum Brevis (FDB) muscles show abnormalities from 4 months of age, which increase after 6 months to reach 90% of myofibers (30% in FDB) [25]. Adding a level of complexity in the analysis of mdx muscle using transversal sections, both peripheral and central nuclei were observed along the same myofiber in B10-mdx muscle [11, 25].

Fig. 1

Analysis of myofibers in WT and DMD ^mdx ^4Cv muscle. TA muscle sections from 1- to 24-month-old WT and DMD ^mdx ^4Cv were immunolabelled for laminin (green) and stained with Hoechst (nuclei) (A). From this immunolabeling, the number and location of nuclei within myofibers (B), the number of myofibers per mm² (C), mean CSA (D), and CSA distribution (E-F) were evaluated. Values are given in means±SEM of 5 experiments (one black circle represents one mouse). P value of Anova analysis is provided on the upper left corner of the graph. Post-hoc comparisons show significant differences. In B, $ vs mdx 1 mo for both central and peripheral myonuclei, £ vs mdx 24 mo for central myonuclei only. In B, the number of myonuclei in both central and peripheral positions differs in WT vs mdx for all ages except peripheral myonuclei at 1 mo. In C,D, ^* vs WT 1 mo, φ vs WT 12 mo, $ vs mdx 1 mo, £ vs mdx 24 mo, § mdx vs WT at same age. Bar = 50μm.

Next, the number and size of myofibers were evaluated on entire muscle sections. In both WT and DMD ^mdx ^4Cv muscles, the number of myofibers per unit area decreased from 1 to 3 months. Then, it did not vary in WT while in DMD ^mdx ^4Cv muscle, the number of myofibers increased at 24 months (Fig. 1C). Similarly, a previous study indicated no high variation between 3 and 12 months of age in B10-mdx hindlimb muscles [17].

Myofiber size is a popular feature to assess skeletal muscle regeneration. We used a semi-automated tool to measure myofiber CSA in regenerating conditions on entire muscle sections [23]. Results of the mean CSA (Fig. 1D) and CSA distribution (Fig. 1E,F) showed that in both WT and DMD ^mdx ^4Cv muscles, CSA increased at 3 months (+ 157 and 182% vs 1 month, respectively). Thereafter, the mean myofiber CSA did not vary until a very late time point, i.e. at 24 months (–59% vs 12 months), where the smallest CSA was observed in DMD ^mdx ^4Cv muscles, in accordance with the increased number of myofibers at this last time-point (Fig. 1C). This decrease was not observed in WT muscles, although the distribution of myofiber CSA showed an increased number of smaller myofibers at 24 months (Fig. 1E). In DMD ^mdx ^4Cv muscles, the distribution of myofiber CSA varied according to the CSA mean, with the smallest myofibers being observed at 1 and 24 months, while bigger myofibers were observed in 6-month-old animals (Fig. 1F). Previous studies reported myofiber hypertrophy during the first months of life of B10-mdx, likely in response to the huge degenerating process occurring at 3–5 weeks of age [5 , 26]. Another study showed an increase of the myofiber CSA at 12 months [17]. However, other analyses indicated that at 10 months of age, B10-mdx muscles exhibit an increase of both small and large myofibers [11, 27]. It is likely that this great heterogeneity is due to myofiber branching, which has been repeatedly reported [7, 25] to increase with age [28] and to be very important after 2 years [11, 29]. Myofiber branching is associated with an alteration of calcium signaling and of excitation/contraction coupling, leading to defects in myofiber function [30, 31]. Myofiber branching is observed in aged normal skeletal muscle, with about 15% of myofibers exhibiting 2 branches [32]. In contrast, myofiber branching is a frequent event in B10-mdx muscle, since 100% of myofibers of EDL muscle are branched at 17 months of age [33], and this may explain the small myofiber CSA that we observed at 24 months.

Muscle stem cells

MuSCs have been particularly investigated in the dystrophic context. We performed Pax7 immunolabeling to count total (both quiescent and activated) MuSCs in WT and DMD ^mdx ^4Cv muscles (Fig. 2A). When counting the number of MuSCs per area unit, no variation was observed from 1 to 24 months of age in both strains, although the number of MuSCs was always much higher in DMD ^mdx ^4Cv than in WT muscle (Fig. 2B, Figure Supplement 2A,B). When counting the number of MuSCs per myofiber, the only significant difference was observed at 24-month-old in DMD ^mdx ^4Cv muscles (–41%) (Fig. 2C, Figure Supplement 2A,B), when muscles exhibited the most myofibers. These results suggest that the number of MuSCs did not vary with age in TA DMD ^mdx ^4Cv muscle. Studies using the same Myf5^nlacZ lineage tracing model and similar isolated EDL single fiber technique reported opposite results with either an increase [33] or a decrease of the number of satellite cells with age [34]. Thus, the evolution of the number of MuSCs in mdx strains should be carefully monitored depending on the technique used, genetic background and sex of the animal.

Fig. 2

Analysis of satellite cells in WT and DMD ^mdx ^4Cv muscle. TA muscle sections from 1- to 24-month-old WT and DMD ^mdx ^4Cv were immunolabelled for Pax7 (green) and for laminin (red) and stained with Hoechst (nuclei) (A). From this immunolabeling, the number of Pax7^pos cells/mm² (B) and the number of Pax7^pos cells/myofiber (C) were evaluated. Values are given in means± SEM of 5 experiments (one black circle represents one mouse). P value of Anova analysis is provided on the upper left corner of the graph. Post-hoc comparisons show significant differences for £ vs mdx 24 mo and § mdx vs WT at same age. Bar = 50μm.

Endothelial cells

Skeletal muscle is highly vascularized. Moreover, endothelial cells exert specific effects on MuSC differentiation and myogenesis [35]. Immunolabeling for CD31 (PECAM1) allowed to evaluate the number of blood vessels and capillaries (Fig. 3A). The number of vessels per surface unit was not significantly altered from 2 to 24 months (Fig. 3B). However, when reporting the number of capillaries per myofiber, there was an increase until 6 months in myofiber capillarization in DMD ^mdx ^4Cv muscles, as compared with WT, followed by an important drop between 6 and 12 months of age in DMD ^mdx ^4Cv animals (–47%) while no variation was observed in WT muscle (Fig. 3C and Figure Supplement 2C,D). These results are consistent with previous studies showing a reduced number of vessels irrigating each myofiber, anatomical alterations with anastomosis, that is associated with a defect in perfusion in one-year-old DMD ^mdx ^4Cv [36] and with a low number of capillaries per myofiber at 24 months [26]. No specific explanation is available to explain the capillarization drop from 12 months.

Fig. 3

Analysis of endothelial cells and macrophages in WT and DMD ^mdx ^4Cv muscle. TA muscle sections from 1 to 24-month-old WT and DMD ^mdx ^4Cv were immunolabelled for CD31 (red in A) or F4/80 (red in D) and for laminin (green) and stained with Hoechst (nuclei) (A,D). From this immunolabeling, the number of CD31^pos cells/mm² (B), the number of CD31^pos cells/ myofiber (C), the number of F4/80^pos cells/mm² (E) and the number of F4/80^pos cells/myofiber (F) were evaluated. Values are given in means± SEM of 5 experiments (one black circle represents one mouse). P value of Anova analysis is provided on the upper left corner of the graph. Post-hoc comparisons show significant differences for # vs mdx 3 mo, Δ vs mdx 6 mo, € vs mdx 12 mo, £ vs mdx 24 mo, and § mdx vs WT at same age. Bar = 50μm.

Macrophages

Macrophages have been shown to be present in dystrophic B10-mdx muscles from early stages (5 weeks) to advanced age (12 months) [5, 37]. A chronic inflammatory response signature was detected in 8-week-old B10-mdx muscles using microarrays [38]. Macrophages are important cells during muscle regeneration but were shown to play both beneficial and adverse roles in mdx muscle of various backgrounds [22 , 40]. Immunolabeling for the pan-macrophage marker F4/80 (Fig. 3D) showed that the number of macrophages per unit area was not altered from 2 to 24 months of age in both strains (Fig. 3E). However, the number of macrophages was much higher in DMD ^mdx ^4Cv muscles than in WT muscles, all life long (from 41 to 91 fold) (Fig. E,F). When counting the number of macrophages per myofiber, no change was observed in WT muscle (Fig. 3F). In DMD ^mdx ^4Cv muscles, an increase was observed at 6 months where macrophages were at least x1.35 fold more numerous than at any other time point (Fig. 3F, Figure Supplement 2E,F). Apart this time point, the number of macrophages remained constant, notably at advanced age, indicating that the number of macrophages did not increase with age.

Necrosis

In DMD, myofibers undergo chronic cycles of damage/degeneration and regeneration. Muscle sections were labelled with anti-mouse IgGs allows to detect leaky myofibers that uptake serum proteins [41]. It was shown that myofibers that appear necrotic in HE staining are positive for plasma protein labelling or for Evans blue dye, although some positively labelled myofibers may appear intact in HE staining [42, 43]. Only a few reports quantified myonecrosis in mdx muscles. While important necrosis is observed at 3 weeks, its extent is dramatically reduced few days later [44]. Fig. 4 shows that the number of myofibers labeled with anti-mouse IgGs was very low, never reaching more that 0.25% of the total WT muscle area (Fig. 4A,B). In DMD ^mdx ^4Cv muscles, the number of positive myofibers was higher but still did not represent more than 1% of the area of the whole muscle section (Fig. 4A,B). Similarly, Pastoret et al. previously showed that only very few degenerating myofibers are present in B10-mdx muscles from 1 to 24 months [19].

Fig. 4

Evaluation of the muscle tissue structure of WT and DMD ^mdx ^4Cv muscle. Necrosis was evaluated after staining with anti-mouse IgGs (A), as the percentage of total muscle section area (B). Lipid deposition was evaluated after staining with Sudan Black (C), as the percentage of total muscle section area (D). Fibrosis was evaluated after immunolabeling for Collagen I (E), as the percentage of total field area (F). Values are given in means±SEM of 5 experiments (one black circle represents one mouse). P value of Anova analysis is provided on the upper left corner of the graph. Post-hoc comparisons show significant differences for ^* vs WT 1 mo, $ vs mdx 1 mo, £ vs mdx 24 mo, φ vs WT 12 mo and § mdx vs WT at same age. Blue = Hoechst. Bars in A, C = 100μm, in E = 50μm.

Lipid deposition

Lipid deposition due to adipocyte infiltration is, with fibrosis, a hallmark of muscle degeneration in DMD, where the parenchyma, i.e., the myofibers, is replaced by fatty-fibrotic tissue, leading to muscle weakness. Sudan Black stains lipid deposits (Fig. 4C). Overall, the extent of lipid deposition in WT and DMD ^mdx ^4Cv muscles was extremely low, accounting for no more than 0.35% of the whole muscle section area (Fig. 4D). An increase of lipid deposits was observed in DMD ^mdx ^4Cv muscles at 24 months as compared with the other time points and WT (Fig. 4D and Figure Supplement 3A). As previously shown in the B10 background, no lipid deposition is present in the mdx muscle at any age, making an important difference with human DMD muscles [5 , 45].

Fibrosis

Fibrosis is a major adverse process in DMD since excess of collagen deposit hampers muscle function. It was shown that at the time of diagnosis, endomysial fibrosis is a bad prognosis of motor outcome in DMD patients years later [46]. However, B10-mdx hindlimb muscles present no or little fibrosis, contrary to the diaphragm [47, 48]. While some studies reported no fibrosis in hindlimb muscles until 9–11 months of age [13, 45], others mentioned an increase of fibrosis at 23 months but it was not quantified [18]. Evaluation of the area of collagen deposition in the TA muscle (Fig. 4E and Figure Supplement 3B) showed that in DMD ^mdx ^4Cv muscles, the amount of “fibrosis” accounted for about 14 to 18% of the muscle field area, and remained stable from 1 month to 24 months of age (Fig. 4F), with no evidence of increased fibrosis with age in the whole muscle section (Figure Supplement 3). The amount of fibrosis was higher in DMD ^mdx ^4Cv than in WT muscles at 6 and 12 months of age, but was not significantly different in 24-month-old animals, since collagen area also increased in WT muscle at that time point (+184% vs 12 months) (Fig. 4F).

CONCLUSION

Altogether, these results show that the histology of the TA muscle of DMD ^mdx^4Cv mice on C57/BL6 background showed modest variations in absolute numbers of the various muscle cell types with age. Myofiber CSA increased at 3 months to remain stable until 24 months when it was strongly reduced. None of the cell types evaluated in this study showed a variation in their density within the muscle tissue. However, analyzing their number relative to the number of myofibers indicated that the number of MuSCs decreased at 24 months, that myofiber capillarization was high until 6 months before dropping at later ages. Necrosis and lipid deposition were very rare in the DMD ^mdx ^4Cv muscle tissue, even at late time points. Finally, collagen I deposition did not increase with age. Overall, the main variations were observed at 24 months of age, when the number of myofibers was strongly increased, probably due to increased branching, inducing a mathematical decrease of all cellular parameters. These results are in accordance with the DMD ^mdx ^4Cv mice behavior that move, behave and breed as well as normal mice when maintained in normal conditions. These results also indicate that the use of the mdx model has limitations in mimicking DMD since there is no obvious worsening of the muscle histology with time. This should be taken into account depending on the type of biological investigations and the purpose of the study. Studies aiming at investigating degenerative myopathies in general may prefer other models, such as sarcoglycan deficient mice, that show a natural evolution of the pathology [49], or, for DMD pathology the more recently described DBA/2-mdx, where stronger features of degenerative myopathies are observed [50, 51]. In this model, a link between TGF-β pathway, fibrotic areas and myogenesis deficit was evidenced in muscles from young adults [3] while some pathological features appear less severe in older animals [50, 52], suggesting that in the DBA/2 model also, the age of the animals should be considered depending on the purpose of the study.

Footnotes

ACKNOWLEDGMENTS

This work used samples that were established for previous studies that were funded by grants from the Framework Programme FP7 Endostem (under grant agreement 241440), AFM-Telethon (grant 16029 and MyoNeurAlp Alliance), Fondation pour la Recherche Médicale (Equipe FRM DEQ20140329495).

CONFLICT OF INTEREST

The authors have no conflict of interest to report.

The supplementary material is available in the electronic version of this article: .

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