Stimulation of EGFR signaling in fibro-adipogenic precursors decreases adipogenesis in Duchenne muscular dystrophy

Abstract

Background

Duchenne Muscular Dystrophy (DMD) is characterized by the formation of fibrosis and fat deposits that progressively replace muscle fibers, resulting in the loss of muscle function. Both fibrosis and adipogenesis are operated by fibroadipogenic precursors (FAPs), but the molecular regulation and interactions between the two processes are not fully understood.

Methods

Adipogenesis was investigated in vivo in the D2-mdx mouse, and in vitro using FAPs isolated from WT (DBA/2) and D2-mdx muscles. Epithelial Growth Factor (EGF) was overexpressed in the D2-mdx muscle via electroporation of an expression plasmid.

Results

We found that the D2-mdx gastrocnemius muscle showed fat deposition from 10 weeks of age and increased until 18 weeks of age, coinciding with fibrosis. Fat deposition was exclusively found within fibrotic areas. In vitro, D2-mdx FAPs proliferated more, and were more prone to adipogenesis than WT FAPs. Cells from both genotypes showed equal fibrogenesis. Analysis of normal muscle snRNAseq data showed that the Epithelial Growth Factor Receptor (EGFR) was primarily expressed by FAPs. Both EGFR expression and EGFR-phosphorylation were decreased in D2-mdx FAPs as compared with WT FAPs. Stimulating FAPs with EGF decreased adipogenesis, more efficiently in D2-mdx FAPs than in WT FAPs. However, EGF stimulation of EGFR had no effect on their fibrogenic differentiation. Finally, in vivo overexpression of EGF in D2-mdx gastrocnemius muscles reduced both adipogenesis and fibrosis, and was associated with an increased muscle force.

Conclusions

In a DMD context, FAPs are more likely to differentiate into adipocytes than in normal muscle, which is associated with decreased EGFR signaling. Stimulating EGFR signaling decreased adipogenesis in vitro and fat deposition in vivo. The impact of EGFR signaling on fibrogenesis is unclear, the reduced fibrosis observed in vivo may be due to indirect mechanisms. This study identifies EGFR signaling as a new molecular mechanism for controlling adipogenesis in skeletal muscle FAPs.

Supplementary Information

The online version contains supplementary material available at 10.1186/s13395-026-00429-2.

Background

In Duchenne Muscular Dystrophy (DMD), the absence of the dystrophin protein causes repeated disruption of sarcolemma of the myofibers, triggering asynchronous muscle injuries. This leads to chronic inflammation in various areas of the muscle and is eventually associated with fibrosis and fat deposition. The mdx mouse, which lacks dystrophin, exhibits a much less severe phenotype than patients with the disease. Specifically, the mdx muscle does not show endomysial fibrosis except in the diaphragm, nor fat deposition [1, 2]. A new mdx model has recently emerged through the crossbreeding of mdx B6 mice with the DBA/2 genetic background (referred to here as D2-mdx) [3–5]. This model exhibits at least 22 polymorphisms, including annexin A6, which is involved in sarcolemma repair, and LTBP4, which is involved in TGFβ secretion [6, 7]. The initial description of the D2-mdx model reported areas of adipose deposition in 6- to 8-week-old animals, albeit at a low level [4]. Interestingly, a peak of muscle damage, notably fibrosis, occurs in D2-mdx muscle at 10–20 weeks of age, followed by an improvement of muscle homeostasis [8].

Both adipogenesis and fibrogenesis originate from skeletal muscle mesenchymal stem cells, called fibroadipogenic precursors (FAPs). Depending on the cues they receive, FAPs can differentiate into fibroblasts/myofibroblasts, adipocytes or osteocytes [9]. FAPs are necessary for muscle regeneration after injury because they establish interactions with muscle stem cells and immune cells, and actively remodel the extracellular matrix [9]. FAP homeostasis is strongly perturbed in degenerative myopathies such as Duchenne Muscular Dystrophy (DMD), where FAPs show high rates of proliferation and differentiation into fibroblasts and adipocytes [10]. While fibrogenic differentiation has been extensively studied, adipogenic differentiation is less well understood, partly due to the lack of available models and because adipogenesis is barely induced by toxin-induced injury, the widely used model for studying muscle regeneration. Nevertheless, a series of effectors, pathways and transcription factors have been identified as regulators of adipogenesis by FAPs, mainly using the glycerol induced muscle injury model. Some pathways activate the adipogenic program in FAPs, such as TGFβ [11], Annexin A2 [12], MMP14/KLF6 [13], FGF2/mir29a/SPARC [14], thyroid hormone [15], ADAMTLS2 [16]. Other pathways work as inhibitors of adipogenesis, including interleukin-15 [17], mir206/runx1 [18], MMP13 [19], Wna7a/βcatenin/TAZ [20], MST1-2/YAP [21] and DHH/TIMP3 [22]. In the mdx muscle, the Wnt5a/GSK-3/βcatenin axis activate adipocyte formation [23], while nitric oxide [24], Notch and TNF⍺ [25] limit adipogenesis.

Here, we identified significant adipogenesis in the gastrocnemius muscle of D2-mdx mice between 10 and 18 weeks of age. Adipogenesis was always located within fibrotic areas. Using available omics data, we determined that FAPs express EGFR. We investigated EGFR signaling in FAPs and found that it was lower in D2-mdx than in WT FAPs. Inducing EGFR signaling reduced adipogenesis but not fibrosis in vitro. However, in vivo reexpression of EGF in D2-mdx muscle decreased both adipogenesis and fibrogenesis, and improved muscle strength.

Methods

Animals

Experiments were performed on 10 to 18 week-old D2-mdx (D2.B10-Dmdmdx/J) and DBA2 (WT) male mice. Mice were housed in group cages in a controlled environment facility with a 12-h light/dark cycle and had access to food and water ad libitum. All experiments and procedures were performed in accordance with French and European legislation on animal experimentation.

Electroporation

Gastrocnemius muscles of 12 week-old D2-mdx mice were electrotransferred by pTT3-EGF-CD4d3 + 4-bio-His plasmid encoding EGF (Addgene #53340) or an empty plasmid (D4d3 + 4-bio, Addgene #32402) kindly given by Michael Rudnicki (OHRI, Ottawa, Canada) [26] as previously described [27]. Briefly, muscles were injected with 100 µl of Hyaluronidase (0.4 U/ml) in 0.9% saline solution 2 h before muscle electrotransfer. Plasmid at 10 µg/µl were injected (50 µl/muscle) and 2 × 4 pulses of 50 V/cm pulses, lasting 60 ms each with a 100-ms interval, were then applied. Four weeks later, mice were prepared for force measurement.

Maximal isometric torque measurement

Was performed as previously described [28]. Briefly, mice were maintained anesthetized with 1.5-2% isoflurane and the right foot was positioned on an ergometer pedal. Plantar flexor muscles were stimulated using two surface electrodes located below the knee and at the level of the Achilles tendon. Maximal current intensity was determined by progressively increasing the current intensity until no further increase in twitch torque occurred. This maximal intensity was used to assess torque production in response to a 250-ms stimulation train delivered at frequencies ranging from 1 to 150 Hz.

Histology preparation

Gastrocnemius muscles were mounted on a cork stopper with tragacanth gum (ThermoFisher scientific #11458417) and snapped frozen in liquid nitrogen-cooled isopentane (Sigma-Aldrich #277258). The samples were then stored at -80 °C. Cryosections (10 μm) were prepared for histological analysis using a cryostat (Epredia™ CryoStar™ Cryostat NX50).

Immuno-histology

All steps were done at room temperature unless indicated. Cryosections were fixed in formaldehyde 4% (Sigma-Aldrich #1004965000) for 15 min, before permeabilization for 10 min in Triton-X100 0.5% (Sigma-Aldrich #T7878). Sections were incubated with Bovin Serum Albumin (BSA) (Sigma-Aldrich #F3385) 4% in PBS for 1 h. Primary antibodies were diluted in BSA 2% and were added to the sections for overnight incubation at 4 °C (goat anti-Col1a antibody [SouthernBiotech #1310, 1/200], rabbit anti-Perilipin antibody [Abcam #AB3526, 1/200]), anti-PDGFRα [R&D Systems AF1062, 1/200], anti-phospho-EGFR [Cell Signaling #3777, 1/400]. Secondary antibodies (diluted in PBS) were incubated for 1 h at 37 °C (Cya3 anti-goat [Jackson ImmunoResearch #705-165-147, 1/200], FITC anti-rabbit [Jackson ImmunoResearch #711-095-152, 1/200]). Sections were counterstained with Hoechst (ThermoFisher Scientific #62249) and mounted in fluoromount G medium (Invitrogen #00-4958-02). Image acquisition was done with an Axio Observer 7 (Zeiss) connected to an ORCA-Flash4.0 LT3 Digital CMOS camera using a 20x objective. For the analysis, whole sections were analyzed using a FIJI macro that was specially developed for this purpose (https://github.com/benedictechazaud/research-papers-code). High fluorescent zones and tendons were removed from the images. Fibrosis areas and the entire muscle were manually selected to measure the section area. Channels were then split, and only the red and green channels were analyzed, as follows: the background was removed, and a threshold was applied. A particle analysis was performed on either the entire section or the area (for perilipin labeling only). All results were given in % to the section area.

FAP culture

Hindlimb muscles were harvested, were separated from skin, fat, and tendons and were finely minced. Muscles were digested in 10 ml of digestion medium (2 mg/ml collagenase B (Roche #11088831001) and 3 mg/ml dispase (Merck D4693-6) in FAP culture medium (DMEM [Sigma-Aldrich #D6429] containing 10% heat-inactivated FBS) for 1 h at 37 °C under gentle stirring. The cell suspension was filtered through a 70 μm cell strainer and was centrifuged for 5 min at 450 g. The supernatant was discarded and cells were resuspended in FAP culture medium for 3–4 h of culture, the supernatant was discarded, leaving to FAP population. Cell purity was assessed just after the preplating by harvesting the cell with trypsin and cytospining them for PDGFRα immunolabeling. FAPs were cultured for 8 days. For adipogenic differentiation, FAPs were seeded at 15000 cells/cm² in Adipogenic Differentiation Medium (DMEM, 10% FBS, 0.5 mM IBMX [Sigma-Aldrich #I5879], 0.25 mM Dexamethasone [Sigma-Aldrich #D2915], 10 mg/ml insulin [Sigma-Aldrich #I1882], 5 mM Rosiglitazone [Sigma-Aldrich #R2408]) renewed every day for 5–8 days. For fibrogenic differentiation, FAPs were seeded at 5000 cells/cm² in Fibrogenic Differentiation Medium (DMEM, 10% FBS, 2 ng/ml TGFβ [Biolegend #763102]) for 24 h. In some experiments, FAPs were treated with recombinant mouse EGF (ThermoFisher Scientific #PMG8041) at 100 ng/ml and differentiated (vehicule was PBS).

FAP/macrophage coculture

FAPs were seeded at 15,000 cells/cm² into 24 well plates in FAP culture medium for 6 h to adhere. Meanwhile, macrophages were isolated from D2-mdx mice as described in [29] and were seeded into inserts (macrophages: FAPs 2:1 ratio) in FAP culture medium. Medium in the wells was changed for adipogenic differentiation medium containing only 1% FBS, inserts were added and cells cultured for 8 days.

Cell immunofluorescence

FAPs were fixed for 10 min in 4% formaldehyde, permeabilized for 10 min in Triton X-100 0.5%. BSA 4% was added for 1 h before primary antibodies were added. FAP were incubated overnight at 4 °C with primary antibodies (mouse anti-αsmooth muscle actin antibody [Sigma-Aldrich, #A5228, 1/500], goat anti-Col1a antibody [1/500], rabbit anti-Perilipin antibody [1/200]), anti-PDGFRα [R&D Systems AF1062, 1/200] then with the secondary antibodies (Cya3 anti-goat [1/200], FITC anti-mouse [Jackson ImmunoResearch, #715-095-150, 1/200], FITC anti-Rabbit [1/200]) for 2 h at room temperature in BSA 4%. In some experiments, FAP were incubated for 1 h at 37 °C with Phalloidin (Sigma-Aldrich #19083, 1/200). Cells were soaked for 1 min in Hoechst and mounted in fluoromount G medium (ThermoFisher Scientific #00-4958-02). Image acquisition was performed using a Zeiss axio observer z1 inverted fluorescence motorized microscope. Five pictures were taken for each sample at 10x magnification. Quantification was made manually using ImageJ software.

Western-blot

FAPs were starved in starved medium (SV) (Sigma-Aldrich #I7633, 0.1% FBS) for 18 h. Medium was changed and FAPs were incubated at 37 °C in SV medium without or with recombinant mouse EGF at 100 ng/ml for 1, 5, 30 min. Activation was stopped by adding 10 ml of cold PBS, cells were centrifuged and pelleted for protein extraction. Proteins were extracted in ice-cold lysis buffer (Tris-HCl pH7.4 50 mM, NaCl 150 mM, EDTA 1 mM, IGEPal 1%, SDS 1%, SDC 1%, Na3VO4 1 mM, Na-Pyrophosphate 1 mM, PMSF 1 mM, beta-glycerophosphate 1 mM) supplemented with protease and phosphatase inhibitor cocktail (Sigma-Aldrich #P8340). Insoluble materials were removed by centrifugation at 14,000 g for 20 min at 4 °C and protein concentration was determined using Pierce BCA assay (ThermoFisher Scientific #23227). Samples were boiled for 5 min in Laemmli buffer and run on 8% Bis-Tris Protein gels and then transferred using the wet system from ThermoFisher Scientific. Membranes were blocked for 2 h in BSA 5% and were incubated overnight with primary antibodies (rabbit anti-phospho-EGFR (Tyr1068) [Cell Signaling #3777, 1/1000], rabbit anti-EGFR [Cell Signaling #4267, 1/1000], mouse anti-actin [Cell Signaling #A222R, 1/2000]). Membranes were then incubated for 1 h with HRP-conjugated secondary antibodies (goat anti-rabbit HRP [Sigma-Aldrich #A9040, 1/10000], goat anti-mouse HRP [Sigma-Aldrich #A9044, 1/10000]). Proteins were visualized by chemiluminescence (ThermoFisher Scientific #32109,) and quantified using ChemiDocTM MP imaging system (BioRad). Spectra Multicolor High Range Protein ladder standards were used as markers of molecular weight (ThermoFisher Scientific #2665).

Transcriptomic analysis

For the GSE297388 data set (token elghwqumptqjzih): cell barcodes were filtered using a multi-step strategy. We used Seurat v4 package [30]. Sample-specific QC thresholds were defined using elbow points identified across metric distributions with akmedois::elbow_point. Doublets were detected using scDblFinder (cluster-based, dbr = 0.1) and features with mean expression > 0.1 were retained. After filtering, 1,476 cells (21,026 features) in D2.wt and 2,164 cells (21,045 features) in D2.mdx were used for downstream analyses. Data were normalized using SCTransform, regressing out mitochondrial content. PCA was performed on the SCT assay. Clustering was performed using the Leiden algorithm. For integrative analyses, datasets were merged and split according to a composite group variable. Integration was performed using HarmonyIntegration on the SCT assay. Visualization was performed using Fit-SNE. Cluster identities were assigned based on canonical marker gene expression and label transfer from a published reference dataset [31] using FindTransferAnchors and TransferData. Seven major cell populations were identified: muscle stem cells, fibro-adipogenic progenitors, endothelial cells, mural cells, immune cells, tenocytes, and Schwan.

For the dataset GSE305185 : we used Seurat v4 package [30]. Sample-specific QC thresholds were defined using elbow points identified across metric distributions with akmedois::elbow_point. Data were normalized using LogNormalize and integrate with Seurat. For integrative analyses, datasets were merged and split according to a composite group variable (condition). Integration was performed Cluster identities were assigned based on canonical marker gene expression. We identified 12 cell populations.

To visualize the expression of the Egfr receptor, we used the Seurat v5 package [30] (version 5.3.0) and R version 4.4.1. The figures were created using the Dimplot, FeaturePlot, VlnPlot, and Dotplot functions from the Seurat package.

Statistics

For both in vivo and in vitro assays, each experiment was repeated at least 3 times using independent animal. Statistics were done using Prism software. Two means comparison was done using Student’s t-test and multiple comparisons were done using one-way or two-way ANOVA with Holm-Sidak tests. The number of independent experiments are given in the figure captions.

Results

D2-mdx muscles present fat deposits

As previously reported [4], we observed that the gastrocnemius muscle of 10-week-old D2-mdx mice showed 8% of the whole section area positive for endomysial collagen 1 staining, that raised up by 2-fold to 18% in 18-week-old animals (Fig. 1, Suppl. Figure 1 A), despite important interindividual heterogeneity among the animals (Suppl. Figure 1B). Interestingly, the D2-mdx muscle exhibited spontaneous fat deposition, as revealed by perilipin-1 immunostaining. The area of adipose deposition was much smaller, accounting for about 1/10 of the fibrotic area in 10-week-old muscles, but increasing 4-fold in 18-week-old muscles (from 0.13 to 0.44% of total area, respectively) (Fig. 1). Notably, all the fat deposition areas were embedded within fibrotic areas, suggesting a link between fibrogenesis and adipogenesis in situ. In the DBA/2 wild-type (WT) muscle, a weak collagen staining was observed in the interstitial space while no lipid staining was observed (Fig. 1).

Fig. 1.

Fibrotic and fat deposition areas in D2-mdx muscles. Gastrocnemius muscles from 10- and 18-week-old WT and D2-mdx male mice were processed for immunostaining of collagen I and perilipin. Nuclei were stained with Hoechst. Fibrotic areas, as positive for collagen I, and fat deposition, as perilipin positive areas, were quantified as % of total muscle section area. Results are means +/- SEM using 4 animals. * p < 0.05 *** p < 0.001 using two-way ANOVA and Holm-Sidak test for multiple comparisons. Bars = 100 μm

D2-mdx FAPs are activated and differentiate into adipocytes

FAPs were isolated from D2-mdx and WT muscles and grown in their regular medium. The preplating method allowed to obtain FAP population that were over 95% positive for PDGFR⍺ at the end of the isolation procedure (Suppl. Figure 2A). The cell density of D2-mdx FAPs increased much faster than that of the DBA/2 FAPs, reaching a density that was 4.2 fold higher by day 4 of culture (Fig. 2A). This higher in vitro expansion capacity suggests that D2-mdx FAPs are more activated within the muscle. We then investigated the differentiation capacities of the FAPs. We induced their fibrogenic differentiation by adding TGFβ for 24 h, as previously reported [32]. We observed that WT FAPs slightly increased their Collagen I expression (+ 15%, p = 0.08 using paired t-test) upon TGFβ treatment, while maintaining their αSMA expression level (Suppl. Figure 2B). In contrast, D2-mdx FAPs did not increase their expression of either fibrogenic markers (Suppl. Figure 2B). These results indicate that TGFβ has a limited impact on FAPs that are already predisposed to fibrogenic differentiation. Next, we assessed the adipogenic potential of FAPs by culturing them in an adipogenic medium for 5 days. Perilipin immunostaining, which indicates differentiation into adipocytes, was positive in 26% of WT FAPs, increasing by 26% to reach 33% in D2-mdx FAPs (Fig. 2B). This suggests an increased adipogenic potential in the latter.

Fig. 2.

Characteristics of FAPs from WT and D2-mdx muscles. FAPs were isolated from WT and D2-mdx muscles. A FAPs were cultured in regular medium and the density of cells was evaluated. B FAPs were cultured in adipogenic medium for 5 days and were processed for immunostaining of perilipin. Nuclei were stained with Hoechst. Results are means +/- SEM using 3–5 independent cultures. * p < 0.05 ** p < 0.01 **** p < 0.0001 using two-way ANOVA and Holm-Sidak test for multiple comparisons (A) and unpaired t-test (B). Bars = 50 μm

Due to the sustained presence of macrophages in mdx muscles [29], we investigated whether macrophages derived from D2-mdx muscle affect FAP adipogenesis. Using inserts, we performed coculture experiments and observed that D2-mdx macrophages stimulated adipogenesis by 30% in WT FAPs but had no effect on D2-mdx FAPs (Suppl. Figure 2 C). These results may suggest that the adipogenic potential of D2-mdx FAPs is already at the maximum. Therefore, we examined intrinsic FAP signaling.

EGFR signaling is dampened in D2-mdx FAPs

To investigate the signaling pathways involved in adipogenesis in FAPs, we analyzed published snRNAseq data of normal WT muscle (in hypertrophied and non-hypertrophied conditions). Screening the expression of various receptors, we found that FAPs were the main cell type expressing EGFR, together with tenocytes, while MuSCs did also express the receptor (Fig. 3A). Using scRNAseq, we compared EGFR expression in D2-mdx FAPs and WT FAPs and we observed a decrease in the former (Suppl. Figure 2D). However, this decrease was not significant at the protein level, using immunoblotting from cultured FAPs (Fig. 3B). To investigate the status of the EGFR signaling pathway in WT and D2-mdx FAPs, we performed immunoblotting of the phosphorylated form of EGFGR on Tyr1068, one of the phosphorylation sites required for its activation [33]. Short treatment were tested using EGF as a ligand at 100 ng/ml (1 to 30 min), a concentration previously used in skeletal muscle [26, 34]. After stimulation with EGF, we found that the ratio of P-EGFR to total EGFR was lower in D2-mdx FAPs than in WT FAPs (e.g. -60% at 5 min) (Fig. 3C). This indicates a reduced EGFR signaling in D2-mdx FAPs upon ligand binding. Consequently, the overall EGFR phosphorylation was lower in D2-mdx than in WT FAPs (e.g. -67% at 5 min) (Fig. 3D) (Suppl. Figure 2E). This result suggests that D2-mdx FAPs have a lower EGFR intracellular signaling than WT FAPs.

Fig. 3.

EGFR signaling in FAPs. A Sn-RNAseq from published data shows that FAPs are the main population that expresses EGFR. B Immunoblotting semi-quantification of EGFR expression by WT and D2-mdx FAPs. C-D Immunoblotting semi-quantification of normalized P-EGFR expression by WT and D2-mdx FAPs treated with EGF (100 ng/ml), relative to total normalized EGFR expression (C) and to total actin (D). E, F WT and D2-mdx FAPs were cultured in adipogenic medium for 8 days with or without EGF (1-100 ng/ml) and were processed for perilipin immunostaining. Nuclei were stained with Hoechst. G WT and D2-mdx FAPs were cultured in fibrogenic medium for 1 day with or without EGF (100 ng/ml) and were processed for collagen I and αSMA immunostaining. Results are means +/- SEM of 3 independent cultures. *,^# p < 0.05 **^,## p < 0.01 **** p < 0.0001 using two-way ANOVA and Holm-Sidak test for multiple comparisons. Bar = 50 μm

To investigate the functional effect of EGFR signaling in FAP differentiation, we treated the cells with EGF and induced their adipogenesis differentiation (for 8 days in adipogenic medium) or fibrogenic differentiation (for 1 day in the presence of TGFβ). We found that EGF, from 1 to 100 ng/ml, did not change the adipogenic differentiation of WT FAPs whereas D2-mdx adipogenesis was decreased by 46 to 54% without a dose-dependent effect (Fig. 3E, F). EGFR stimulation with EGF at 100 ng/ml had no impact on fibrogenic differentiation in both WT and D2-mdx FAPs (Fig. 3G). These results show that D2-mdx FAPs have a decreased EGFR signaling pathway, the latter being an inhibitor of adipogenesis. This suggests that a lower EGFR signaling in D2-mdx FAPs is associated with an increased capacity to form adipocytes.

Promoting EGFR signaling decreases fat deposition in vivo and improves muscle function

To establish the physiological relevance of the anti-adipogenic effect of EGF on FAPs, we electroporated the gastrocnemius muscle of 12-week-old D2-mdx mice with a plasmid encoding for EGF [26] (Fig. 4A). Four weeks later, the treatment led to an 33% increase of the presence of P-EGFR in FAPs in the EGF electroporated muscle (vs. empty vector) (Fig. 4B). The muscles that have been electroporated with the EGF expression plasmid showed a 68% reduction in fat deposition as compared with muscles that have been electroporated with an empty plasmid (0.2 vs. 0.65%, respectively) (Fig. 4C). Moreover, the collagen I-positive area was also reduced by 51% in the treated animals (15 vs. 32% in the control) (Fig. 4D). Torque production was measured at increasing frequencies and was found to be increased by EGF expression in the D2-mdx muscle between 75 Hz and 150 Hz (by 28% at 100 Hz) (Fig. 4E). These results show that EGF reduces both adipogenesis and fibrosis, via EGFR signaling, thereby improving muscle function.

Fig. 4.

EGFR signaling and adipogenesis in vivo. 12-week-old D2-mdx mice gastrocnemius muscles were electroporated with empty or EGF expression plasmids and muscles were harvested 4 weeks later. A Two image sets of representative examples of collagen I and perilipin immunostaining for each condition. Nuclei were stained with Hoechst. B The number of FAPs (PDGFR⍺^pos) positive for P-EGFR was quantified. C Fat deposit areas, as perilipin positive were quantified as % of total muscle section. D Fibrotic areas as collagen I positive were quantified as % of total muscle section. E Torque produced by the plantar flexor muscles recorded at incremental stimulation frequencies. Results are means +/- SEM of 3 (A-D) and 6 (E) independent experiments. * p < 0.05 ** p < 0.01 *** p < 0.001 using unpaired t-test (B, D) and using two-way ANOVA and Holm-Sidak test for multiple comparisons (D)

Discussion

Here, we investigated the role of EGFR signaling in adipogenesis in FAPs in the DMD context. Although the B10/B6-mdx DMD mouse model shows no fibrosis in hindlimb muscles [1], the D2-mdx model is an interesting model for studying fibrosis within the DMD framework. At 6/7 months of age, about 20% of the muscle area is occupied by collagen material [4, 35]. Another hallmark of muscle deterioration in DMD patients is the appearance of fat deposits, which replace myofibers alongside fibrosis and accelerate muscle loss [36]. No adiposis is observed in B10/B6-mdx muscle; however, up to 2% of the muscle area in 6-month-old D2-mdx hindlimb muscles is stained by the lipid marker Oil Red O [4]. We investigated the appearance of fat deposition from 10 weeks of age, when fibrosis is already present in the gastrocnemius muscle [8] and associated with impaired local myogenesis by MuSCs [37]. The presence of both fibrosis and adipose deposition in the same areas, increasing from 10- to 18-week-old animals, suggests that the two processes are linked in vivo. Both fibrogenesis and adipogenesis are driven by FAPs in skeletal muscle [38, 39]; however, knowledge of how FAPs enter one program or the other is limited. Only a few in vitro studies have reported the, more often antagonistic, impact of effectors on adipogenesis versus fibrogenesis including TGFβ, BMP7 [19], ADAMTSL2 [16] or coculture with myogenic cells [40].

FAP properties in DMD

We found that FAPs isolated from D2-mdx mice expand more efficiently and form adipocytes more efficiently than FAPs derived from WT mice. Similar findings have been reported for FAPs derived from B10-mdx muscle [23, 41]. Although their molecular interactions have yet to be identified, macrophages and FAPs are in close proximity in regenerating muscle, including in the DMD context [42–45]. In mdx muscle, macrophages exhibit a prominent signature characterized by the high expression of fibrotic factors, such as galectin-3 and osteopontin [46]. Galectin-3, which is primarily expressed by immune cells, promotes adipogenesis in regenerating skeletal muscle in an high-fat-diet context [47]. Galectin 3-positive macrophages stimulate FAPs to express collagen I [46]. In mdx muscle deficient for osteopontin in macrophages, subsets of FAPs associated with adipogenesis gene expression were reduced. This suggests a direct involvement of macrophage-derived osteopontin in promoting adipogenesis [48]. Here, we showed that coculturing FAPs with macrophages derived from D2-mdx muscles induces adipogenesis in WT FAPs, but not in D2-mdx FAPs. One explanation would be that in D2-mdx muscle, FAPs are already activated and do not further respond to macrophage-derived cues.

EGFR in skeletal muscle

The EGFR signaling pathway has been shown to be involved in skeletal muscle regeneration, particularly through its effects on MuSCs. Indeed, EGF stimulates myoblast and MuSC proliferation and differentiation [49, 50], Tenascin-C activates EGFR via its EGF-like domain to promote MuSC proliferation [51, 52], and Amphiregulin, another EGFR ligand, stimulates MuSC differentiation [53]. The EGFR/Aurora kinase A axis controls the EGF-stimulated asymmetric division, which is absent in the mdx MuSCs [26]. Conversely, the impact of EGFR signaling on FAPs has been poorly investigated. One study indicates that EGF promotes FAP proliferation in vitro [54]. Here we found a decrease in EGFR signaling in D2-mdx FAPs vs. WT FAPs. Since EGFR was also expressed by MuSCs and tenocytes, the benefit of EGF electroporation on D2-mdx muscle homeostasis and function may also account for an effect on these two cell types, notably on MuSCs, through a stimulation of their proliferation and differentiation. Indeed, P-EGFR immunolabeling on the sections of electroporated muscle showed non-FAP positive cells, indicative of EGFR signaling in other cell types.

EGFR and adipogenesis

Similarly, the effects of EGFR signaling on adipogenesis have only been studied using the 3T3-L1 preadipocyte cell line. One study found that EGF enhances adipogenesis in a dose- and time-dependent manner in differentiated cells but not in preadipocytes [55]. At concentrations above 10 nM (like in the present study), EGF inhibits preadipocyte differentiation via EGFR dimerization. However, at lower concentrations, EGF supports adipocyte differentiation via the EGFR/EB2R heterodimer [56]. Furthermore, the anti-adipogenic effect of evodiamine occurs through the phosphorylation of EGFR (and its downstream targets PKCalpha and ERK). Similarly, we found that adding EGF induces an anti-adipogenic effect in FAPs, particularly in D2-mdx FAPs, after 8 days in an adipogenic medium. In short-time experiments, we observed reduced EGFR signaling in D2-mdx FAPs, as compared with WT FAPs. These results suggest that the EGFR pathway is decreased in FAPs in situ in the DMD muscle, leading to increased adipogenesis. This was reversed by overexpressing EGF in vivo, which decreased adipogenesis and increased muscle force in the D2-mdx muscle.

EGFR and fibrosis

The EGFR signaling pathway has been associated with fibrosis in various contexts. Essentially, the EGFR signaling pathway is associated with promoting fibrosis in various organs including the lungs [57], kidney [58] and skin [59]. However, we did not observe any effect of EGFR signaling on FAP conversion into fibroblasts in vitro. One limitation of this experiment is that the FAPs were already highly differentiated in this assay. Moreover, our in vivo results show that overexpressing EGF, therefore stimulating the EGFR pathway, reduced the fibrotic area. While the involvement of the EGFR pathway in fibrosis is established in various contexts, particularly its cross-activation and/or synergy with TGFβ [60–63], the regulation of the EGFR pathway is highly complex. It includes transactivation via neighboring receptors, notably via G protein-coupled receptors (GPCR) [64]. Recently, the process of EGFR transinhibition has emerged [65], adding a layer of complexity to its regulation. For example, some GPCR (GPR27) can dephosphorylate EGFR [66]. Additionally, some enzymes, such as glucuronyl C5-epimerase (which catalyzes the biosynthesis of heparan sulfate) can inactivate EGFR signaling, impeding TGF-β signaling pathway, leading to reduced renal fibrosis [67]. TGF-β is highly present in the D2-mdx skeletal muscle; thus, further investigations are required to explain the underlying mechanisms of anti-fibrotic EGF overexpression in DMD muscle.

Fibrogenesis versus adipogenesis

Our histology results show that fat deposits are exclusively localized within fibrosis areas, suggesting a link between the two differentiation processes. A still-unresolved question is understanding the molecular mechanisms controlling the commitment of FAPs into differentiation (adipocyte or fibroblast). A recent study has shown that the senescence induced by muscle injury has a beneficial paracrine effect on cell reprogramming, particularly in MuSCs, to promote regeneration [68]. It has been demonstrated that the EGFR ligand Amphiregulin is involved in this cellular reprogramming in skeletal muscle in vivo [69]. Notably, this study examined MuSC reprogramming in vivo and showed in vitro that amphiregulin can also reprogram mouse embryonic fibroblasts, which are mesoderm-derived cells like FAPs. Another interesting aspect of FAP homeostasis is the presence of a FAP subset in mdx muscle, that exhibits high Sca-1 expression and a high proliferation rate. This subset differentiates easily into adipocytes whereas Sca-1^low FAPs are more likely to differentiate into fibroblasts [70]. Although phospho-EGFR expression does not correlate with Sca-1 expression levels [70], the response to the EGFR ligands in both mdx FAP subsets could provide insight into the impact of EGFR signaling on FAP regulation in the DMD muscle.

Limitations of the study

The results in the present study do not fully explain the beneficial effects of the EGF overexpression in the d2-mdx muscle. In particular, the EGF-driven decrease of fibrosis observed in vivo is not sustained by a direct effect on cultured FAPs under TGFβ treatment. Moreover, the decreased EGFR signaling in FAPs from DMD muscle requires further investigation to identify the molecular process at work, which does not rely on a strong decreased expression of the receptor by the cells.

Conclusion

The present work provides a new line of FAP biology in the DMD context, and a new anti-adipogenic function of the EGFR signaling in FAPs.

Authors’ contributions

JGu: Investigation, Methodology, Software, Visualization, Writing – original draft, Writing – review and editing. GP: Investigation, Writing – review and editing. LD: Formal analysis, Writing – review and editing. WJ: Formal analysis, Writing – review and editing. RM: Investigation, Resources, Writing – review and editing. JGo: Investigation, Resources, Validation, Writing – review and editing. SG: Investigation, Methodology, Writing – original draft, Writing – review and editing. BC: Conceptualization, Funding acquisition, Project administration, Resources, Supervision, Validation, Visualization, Writing – original draft, Writing – review and editing.

Funding

This study was supported by AFM-Telethon (MyoNeurALP 2 Alliance) and by Agence Nationale de la Recherche (ANR-19-CE14-0008 and ANR-19-RAR4-0015).

Data availability

The datasets analyzed during the current study are available in the GEO repository, at [https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi? acc=GSE305185](https:/www.ncbi.nlm.nih.gov/geo/query/acc.cgi? acc=GSE305185) and [https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi? acc=GSE297388](https:/www.ncbi.nlm.nih.gov/geo/query/acc.cgi? acc=GSE297388) . All other data used during the current study are available from the corresponding author on reasonable request.

Declarations

Ethics approval and consent to participate

The protocols used in this study have been approved by the local ethical committee and French Ministry of Research #10463-2017062617107339.

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.