Duchenne muscular dystrophy (DMD) is a progressive muscle-wasting disease that causes respiratory and cardiac failure. Inflammation is a key pathological characteristic of dystrophic muscle lesion formation, but its role and regulation in the disease time course has not been sufficiently examined. In the present study, we used IL-10−/−/mdx mice lacking both dystrophin and the anti-inflammatory cytokine, interleukin-10 (IL-10), to investigate whether a predisposition to inflammation affects the severity of DMD with advancing age. The IL-10 deficiency caused a profound DMD phenotype in the dystrophic heart such as muscle degeneration and extensive myofiber loss, but the limb muscle and diaphragm morphology of IL-10−//mdx mice was similar to that of mdx mice. Extensive infiltrates of pro-inflammatory M1 macrophages in regeneration of cardiotoxin-injured muscle, altered M1/M2 macrophage phenotype and increased pro-inflammatory cytokines/chemokines production were observed in the diaphragm and heart of IL-10−/−/mdx mice. We characterized the IL-10−/−/mdx mice as a dystrophic model with chronic inflammation and severe cardiorespiratory dysfunction, as evidenced by decreased percent fractional shortening (%FS) and ejection fraction percent (EF%) on echocardiography, reduced lower tidal volume on whole-body plethysmography. This study suggests that a predisposition to inflammation is an important indicator of DMD disease progression. Therefore, the development of anti-inflammatory strategies may help in slowing down the cardiorespiratory dysfunction on DMD.

INTRODUCTION

Duchenne muscular dystrophy (DMD) is a severe X-linked muscle disease in which mutations in the gene encoding the cytoskeletal protein, dystrophin (1,2). The altered mechanical and signaling functions contribute to membrane fragility, necrosis and inflammation and result in progressive degeneration of striated muscle, manifesting as muscle weakness and, eventually, skeletal muscle atrophy (3). As the disease progresses, wheelchair and ventilatory assistance are required, and patients often succumb to cardiac dysfunction and respiratory failure (4).

Inflammation is a large component of the muscle pathology in DMD. Anti-inflammatory glucocorticoids are widely used to improve muscle strength of DMD patients (57), although the beneficial effects of glucocorticoids vary from patient to patient. Furthermore, long-term as well as short-term steroid treatment induces side effects. Therefore, new and improved therapeutic approaches to the treatment of DMD are needed. Moreover, it is important to determine the effect of chronic inflammation on DMD progression.

Activated immune-cell infiltrates (e.g. T lymphocytes and macrophages) are observed during early disease stages in dystrophic muscle and play critical roles in muscle wasting (813). Depletion or inhibition of these cells significantly improves dystrophic muscle pathology (11,14,15). The findings of these studies suggest that much of the muscle damage in dystrophin deficiency is caused by inflammatory cells as well as by direct mechanical damage, although mechanical injury and membrane defects induced by infiltration of inflammatory cells are also known to promote dystrophic pathology (16,17). Mdx mice are widely used as a model of DMD to evaluate various therapeutic strategies, but the pathological features in these mice are relatively mild compared with human DMD (18). In addition, the detailed role of chronic inflammation in disease progression has not been sufficiently demonstrated in these mice.

Interleukin-10 (IL-10) modulates the inflammatory immune response and reduces M1 macrophage activation and production of pro-inflammatory cytokines, such as IFN-γ, TNF-α, IL-1β and IL-6 in inflamed tissues. Macrophage populations in mdx muscle switch from a pro-inflammatory M1 phenotype to an anti-inflammatory M2 phenotype (19,20). IL-10 plays a particularly important role in mediating this switch of macrophage phenotype in dystrophic muscle, resulting in muscle damage repair (19,20). Although the histological aspects of dystrophic muscle pathology have been revealed, it remains unclear whether inflammation in the dystrophic muscle affects clinical disease such as cardiac and respiratory dysfunction. IL-10 suppresses pro-inflammatory responses in tissues, and low levels of IL-10 expression might impact the severity of inflammatory disease and immunopathology. Indeed, IL-10-deficient mice display several features of inflammatory bowel disease, Crohn's disease (21) and increased susceptibility to helicobacter hepaticus-induced colitis (22). Therefore, we postulated that reduced anti-inflammatory activity causes chronic inflammation and results in more severe dystrophic pathology. To investigate this hypothesis, we generated mice lacking both IL-10 and dystrophin (IL-10−/−/mdx) and examined the cardiac and respiratory dysfunction in mdx muscle.

RESULTS

Muscle histopathology associated with the inflammatory response in IL-10-deficient mdx mice

IL-10−/− mice with C57BL/6 background were crossed with mdx mice with C57BL/6 background to generate IL-10−/−/mdx mice. First, we examined whether the defect in IL-10 affects the histological appearance of dystrophic muscle (Fig. 1). Histopathological analysis of the limb muscle and diaphragm from IL-10−/−/mdx mice showed typical dystrophic pattern including size variation among individual muscle fibers, fiber splitting, small regenerated basophilic fibers, numerous fibers with centrally located myonuclei and significant connective tissue replacement at 8 months of age (Fig. 1, and data not shown), like the muscle morphology of mdx mice. We next investigated the changes in inflammation and severity of dystrophic phenotype in limb muscle. Inflammation is a key response to muscle injury and is critical for muscle regeneration (23). Macrophages that infiltrate the damaged tissue produce pro-inflammatory cytokines and chemokines at early stages, and trophic and anti-inflammatory cytokines at later stages, of the injury/repair process. We analyzed the injury/repair process in IL-10-deficient dystrophic muscle by using cardiotoxin, which is known to damage the plasma membrane of myofibers (24). IL-10−/−/mdx mice showed strong infiltration of CD68+ macrophage in limb muscle (Supplementary Material, Fig. S1A), like previous report (20). In the regenerating stage, the macrophage population shifts toward a M2 phenotype causing attenuation of muscle damage (25). The shift in favor of M2 phenotype was not detectable in IL-10-deficient mice, and continuous M1 activation was observed in post-injured muscle (Supplementary Material, Fig. S1A), associated with a size variation of myofibers with many degenerating and necrotic myofibers (Supplementary Material, Fig. S1B).

Figure 1.

Histological lesion associated with inflammation in dystrophic muscle. H&E staining of TA muscle, diaphragm and heart sections from 8-month-old mdx and IL-10−/−/mdx mice. Bar = 100 μm.

Figure 1.

Histological lesion associated with inflammation in dystrophic muscle. H&E staining of TA muscle, diaphragm and heart sections from 8-month-old mdx and IL-10−/−/mdx mice. Bar = 100 μm.

Extensive macrophage infiltrates and altered M1/M2 macrophage phenotype shift in the diaphragm and heart of IL-10−/−/mdx mice

The diaphragm muscle from IL-10−/−/mdx mice showed a similar amount of necrotic fibers compared with those seen in mdx mice without chemical stimulation (Fig. 1). We used an immunohistochemical approach to determine the degree of infiltrating macrophages displaying M1/M2 (CD68+) or M2 (CD207+) phenotypes. IL-10−/−/mdx mice showed severe cardiac muscle degeneration and extensive myofiber loss with increased cell infiltrations compared with mdx mice (Figs 1 and 2). IL-10−/−/mdx mice showed higher levels of CD68+ macrophage infiltration in the diaphragm and heart muscle than age-matched mdx mice (Fig. 2A and B). The infiltration in the heart of IL-10−/−/mdx mice increased with age, without the alternative activation of M2 (CD207+) phenotypes observed in the mdx heart (Fig. 2B). Our immunohistochemical data also showed that IL-10 ablation in dystrophic muscle increased the level of macrophage infiltration and altered the population of M1 to M2 phenotype.

Figure 2.

Macrophage infiltration and altered M1/M2 phenotype shift in the diaphragm and heart of aged mdx and IL-10−/−/mdx mice. Immunofluorescence staining of (A) diaphragm, and (B) heart tissue samples from 3- and 8-month-old (3 and 8 m) mdx and IL-10−/−/mdx mice with anti-CD68 antibodies (green), anti-CD206 (red) antibodies and corresponding DAPI (blue) staining to identify macrophage (Mφ) populations. Bar = 100 μm.

Figure 2.

Macrophage infiltration and altered M1/M2 phenotype shift in the diaphragm and heart of aged mdx and IL-10−/−/mdx mice. Immunofluorescence staining of (A) diaphragm, and (B) heart tissue samples from 3- and 8-month-old (3 and 8 m) mdx and IL-10−/−/mdx mice with anti-CD68 antibodies (green), anti-CD206 (red) antibodies and corresponding DAPI (blue) staining to identify macrophage (Mφ) populations. Bar = 100 μm.

Increased pro-inflammatory cytokines and chemokines production in the diaphragm and heart of IL-10−/−/mdx mice

Cytokine and chemokine levels in the diaphragm and heart extracts from wild-type (WT), IL-10−/−, mdx and IL-10−/−/mdx mice were determined by multiplex cytokine/chemokine array analysis (Fig. 3 and Supplementary Material, Fig. S2). The cytokines IL-1α, IL-1β, IL-1ra, IL-16 and chemokines such as regulated upon activation normal T cell expressed and presumably secreted (RANTES), macrophage colony stimulating factor (M-CSF), monokine induced by gamma interferon (MIG) and tissue inhibitors of metalloproteinase-1 (TIMP-1) were evaluated in the diaphragm of IL-10−/−/mdx mice at 8 months of age and compared with WT, IL-10−/− and mdx mice. High levels of expressed monocyte chemoattractant protein-1 (JE/MCP-1) were observed in the diaphragm, but not significant different among IL-10−/−, mdx and IL-10−/−/mdx mice. The levels of these cytokines and chemokines are presented in Figure 3C and D. IL-1ra and IL-2 were similarly elevated in the heart from IL-10−/−/mdx mice (Fig. 3B and D). TIMP-1 and JE/MCP-1 were increased in both the heart from mdx, IL-10−/− and IL-10−/−/mdx mice compared with WT. There were no significant differences in the other cytokines and chemokines including IL-1α, IL-1β, RANTES, M-CSF, MIG and IL-6 between the four groups of mice. The array analysis demonstrated increased levels of pro-inflammatory cytokines and chemokines in severely damaged muscle.

Figure 3.

Differential cytokine and chemokine expression patterns in the diaphragm and heart of wild-type, mdx, IL-10−/− and IL-10−/−/mdx mice. Relative expression of cytokines and chemokines in muscle lysates was quantified using the Proteome ProfilerTM Array. Array images of diaphragm (A) and heart (B) (n = 8/group). (C and D) Quantification of representative data presented in (A) and (B).

Figure 3.

Differential cytokine and chemokine expression patterns in the diaphragm and heart of wild-type, mdx, IL-10−/− and IL-10−/−/mdx mice. Relative expression of cytokines and chemokines in muscle lysates was quantified using the Proteome ProfilerTM Array. Array images of diaphragm (A) and heart (B) (n = 8/group). (C and D) Quantification of representative data presented in (A) and (B).

The serum creatine kinase levels, as an index of ongoing muscle membrane instability, were not significantly different between IL-10−/−/mdx and mdx mice at 3 months of age (mean ± SD, n = 5 animals per group, mdx; 6717 ± 1352 IU/l, IL-10−/−/mdx; 5874 ± 564 IU/l, WT; 166 ± 153 IU/l), and at 8 months of age (mdx; 1295 ± 775 IU/l, IL-10−/−/mdx; 1773 ± 443 IU/l).

Elevated fibrosis in dystrophic diaphragm and heart of aged IL-10−/−/mdx mice

Initially, damage in dystrophic muscle is followed by regeneration, but eventually the muscle is replaced by an accumulation of collagen and adipose tissue in the extra cellular matrix (26). The large amount of damaged fibers were replaced by fibrotic connective tissue in an age-related fashion in dystrophic skeletal muscle (26), and in cardiomyopathy (27,28). Therefore, the diaphragm and cardiac muscle sections from WT, mdx, IL-10/− and IL-10/−/mdx mice were stained with sirius-red and Masson's trichrome to examine collagen deposition and distribution (Fig. 4). The staining displayed consistently intense fibrillar collagen in the diaphragm from IL-10−/−/mdx and mdx mice at 8 months of age, while IL-10/− mice showed statistically equivalent to that of WT mice (Fig. 4A). The cardiac sections from IL-10/−/mdx mice showed widely distributed fibrosis compared with mdx mice (Fig. 4B and C). In addition to myocyte disarray described in Figure 1, histological examination showed left and right ventricular (LV, RV) dilatation with those posterior wall thickness and interventricular septal thickness in the heart of IL-10−/−/mdx mice compared with mdx mice (Fig. 4C).

Figure 4.

Extensive fibrosis in diaphragm and pathological diagnosis of dilated cardiomyopathy. Fibrosis in (A) diaphragm and (B) heart of WT, mdx, IL-10−/− and IL-10−/−/mdx mice was detected by sirius-red staining. Red areas in the unpolarized views (upper panels) and bright areas in the polarized views (lower panels) represent fibrosis. (C) H&E and Masson's trichrome staining of transverse heart sections from 10-month-old mdx and IL-10−/−/mdx mice. The high-magnification images (lower panels) represent the boxed regions in upper panels. Bar = 1.0 mm (upper panels) and 100 μm (lower panels).

Figure 4.

Extensive fibrosis in diaphragm and pathological diagnosis of dilated cardiomyopathy. Fibrosis in (A) diaphragm and (B) heart of WT, mdx, IL-10−/− and IL-10−/−/mdx mice was detected by sirius-red staining. Red areas in the unpolarized views (upper panels) and bright areas in the polarized views (lower panels) represent fibrosis. (C) H&E and Masson's trichrome staining of transverse heart sections from 10-month-old mdx and IL-10−/−/mdx mice. The high-magnification images (lower panels) represent the boxed regions in upper panels. Bar = 1.0 mm (upper panels) and 100 μm (lower panels).

Since fibrosis is regulated by signaling through transforming growth factor-β (TGF-β), which is a fibrotic cytokine, we analyzed the expression levels of TGF-β. TGF-β1 and collagen-I production were assayed by enzyme-linked immunosorbent assay (ELISA). As shown in Figure 5A, the level of mature TGF-β was elevated in IL-10−/−/mdx mice compared with IL-10−/− and mdx mice. Similar to these data, western blot analysis showed the increased expression of mature TGF-β in IL-10−/−/mdx mice (Supplementary Material, Fig. S3). As expected, IL-10−/−/mdx mice demonstrated significantly higher expression of collagen-I in diaphragm and heart compared with other mice (Fig. 5B). On the other hand, the replacement of necrotic fibers by adipose tissue in diaphragm muscle was not significantly different between IL-10−/−/mdx and mdx mice (data not shown). These observations suggested that the absence of IL-10 gene induced the extensive fibrosis in the dystrophic diaphragm and heart. Although dystrophic muscle pathology in the diaphragm and heart has been described histologically, it remains unclear how chronic inflammation modulates disease progression, especially cardiac and respiratory dysfunction. We next investigated whether the genetic depletion of IL-10 was related to the cardiorespiratory dysfunction in mdx mice.

Figure 5.

Elevated TGF-β and collagen production in IL-10-deficient dystrophic mice. Quantification of (A) TGF-β1 (** P < 0.01) and (B) type-I collagen (*P < 0.05 and #P < 0.01; IL-10−/−/mdx versus WT, mdx and IL-10−/− mice) in diaphragm and heart lysate from 8-month-old mice by ELISA (n = 4–6, each group).

Figure 5.

Elevated TGF-β and collagen production in IL-10-deficient dystrophic mice. Quantification of (A) TGF-β1 (** P < 0.01) and (B) type-I collagen (*P < 0.05 and #P < 0.01; IL-10−/−/mdx versus WT, mdx and IL-10−/− mice) in diaphragm and heart lysate from 8-month-old mice by ELISA (n = 4–6, each group).

Severe respiratory dysfunction in IL-10−/−/mdx mice

To evaluate the progression of respiratory dysfunction, sensitive respiratory parameters were monitored by whole-body plethysmography, which provides extensive pulmonary analysis (Fig. 6). Although a nearly identical respiratory rate (RR) was observed among all groups of mice (Fig. 6A), Figure 6B shows that the tidal volume (TV) significantly decreased in the 8-month-old IL-10−/−/mdx mice compared with the other age-matched mice (mean ± SD, P < 0.01; WT, 22.6 ± 0.8 ml; IL-10−/−, 22.5 ± 0.7 ml; mdx, 28.3 ± 3.9 ml; IL-10−/−/mdx, 17.1 ± 3.0 ml). In addition, transient apnea was sporadically detected in IL-10−/−/mdx mice, but not in the other mice (Supplementary Material, Fig. S4). These data suggested that IL-10−/−/mdx mice have more severe dysfunction of respiratory muscles and diaphragm compared with mdx mice.

Figure 6.

Severe respiratory dysfunction in IL-10−/−/mdx mice. (A) Respiratory function was evaluated by whole-body non-invasive plethysmography in 8-month-old WT, mdx, IL-10−/− and IL-10−/−/mdx mice. (A) Respiratory rate and (B) tidal volume were assessed using a respiration monitoring system.

Figure 6.

Severe respiratory dysfunction in IL-10−/−/mdx mice. (A) Respiratory function was evaluated by whole-body non-invasive plethysmography in 8-month-old WT, mdx, IL-10−/− and IL-10−/−/mdx mice. (A) Respiratory rate and (B) tidal volume were assessed using a respiration monitoring system.

Significant decrease in cardiac function caused by IL-10 deficiency

In addition of the LV, RV dilatation and patchy fibrosis (Fig. 4C), to further investigate the effects of IL-10 signaling on the development of cardiac dysfunction in DMD, we performed echocardiograms. M-mode echocardiography showed decreased percent fractional shortening (%FS) and ejection fraction percent (EF%) in mdx mice at 8 months of age compared with age-matched WT and IL-10−/− mice (Table 1). The decrease in %FS and EF% was significantly larger in IL-10−/−/mdx mice compared with mdx mice, suggesting that IL-10−/−/mdx mice developed dilated cardiomyopathy with cardiac inflammation.

Table 1.

Conventional echocardiographic parameters

 WT mdx IL-10−/− IL-10−/−/mdx 
HR (bpm) 185 ± 10 209 ± 9 189 ± 19 189 ± 19 
EF (%) 79.4 ± 1.2 74.7 ± 3.7a 80.2 ± 0.1 65.6 ± 2.8b 
FS (%) 42.0 ± 1.1 37.8 ± 3.2a 42.7 ± 0.9 30.9 ± 1.9b 
 WT mdx IL-10−/− IL-10−/−/mdx 
HR (bpm) 185 ± 10 209 ± 9 189 ± 19 189 ± 19 
EF (%) 79.4 ± 1.2 74.7 ± 3.7a 80.2 ± 0.1 65.6 ± 2.8b 
FS (%) 42.0 ± 1.1 37.8 ± 3.2a 42.7 ± 0.9 30.9 ± 1.9b 

Echocardiography was performed on 8–10-month-old mice. Comparisons of echocardiography values (mean ± SD) for wild-type (WT), IL-10−/−, mdx and IL-10−/−/mdx mice aged 8 months.

HR, heart rate; FS, fractional shortening; EF, ejection fraction.

aDifference (P< 0.05) between WT and IL-10−/− mice.

bDifference (P<0.01) between WT, IL-10−/− and mdx mice.

Reduced early-life growth and survival of dystrophic mice

We examined whether aberrant IL-10 affects the lifespan of mdx mice. Kaplan–Meier survival curves revealed that 50% of IL-10−/−/mdx mice died around the first 50 weeks after birth (Fig. 7A), whereas mdx mice survived up to 2 years. We also examined progressive changes in body weight. IL-10−/−/mdx mice revealed smaller body mass by the age of 14 weeks (mean ± SD, IL-10−/−/mdx; 25.1 ± 2.1 g, versus mdx, 9.0 ± 1.6 g) (Fig. 7B), but IL-10−/−/mdx mice gained weight with age similar to mdx mice. These data showed that aberrant IL-10 affects the lifespan and growth rate of dystrophic mice.

Figure 7.

Reduced survival and early-life growth in IL-10−/−/mdx mice. (A) Kaplan–Meier survival curves for wild-type, mdx and IL-10−/−/mdx mice (n = 7–37, each). (B) Growth curves for WT, mdx and IL-10−/−/mdx mice (n = 4–6, each group). Data are expressed as means ± SD. *P < 0.05 and ** P < 0.01.

Figure 7.

Reduced survival and early-life growth in IL-10−/−/mdx mice. (A) Kaplan–Meier survival curves for wild-type, mdx and IL-10−/−/mdx mice (n = 7–37, each). (B) Growth curves for WT, mdx and IL-10−/−/mdx mice (n = 4–6, each group). Data are expressed as means ± SD. *P < 0.05 and ** P < 0.01.

DISCUSSION

In this study, we demonstrated that genetic ablation of IL-10 in dystrophic mdx mice with C57BL/6 background caused more severe phenotype in heart and respiratory function, as evidenced by increased macrophage infiltration, high levels of inflammatory factors in the muscle and also progressive cardiorespiratory dysfunction. IL-10 might be an important immune-modulator in dystrophic muscle.

Previous immune-histological data suggested that the early phase of dystrophic limb muscle pathology involves pro-inflammatory M1 macrophages, and during the later progressive phase, the macrophage subpopulation shifts toward an M2a phenotype regulated by IL-10 (19,20,29). In this study, we demonstrated that IL-10 ablation in mdx mice causes an increase in muscle injury, an effect probably mediated through the observed imbalance of M1/M2 phenotypes, especially in the diaphragm and heart of mdx mice and also delayed muscle repair in acute injured muscle (Figs 1 and 2 and Supplementary Material, Fig. S1). An imbalance of inflammatory factors could result from a deficiency in suppression by anti-inflammatory M2 macrophages or strong activation of pro-inflammatory M1 macrophages (Fig. 3).

The expression of pro-inflammatory factors (e.g. TNF-α, IL-1, TGF-β and MCP-1) has been reported prior to onset of muscle degeneration in DMD patients and/or mdx mice (28,30,31). TNF-α-deficient mdx mice demonstrated altered pathological progression of diaphragm and quadriceps muscle (30). Ablation of IFN-γ reduced muscle damage in mdx mice, showing significantly lower pathological markers such as macrophage/neutrophil infiltration and necrosis of myofibers (32). Therefore, these pro-inflammatory cytokines were described as key factors for the muscle damage induction caused by M1 macrophages (28). In contrast, we characterized that IL-10−/−/mdx mice closely recapitulate the DMD phenotype, as evidenced by the characteristic progressive muscle dysfunction associated with severe inflammation.

Increased levels of IL-1α, IL-1β, IL-1ra, IL-16, RANTES, M-CSF, MIG, JE/MCP-1 and TIMP-1 were detectable in the diaphragm and/or heart of aged IL-10−/−/mdx mice. Nuclear factor κB (NF-κB) activity is thought to contribute the up-regulation of pro-inflammatory factors, because IL-10 inhibits NF-κB activation rather than several other transcriptional factors including NF-IL-6 and AP-1 (33). IL-16, a potent chemoattractant for several immune cells such as monocyte, and CD4+T cells (34), would mediate further immune cell infiltration and activation in damaged muscle, similarly IL-1 and IL-1 receptor. Since IL-1β and IL-1ra are produced by M1 macrophages, our results suggest strong activation of M1 macrophages in IL-10-deficient dystrophic muscle. Whereas high level of TNF-α expression was known in the early phase of mdx mice (28), the TNF-α elevation in aged mice was not detectable in our immunoblot analysis (Fig. S2). However, up-regulation of TNF-α was confirmed in aged IL-10−/−/mdx mice compared with mdx mice by ELISA analysis (Supplementary Material, Fig. S5). We also demonstrated that elevated TGF-β signaling and type-I collagen expression resulted in widespread fibrosis in the diaphragm and heart of IL-10−/−/mdx mice lather than that of mdx mice (Figs 4 and 5). As previously reported, TGF-β is produced in response to fibrinogen stimulation of M1 macrophages via the intermediate induction of IL-1β (35). Since TGF-β was also reported to promote the TIMP-1 expression (36), up-regulated TIMP-1 may result in increased deposition of extracellular matrix through the TGF-β-TIMP-1 pathway and lead to tissue fibrosis. In the IL-10 regulated DMD muscle, active production of these inflammatory factors from infiltrating macrophages is associated with chronic inflammation to lead disease progression.

We demonstrated that the lack of IL-10 leads to respiratory dysfunction through severe inflammation and marked fibrosis in dystrophic diaphragm, without lung tissue damage (data not shown). The decreased tidal volume in IL-10−/−/mdx mice (Fig. 6) might to be due to the apnea followed by hyperpnea (Supplementary Material, Fig. S4). Mice lacking both utrophin and dystrophin (utrn−/−/mdx) exhibit a more severe DMD-like dystrophy with cardiomyopathy (37,38). IL-10−/−/mdx mice exhibited similar respiratory dysfunction to utrn−/−/mdx mice (data not shown), implying that respiratory dysfunction may be less dependent on membrane structure. Utrn−/−/mdx mice died by 16–20 weeks due to severe DMD phenotypes (37,39), while 80% of IL-10−/−/mdx mice were still alive at the same age (Fig. 7A). Furthermore, for breeding these mice, heterozygous for utrophin and either heterozygous for the dystrophin mutation (females) or hemizygous for the dystrophin mutation (males) are used to produce the utrn−/−/mdx mice (37). On the other hand, IL-10-homozygous deficient mdx mice, homozygous females and hemizygous male mice, can be utilized to generate the littermates. In these contexts, IL-10−/−/mdx mouse must be a quite useful model of DMD for long-term observation of disease phenotype under various therapeutic studies.

The most common manifestation of heart disease in DMD patient and mdx mice is dilated cardiomyopathy, which is characterized by a progressive decline in cardiac contractility, ventricular dilation and cardiac arrhythmias (40,41). A recent study showed that old IL-10 knockout mice develop left ventricle end-systolic diameter dilatation and heart enlargement with a reduction of EF% (42). Cardiac inflammation induced by IL-10 ablation induced cardiac dysfunction as the decreased LV function with LV and RV dilatation in IL-10−/−/mdx mice (Table 1 and Fig. 4C).

In this study, we confirmed our hypothesis that a predisposition to inflammation causes chronic inflammation and results in more severe cardiorespiratory dysfunction by pathological analysis using IL-10−/−/mdx mice. Our findings are especially important for the development of effective therapies using anti-inflammatory drugs and/or immunomodulatory stem cells, such as mesenchymal stromal cells, to improve muscle and cardiorespiratory dysfunction.

MATERIALS AND METHODS

Animal care and sampling

All experimental procedures were approved by the Experimental Animal Care and Use Committee at the National Center of Neurology and Psychiatry (NCNP, Tokyo, Japan). C57BL/6-background mdx mice were a generous gift from Dr T. Sasaoka (National Institute for Basic Biology, Aichi, Japan) and were maintained in our animal facility. C57BL/6J (WT) mice were purchased from Nihon CLEA (Tokyo, Japan). Mice carrying null mutation for IL-10 that had been back-crossed onto a C57/BL6 background, B10.129P2(B6)-Il10tm1Cgn/J mice, were purchased from The Jackson Laboratory (Bar Harbor, ME, USA). IL-10 knockout (B10.129P2(B6)-Il10tm1Cgn/J) mice were crossed with mdx mice to generate mice that lacked both IL-10 and dystrophin. All animals were maintained according to the standard protocol for animal care at the NCNP. Genotyping to detect the mutated alleles of IL-10 and dystrophin was performed by PCR. Null mutation of the dystrophin gene was confirmed using mdx-amplification-resistant mutation system PCR (43). Age-matched littermate mice (3-, 8-, and12-month-old) were used for experiments. Samples from heart, diaphragm and TA muscle were taken and immediately frozen in liquid nitrogen-cooled isopentane. Six to eight mice from each group were used for analysis at each time point.

Histopathology and immunohistochemistry

Transverse cryosections (10 μm thick) prepared from heart, diaphragm and TA muscle were stained with hematoxylin and eosin (H&E) using standard procedures. For immunohistochemistry, 8 μm thick cryosections were fixed in 1% paraformaldehyde–PBS for 30 min at 4°C, treated with Triton X-100 and then blocked with 3% bovine serum albumin (BSA) in PBS. The following antibodies were used at 1:50–1:100 dilutions for macrophage detection: rabbit anti-mouse CD68 antibodies (Abcam, Cambridge, UK), rat anti-mouse CD206 antibodies (Bio-Rad Laboratories, Inc., Hercules, CA, USA). All antibodies were diluted with PBS containing 0.5% BSA and incubated with the tissue sections for either 1 h at room temperature or overnight at 4°C. The tissue sections were washed with PBS and then incubated with Alexa 488- or Alexa 594-conjugated anti-rabbit or rat IgG antibodies (Life Technologies, Carlsbad, CA, USA), at 1:500 dilution for 1 h at 4°C according to the MOM procedure (Vector Laboratories, Burlingame, CA, USA). Glass slides were washed with PBS and mounted in Vectashield (Vector Laboratories) with 4′,6′-diamidino-2-phenylind ole (DAPI). Immunofluorescence was visualized using an IX71 fluorescence microscope (Olympus, Tokyo, Japan) and BZ-X700, BZ-9000 (KEYENCE, Osaka, Japan). For estimation of fibrosis, sections taken from frozen heart and diaphragm samples were stained in sirius-red solution or Masson's trichrome solution, as previously described (44).

Proteome profiler cytokine array

Muscle tissue samples were disrupted in a Multi-Beads Shocker (Yasui Kikai, Osaka, Japan) and the protein concentrations were determined using the Pierce® BCA Protein Assay Kit (Thermo Fisher Scientific Inc., IL, USA). The relative expression of cytokines and chemokines in muscle lysates was quantified using the Proteome Profiler™ Array (Mouse Cytokine Array, Panel A; R&D Systems Inc., Minneapolis, MN, USA). The array was performed according to the manufacturer's exact specifications using 250 μl plasma (n = 8). To achieve maximum assay sensitivity, the blots were incubated with plasma for overnight. ECL incubation was performed for 5 min using the Super Signal West Femto Chemiluminescent Kit (Thermo Scientific Pierce, Rockford, IL, USA), and then images were captured and analyzed using Image Quant LAS 4000 coupled with Image Quant TL software (GE Healthcare).

ELISA

The amount of TGF-β1 and type I collagen in muscle lysate obtained from diaphragm and heart was determined by the Quantikine ELISA mouse TGF-β1 immunoassay (R&D Systems, Inc.) and the mouse collagen type 1 (Col1) ELISA kit (Cloud-Clone Corp., TX, USA) according to the manufacturer's recommendations. Final values were normalized for protein concentration.

Assessment of respiratory function

Mouse respiratory function was evaluated by whole-body plethysmography. The plethysmograph was composed of two Plexiglas cylinders, one serving as the animal chamber and the other as reference chamber, and each unrestrained conscious 3- and 8-month-old mouse was placed in a ‘free moving’ chamber. RR and TV were assessed by a respiration monitoring system (Model RM-80, Columbus Instruments, Columbus, OH, USA). The signal was digitized using a Power Lab data acquisition system with Chart-Pro 6 software (AD Instruments, Dunedin, New Zealand) for recording and analysis.

Echocardiography

Mice were anesthetized by 2% isoflurane inhalation (Univentor 400 Anaesthesia unit, Univentor Limited, Malta). After shaving their chest, the mice were imaged using Vivid-i Dimensions (GE Vingmed, Horten, Norway) equipped with a transducer (i12-RS) transmitting at 10 MHz echocardiographic probe. Standard gray scale M- and B-mode images were acquired using a parasternal long- and short-axis view. For the basic measurements of left ventricular function, %FS and EF% were examined using M-mode echocardiography.

SUPPLEMENTARY MATERIAL

Supplementary Material is available at HMG online.

FUNDING

This work was supported by the research grant from JCR Pharmaceuticals Co., Ltd., Health Sciences Research Grants for Research on Human Genome and Gene Therapy from the Ministry of Health, Labor and Welfare of Japan (grant number: 21A-3), and a Grant-in-Aid for Scientific Research from the Ministry of Education, Culture, Sports, Science and Technology (grant number: 22390284, 22-40134).

ACKNOWLEDGMENTS

The authors thank Akinori Nakamura, Tetsuya Nagata, Jun Tanihata, Kazumi Motoki, Michihiro Imamura, Naoki Itoh, Kazuhiro Yamamoto, Masayuki Sekiguchi, Hiroshi Takano and Yoko Fujii for technical advice, support and helpful discussions; Ryoko Nakagawa, Shota Saka and Nana Tsumita for technical assistance. We also thank for the research support from JCR Pharmaceuticals Co., Ltd.

Conflict of Interest statement. We received the research support from JCR Pharmaceuticals Co., Ltd. and TaKaRa Bio Inc.

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