CCR2 deficiency does not provide sustained improvement of muscular dystrophy in mdx^5cv mice

Animals

Mdx^5cv were derived from The Jackson Laboratory (Bar Harbor, ME, USA). Ccr2^−/− mice were kindly provided by R.M.R. Mdx^5cv mice were used instead of mdx mice because mdx^5cv mice are in the C57BL/6J background, the same as the Ccr2^−/− mice, while mdx mice are in a different background. Mdx^5cv mice showed muscle pathology comparable to mdx mice (30). mdx^5cv-Ccr2^−/− mice were generated by crossbreeding mdx^5cv mice with Ccr2^−/− mice. ColI-GFP transgenic mice with green fluorescent protein (GFP) expression driven by collagen I (α1) gene promoter to label with high sensitivity and specificity collagen I α1–producing cells were kindly provided by D. Brenner (University of San Diego, La Jolla, CA, USA) (31–33). Mdx^5cv/ColI-GFP and mdx^5cv-Ccr2^−/−/ColI-GFP mice were generated by crossbreeding ColI-GFP with mdx^5cv and mdx^5cv-Ccr2^−/− mice, respectively, for identifying and studying ColI (GFP)⁺ fibroblasts. Our study protocols were approved by the Institutional Animal Care and Use Committee at Icahn School of Medicine at Mount Sinai.

Immunostaining

Frozen muscle sections were fixed with 4% paraformaldehyde for 30 min, blocked in 5% serum for 2 h, and incubated overnight with the primary collagen III antibody (Southern Biotech, Birmingham, AL, USA) or FITC-conjugated mouse anti-IgG antibody (Vector Laboratories, Burlingame, CA, USA) for 2 h. For collagen III immunostaining, the sections were then washed with PBS and incubated with FITC-conjugated secondary antibody for 1 h. After stained with DAPI, slides were mounted with mounting medium, and antibody binding was visualized under a fluorescent microscope.

Quantification of muscle damage

Muscle damage was quantified by measuring the number and calculating the percentage of IgG⁺ necrotic/degenerating fibers. Briefly, 5 sections from each specimen separated by at least 40 μm were immunostained with the anti-IgG antibody, and 4 nonoverlapping areas of each section were digitally captured. The number of IgG⁺ fibers and the total number of muscle fibers were quantified blinded to the genotype. The percentage of IgG⁺ fibers was calculated.

Hydroxyproline assay

Muscles were snap frozen and grounded in liquid nitrogen, followed by homogenization in distilled water. The muscle homogenates were then used for hydroxyproline assay using a kit purchased from Sigma-Aldrich (St. Louis, MO, USA) following the manufacturer’s instructions.

Single-cell suspension preparation from injured muscle, flow cytometry analysis, and cell sorting

Muscle single-cell suspension was prepared by collagenase/dispase digestion. Briefly, each muscle was minced in 2.5 ml of digestion solution containing 1 U/ml of collagenase B and 1 U/ml of dispase II (Roche Diagnostics, Indianapolis, IN, USA) in PBS and incubated at 37°C for 1 h. The reaction was terminated by addition of 10 ml of PBS with 10% of fetal bovine serum. The mixture was then filtered through 70-μm cell strainer and subjected to centrifugation twice at 250 g for 5 min. The pellets were combined, washed with PBS, and centrifuged at 670 g for 10 min. The pallet was resuspended in 3 ml of PBS, filtered through a 40-μm cell strainer, layered on an equal volume of Lymphocyte-M solution (Cedarlane, Burlington, NC, USA), and centrifuged at 2095 g for 45 min. Cells at the interface were collected, centrifuged at 670 g for 10 min, and resuspended in fluorescence-activated cell sorting staining buffer [PBS with 2% of normal mouse serum (Thermo Fisher Scientific, Waltham, MA, USA) and 2% of bovine serum albumin (Sigma-Aldrich)]. The following antibodies were used for flow cytometry. PerCP-Cy5.5-anti-CD45 and PE-Cy7-anti Sca-1 were from BD Biosciences (San Jose, CA, USA). PE-anti-F4/80 and PE-Cy7-anti-Ly6C were from BioLegend (San Diego, CA, USA). PE-anti-CD11b, PE-anti-CD31, and APC-anti-bromodeoxyuridine (BrdU) were from eBioscience (San Diego, CA, USA). PerCP-Cy5.5-anti–α7 integrin was from R&D Systems (Minneapolis, MN, USA). We used Sca-1⁻/CD45⁻/CD31⁻/CD11b⁻/α7⁺ to identify satellite cells and CD45⁻/GFP⁺ to identify fibroblast. All flow analyses were done using LSR II (BD Biosciences, San Jose, CA, USA), and data were analyzed using FlowJo 8.2.6 (Tree Star, Ashland, OR, USA). Cell sorting was performed by the Flow Cytometry Core of the Icahn School of Medicine at Mount Sinai.

Hind-limb grip-strength measurements

Hind-limb grip strength was measured in wild-type (WT), mdx^5cv, mdx^5cv-Ccr2^−/− mice at age 14 wk and 6 mo using an automated grip-strength meter (Columbus Instruments, Columbus, OH, USA) (7). The grip-strength meter provides a validated measurement of motor function. The instrument uses an electronic digital force gauge that measures the peak force exerted on it by the action of the animal. For each time point, we measured grip strength 20 times in each individual mouse and repeated the measurements on a separate day within 3 d, and the average was used as the outcome value.

Respiratory function analysis

Respiratory function in conscious and unrestrained mice was assessed by whole-body noninvasive plethysmography (Buxco Research Systems, Sharon, CT, USA) using Biosytems XA software (7). Unrestrained plethysmograph was calibrated according to the manufacturer’s instructions. The bias flow regulator was calibrated at room temperature and humidity, and the flow was set at 1.2 L/min. Mice were placed in the free moving plethysmograph chamber and allowed to acclimate for 5 min. Then measurements were taken of airway mechanics, including respiratory frequency, tidal volume, minute volume, peak inspiratory flow (PIF), and peak expiratory flow over a 10-min interval. Values measured during the 10-min sequence were then averaged.

Real-time quantitative RT-PCR

Sorted cells were lysed in Trizol reagent (Ambion, Grand Island, NY, USA). Total RNA was then purified and further cleaned up using the RNeasy Micro Kit (Qiagen, Germantown, MD, USA). Reverse transcription was performed using SuperScript II kit (Thermo Fisher Scientific) following the manufacturer’s instructions. The cDNA samples were then subjected to real-time quantitative PCR (qPCR) using SYBR Green reagent and an Eppendorf Realpix4 cycler. The gapdh expression was used as an internal control. Reaction specificity was determined by product melting curves. The PCR products were verified by running 3% agarose gels. Data were analyzed by ΔΔC_t method and presented as fold changes. The primer sequences are listed in Table 1.

BrdU labeling

Mice were injected intraperitoneally with 10 mg/ml BrdU (Sigma-Aldrich) twice a day for 3 d before the experiments. BrdU (0.8 mg/ml) was also added to the drinking water.

Microphage and fibroblast coculture

Intramuscular MPs (CD45⁺F4/80⁺) sorted from diaphragm of 3-mo-old mdx^5cv and mdx^5cv-Ccr2^−/− mice were plated together with NIH3T3 fibroblasts at a ratio of 1:1 in DMEM supplemented with 10% of fetal bovine serum (1 × 10⁵ cells/2 ml medium per well for each cell type). After 48 h of coculture, cells were trypsinized, and microphages were depleted by negative selection using the anti-CD45 MACS system (Miltenyi Biotec, San Diego, CA, USA) following the manufacturer’s instructions. Purified NIH3T3 fibroblasts were then counted and lysed in Trizol reagent. Collagen gene expression by NIH3T3 cells was then analyzed by real-time qPCR using the method previously described.

Statistical analyses

GraphPad (GraphPad Software, La Jolla, CA, USA) was used for statistical analyses. All data were presented as means ± sem. A 2-tailed Student’s t test was used for comparisons between experimental groups. A value of P < 0.05 was considered statistically significant.

CCR2 deficiency markedly reduced Ly6C^hi MPs at all stages but reduced Ly6C^low MPs only at early stages in mdx^5cv-Ccr2^−/− quadriceps and diaphragm muscles

Murine blood MOs mainly consist of 2 subsets, CX3CR1^lowCCR2⁺Ly6C^hi and CX3CR1^hiCCR2⁻Ly6C^low cells (34). CX3CR1^lowCCR2⁺Ly6C^hi cells are inflammatory MOs, which enter tissues in response to injury or infection and differentiate into inflammatory MPs or dendritic cells within inflamed tissues. CX3CR1^hiCCR2⁻Ly6C^low MOs patrol the vascular space and contribute to resident MPs (34, 35). We first addressed whether CCR2 deficiency affected the intramuscular Ly6C^hi and Ly6C^low subsets of MPs. We performed flow cytometry analysis using single-cell suspension of quadriceps and diaphragm muscles from mdx^5cv-Ccr2^−/− mice at 4, 9, and 14 wk, and 6 mo of age, and compared them to mdx^5cv controls (Fig. 1A). The number of F4/80⁺/Ly6C^hi MPs was markedly reduced in the quadriceps and diaphragm muscles of mdx^5cv-Ccr2^−/− mice at all stages compared to mdx^5cv controls (Fig. 1A). The number of F4/80⁺/Ly6C^low MPs was also significantly lower in the quadriceps and diaphragm muscles of mdx^5cv-Ccr2^−/− mice than that in mdx^5cv controls at 4 wk (Fig. 1Aa) and 9 wk (Fig. 1Ab), but it was similar to mdx^5cv controls at 14 wk (Fig. 1Ac) and 6 mo (Fig. 1Ad). Likewise, the total number of intramuscular MPs was also low in mdx^5cv-Ccr2^−/− mice compared to mdx^5cv controls at 4 wk (Fig. 1Aa) and 9 wk (Fig. 1Ab), but similar at 14 wk (Fig. 1Ac) or 6 mo (Fig. 1Ad). The majority (over 90%) of the intramuscular MPs were Ly6C^low in mdx^5cv-Ccr2^−/− mice at all time points studied. To address whether cell proliferation contributed to the increase of the Ly6C^low MP number in mdx^5cv-Ccr2^−/− quadriceps and diaphragm muscles to reach the same level as in mdx^5cv controls at the late stages, we used BrdU to label proliferating cells. There were more BrdU⁺ Ly6C^low MPs (Fig. 1B) in the quadriceps and diaphragm muscles of mdx^5cv-Ccr2^−/− mice than in mdx^5cv controls at 10 wk. Therefore, the expansion of intramuscular Ly6C^low MPs at the late stages could be partly due to the increased cell proliferation.

CCR2 deficiency reduced quadriceps and diaphragm muscle damage at 14 wk but not 6 mo

To address whether less inflammation at the early stages could lead to reduced muscle damage at the late stages, we did IgG staining to identify (Fig. 2A) and quantify (Fig. 2B) degenerating/necrotic muscle fibers at 14 wk and 6 mo. While the percentage of IgG⁺ muscle fibers was reduced in both quadriceps and diaphragm muscles of mdx^5cv-Ccr2^−/− mice at 14 wk compared to mdx^5cv controls, it was not different at 6 mo. Therefore, the muscle fiber damage reduction associated with CCR2 deficiency did not last to an advanced stage.

CCR2 deficiency reduced endomysial fibrosis of mdx^5cv-Ccr2^−/− quadriceps and diaphragm muscles at 14 wk but not 6 mo

Both mdx^5cv quadriceps and diaphragm muscles display mild fibrosis with increased extracellular matrix (ECM) protein deposition at 14 wk. After that point, inflammation gradually subsides in quadriceps with no progressive fibrosis. But mdx^5cv diaphragm shows persistent inflammation and progressive fibrosis. The muscle fibrosis in mdx and mdx^5cv diaphragm is mild at 14 wk and moderately severe at 6 mo (6, 30). We next addressed whether less inflammation and reduced tissue damage at the early stages by CCR2 deficiency could lead to continuous reduction of muscle fibrosis at the late stages. We did collagen III immunostaining to assess endomysial collagen deposition, and hydroxyproline assay to measure collagen synthesis in the quadriceps and diaphragm muscles of mdx^5cv-Ccr2^−/− mice at 14 wk (Fig. 3A, C) and 6 mo (Fig. 3B, D), and compared them to mdx^5cv controls. We also did flow cytometry to quantify and compare the number of fibrogenic effector cells, intramuscular fibroblasts (Fig. 3E). While the hydroxyproline level was reduced in both quadriceps and diaphragm muscles of mdx^5cv-Ccr2^−/− mice at 14 wk compared to mdx^5cv controls (Fig. 3C), no difference was seen at 6 mo (Fig. 3D). The number of fibroblasts was similar in the quadriceps and diaphragm muscles of mdx^5cv-Ccr2^−/− mice compared to mdx^5cv controls at 14 wk (Fig. 3E). Therefore, CCR2 deficiency reduced muscle fibrosis at 14 wk, but the effect did not last to 6 mo.

CCR2 deficiency improved muscle regeneration in diaphragm at 14 wk but not at 6 mo

CCR2 deficiency blocked inflammatory MO/MP infiltration with resultant impaired muscle regeneration after acute skeletal muscle injury (19, 21, 23). But CCR2 deficiency reduced inflammatory MP infiltration, accompanied by improved muscle regeneration in mdx diaphragm at 12 wk (28). We next addressed whether CCR2 deficiency provided sustained improvement of muscle regeneration at the late stages in mdx^5cv-Ccr2^−/− quadriceps and diaphragm muscles. We measured a cross-sectional area (CSA) of muscle fibers and quantified satellite cell numbers in mdx^5cv-Ccr2^−/− quadriceps and diaphragm muscles, and compared them to mdx^5cv controls (Fig. 4). The mean CSA was larger in the mdx^5cv-Ccr2^−/− diaphragm than in mdx^5cv controls at 14 wk (Fig. 4A, C). But the myofiber CSA was not different in the mdx^5cv-Ccr2^−/− quadriceps at 14 wk (Fig. 4A, C) or in the mdx^5cv-Ccr2^−/− diaphragm or quadriceps at 6 mo (Fig. 4B, D). The number of satellite cells was similar in the mdx^5cv-Ccr2^−/− quadriceps and diaphragm muscles at 14 wk compared to mdx^5cv controls (Fig. 4E). Therefore, CCR2 deficiency only provided a transient benefit to the diaphragm muscle regeneration.

CCR2 deficiency improved respiratory function at 14 wk but not 6 mo, and did not improve hind-limb grip strength at 14 wk or 6 mo

CCR2 deficiency reduced muscle damage and improved diaphragm function in mdx mice at 12 wk of age (28). We next addressed whether the functional improvement could be detected mdx^5cv-Ccr2^−/− diaphragm and quadriceps and persisted to the late stages. We measured hind-limb grip strength and respiratory function by plethysmography (7, 14) at 14 wk and 6 mo (Fig. 5). While the hind-limb grip strength in mdx^5cv and mdx^5cv-Ccr2^−/− mice was significantly reduced compared to the WT controls, it was not different between mdx^5cv and mdx^5cv-Ccr2^−/− mice (Fig. 5). PIF, which is the main parameter for gauging the diaphragm function (14), was reduced in mdx^5cv and mdx^5cv-Ccr2^−/− mice compared to WT controls at both time points. PIF was improved in mdx^5cv-Ccr2^−/− mice at 14 wk but not at 6 mo compared to mdx^5cv mice. In keeping with the loss of the beneficial effect of CCR2 deficiency on diaphragm muscle damage and fibrosis at the advanced stage, CCR2 deficiency did not provide a sustained improvement of the diaphragm function either.

CCR2-deficient diaphragm MPs expressed low levels of proinflammatory cytokines but similar levels of profibrotic growth factors compared to mdx^5cv controls

Over 90% of intramuscular MPs were Ly6C^low in mdx^5cv-Ccr2^−/− mice, and the number of Ly6C^low MPs climbed up and reached the same level in mdx^5cv-Ccr2^−/− diaphragm as in mdx^5cv controls at 14 wk (Fig. 1). Because CCR2 deficiency did not lead to a sustained improvement of muscle pathology or function in the diaphragm of mdx^5cv-Ccr2^−/− mice, we next addressed whether Ly6C^low MPs contributed to the progressive diaphragm fibrosis at the late stages in mdx^5cv-Ccr2^−/− mice. To this end, we sorted out MPs from the mdx^5cv-Ccr2^−/− and mdx^5cv diaphragm muscles at 14 wk and compared their gene expression of proinflammatory cytokines (Fig. 6A), alternatively activated MP type 2 (M2) cell markers and cytokines (Fig. 6B), and profibrotic growth factors (Fig. 6C) by real-time qPCR. The gene expression of IFN-γ, IL-1β, and IL-6 was reduced in the CCR2-deficient diaphragm MPs, but the gene expression of TNF-α was not different (Fig. 6A). The CCR2-deficient MPs expressed a low level of CD206, arginase 1, and Ym-1 genes, a similar level of CD68, CD163, and IL-10 genes, and a high level of osteopontin (OPN) gene compared to the CCR2-normal MPs (Fig. 6B). The gene expression of IGF-1 was also increased in the CCR2-deficient MPs, but the gene expression of transforming growth factor β1 (TGF-β1) and platelet-derived growth factor α (PDGF-α) was not different (Fig. 6C). The gene expression of iNOS, PDGF-β, and connective tissue growth factor was low in intramuscular MPs (data not shown). Therefore, the CCR2-deficient MPs appeared less inflammatory, but they expressed a similar level of profibrotic growth factor genes and a high level of IGF-1 and OPN genes.

Activated tissue fibroblasts are the effector cells of fibrogenesis. Thus, we next compared the effector function of intramuscular fibroblasts in mdx^5cv and mdx^5cv-Ccr2^−/− mice. We sorted out the fibroblasts from the diaphragm muscles of mdx^5cv-Ccr2^−/− and mdx^5cv mice at 14 wk and compared their gene expression of profibrogenic growth factors (Fig. 6C) and ECM proteins (Fig. 6D). The gene expression of TGF-β1, PDGF-α, connective tissue growth factor, collagen I, and collagen III was not different, but the gene expression of collagen VI and fibronectin was increased in the CCR2-deficient diaphragm fibroblasts (Fig. 6C, D). Therefore, the CCR2-deficient diaphragm fibroblasts in mdx^5cv-Ccr2^−/− mice at 14 wk exerted a slightly high fibrogenic function compared to mdx^5cv controls in terms of the ECM production. To further address whether the diaphragm MPs from mdx^5cv-Ccr2^−/− and mdx^5cv mice at 14 wk can promote the fibrogenic function of fibroblasts, we performed coculture experiments using intramuscular diaphragm MPs and NIH3T3 fibroblasts (Fig. 6E). The diaphragm MPs from both mdx^5cv and mdx^5cv-Ccr2^−/− mice stimulated the gene expression of collagens in the 3T3 fibroblasts, and the fibroblast expression was low in collagen I, the same in collagen III, and high in collagen VI when cultured with the mdx^5cv-Ccr2^−/− diaphragm MPs compared to the mdx^5cv controls.