Early–disease stage muscle histopathology of mild and severe dystrophic muscle.

We noted that D2-mdx mouse muscles exhibited severe alterations at the macroscopic level as early as 3–4 weeks of age, where the triceps and pectoral muscles showed clearly visible whitened muscle bundles along their length, which continued to worsen even beyond 5 weeks of age (Figure 2A). In contrast, such gross abnormalities were not detected in any of the age-matched B10-mdx mice (Figure 2A). Based on our previous observations of heightened fibrosis and inflammation in the D2-mdx model (13), we first examined collagen deposition in muscles at this early disease stage. We monitored the presence of fibrous collagen in triceps of 38-day-old D2-mdx mice using Masson’s trichrome stain in conjunction with second harmonic generation microscopy (Figure 2, B and C). Both approaches clearly identified deposition of fibrous collagen within the fascicles of D2-mdx muscles (Figure 2, B and C). Additionally, using alizarin red staining, we determined that the macroscopic muscle whitening in D2-mdx mice was due to muscle degeneration where myofibers were lost to calcification (Figure 2D). This was not seen at 14 days of age but became apparent at 21 days of age in D2-mdx muscle, but not in B10-mdx muscles of the same age (Figure 2, D and E). From 21 to 38 days of age, the areas of calcification increased cumulatively in D2-mdx muscles, doubling from under 5% at 21 days to covering over 10% of the entire muscle cross-sectional area by 38 days (Figure 2F; and Supplemental Figure 2). In contrast, B10-mdx muscles showed relatively small regions of myonecrosis without the presence of calcification even at 38 days of age compared with D2-mdx (Figure 2, E and F; and Supplemental Figure 2). Interestingly, by 38 days of age, myofiber regeneration, as indicated by the presence of CNFs, was over 3-fold higher in B10-mdx (>40% of all fibers) as compared with approximately 12% in D2-mdx (Figure 2G). Thus, at the onset of spontaneous muscle degeneration at approximately 3 weeks of age, the D2-mdx muscle is not accompanied by the vigorous regeneration seen in the B10-mdx mouse, and necrotic spaces are replaced instead by patchy fibrosis and extensive calcification.

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Figure 2

Muscles from D2-mdx mice show evidence of muscle degeneration and calcification starting at the onset of disease.

(A) Macroscopic images of 14-, 21-, and 38-day-old D2-mdx mice, 38-day-old D2-WT mice, and 38-day-old B10-mdx mice. D2-mdx mice show development of conspicuous pale stripes in the pectoral muscles between 21 and 38 days of age accompanied by similar changes in other muscles (e.g., triceps, black arrowhead) corresponding to areas of calcification identified in sections by alizarin red staining (D). No abnormal changes were seen in either D2-WT or B10-mdx mice (white arrowhead). (B and C) Signs of endomysial fibrosis in and around areas of degeneration in the triceps muscle were identified by second harmonic generation (B) and trichrome stain (C) in muscles of D2-mdx mice at 38 days of age. (D and E) H&E and alizarin red staining of triceps muscle sections from D2-mdx and B10-mdx mice at 14, 21, and 38 days of age. (F and G) Quantification of the percentage of calcified area and centrally nucleated fibers (CNFs) in triceps muscles from B10-mdx and D2-mdx at 21 days and 38 days old. (F) Quantification of percentage of calcification from 21-day-old D2-mdx mice (n = 4), 38-day-old D2-mdx mice (n = 7), and 38-day-old B10-mdx mice (n = 4). (G) Quantification of CNF percentage from 21-day-old D2-mdx mice (n = 4), 38-day-old D2-mdx mice (n = 8), and 38-day-old B10-mdx mice (n = 4). For F and G, data are expressed as mean ± SD. One-way ANOVA with Tukey’s post hoc comparison. **P < 0.01; ***P < 0.001; ****P < 0.0001. Scale bars: 50 μm (B and C), 20 μm (D and E).

Failed myogenesis and FAP osteogenesis mark spontaneous muscle degeneration in young D2-mdx mice.

To determine whether the low frequency of CNFs in D2-mdx muscles are indicative of selective loss of regenerating myofibers or a failure of satellite cell function, we first evaluated the expression of myogenic transcriptional factors, i.e., paired box 7 (Pax7), myoblast determination protein 1 (MyoD), and myogenin, at 2 ages around the onset of pathogenesis in the mild (B10-mdx) and severe (D2-mdx) models (Figure 3, A–C). Although there were no significant differences in gene expression at 21 days of age between these models, expression of Pax7, MyoD, and myogenin were all significantly lower in D2-mdx muscles at 38 days of age than in B10-mdx muscles (Figure 3, A–C).

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Figure 3

Satellite cells from D2-mdx mice show limited myogenic capacity during spontaneous muscle regeneration.

(A–C) Gene expression of Pax7, MyoD, and myogenin from 21- and 38-day-old D2-WT (n = 3 per age), D2-mdx (n = 4 per age), and B10-mdx (n = 3 per age) mice. (D) Immunofluorescence (IF) images stained for labeling Pax7⁺ satellite cells in intact areas of the triceps muscle from 38-day-old B10-mdx and D2-mdx mice. White arrowheads, Pax7⁺ cells. (E) Quantification of the total number of Pax7⁺ cells per area (mm²) in B10-mdx (n = 8) and D2-mdx (n = 8) muscles. (F) IF images stained for labeling Pax7⁺ satellite cells in damaged areas of the triceps muscle from 38-day-old B10-mdx and D2-mdx mice. White arrowheads, Pax7⁺ cells. (G) Quantification of the total number of Pax7⁺ cells per area (mm²) in muscles from B10-mdx (n = 6) and D2-mdx (n = 9). (H) Schematic figure demonstrating the pattern for pulsed 5′-bromo-2′-deoxyuridine (BrdU) labeling of proliferating satellite cells. (I) Quantification of BrdU⁺ CNFs relative to total fibers per muscle section (n = 3 for each labeling period per genotype). (J) IF images from the triceps muscle from 38-day-old B10-mdx and D2-mdx mice labeled with BrdU for 9 days. White arrowheads, BrdU⁺ CNFs. (K) Quantification of the total number of BrdU⁺ CNFs after 6–9 days of BrdU labeling in muscle sections from 38-day-old B10-mdx (n = 6) and D2-mdx (n = 6) mice. For A, B, and C, *P < 0.05, ***P < 0.001, and ****P < 0.000; 1-way ANOVA with Tukey’s post hoc comparison; for E, G, I, and K, **P < 0.01, ***P < 0.001, and ****P < 0.0001; 2-tailed, nonparametric Mann-Whitney U test. Scale bars: 50 µm (D, F, and J).

Simple visual inspection of muscle cross sections showed a clear difference between 38-day-old B10-mdx and D2-mdx mice regarding the way that areas of muscle fiber damage were resolved. In B10-mdx mice there was clear evidence of rapid myofiber regeneration with a high incidence of small CNFs with occasional inflammatory cells and damaged myofibers (Figure 2D, Supplemental Figure 2A). In contrast, areas of damage in D2-mdx muscles did not resolve in this way and remained largely devoid of regenerated myofibers, the space being occupied by inflammatory cells, acellular debris, and calcified areas between the muscle fibers (Figure 2E, Supplemental Figure 2B). The areas in each muscle that were free of inflammatory cells, damaged myofibers, and small-caliber CNFs were classified as “intact” while areas characterized by the presence of 1 or more of these features were classified as “damaged” (Figure 3). In both strains, the damaged areas were readily distinguished from intact areas because the latter contained closely packed polygonal fibers, with peripherally placed myonuclei and few inflammatory cells (Figure 3). Intact muscle areas contained similar numbers of Pax7⁺ cells in B10-mdx and D2-mdx (Figure 3, D and E), while damaged areas contained 7-fold fewer Pax7⁺ cells in D2-mdx than B10-mdx muscles (Figure 3, E and G).

To further analyze satellite cell function in vivo, we used the thymidine analog BrdU to label recently generated myofiber nuclei. We labeled proliferating myogenic cells by administering BrdU for 3, 6, or 9 consecutive days starting at 25 days of age and counted those myofibers that contained a BrdU-labeled central nucleus as an indicator of the frequency of myoblasts that had proliferated during that labeling period and subsequently fused into regenerating myofibers (Figure 3, H and I). B10-mdx mice in the 9-day labeled group displayed a progressive increase in the proportion of BrdU-labeled fibers, reaching up to 20% of the total fibers. In contrast, only 5% of total fibers in D2-mdx muscles were labeled with BrdU after the same 9-day labeling period (Figure 3I). Quantification of BrdU-labeled CNFs indicated 6-fold more muscle regeneration in the B10-mdx than in D2-mdx muscles (Figure 3, J and K). Collectively, these data indicate that the phenotypic differences between the 2 models at the onset of disease are largely attributable to substantial muscle damage in both but with a markedly limited regeneration in D2-mdx mice.

Myonecrotic muscle diseases, such as DMD and limb girdle muscle dystrophy 2B, feature chronic overactivation of FAPs, marked by platelet-derived growth factor receptor–α (PDGFRα), which contributes to disease pathology by promoting fibrosis and adipogenesis (35, 36). We thus investigated the involvement of these mechanisms in the prominent muscle degeneration in D2-mdx muscle. Immunostaining revealed FAPs in intact and spontaneously damaged areas of triceps muscles in both B10-mdx and D2-mdx mice (Figure 4, A and B; and Supplemental Figure 3). However, FAP-containing areas in D2-mdx mice were more extensive than in B10-mdx mice (Figure 4, A–C; and Supplemental Figure 3).

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Figure 4

Increased FAP proliferation is responsible for early calcification in muscles from D2-mdx mice.

(A and B) IF images stained for PDGFRα⁺ cells in intact (A) and damaged (B) areas in triceps from 38-day-old B10-mdx and D2-mdx mice. Note the higher accumulation of PDGFRα⁺ cells in muscles from D2-mdx mice, showing increased presence of FAPs. (C) Quantification of PDGFRα⁺ areas is shown as a percentage of the total triceps muscle section area from B10-mdx (n = 5) and D2-mdx mice (n = 6). (D) Alkaline phosphatase (ALP) staining of primary FAPs isolated from B10-mdx and D2-mdx muscle after 11 days in culture. (E) Quantification of ALP⁺ area as a measure of osteogenic differentiation of B10-mdx and D2-mdx FAPs (n = 3 replicates/genotype). (F and G) Images and quantification of in vitro FAP osteogenesis following stimulation with 25 ng/mL recombinant BMP2. For C, *P < 0.05; 2-tailed, nonparametric Mann-Whitney U test. For E and G, ***P < 0.001; ****P < 0.0001; 2-way ANOVA, Holm-Šídák’s multiple-comparisons test. Scale bars: 50 μm (A and B), 100 μm (D and F).

The conspicuous calcified degeneration in D2-mdx and the known role of FAPs in heterotopic ossification of muscles (37) prompted us to investigate whether increased disease severity in the D2-mdx FAPs was associated with pathways favoring ossification, indicated by increased alkaline phosphatase (ALP) activity. We found that primary PDGFRα⁺ FAPs isolated from the 28-day-old D2-mdx muscle underwent extensive spontaneous osteogenic differentiation in vitro (Figure 4, D and E), marked by positive staining for ALP. In contrast to FAPs isolated from D2-mdx, FAPs from B10-mdx showed extremely low levels of spontaneous osteogenesis, suggesting that a difference in the cellular niche drives FAP proliferation and adoption of an osteogenic fate as early as 28 days of age. Bone morphogenetic protein 2 (BMP2) has been shown to induce FAP osteogenesis (37). Thus, next we examined D2-mdx FAPs for enhanced sensitivity for osteogenic differentiation by BMP2. Again, D2-mdx FAPs showed increased osteogenesis in response to BMP2 induction, while B10-mdx FAPs were unresponsive to osteogenic differentiation even in response to BMP2 stimulation (Figure 4, F and G). Together, the above results indicate that the severe muscle pathology in young D2-mdx mice is attributable to a combination of conspicuous muscle degeneration and failed regenerative response of satellite cells in the context of increased FAP accumulation and osteogenesis.

Injury triggers poor regeneration and excessive degeneration of muscle of D2-mdx mice.

Our observation of limited regenerative capacity in D2-mdx muscles in response to spontaneous damage (Figure 3) prompted us to examine whether the deficit in regenerative competence is a more generalized feature of satellite cells in this model. Accordingly, we induced muscle damage in D2-mdx mice, choosing the tibialis anterior (TA) muscle, in which spontaneous damage is limited at this age. Following a single injection of the myotoxin notexin (NTX), at 24 days of age, we monitored myogenic response by BrdU labeling on successive single days, starting 1day postinjury and continuing daily up to 6 days postinjury. Animals were euthanized 3 days after the end of BrdU administration to allow full nuclear incorporation into myofibers and help delineate the timing and extent of satellite cell proliferation responsible for regeneration of these damaged muscles. Just 1 day following NTX injury (i.e., 1-day BrdU labeling period; Figure 5, A and B), over 30% of fibers contained a BrdU⁺ myonucleus in the NTX-injured B10-mdx mice, while only 6% of total fibers were BrdU⁺ in NTX-injured D2-mdx muscles (Figure 5B). In muscles receiving BrdU at 2 days postinjury, 13% of fibers were BrdU⁺ in B10-mdx muscles, while only 5% of D2-mdx fibers were BrdU⁺. At the 3-day time point, the level of labeling in NTX-injured D2-mdx muscles was indistinguishable from the contralateral muscle, where it probably reflects the low background regeneration of spontaneous dystrophic lesions. On subsequent days, BrdU incorporation had returned to the baseline levels seen in noninjured contralateral muscles in both models. These data indicate that the decreased BrdU incorporation in D2-mdx muscle was not the result of a simple delay in myogenesis but a deficit in satellite cell expansion and fusion in D2-mdx muscles following injury, which is consistent with our results for spontaneous damage by the disease process.

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Figure 5

Staggered BrdU labeling after synchronized NTX-induced injury in TA muscles from B10-mdx and D2-mdx mice.

(A) IF images from B10-mdx and D2-mdx NTX-injected TAs labeled with BrdU from 24 to 48 hours postinjury (day 1 labeling). White arrowheads, BrdU⁺ CNFs. (B) Percentage of BrdU⁺ CNFs across the 6-day labeling period after NTX-injury: B10-mdx (n = 5 for day 1; n = 5 for day 2; n = 4 for days 3 and 4; n = 4 for days 5 and 6 post-BrdU) and D2-mdx (n = 4 for day 1; n = 4 for day 2; n = 4 for days 3 and 4; n = 4 for days 5 and 6 post-BrdU). (C) IF images stained for labeling Pax7⁺ cells in NTX-injured TAs from B10-mdx and D2-mdx mice. White arrowheads, Pax7⁺ cells. (D) Quantification of the total number of Pax7⁺ cells per area (mm²) at 5–6 days postinjury in NTX-injured TAs from B10-mdx (n = 7) and D2-mdx (n = 8). (E–G) Percentage of CNFs, calcification, and total area of degeneration from B10-mdx (n = 4 for CTL, and n = 5 for NTX TAs) and D2-mdx (n = 4 for CTL, and n = 5 for NTX TAs). (H–J) Quantification and IF images for PDGFRα⁺ cells 14 days after NTX injury of TA muscles from B10-mdx and D2-mdx mice (injury was performed at 24 days of age). (H) Quantification of PDGFRα⁺ areas shown as a percentage of total muscle section from B10-mdx (n = 6) and D2-mdx mice (n = 5). For B and D–H, *P < 0.05, **P < 0.01, and ***P < 0.001; 2-tailed, nonparametric Mann-Whitney U test (B, D, and H); **P < 0.01, and ****P < 0.0001; 1-way ANOVA with Tukey’s post hoc comparison (E–G). Scale bars: 50 μm.

A count of Pax7⁺ cells was conducted to determine whether the limited myoblast fusion observed following NTX injury of D2-mdx muscle was accompanied by a reduction in the number of satellite cells. The numbers of Pax7⁺ cells were nearly 2-fold lower in D2-mdx muscles than in B10-mdx muscles (Figure 5, C and D), mirroring the satellite cell proliferation and fusion deficit observed in spontaneously degenerating D2-mdx muscles (Figure 3, E and G). To determine whether there was any association between the limited myogenesis in NTX-injured D2-mdx muscles and the amount of muscle damage, we counted CNFs and measured both calcification and total degeneration per cross section of NTX-injured TA muscles (Figure 5, E–G). The percentage of CNFs was 3-fold lower in NTX-injured D2-mdx muscles than in B10-mdx muscles (Figure 5E). Further, the degenerated areas that were calcified were 10-fold greater (Figure 5F) while the total degenerated areas were 3-fold greater in D2-mdx as compared with B10-mdx (Figure 5G). These findings indicate that the lower regenerative myogenesis and greater degeneration in the D2-mdx muscle occur independent of the cause of muscle damage and are inherent deficits in the myogenic maintenance mechanisms of the D2-mdx muscle.

Our earlier observation of FAP expansion in response to spontaneous damage (Figure 4, A–C) prompted us to investigate the underlying degenerative process after NTX injury (Figure 5, H–J; and Supplemental Figure 4). NTX was administered intramuscularly into the TAs of both B10-mdx and D2-mdx, and FAPs were assessed by PDGFRα labeling. At 14 days postinjury, the percentages of muscle area occupied by PDGFRα⁺ cells were 1.5-fold higher in D2-mdx than in B10-mdx muscles (Figure 5H). This increase was even greater in the NTX-damaged regions than in adjacent undamaged regions (Figure 5, I and J; and Supplemental Figure 4). D2-mdx NTX-injured muscles showed strong and persistent PDGFRα staining within the inflamed interstitium that remained in a state of active inflammation even at 14 days postinjury (Figure 5J and Supplemental Figure 4). This implicates dysregulation of FAPs in D2-mdx muscle as a source of prolonged state of degeneration and perhaps accentuation of the myogenic deficit in this model.

Myofiber isolation, staining, and imaging.

EDL muscle fibers were isolated following our previously described protocol (48, 49). EDL muscle was carefully dissected and incubated in 0.2% collagenase type I (Sigma-Aldrich, C0130) for approximately 2 hours. After collagenase digestion, single fibers were isolated by gentle trituration in DMEM (Lonza, 12-604F), transferred to 2-mL Eppendorf vials containing 30% sucrose (Sigma-Aldrich, S0389), and stored at –80°C. On the day of staining, Eppendorf vials were thawed at 4°C, and sucrose solution was removed and replaced with blocking buffer (0.5% Triton X-100, 0.1% Tween 20, 2% BSA, and 20% goat serum in TBS-Tween buffer) and incubated at 4°C overnight. After blocking, fibers were washed and resuspended in/with Alexa Fluor 594–conjugated phalloidin for 20 minutes and left overnight at 4°C in TBS with Tween buffer. On the following day fibers were mounted with ProLong Gold Antifade with DAPI (P36931, Life Technologies).

BrdU labeling.

BrdU (Sigma-Aldrich, B9285) was administered ad libitum in sterile drinking water at a concentration of 0.8 mg/mL and kept protected from light during administration (50). BrdU was administered from 25 to 28 days (3 days), 25 to 31 days (6 days), and 25 to 34 days (9 days); all animals were euthanized at 38 days of age. For single-day BrdU administration after timed NTX-induced injury, mice were administered BrdU in drinking water at specific time intervals — either 24 or 48 hours (day 1 BrdU), 48–72 hours (day 2 BrdU), 72–96 hours (day 3 BrdU), 96–120 hours (day 4 BrdU), 120–144 hours (day 5 BrdU), and 144–168 hours (day 6 BrdU) after injury — and animals were euthanized and tissues harvested 3 days after cessation of BrdU administration. BrdU was administered between 24 and 72 hours afterward in experiments involving IM injections of ITD-1 and NTX.

NTX-induced injury and ITD-1 IM injection.

Animals that were 24 days old were anesthetized with isoflurane, and the anterior portion of their hind limb was shaved before IM injection of 40 μL of NTX (10 μg/mL, Latoxan, L8104) with a 28-gauge insulin syringe (BD Biosciences 329461). The needle was dipped in green tattoo dye (Harvard Apparatus, 72-9384) to track the site of muscle injury. For experiments involving IM injections of ITD-1 and NTX, 2000 ng of ITD-1 (Tocris, 5068) was added to 30 μL of NTX (10 μg/mL, Latoxan, L8104) and injected into the TA muscle; the contralateral TA was injected with 30 μL of NTX only. At 2 and 4 days after the initial IM injection, one TA received injections of 20 μL of ITD-1 (2000 ng) whereas the contralateral TA was injected with an equal volume of saline.

Muscle histology.

Frozen muscles were removed from the freezer and thawed in a Leica CM1950 cryostat at –20°C for 15–20 minutes. The muscles were then sectioned at 8 μm each, mounted on microscope slide, and stained for H&E, alizarin red, and Masson’s trichrome.

Microscopy.

We used Olympus BX61 VS120-S5 Virtual Slide Scanning System with UPlanSApo 40×/0.95 objective, Olympus XM10 monochrome camera, and Olympus VS-ASW FL 2.7 imaging software. Analysis was performed using Olympus CellSens 1.13 and ImageJ software (National Institutes of Health). Two-photon and second harmonic images were collected on an Olympus FVMPE-RS multiphoton laser scanning microscope with a 3W Mai Tai HP Ti:Sapphire laser set at an excitation wavelength of 910 nm. Using a 25×/1.05 NA water immersion objective, the SHG and 2-photon signal were collected using band-pass filters BP410-455 and BP495-540, respectively. Images were acquired with Olympus FV30S-SW software.

Immunofluorescence.

Muscle sections were stained with anti–laminin-α2 (4H8-2, 1:100, Enzo), anti–BrdU-biotin conjugate (B35138, 1:100, Life Technologies), DAPI nuclear stain (ProLong Gold Antifade with DAPI), anti-PDGFRα (D1E1E, 1:250, Cell Signaling Technology), p-SMAD3 (ab52903, 1:200, Abcam), and anti-PAX7 (PAX7 concentrate, 1:20, Developmental Studies Hybridoma Bank, University of Iowa). Unless indicated otherwise, muscle sections were fixed in ice-cold acetone for 10 minutes, washed in PBS (0.1% Tween-20), and blocked for 1 hour in PBS supplemented with 10% goat serum and 10% horse serum (GeneTex), 0.1% Tween-20 (Sigma-Aldrich), and 10 mg/mL BSA (Sigma-Aldrich). Muscle sections were incubated with primary antibodies overnight at 4°C and subsequently probed with Alexa Fluor secondary antibodies, including goat anti–mouse IgG1 Alexa Fluor 488/568 (A-21121/A-21124, Thermo Fisher Scientific), goat anti–rat IgG (H+L) Alexa Fluor 488/647 (A-11006/A-21247, Thermo Fisher Scientific), goat anti–rabbit IgG (H+L) Alexa Fluor 488/594 (A-11008/A-11012, Thermo Fisher Scientific), and streptavidin Alexa Fluor 488/568 (S32354/S11226, Thermo Fisher Scientific). Sections were mounted with ProLong Gold Antifade with DAPI for nuclear staining. For Pax7 immunostaining, sections were instead fixed in ice-cold 4% PFA (Electron Microscopy Sciences, 15710), permeabilized in 0.3% Triton X-100 in PBS, and blocked overnight with mouse on mouse blocking reagent (Vector Laboratories, MKB-2213). Sections were washed in 0.05% Triton X-100 in PBS, then blocked for 30 minutes in 0.05% Triton X-100 in PBS supplemented with 10% goat serum and 10 mg/mL fat-free milk. After blocking, sections were incubated overnight at 4°C in a humidified chamber with Pax7 primary antibody. Sections were then incubated with the appropriate antibodies as described above. For BrdU immunostaining, sections were fixed in ice-cold acetone for 10 minutes, incubated in 2N HCl at 37°C for 30 minutes, and briefly neutralized with 0.15 M sodium tetraborate (Sigma-Aldrich).

TGF-β1 ELISA.

The amount of active TGF-β1 in the triceps muscle was determined using the Quantikine ELISA mouse TGF-β1 immunoassay (R&D Systems, Bio-Techne, MB100B) according to the manufacturer’s recommendations. Final values were normalized to total protein concentration.

Gene expression.

Triceps muscles from 21- and 38-day-old B10-WT, B10-mdx, D2-WT, and D2-mdx mice were used to perform gene expression analysis. Specific mRNAs were quantified using individual TaqMan assays specific for each mRNA according to the manufacturer’s protocol (Pax7, assay ID: Mm01354484_m1; myogenin, assay ID: Mm00446194_m1; and MyoD, assay ID: Mm00440387_m1). Total RNA was converted to cDNA using Random Hexamers and High Capacity cDNA Reverse Transcription Kit (Thermo Fisher Scientific, 4368814). The mRNAs were then quantified using individual TaqMan assays on an ABI QuantStudio 7 Real-Time PCR machine (Applied Biosystems) using TaqMan Fast Advanced MasterMix (Thermo Fisher Scientific, 4444556).

Isolation and in vitro osteogenesis of FAPs.

FAPs were isolated from the hind limb muscles of 4-week-old B10-mdx and D2-mdx mice according to previously published protocols (35). Mice were euthanized and the TA, EDL, gastrocnemius, soleus, quadriceps, and psoas muscles were immediately dissected. Muscles were minced finely in a sterile dish and incubated in collagenase type II (2.5 U/mL, Thermo Fisher Scientific, 17101015) in PBS for 30 minutes at 37°C. The resulting slurry was washed with sterile PBS before further digestion in Collagenase D (1.5 U/mL, Sigma-Aldrich, COLLD-RO) and Dispase II (2.4 U/mL, Sigma-Aldrich, D4693) in PBS for 60 minutes at 37°C. Resulting slurries were passed through 100-μm and 40-μm strainers, and primary cells were resuspended in 1 mL PBS with 2% FBS and 2 nM EDTA. The primary cell suspension was stained with anti–PDGFRα-APC (1.0 μg per 106 cells in 100 μL, BioLegend, 135908) and isotype control (1.0 μg per 106 cells in 100 μL, BioLegend, 400512) for FACS. Cells stained with isotype control were used as a control for gating positive and negative events. Cells stained with anti–PDGFRα-APC were then sorted on an Influx cell sorter (BD, 646500) using 2-way sorting to enrich for positive PDGFRα-expressing cells for further analysis.

Freshly isolated primary PDGFRα⁺ population (FAPs) were plated in Matrigel-coated Nunc Lab-Tek chamber slides (Thermo Fisher Scientific, 154534) at a density of 10,000 cells/well, with 3 replicates per condition. Cells were cultured in DMEM (Lonza, 12-604F) supplemented with 20% fetal bovine serum, 1% penicillin-streptomycin, and 2.5 ng/mL basic FGF (BioLegend, 579604) for 3 days. Osteogenic differentiation was induced by exposure to DMEM with 5% FBS, 1% penicillin-streptomycin, and 25 ng/mL recombinant BMP2 (R&D Systems, Bio-Techne, 355-BM) for 3 days. Following this, cells were cultured in growth medium (DMEM with 5% FBS and 1% penicillin-streptomycin) for a further 4 days. Uninduced cells were not exposed to the osteogenic differentiation medium but instead cultured for 7 days in the growth medium, with the medium changed after 3 days. At the beginning of day 11, cells were fixed in chilled 4% PFA before staining with Alkaline Phosphatase Detection Kit (Sigma-Aldrich, SCR004) to mark cells undergoing osteogenic differentiation.

Statistics.

GraphPad Prism 7.03 was used for all statistical analyses of data. Statistical analysis was performed using the following: linear regression models clustered by mouse group with Tukey’s post hoc multiple-comparisons adjustment, 1-way ANOVA with Tukey’s post hoc comparison; 2-tailed, nonparametric Mann-Whitney U test; or 2-way ANOVA, Holm-Šídák’s multiple-comparisons test. The precise statistical test used varied depending on the nature of the analysis and is listed in the figure legend for each figure. Data normality was assessed for all statistical comparisons. All P values less than 0.05 were considered significant. *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001. Data represent mean ± SD.

We thank Dr. Shin’ichi Takeda of the National Center of Neurology and Psychiatry in Tokyo, Japan, for providing B6-mdx mice (47). We acknowledge Shivaprasad Bhuvanendran of Children’s National Hospital for imaging muscle sample using second harmonic generation. Funding for this project was obtained by DAGM, JSN, JKJ, and TAP. This work was supported by the National Institutes of Health Institute National Institute of Arthritis and Musculoskeletal and Skin Diseases (NIAMS) T32 Genetics and Genomics of Muscle Postdoctoral Training Grant (T32AR056993 to DAGM, JSN, JKJ, and TAP), Foundation to Eradicate Duchenne (to DAGM and JSN), National Institutes of Health NIAMS (R01AR055686 to JKJ), Muscular Dystrophy Association (MDA295203 to TAP, MDA480160 to JSN, MDA 477331 to MWH), Parent Project Muscular Dystrophy (to TAP), and Duchenne Parent Project – Netherlands (to JSN). Microscopy imaging was performed at the Children’s National Medical Center/Children’s Research Institute (CRI) Cellular Imaging Core, which is supported by funds from CRI and the National Institutes of Health Eunice Kennedy Shriver National Institute of Child Health and Human Development (U54HD090257 to JKJ).